Nitric Oxide, Cell Signaling, and Gene Expression (Oxidative Stress and Disease)

Nitric Oxide, Cell Signaling, and Gene Expression OXIDATIVE STRESS AND DISEASE Series Editors LESTER PACKER, PH.D. E...

Author: Santiago Lamas (Editor) | Enrique Cadenas (Editor)

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Nitric Oxide, Cell Signaling, and Gene Expression

OXIDATIVE STRESS AND DISEASE Series Editors

LESTER PACKER, PH.D. ENRIQUE CADENAS, M.D., PH.D. University of Southern California School of Pharmacy Los Angeles, California

1. Oxidative Stress in Cancer, AIDS, and Neurodegenerative Diseases, edited by Luc Montagnier, René Olivier, and Catherine Pasquier 2. Understanding the Process of Aging: The Roles of Mitochondria, Free Radicals, and Antioxidants, edited by Enrique Cadenas and Lester Packer 3. Redox Regulation of Cell Signaling and Its Clinical Application, edited by Lester Packer and Junji Yodoi 4. Antioxidants in Diabetes Management, edited by Lester Packer, Peter Rösen, Hans J. Tritschler, George L. King, and Angelo Azzi 5. Free Radicals in Brain Pathophysiology, edited by Giuseppe Poli, Enrique Cadenas, and Lester Packer 6. Nutraceuticals in Health and Disease Prevention, edited by Klaus Krämer, Peter-Paul Hoppe, and Lester Packer 7. Environmental Stressors in Health and Disease, edited by Jürgen Fuchs and Lester Packer 8. Handbook of Antioxidants: Second Edition, Revised and Expanded, edited by Enrique Cadenas and Lester Packer 9. Flavonoids in Health and Disease: Second Edition, Revised and Expanded, edited by Catherine A. Rice-Evans and Lester Packer 10. Redox–Genome Interactions in Health and Disease, edited by Jürgen Fuchs, Maurizio Podda, and Lester Packer 11. Thiamine: Catalytic Mechanisms in Normal and Disease States, edited by Frank Jordan and Mulchand S. Patel 12. Phytochemicals in Health and Disease, edited by Yongping Bao and Roger Fenwick 13. Carotenoids in Health and Disease, edited by Norman I. Krinsky, Susan T. Mayne, and Helmut Sies

14. Herbal and Traditional Medicine: Molecular Aspects of Health, edited by Lester Packer, Choon Nam Ong, and Barry Halliwell 15. Nutrients and Cell Signaling, edited by Janos Zempleni and Krishnamurti Dakshinamurti 16. Mitochondria in Health and Disease, edited by Carolyn D. Berdanier 17. Nutrigenomics, edited by Gerald Rimbach, Jürgen Fuchs, and Lester Packer 18. Oxidative Stress, Inflammation, and Health, edited by Young-Joon Surh and Lester Packer 19. Nitric Oxide, Cell Signaling, and Gene Expression, edited by Santiago Lamas and Enrique Cadenas


edited by

Santiago Lamas Enrique Cadenas

Boca Raton London New York

A CRC title, part of the Taylor & Francis imprint, a member of the Taylor & Francis Group, the academic division of T&F Informa plc.

Published in 2006 by CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2006 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-10: 0-8247-2960-9 (Hardcover) International Standard Book Number-13: 978-0-8247-2960-8 (Hardcover) Library of Congress Card Number 2005044018 This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe.

Library of Congress Cataloging-in-Publication Data Nitric oxide, cell signaling, and gene expression / edited by Santiago Lamas and Enrique Cadenas. p. cm. -- (Oxidative stress and disease ; 19) Includes bibliographical references and index. ISBN 0-8247-2960-9 (alk. paper) 1. Nitric oxide--Physiological effect. 2. Nitric oxide--Pathophysiology. 3. Cellular signal transduction. 4. Genetic regulation. I. Lamas, Santiago. II. Cadenas, Enrique. III. Series. QP535.N1N5475 2005 616'.0473--dc22

2005044018

Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com Taylor & Francis Group is the Academic Division of T&F Informa plc.

and the CRC Press Web site at http://www.crcpress.com

Series Introduction Oxygen is a dangerous friend. Through evolution, oxygen—itself a free radical— was chosen as the terminal electron acceptor for respiration. The two unpaired electrons of oxygen spin in the same direction; thus, oxygen is a biradical. Other oxygen-derived free radicals, such as superoxide anion or hydroxyl radicals, formed during metabolism or by ionizing radiation are stronger oxidants (i.e., endowed with a higher chemical reactivity). Oxygen-derived free radicals are generated during oxidative metabolism and energy production in the body, and are involved in regulation of signal transduction and gene expression; activation of receptors and nuclear transcription factors; oxidative damage to cell components; the anti-microbial and cytotoxic action of immune system cells, neutrophils, and macrophages; and in aging and age-related degenerative diseases. Overwhelming evidence indicates that oxidative stress can lead to cell and tissue injury; however, the same free radicals that are generated during oxidative stress are produced during normal metabolism and, as a corollary, are involved in both human health and disease. In addition to reactive oxygen species, research on reactive nitrogen species has been gathering momentum to develop an area of enormous importance in biology and medicine. Nitric oxide or nitrogen monoxide (NO) is a free radical generated by nitric oxide synthase (NOS). This enzyme modulates physiological responses in the circulation, such as vasodilation (eNOS) or signaling in the brain (nNOS). During inflammation, however, a third isoenzyme is induced—iNOS—resulting in the overproduction of NO and causing damage to targeted infectious organisms and to healthy tissues in the vicinity. More worrisome, however, is the fact that NO can react with superoxide anion to yield a strong oxidant—peroxynitrite. Oxidation of lipids, proteins, and DNA by peroxynitrite increases the likelihood of tissue injury. Both reactive oxygen and nitrogen species are involved in the redox regulation of cell functions. Oxidative stress is increasingly viewed as a major upstream component in the signaling cascade involved in inflammatory responses and stimulation of adhesion molecule and chemoattractant production. Hydrogen peroxide decomposes in the presence of transition metals to the highly reactive hydroxyl radical, which by two major reactions—hydrogen abstraction and addition—accounts for most of the oxidative damage to proteins, lipids, sugars, and nucleic acids. Hydrogen peroxide is also an important signaling molecule that, among others, can activate NF-κB, an important transcription factor involved in inflammatory responses. At low concentrations, hydrogen peroxide regulates cell signaling and stimulates cell proliferation; at higher concentrations, it triggers apoptosis and, at even higher levels, necrosis. Virtually all diseases thus far examined involve free radicals. In most cases, free radicals are secondary to the disease process, but in some instances, free radicals are causal. Thus, a delicate balance exists between oxidants and antioxidants in health and disease. Their proper balance is essential for ensuring healthy aging. The term oxidative stress indicates that the antioxidant status of cells and tissues is altered by exposure to oxidants. The redox status is thus dependent on the degree

to which cells’ components are in the oxidized state. In general, the reducing environment inside cells helps to prevent oxidative damage. In this reducing environment, disulfide bonds (S–S) do not spontaneously form because sulfhydryl groups are maintained in the reduced state (SH), thus preventing protein misfolding or aggregation. This reducing environment is maintained by oxidative metabolism and by the action of antioxidant enzymes and substances, such as glutathione, thioredoxin, vitamins E and C, and enzymes such as superoxide dismutases, catalase, and the selenium-dependent glutathione reductase, as well as glutathione and thioredoxin hydroperoxidases, which serve to remove reactive oxygen species (hydroperoxides). Changes in the redox status and depletion of antioxidants occur during oxidative stress. The thiol redox status is a useful index of oxidative stress mainly because metabolism and NADPH-dependent enzymes maintain cell glutathione (GSH) almost completely in its reduced state. Oxidized glutathione (glutathione disulfide [GSSG]) accumulates under conditions of oxidant exposure and this changes the ratio GSSG/GSH; an increased ratio is usually taken as indicating oxidative stress. Other oxidative stress indicators are ratios of redox couples such as NADPH/NADP, NADH/NAD, thioredoxinreduced/thioredoxinoxidized, dihydrolipoic acid/α-lipoic acid, and lactate/pyruvate. Changes in these ratios affect the energy status of the cell, largely determined by the ratio ATP/ADP + AMP. Many tissues contain large amounts of glutathione, 2–4 mM in erythrocytes or neural tissues, and up to 8 mM in hepatic tissues. Reactive oxygen and nitrogen species can oxidize glutathione, thus lowering the levels of the most abundant non-protein thiol, sometimes designated as the cell’s primary preventative antioxidant. Current hypotheses favor the idea that lowering oxidative stress can have a health benefit. Free radicals can be overproduced or the natural antioxidant system defenses weakened, first resulting in oxidative stress, and then leading to oxidative injury and disease. Examples of this process include heart disease, cancer, and neurodegenerative disorders. Oxidation of human low-density lipoproteins is considered an early step in the progression and eventual development of atherosclerosis, thus leading to cardiovascular disease. Oxidative DNA damage may initiate carcinogenesis. Environmental sources of reactive oxygen species are also important in relation to oxidative stress and disease. A few examples include: UV radiation, ozone, cigarette smoke, and others are significant sources of oxidative stress. Compelling support for the involvement of free radicals in disease development originates from epidemiological studies demonstrating that an enhanced antioxidant status is associated with reduced risk of several diseases. Vitamins C and E, in the prevention of cardiovascular disease, are a notable example. Elevated antioxidant status is also associated with decreased incidence of cataracts, cancer, and neurodegenerative disorders. Some recent reports have suggested an inverse correlation between antioxidant status and the occurrence of rheumatoid arthritis and diabetes mellitus. Indeed, the indications in which antioxidants may be useful in the prevention or the treatment of disease are increasing in number. Oxidative stress, instead of being the primary cause of disease, is more often a secondary complication in many disorders. Oxidative stress diseases include inflammatory bowel diseases, retinal ischemia, cardiovascular disease and restenosis, AIDS, adult respiratory distress syndrome, and neurodegenerative diseases such as

stroke, Parkinson’s disease, and Alzheimer’s disease. Such indications may prove amenable to antioxidant treatment (in combination with conventional therapies) because a clear involvement of oxidative injury exists in these disorders. In this series of books, the importance of oxidative stress and disease associated with organ systems of the body is highlighted by exploring the scientific evidence and the medical applications of this knowledge. The series also highlights the major natural antioxidant enzymes and antioxidant substances such as vitamins E, A, and C, flavonoids, polyphenols, carotenoids, lipoic acid, coenzyme Q10, carnitine, and other micronutrients present in food and beverages. Oxidative stress is an underlying factor in health and disease. More evidence indicates that a proper balance between oxidants and antioxidants is involved in maintaining health and longevity, and that altering this balance in favor of oxidants may result in patho-physiological responses that cause functional disorders and disease. This series is intended for researchers in the basic biomedical sciences and clinicians. The potential of such knowledge for healthy aging and disease prevention warrants further knowledge about how oxidants and antioxidants modulate cell and tissue function. Lester Packer Enrique Cadenas Series Editors

Preface The role of nitric oxide (NO) as a physiological mediator was established with the discovery in the early 1980s of its capacity to regulate the vascular tone through cyclic GMP. Over the past 10 years, newer roles for NO have emerged, related to the ability of NO to interact with and modify a wide variety of other molecules, such as the free radical superoxide anion, key redox regulators such as glutathione, and macromolecules such as DNA and proteins. This forms the basis for the possibility of NO to influence crucial processes within the cell, such as the response to redox perturbations, protein function, and gene expression through non-enzymatic modifications. Among the mechanisms that underlie these effects, S-nitrosylation of proteins has attracted increasing interest in recent years, and it has been postulated as a possible new paradigm of signal transduction. The capacity of NO to interact with crucial mitochondrial enzymes, such as cytochrome oxidase, and the discovery of mitochondrial NO add even more relevance to the wide array of cellular functions on which NO may have an influence. Nitric Oxide, Cell Signaling, and Gene Expression is a collection of chapters written by experts on various aspects of NO functions: regulation of mitochondrial respiration by NO; mitochondrial NO signaling in redox modulation of cell behavior, synaptic plasticity, and cell death; and deleterious effects of NO on mitochondria, partly caused by peroxynitrite. The importance of NO in hypoxia is exemplified and analyzed in a chapter where the relationship between NO and the hypoxia-sensor HIF-1 is described in detail. Other chapters address modulation of cell metabolism by NO, regulation of cell signaling by cGMP, protein nitrosylation/denitrosylation, the Ras superfamily GTPases, and ceramide, as well as the involvement of NO in apoptosis through activation of caspases. A subset of chapters is devoted to the role of NO in gene expression and the post-transcriptional control of gene expression as well as a role for NO in tumor biology. The editors are grateful to the contributors for having shared their expertise in the completion of this work. Santiago Lamas Enrique Cadenas

About the Editors Enrique Cadenas is professor and chairman of molecular pharmacology and toxicology at the University of Southern California School of Pharmacy in Los Angeles. He earned his M.D. and Ph.D. in biochemistry from the University of Buenos Aires, Argentina. His research programs focus on mitochondrial oxidative/nitrosative stress, with implications for neurodegeneration and aging. Dr. Cadenas has authored 200 publications, edited numerous books, and serves on the editorial boards of several prestigious journals. Santiago Lamas is professor of research at the Centro de Investigaciones Biológicas from the Consejo Superior de Investigaciones Científicas (CSIC) in Madrid, Spain. He is also a group leader in the National Center for Cardiovascular Research (CNIC), working on the molecular patho-physiology of the vascular wall. His M.D. and Ph.D. degrees were earned at the Universidad Autónoma de Madrid in Spain. He has contributed to many publications on the field of nitric oxide and vascular biology, and is currently a member of the editorial boards of several journals on cardiovascular and free radical research.

Contributors Angeles Almeida Hospital Universitario de Salamanca Salamanca, Spain

Enrique Cadenas University of Southern California Los Angeles, California

Karl-Friedrich Beck Klinikum der Johann Wolfgang Goethe Universität Frankfurt, Germany

Sharon L. Campbell University of North Carolina Chapel Hill, North Carolina

Juan P. Bolaños Universidad de Salamanca Salamanca, Spain Cécile Bouton Institut de Chimie des Substances Naturelles, CNRS Gif-sur-Yvette, France Alberto Boveris University of Buenos Aires Buenos Aires, Argentina Bernhard Brüne University of Frankfurt Medical School Frankfurt, Germany Maurizio Brunori University of Rome “La Sapienza” and Consiglio Nationale delle Ricerche Rome, Italy Juanita Bustamante University of Buenos Aires Buenos Aires, Argentina

María Cecilia Carreras University of Buenos Aires Buenos Aires, Argentina Adriana Cassina Universidad de la República Montevideo, Uruguay Laura Castro Universidad de la República Montevideo, Uruguay Pilar Cidad Universidad de Salamanca Salamanca, Spain Emilio Clementi San Raffaele Scientific Institute Milano, Italy Daniela P. Converso University of Buenos Aires Buenos Aires, Argentina Analía Czerniczyniec University of Buenos Aires Buenos Aires, Argentina

María Delgado-Esteban Universidad de Salamanca Salamanca, Spain Clara De Palma University of Calabria Rende, Italy Stefanie Dimmeler University of Frankfurt Frankfurt, Germany Jean-Claude Drapier Institut de Chimie des Substances Naturelles, CNRS Gif-sur-Yvette, France Sestina Falcone University of Milano Milano, Italy Olivier Feron UCL Medical School Brussels, Belgium Soledad Galli University of Buenos Aires Buenos Aires, Argentina Paula García-Nogales Universidad de Salamanca Salamanca, Spain Benjamin Gaston University of Virginia Health System Charlottesville, Virginia Pedram Ghafourifar Marshall University Huntington, West Virginia

Alessandro Giuffrè University of Rome “La Sapienza” and Consiglio Nationale delle Ricerche Rome, Italy Judith Haendeler University of Frankfurt Frankfurt, Germany Jongyun Heo University of North Carolina Chapel Hill, North Carolina Lars-Oliver Klotz Heinrich Heine Universität Düsseldorf Düsseldorf, Germany Joshua Krumenacker University of Texas Health Science Center Houston, Texas Santiago Lamas Fundación Centro Nacional de Investigaciones Cardiovasculares (CNIC) Centro de Investigaciones Biológicas (CIB, CSIC) Instituto Reina Sofía de Investigaciones Nefrológicas Madrid, Spain Silvia Lores-Arnaiz University of Buenos Aires Buenos Aires, Argentina Joan B. Mannick University of Massachusetts Medical School Worcester, Massachusetts

Emil Martin University of Texas Health Science Center Houston, Texas Antonio Martínez-Ruiz Fundación Centro Nacional de Investigaciones Cardiovasculares (CNIC) Centro de Investigaciones Biológicas (CIB, CSIC) Instituto Reina Sofía de Investigaciones Nefrológicas Madrid, Spain Ferid Murad University of Texas Health Science Center Houston, Texas Leonor Oliveira Institut de Chimie des Substances Naturelles, CNRS Gif-sur-Yvette, France Lisa A. Palmer University of Virginia Health System Charlottesville, Virginia

Celia Quijano Universidad de la República Montevideo, Uruguay Rafael Radi Universidad de la República Montevideo, Uruguay Marianela Rodriguez Universidad de la República Montevideo, Uruguay Alfredo Saavedra-Molina Universidad Michoacana de San Nicolás de Hidalgo Morelia, México Paolo Sarti University of Rome “La Sapienza” and Consiglio Nationale delle Ricerche Rome, Italy Aurora Rachel Seminara University of Texas Health Science Center Houston, Texas

Cristiana Perrotta University of Calabria Rende, Italy

Iraida Sharina University of Texas Health Science Center Houston, Texas

Josef Pfeilschifter Klinikum der Johann Wolfgang Goethe Universität Frankfurt, Germany

Pierre Sonveaux UCL Medical School Brussels, Belgium

Juan José Poderoso University of Buenos Aires Buenos Aires, Argentina

Khalequz Zaman University of Virginia Health System Charlottesville, Virginia

Carlos Zaragoza Fundación Centro Nacional de Investigaciones Cardiovasculares (CNIC) Centro de Investigaciones Biológicas (CIB, CSIC) Instituto Reina Sofía de Investigaciones Nefrológicas Madrid, Spain

Jie Zhou University of Frankfurt Medical School Frankfurt, Germany

Table of Contents Chapter 1

Nitric Oxide Controls Cell Respiration by Reacting with Mitochondrial Complex IV ............................................................1

Paolo Sarti, Alessandro Giuffrè, and Maurizio Brunori Chapter 2

Mitochondrial Nitric Oxide Signaling in Synaptic Plasticity and Cell Death ..............................................................................29

Alberto Boveris, Silvia Lores-Arnaiz, Juanita Bustamante, and Analía Czerniczyniec Chapter 3

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior .......................................................45

María Cecilia Carreras, Soledad Galli, Daniela P. Converso, Juan José Poderoso, and Enrique Cadenas Chapter 4

Functions of Mitochondrial Nitric Oxide Synthase ....................77

Pedram Ghafourifar and Alfredo Saavedra-Molina Chapter 5

Peroxynitrite: A Mediator of Nitric-Oxide-Dependent Mitochondrial Dysfunction in Pathology ....................................99

Celia Quijano, Adriana Cassina, Laura Castro, Marianela Rodriguez, and Rafael Radi Chapter 6

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons .........................................................145

Juan P. Bolaños, María Delgado-Esteban, Pilar Cidad, Paula García-Nogales, and Angeles Almeida Chapter 7

Nitric Oxide Cell Signaling Mediated by cGMP ......................167

Emil Martin, Iraida Sharina, Aurora Rachel Seminara, Joshua Krumenacker, and Ferid Murad

Chapter 8

Regulation of Cell Signaling by Protein Nitrosylation/Denitrosylation .....................................................217

Joan B. Mannick Chapter 9

Nitric Oxide and Caspase Activation ........................................231

Judith Haendeler and Stefanie Dimmeler Chapter 10 Signaling Effects of Peroxynitrite in Mammalian Cells ...........245 Lars-Oliver Klotz Chapter 11 Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases .....................................................263 Jongyun Heo and Sharon L. Campbell Chapter 12 Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System ...................................................................291 Jie Zhou and Bernhard Brüne Chapter 13 The Cross Talk between Nitric Oxide and Ceramide: Coordinate Interactions among Signaling Pathways Regulating Cell Death, Survival, and Differentiation ...............311 Cristiana Perrotta, Clara De Palma, Sestina Falcone, and Emilio Clementi Chapter 14 S-Nitrosothiol Signaling and Gene Regulation in Pulmonary Pathophysiology .........................................................................321 Khalequz Zaman, Lisa A. Palmer, and Benjamin Gaston Chapter 15 Nitric Oxide and Gene Expression ............................................331 Josef Pfeilschifter and Karl-Friedrich Beck Chapter 16 Nitric Oxide as a Modifier of Gene Expression .......................353 Santiago Lamas, Antonio Martínez-Ruiz, and Carlos Zaragoza Chapter 17 Nitric Oxide and Post-Transcriptional Control of Gene Expression by the IRE/IRP System .................................371 Leonor Oliveira, Cécile Bouton , and Jean-Claude Drapier

Chapter 18 Nitric Oxide and Tumor Biology ...............................................395 Pierre Sonveaux and Olivier Feron Index ................................................................................................................ 421

Oxide Controls Cell 1 Nitric Respiration by Reacting with Mitochondrial Complex IV Paolo Sarti, Alessandro Giuffrè, and Maurizio Brunori University of Rome ”La Sapienza” and Consiglio Nationale delle Ricerche, Rome, Italy

CONTENTS 1.1 1.2

The Mitochondrial Production of NO .......................................................2 The Functional Relevance of the Reactions between NO and Mitochondria ................................................................................3 1.3 The Fast-Responding Mitochondrial Target of NO is Cytochrome c Oxidase ..............................................................................4 1.4 Two Mechanisms of the Inhibition of CcOX by NO ...............................8 1.5 The Reaction of NO with Oxidized CuB Yields the Nitrite-Inhibited CcOX, Rapidly Recovering Function ........................................................9 1.6 The Reaction of NO with the Fully Reduced (R) or the Half Reduced (E) Species Yields the Nitrosyl-Inhibited CcOX, Slowly Recovering Function ...................................................................10 1.7 The Reaction between NO and CcOX in Turnover ................................12 1.8 Experimental Designs ..............................................................................17 1.9 Persistence of NO in the Mitochondrion ................................................20 1.10 Acknowledgments ....................................................................................22 References .................................................................................................22 Over the past 10 years, evidence has been collected that suggests a role for nitric oxide (NO) in cell bioenergetics, and many reviews have highlighted the relevance and the multiple aspects of the issue [1–12]. This chapter summarizes some of the experimental information available, particularly focusing on the structural basis and the mechanisms of the reactions between NO and mitochondrial complex IV, and their functional relevance. 1

2


1.1 THE MITOCHONDRIAL PRODUCTION OF NO Nitric oxide (i.e., the nitrogen monoxide NO) has been recognized as ubiquitous, its presence inducing a variety of intra- and intercellular physiological actions [13–17], all virtually prone to become of pathological relevance due to the high reactivity of NO. In the cell as well as in vivo, NO is enzymatically produced by the NO-synthases (NOSs), converting L-arginine to L-citrulline in the presence of NADPH, O2 and other co-factors [18–20]. Three NOS isoforms have been isolated; these are almost ubiquitously expressed, although more typically by different cell lines or under inducible metabolic or experimental conditions (see Table 1.1). The so-called neuronal NOS (nNOS) and the endothelial NOS (eNOS) are the constitutive Ca++-dependent NOSs, whereas the expression of the inducible Ca++-independent NOS (iNOS) can be enhanced in immunocompetent cells such as macrophages [21, and references therein]. The iNOS activity is up-regulated during the inflammatory response, and can be stimulated by effectors, such as the lipopolysaccharide and the cytokines interferon γ, and/or the tumor necrosis factor α (TNF-α). The three types of NOS have been sequenced and characterized [22], whereas the existence of a fourth isoform, initially proposed as a specific mitochondrial NOS (mtNOS), has been ruled out [21]. Kanai et al. [23] proposed that the mtNOS is actually an nNOS, based on the observation that mitochondria of cardiomyocytes from mice knockout for nNOS do not produce NO, contrary to wild-type mice. Later, Elfering et al. [24] reported that the mtNOS most likely is the isoform alfa of the nNOS, excluding the existence of an additional splicing product of the nNOS alternative to the three identified. According to these authors [21, 24], the mtNOS is a constitutive nNOS-alfa bearing two post-translational modifications, namely an acylation and a phosphorylation, accounting for the interplay of this NOS with the mitochondrial inner membrane [21, and references therein]. Thus, the mtNOS is a membrane-bound NOS, localized in the mitochondrion as proposed originally [25–30]. Accordingly, NO has been demonstrated to be produced by mitochondria isolated to a high degree of purity from brain, heart [31], and other organs [23, 31–34]. Purified mitochondria in the presence of L-arginine produced NO and nitrite; thereby, respiration is inhibited, and inhibition is released by the NOS inhibitor L-nitrosoarginine, or similar compounds [35]. It is worth noting that NO like O2 can diffuse very rapidly through biological membranes [36, 37], making the existence of a local mtNOS perhaps less relevant to physiology. Regardless of where NO is produced, more than 10 years of investigation has proven the rapid inhibition of the aerobic mitochondrial respiration by submicromolar NO [11, 38–40]. As the release of mitochondrial inhibition is also rapid, the interaction between NO and the respiratory chain displays the characteristics of a functional control reaction. For this reason, the effect of NO on cell respiration is of particular interest and, depending on circumstances, might be of physiological or pathological relevance.

Nitric Oxide Controls Cell Respiration

3

TABLE 1.1 A Synopsis of the NO Synthases

Isoform

Nickname

Tissue/cell specificya

MW (kDa)

Ca++dependence

NOS-1 NOS-2

nNOSb iNOS

Neurons Macrophages

157 135

+ -

NOS-3

eNOS

Endothelium

140

+

NOS-1

mtNOSc

Mitochondria

130–147

+

a

All NOSs isoforms are widely distributed through most cells and tissues; tissue specificity is herein intended as to where NOSs typically have been predominantly found and historically purified. b Four splice variants of the nNOS-1 (nNOS-α) have been described, namely the nNOS-β, nNOS-γ, nNOS-µ, and nNOS-2. c

The most accredited hypothesis is that the mtNOS is an nNOS-α that is post-translationally modified [21].

1.2 THE FUNCTIONAL RELEVANCE OF THE REACTIONS BETWEEN NO AND MITOCHONDRIA A simple way to directly unveil in cultured cells the inhibition of cell respiration by NO is to perform fluorescence-microscopy experiments aimed at functionally visualizing mitochondria. In such experiments, the cells respiring on physiological substrates and glucose electrophoretically accumulate in the mitochondria a cationic fluorescent dye, typically rhodamine or JC-1. The import is driven by the membrane potential component, ∆ψ, of the proton electrochemical potential gradient, ∆µH+ [41, 42]. Under these conditions, mitochondria rapidly accumulate the probe and light up (Figure 1.1a). The same experiment performed in the presence of NO-donors, or after stimulation of the endogenous NO-synthase, demonstrated a marked decrease of mitochondrial fluorescence, indicative of the respiratory chain inhibition (Figure 1.1b). Mitochondrial inhibition appears rapidly and is reversible upon interruption of the NO flux, or washing the cells, or by specifically inhibiting NOS with 7-nitroindazole [42]. In 1994, clear-cut experiments [43, 44] demonstrated that the time course of NO inhibition of respiration sustained by mitochondria of neuronal synaptosomes is fully compatible with the interaction of NO with the terminal acceptor of the

4


150 120

1

60

4

2

90

3

5

30 0 1 (a)

2

3

4

5

(b)

FIGURE 1.1 Mitochondrial fluorescence microscopy of cells importing rhodamines. Typical images of the mitochondrial network in astrocytes and neurons (left panel). Computeraided image analysis allows quantitation of fluorescence as observed: (1) in the presence of nigericin alone, to fully convert ∆µH+ into ∆Ψ; (2) by inhibiting the NOS (7-nitroindazole, 7-N); (3) by stimulating the NOS (N-methyl-D-aspartate, NMDA); (4) as 3, but in the presence of 7-N, or (5) after collapsing ∆Ψ with valinomycin.

respiratory chain, the mitochondrial complex IV (i.e., cytochrome c oxidase [CcOX]) [43]. Since then, a wealth of evidence has been produced pointing to CcOX as the primary mitochondrial target for NO, and the fast (seconds) phenomenology triggered by the production/supplementation of NO to respiring cells appears to be due to the reaction of NO with CcOX. On a much longer time scale (tens of minutes to hours) and higher NO concentrations (>> µM), other mitochondrial complexes also react and are inhibited [45, 46]. This chapter describes in detail the cell respiratory changes observed in the presence of NO, particularly focusing on the mechanism(s) by which NO reacts with CcOX.

1.3 THE FAST-RESPONDING MITOCHONDRIAL TARGET OF NO IS CYTOCHROME C OXIDASE Cytochrome c oxidase belongs to the heme-copper oxidase superfamily. Ubiquitous in the aerobic organisms, the heme-copper oxidases transfer electrons from reduced cytochrome c or quinols (in some bacteria) to O2 [49]. This redox reaction is coupled to a vectorial proton translocation (pump) across the inner mitochondrial (eukaryotes) or the periplasmic (bacteria) membrane. The free energy release contributes to formation and maintenance of the proton electrochemical gradient ∆µH+ used to synthesize ATP [47]:


5

4 cyt.c2+ + O2 + 8 H+in → 4 cyt.c3+ + 2 H2O + 4 H+out In 1995, the structure of the aa3-type Paracoccus denitrificans and that of the enzyme purified from beef heart were simultaneously published. The latter is a dimer of 200 kDa monomers, each comprising 13 different polypeptides/subunits [49]. In the monomer (Figure 1.2), three Cu and two Fe ions are organized into four redox-active metal centers, namely the CuA (bimetallic), the low-spin heme a, and the CuB and the high-spin heme a3 (the so-called binuclear center). A Zn and a Mg atom (Mn in bacteria) were found to be present in the structure [50]. The Mg/Mn site, located close to the heme a3-CuB site, was suggested to be involved in the exit pathway for protons/water molecules [51]. The role of these additional metals is still obscure; they are redox-silent and, as far as we know, do not participate to the NO or other ligand/substrate binding chemistry. They likely contribute to stabilizing a trans-membrane structure, suitable for electron and proton transfer via pathways and channels of the protein moiety. X-ray structures of the bacterial aa3-type CcOX from Rhodobacter sphaeroides and of the ba3-type CcOX from the thermophilic bacterium Thermus thermophilus have been also reported, together with the structure of a ubiquinol oxidase, the bo3 from Escherichia coli [52]. The bimetallic CuA site of CcOX is the electron-entry door of the enzyme (Figure 1.2) [53]; CuA accepts electrons from reduced cytochrome c, located in the intermembrane space of the mitochondrion (the periplasmic space of bacteria) or from other reducing substrates [47]. CuA is in rapid equilibrium with heme a, and electrons are thereby rapidly transferred intramolecularly to the active binuclear site where O2, NO, and other ligands can bind. Interestingly, O2 and CO only bind to the fully (two-electrons) reduced binuclear site, whereas NO can also bind to the half (one-electron) reduced or even to the oxidized site, as further discussed next [3, and references therein]. The possibility of NO to react with several CcOX intermediates/species is important to understand its peculiar efficacy as inhibitor of respiration. Thus, the reaction with NO occurs at the level of the active site (i.e., in the same site where O2 binds and reacts) [3]. In all heme-copper oxidases, this conserved bimetallic site (Figure 1.3) is constituted by a high-spin heme (a3, b3, o3 depending on the organism) and a copper ion (called CuB). In the beef-heart enzyme, heme a3 is coordinated by H376, whereas CuB is coordinated by H240, H290, and H291 [49, 50]. A tyrosine residue, Y244, is covalently bound to H240 and is highly conserved, being presumably absent only in the cbb3-type oxidases, the most divergent members of the heme-copper oxidases superfamily [54, 55]. Tyrosine 244 has been proposed to become a radical during catalysis [56]; it is, therefore, a putative additional target for NO, although this is, at present, a speculation. By reacting with NO, activated tyrosines and thiols yield, respectively, the nitro- and the nitroso-derivatives. In addition to Tyr244, beef heart CcOX contains the bulk-exposed cys115 in subunit III, a potential additional reaction site for NO [57, 58].

6


e− Cyt c

CuA

CuB heme a3

heme a

FIGURE 1.2 Cytochrome c oxidase. Purified from ox heart as a dimer, each 200 kDa monomer comprises 13 different polypeptides/subunits. In the monomer, three Cu and two Fe ions are organized into four redox-active metal centers; namely a bimetallic CuA center accepting the electrons donated by cytochrome c, heme a in rapid equilibrium with CuA, wherefrom electrons are intramolecularly transferred to heme a3 and CuB, the binuclear active site of the enzyme. Y244

H240

H290 H376

H291

FIGURE 1.3 The active site of cytochrome c oxidase. Notice the tyrosine residue, Y244, covalently bound to H240; this highly conserved residue may possibly react with NO (reaction not reported, so far). (From the Protein Data Bank coordinates deposited by Tsukihara et al., Science (1995) 269: 1069–74.)


7

Despite the existence of several potential reaction sites, at physiological NO concentrations (micromolar or less), NO appears to react with CcOX, only at the level of the redox metals heme a3 and CuB [3]. Peroxynitrite (ONOO−), exogenously added and only in large excess over CcOX (≥ 100-fold), induces protein nitration [59]. Thus, when dealing with the reaction of NO with CcOX, the attention should be focused on the reactions involving the metals in the active site (Fe and Cu). As discussed next, these reactions lead to accumulation of products with an impact on mitochondria metabolism that might be substantially different. The reaction between NO and CcOX has to take into account the fact that, during catalysis, the metals in the active site undergo rapid (micro/milliseconds) redox and ligation changes, forming oxidized, partially reduced, and O2bound species (Figure 1.4). Particularly the so-called half-reduced binuclear site (i.e., a species where only one electron resides on the site) has been recently demonstrated to rapidly react with NO, providing an additional rationale to the advantage that NO appears to have over O2 when reacting with CcOX [60]. When attempting to draw an overall picture of the NO to CcOX interactions, one should keep in mind that: 1. All redox species and intermediates of CcOX react with NO 2. The complete, though schematic, catalytic cycle includes two slow (milliseconds) reductive steps, followed by the diffusion limited (k ≥ 108 M−1s−1) O2 binding and three fast (microseconds) oxidative steps 3. Moreover, at non-limiting O2 concentration (i.e., in a first approximation above KM,O2 (≥ 1– 10 µM)) the fraction of intermediates populated during turnover depends on the concentration of reduced cytochrome c, whereas the balance between the oxidative and the reductive equivalents at the CcOX site becomes relevant when O2 becomes limiting— below KM,O2 [61]. A schematic view of the catalytic cycle is reported in Figure 1.4 to allow a clearer understanding of the reactions of NO with CcOX during turnover. In this respect, it is worth knowing that most of the spectroscopic information on the chemistry of the binuclear site has been gathered by flashing the fully reduced CO-bound derivative of CcOX in the presence of oxygen [62] or, alternatively generating oxygen in situ on a nanosecond (or faster) time scale, by photolysing a synthetic caged O2-carrier [63]. In summary, a catalytic cycle includes a reductive and an oxidative limb [64, 65]; in Figure 1.4, the species populated are labeled with capital letters. Reduction of the oxidized active site O proceeds via two sequential electron donations from CuA via heme a. The first electron leads to formation of the half-reduced intermediate E (E1 and E2, indicating respectively the two half-reduced species with the electron residing on heme a3, or on CuB [66]); the second electron leads to the fully reduced R. After the relatively slow (ms) reduction [56, 67], the much faster (µs) oxidation restores the initial species O through the transient formation of the so-called intermediates P and F; these are both oxo-ferryl adducts [68],

8

Nitric Oxide, Cell Signaling, and Gene Expression Reductive phase

Oxidative phase

a33 NO2− +

NO

NO

e−

1000 µs

k = 107 − 108 s−1

O ms

e−

F

NO

100 µs e−

E1

E2 NO

P

−

ms

300 µs k = 104 − 105 s−1

e

+

a32 NO

R O2

NO

FIGURE 1.4 The catalytic cycle and the reaction of NO with the intermediates of cytochrome c oxidase. This is an oversimplified scheme of the catalytic redox cycle of the active site. O, R, P, and F stand for oxidized, reduced, peroxy, and ferryl state. E identifies the half-reduced active site (see text). The rate constant values are within ms for the slow reductive phase (O Æ R) and hundreds of µs for the fast oxidative phase (R Æ O). NO reacts very rapidly (k = 107 − 108 M−1s−1) with the reduced species R more slowly (k = 104 − 105 M−1s−1) with intermediates O, P and F. Differences in rate constant can be compensated by a higher occupancy in turnover of the O, P, and F species. (Modified from Sarti et al., Free Radic. Biol. Med. (2003) 34:509–20.)

although their detailed chemical identity is still controversial. Regardless of the number of CcOX species that can react with NO, only two adducts have been observed and identified, namely a nitrosyl- [CuB+a32+ NO] and a nitrite[CuB++a33+NO2−] derivative. It is worth mentioning that, in the nitrite-derivative, although the oxidation state of the heme-Fe can be observed and is thus defined, the redox state of CuB is only assumed.

1.4 TWO MECHANISMS OF THE INHIBITION OF CcOX BY NO Although the first report of a reaction between NO and CcOX dates to 1955 [69], the physiological relevance of this reaction was ignored until the late 1980s. Meanwhile, the reaction of NO with the reduced CcOX [70] was investigated, leading to the conclusion that NO is a very efficient reactant for the reduced heme-iron, and thus is a tool alternative to CO to stabilize the heme a3 Fe2+ [71]; however, NO, differently from CO and O2, proved to also react with the oxidized active site of the enzyme [72]. In 1994, however, NO proved to affect mitochondrial respiration via a fully reversible, transient inhibition of CcOX, displaying a competition with O2 [73].


9

With purified mitochondrial CcOX, ligand binding to the fully reduced enzyme in the absence of O2 is not associated to a redox reaction. Combination occurs at the reduced heme a3, following bimolecular kinetics [70, 74, 75]. Under anaerobic conditions, NO reacts very quickly (k ≈ 108 M−1 s−1) and with high affinity (Kaff ≈ 1011 M−1, at 20°C), yielding a typical Fe2+-NO nitrosyl-adduct [74, 75]: CuB+ a32+ + NO

CuB+a32+-NO

On the contrary, the reaction of NO with oxidized CuB (see next paragraph) leads to the oxidative degradation of NO to nitrite, presumably via the transient formation of a nitrosonium ion (NO+); the newly formed nitrite binds momentarily to the active site of the enzyme leading to inhibition [76]: CuB2+a33+ + NO → CuB+ a33+-NO+ + OH− → CuB2+ a33+-NO2− + H+ + e− At a given NO concentration and during turnover, both the reactions just described can occur, although to a different extent depending on the relative occupancy of the intermediates bearing reduced heme a3 or oxidized CuB. A third type of reaction has been proposed [77–79], and according to Pearce et al. [77], may yield the transient formation of peroxynitrite bound to the site and its subsequent reduction to nitrite. For the sake of clarity, we will dissect the catalytic cycle and treat separately the reactions with NO of all CcOX species and intermediates, having either oxidized CuB as in species O, P, F, or reduced heme a3 as in species R and E1. Interestingly, E1 bears at the same time reduced heme a3 and oxidized CuB; whether this peculiar redox state plays a special role in NO binding/degradation is still obscure.

1.5 THE REACTION OF NO WITH OXIDIZED CuB YIELDS THE NITRITE-INHIBITED CcOX, RAPIDLY RECOVERING FUNCTION The ability of the CcOX intermediates to react with NO has been studied in detail by mixing, in a stopped flow, NO with CcOX oxidized, O, or in the R, P, and F state [61, 78]. All these species can be independently generated and react with NO. The reaction of the oxidized CcOX with NO yields the inhibited nitritebound form of the enzyme, but removal of chloride is necessary to observe this reaction [79]. It is worth mentioning, in fact, that chloride is commonly bound to the active site of the oxidized enzyme as prepared and affects its reaction with ligands [80]. Although direct X-ray crystallographic evidence for the presence of Cl− in the site is missing, a wealth of indirect measurements (EXAFS, EPR) suggest that chloride is bound, probably to CuB2+. Thus, to observe the reaction of the oxidized enzyme with NO, CcOX has to be preliminarily stripped from

10


chloride, usually by a reduction/reoxidation cycle in a chloride free medium. In the reaction with oxidized CuB, NO is oxidized to nitrite, which then binds to the binuclear site perturbing the spectrum of the heme a33+ [61, 78]; the nitrite ion, by occupying the site, inhibits the enzyme [81]. The fully oxidized species O is (obviously) the most stable CcOX species, in air equilibrated buffer and at room temperature. Intermediates P and F can also be prepared in a sufficiently pure and stable state to be characterized. Similar to O, intermediates P and F bear oxidized CuB in the active site, and on this basis, Torres et al. [78] proposed a mechanism common to the three intermediates, involving the oxidation of NO to nitrite via formation of the nitrosonium ion NO+ at the level of CuB. Interestingly, the reaction of NO with these species rapidly generates in the absence of reductants a stable spectral perturbation of heme a3 identical to that observed when nitrite is added to the oxidized chloride-free enzyme (Figure 1.5). Taken together, both the optical spectroscopy [61, 78] and the NO amperometry [61] measurements suggest that intermediates O, P, and F react with NO with the same stoichiometry (1:1) and kinetics (kO = 2 × 105 M− 1s−1 to be compared with k 4 5 −1 −1 P,F ≈ 10 ÷ 10 M s , at 20°C). The interesting finding is that the adduct, which accumulates in all cases, is the inhibited nitrite-bound oxidized heme a3, regardless of whether the experiment has been performed with O, P, or F (Figure 1.5). Owing to the lower affinity of the reduced active site for nitrite [61, 81], and relevant to the mitochondrial respiratory chain function, the nitrite CcOX-derivative promptly recovers activity upon reduction by cytochrome c or other artificial electron donors [61]. To better focus on the functional role of the reaction between NO and the oxidized CuB, it is worth noticing that the reaction is two to three orders of magnitude slower than that with R (k ≈ 1 × 108 M−1s−1). Nevertheless the reaction is still relevant when the overall occupancy of the intermediates O, P, and F is predominant, as it happens when turnover is sustained by a slow electron supply to CcOX [61].

1.6 THE REACTION OF NO WITH THE FULLY REDUCED (R) OR THE HALF REDUCED (E) SPECIES YIELDS THE NITROSYL-INHIBITED CcOX, SLOWLY RECOVERING FUNCTION The reaction of the partially or the fully reduced CcOX with NO yields the inhibited nitrosyl form of the enzyme, bearing NO bound to reduced heme a3 (a32+NO). Until the kinetics of the reaction of NO with the oxidized CuB was reexamined and the reaction with the partially reduced active site discovered, the formation of a complex with the fully reduced CcOX was the only pathway considered responsible for CcOX inhibition. As outlined previously, the reaction of NO with R is a fast bimolecular process (k = 0.4 − 1.0 × 108 M−1 s−1 at 20°C, [70, 82]), yielding a tight heme a32+-NO adduct [74, 75]. Inhibition of CcOX is reversible, and a competition between NO and O2 was clearly demonstrated [73], proving that the fully reduced active site is one of the targets of NO. Removal


11

412 O, P, F

∆ OD

0.02

0

432

−0.02

0.04

426

∆ OD

R

0

−0.04

444

400

500 λ (nm)

600

FIGURE 1.5 Spectroscopic features of the nitrosyl- and the nitrite-derivative of cytochrome c oxidase. Difference spectra recorded by mixing intermediates O, P, F (top panel), or R (bottom panel), ∼2 µM CcOX functional unit (i.e., containing 2 hemes), with NO; notice the different position of the peaks. In the top panel, the difference spectra are reported after subtraction of cytochrome a contribution; they display high similarity and closely match the spectrum obtained by mixing oxidized CcOX with excess NO2−. (Modified from Giuffrè et al., Biochemistry (2000) 39: 15446–53.)

of free NO is associated to recovery of activity, by slow dissociation NO from the Fe2+ of heme a3 (k′ = 0.01 s−1 at 37°C [83]). This dissociation process, which is relatively fast for a heme protein [84 and references therein], is compatible with a fairly rapid respiration recovery and preservation of mitochondrial function, but it is still quite slow if compared with dissociation of nitrite from the active site under reducing conditions. Interestingly, from the experimental point of view, the NO dissociation rate is light sensitive (see below) [83, 85]. In the dark at 20°C, the functional recovery of respiration occurs at k′ = 4 × 10−3 s−1, a value that can be dramatically accelerated by white light (e.g., up to 20- to 30fold, by a 150 W xenon lamp, heat filtered, see Figure 1.6). As indicated by amperometric measurements performed anaerobically, in the reaction with the fully reduced enzyme, a single NO molecule binds to heme a3 with no redox changes [86]. No evidence exists for binding of a second NO molecule to CuB+,

12

Nitric Oxide, Cell Signaling, and Gene Expression 0.1

k' (s−1)

0.08

0.06 0.04

0.02 k = 4 × 10−3 s−1

0

0

2000 4000 Light intensity (counts)

6000

FIGURE 1.6 Light-induced dissociation of NO from fully reduced nitrosyl-cytochrome c oxidase. Fully reduced CcOX nitrosylated with stoichiometric NO is mixed with O2 in a photodiode array stopped flow, and the intensity of the incident white light beam is varied. In the dark (back extrapolation), the observed dissociation proceeds at k = 4 × 10−3s−1 (T = 20°C).

up to ∼20 µM NO (the upper limit for amperometric measurements of the type reported in Reference 86). This finding suggests that NO has a low affinity for reduced CuB, and makes the hypothesis of a NO reductase-like activity of CcOX less likely, consistently with the report of Stubauer et al. [86]. The two single-electron reduced species, E1 and E2, have never been produced separately in a stable form so to allow differentiation of their reactivity, thus both have been proposed to react with NO. We tend to believe that E1 (i.e., the species with the reduced heme a3) instead of E2 reacts with NO based on the just mentioned higher affinity of heme a3 for the ligand in the R state of CcOX. Consistently, the reaction of NO with the single-electron reduced intermediate E also apparently yields the nitrosyl-derivative of CcOX [60]. This conclusion has been obtained by studying the reactivity toward NO of a K-channel mutant of Paracoccus denitrificans oxidase (K354M), in which electron transfer to the active site is severely slowed down [60]. In the presence of excess reductants, the E species of the K354M mutant does not display a significant reactivity toward O2, this reaction demanding two electrons in the heme a3-CuB site, whereas it promptly reacts with NO yielding the nitrosyl-derivative; see Figure 1.7.

1.7 THE REACTION BETWEEN NO AND CcOX IN TURNOVER Both the nitrosylated- and the nitrite-bound (native) enzyme recover function by dissociation of the inhibitor (NO or NO2−) from the active site. In the presence of O2, the reduced nitrosylated enzyme recovers activity at the rate of dissociation


13

0.4 1

OD

3

0.2

2

0 400

500 λ (nm)

600

FIGURE 1.7 Absolute spectra of the K354M mutant of the Paracoccus denitrificans cytochrome oxidase. Absolute spectra of cytochrome c oxidase oxidized (1), heme a-reduced (2), and NO bound, fully reduced nitrosylated (3). (Modified from Giuffrè et al., J. Biol. Chem. (2002) 277: 22402–6.)

of NO from Fe2+ of heme a3. Still unclear, though possibly relevant to physiology, appears to be the fate of dissociated NO. According to recent reports [77, 87], NO would be released in the bulk not as such, as originally assumed [3 and references therein] but oxidized to nitrite instead. Pearce et al. [77] claim that in the nitrosylated enzyme CuB+ would react with O2 leading to formation of superoxide, which would react within the active site with NO producing peroxynitrite, thereafter reduced to nitrite before dissociation from the enzyme. If this hypothesis holds, activity recovery of the fully reduced nitrosylated enzyme in the presence of O2 would also be associated with the CcOX-mediated oxidative degradation of NO to nitrite. It is agreed that the NO inhibition occurs in competition with O2. The dependence of inhibition on O2 concentration, however, is a complex issue; both ligands target the same site, but NO appears to react with R and E1, whereas O2 reacts only with R. Moreover, NO reacts also with O, as well as with P and F, these two intermediates being significantly populated when turnover is sustained by low levels of reductants [61]. Finally, when dealing with the NO inhibition under turnover conditions, some attention should be paid also to the bulk reaction of NO and O2 occurring according to the equation: V = k [NO]2 [O2]; k(aq) = 2 × 106 M−2 s−1, at 25°C [88] a process that is faster in hydrophobic environments given the increase in the concentration of both gases [5]. Despite complexity, an apparent KI = 60 nM at [O2] = 30 µM has been measured using respiring synaptosomes [44]. Thus, as

14


outlined by Brown [89], one interesting conclusion is that in the presence of nM concentrations of NO (i.e., a condition that is fully compatible with in vivo physiological NO fluxes), there might be a finite amount of inhibited CcOX. The results of fluorescence microscopy experiments aimed at measuring the mitochondrial membrane potential of cultured neuroblastoma cells support this contention. In the presence of NOSs inhibitors, such as the 7-nitroindazole or the L-nitroso-arginine, the import of rhodamine is significantly higher than in their absence (Figure 1.1), demonstrating that, at least in cultured cells, a measurable NOS-mediated inhibitory effect occurs on cell respiration. Together, these observations account for the fact that the apparent Km of CcOX for O2 measured in tissues (≥ 1 µM) is higher than that determined in vitro (0.1 µM), with the purified enzyme. By using the oversimplified reaction mechanism including CcOX in turnover with reduced cytochrome c and O2, the apparent KM,O2 rises in the range measured in vivo [90]. Another interesting aspect of this issue, discussed by Moncada and Erusalimski [9], is the possible cross talk between CcOX and guanylyl-cyclase (GC). Namely, at 10 µM O2, one may predict that 20 nM NO would inhibit 50% of CcOX; interestingly, half activation of GC by NO is also achieved at 20 nM [91]. Thus, both enzymes sense NO with comparable affinities but opposite effects (i.e., activation for GC and inhibition for CcOx); the cross talk between the two pathways appears likely, but is still obscure and needs to be further investigated. Which one of the two NO inhibition mechanisms may predominate? By probing purified CcOX in turnover at different concentrations of reductants [83], it was found that at high reductants the nitrosyl-derivative accumulates, whereas at low reductants the nitrite-derivative is preferentially formed (Figure 1.8) [83, 92]. Amperometric measurements performed using mammalian CcOX demonstrated that in all cases, and regardless of the redox state of the enzyme, NO binds to the active site in a 1:1 stoichiometry (Figure 1.9) [61], apparently making less likely the possibility of forming N2O. Studies of the chemical modifications induced by NO in tissues/organs as well as in vivo have also been performed. In these cases, the direct assignment to a given chemical change induced by NO into any of the respiratory chain complexes might be difficult, unless a stable type of adduct is formed, such as S-nitrosothiols and nitro-tyrosines, the formation of which has been reported for only complex I so far [93]. Simultaneous measurements of cell respiration and spectral perturbation of the respiratory complexes are most informative [83, 94]. The redox changes of the mitochondrial cytochromes can be detected by multi-wavelength visible spectroscopy and correlated at given concentrations of O2 and exogenous NO (by NO-releasers) to simultaneously measured changes in O2 consumption. This methodological approach appears promising to elucidate the mechanism(s) by which endogenous NO produced by transiently activated constitutive NOS (Ca++ transient) controls cell respiration. Fluorescence microscopy [41, 42] and O2 polarography (respirometry) [83, 92] experiments have been successfully performed to study the functional effects of the reaction between NO and CcOX.

Nitric Oxide Controls Cell Respiration +

+

+ a32 CuB+NO (last)

0.2

400

500 (nm)

600

a33 CuB2+ (first) 0.4 Absorbance

Absorbance

+

a33 CuB2 (first)

0.4

0

15

+

a33 NO2−CuB+(?) (last)

0.2

0

400

500 λ (nm)

600

FIGURE 1.8 Spectral changes induced by NO on cytochrome c oxidase in turnover at high (left panel) and low (right panel) reductant concentration. The experiment was performed by using a sequential mixing stopped-flow, thus avoiding incubation of NO with reducing agents; the sequential mode allows the rapid (∼ 1 ms) premixing of NO with chosen concentrations of reductants, followed (≥ 10 ms) by mixing with oxidized CcOX. At higher reductant concentration, the fully reduced NO-bound enzyme is formed, whereas at low reductant concentration, the nitrite-inhibited enzyme is populated. The question mark on CuB outlines that the redox state of the metal when nitrite is in the site is still unknown. (Modified from Sarti et al., Biochem. Biophys. Res. Comm. (2000) 274:183–7.)

The former approach provides information on the membrane potential set across the mitochondrial membrane; its use is particularly helpful in cytology as it allows measurements on living cells. Respirometry also may provide information on intact cells, but most importantly yields insight into the NO inhibition mechanism prevailing under a given experimental condition; it is simple, powerful and will therefore be illustrated more in detail. In a typical polarographic experiment, purified CcOX, mitochondria, or cells are allowed to respire on reducing substrates in a reaction chamber monitoring amperometrically both O2 and NO. As schematically depicted in Figure 1.10, when low (µM) NO is added to the system, respiration is abolished until NO is either consumed or scavenged (typically by HbO2). In the absence of free NO, two different respiration recovery patterns can be observed. In the presence of reductants and O2 (i.e., as in the cell or in vivo), the nitrite-derivative recovers immediately, whereas the nitrosyl-derivative recovers more slowly at the NO off-rate from the active site (k ≈ 0.004 s−1 at 20°C). Thus, if nitrite-CcOX is formed, upon addition of the NO scavenger respiration reactivates immediately; on the contrary, if the nitrosyl-CcOX predominates, recovery is slow and appears autocatalytic [83, 92]. The light sensitivity of the nitrosyl-derivative of CcOX may add additional information, since illumination by increasing the rate of dissociation of NO from the nitrosylated site induces the prompt recovery of respiration (Figure 1.10). This very simple protocol allows us to distinguish between the two pathways and has been used with detergent-solubilized CcOX [83], mitochondria, and cells [92]. Interestingly, cultured neuroblastoma (SY-5Y) cells revealed a prompt recovery of respiration after addition of HbO2, the NO scavenger, indicative of nitrite

16


O

F

P

[NO] 0.2 µM

R

[NO] 4 µM

5 Time (min)

FIGURE 1.9 Amperometric determination of the NO to cytochrome c oxidase binding stoichiometry. Experiments have been performed anaerobically, using the ISO-NO World Precision Instrument apparatus equipped with a 2-mm NO-sensitive electrode. The protein at suitable concentration (0.2 and 4 µM for O, P, F, and R, respectively) is added (arrows). Notice that, even in the presence of large excess reductants, the binding of approximately 1 NO/CcOX functional unit is followed by a drift with a slope that is identical to that recorded before CcOX addition, ruling out enzymatic NO degradation. (Modified from Giuffrè et al., Biochemistry (2000) 39: 15446–53; Giuffrè et al., Proc. Natl. Acad. Sci. USA (1999) 96: 14718–23.)

formation. On the other hand, in the presence of artificial (externally added) reductants able to increase the electron transfer flux through the respiratory chain, formation of the nitrosyl-derivative was strongly enhanced, with the appearance of the slow and light-sensitive recovery from inhibition (Figure 1.11). This finding has been interpreted as indicative of a particular attitude of these cells to safely dispose of NO at the mitochondrial level, by degrading NO to nitrite. Coupled rat liver mitochondria (RCR = 4 − 6) consuming O2 in the presence of standard concentrations of reductants (mM), such as malate and succinate, also appear to degrade NO to nitrite, particularly if under state 4 conditions. Whether mitochondria are more susceptible to nitrosylation, and thus more severely inhibited by NO, in state 3 than in state 4 still needs to be clarified [92, 95]. In synthesis, two principal reaction pathways can be observed, and the prevalence of one over the other can make the difference in terms of persistence of mitochondrial inhibition by NO. This conclusion is based on the consideration that the scavenging of free NO from the bulk phase leaves CcOX in two different conditions, depending on the prevailing mechanism. The nitrite derivative recovers activity rapidly by dissociating (innocuous) nitrite. In contrast, the nitrosylated CcOX can only recover function at the rate of NO dissociation from Fe2+ of heme a3 (k ∼0.01 s−1, at 37°C [83]), although in this case, it is


17

Reductants CcOX NO

Hb

[O2 ]

Dark

Low e− flux

High e− flux

Light

Time NO

[NO]

Hb

Time

FIGURE 1.10 Investigating the NO inhibition mechanism by polarography. Typical oxygraphic (top) and nitroxygraphic (bottom) profiles observed upon inhibiting purified CcOX (or any other respiring system) by µM NO, and releasing inhibition with oxy-hemoglobin used as NO scavenger. After NO scavenging, the kinetics of recovery is either slow and lags behind NO dissociation (high electron flux through the respiratory chain) or is fast (low electron flux); further supporting this hypothesis, the light facilitates removal of inhibition by accelerating NO dissociation.

debated whether dissociated NO is released into the medium as such or as nitrite [77, 87].

1.8 EXPERIMENTAL DESIGNS The study of the chemical reactions between NO and CcOX has been made possible thanks to the development of protocols allowing the characterization of: 1. The NO/CcOX adduct formed during turnover 2. The final state of NO after reaction, whether NO radical, oxidized to NO2− or reduced to N2O

18

Nitric Oxide, Cell Signaling, and Gene Expression Neuroblastoma cells Purified oxidase

120

120

Light

Activity %

Light 80

80

40

40

Dark

Dark 0

0

5 10 15 Cytochrome c (µM)

20

0

0

1

2 3 TMPD (mM)

4

5

FIGURE 1.11 Effect of the reductant concentration on the NO inhibition mechanism. The increase of reduced cytochrome c, or TMPD (in the presence of ascorbate) favors the accumulation of the nitrosyl light-sensitive CcOX-adduct, in assays employing either purified CcOX (left panel) or cells (right panel).

The oxygraphic assay just described allows for the collection of indirect but meaningful evidence on the accumulation of either the nitrosyl or the nitrite derivative of CcOX, turning over in the presence of NO. As already pointed out this protocol is particularly useful when dealing with cells and mitochondria, which are very difficult to analyze by spectrophotometry. Multi-wavelength stopped-flow spectroscopy proved very useful in identifying the intermediate state (adduct formed) in the reaction between detergentsolubilized CcOX and NO [83]. This approach allowed us to assign transient optical signals to either the nitrosyl- or the nitrite-derivative, by comparing experiments performed under high or low reductive pressure, mostly using ruthenium hexamine, as efficient as cytochrome c in reducing CcOX, but spectroscopically silent. In a typical experiment, oxidized N2-equilibrated CcOX is mixed in a diode-array stopped-flow with a solution containing large excess ascorbate and variable ruthenium hexamine. Figure 1.8 illustrates the spectral changes and the time courses observed at two extreme concentrations (high and low) of ruthenium hexamine. In the absence of NO, the enzyme becomes half-reduced (with heme a reduced) at steady state and eventually becomes fully reduced upon O2 exhaustion [83]. In the presence of NO, in small excess over CcOX, two different behaviors can be detected: at high reductant concentration, the enzyme becomes fully reduced nitrosylated (Figure 1.8, left panel), while at low reductant concentration (Figure 1.8, right panel) after approaching steady state, the enzyme transiently displays the features of the nitrite-bound derivative. The reduction of NO to N2O catalyzed by some bacterial heme-copper oxidases has been clearly detected [96–98]. Contrary to the mammalian (beef heart) enzyme that has no NO reductase activity [86], both the ba3 and caa3 oxidases from Thermus thermophilus and the cbb3 oxidase from Pseudomonas stutzeri proved able to reduce NO to N2O, as also demonstrated by head-space gas chromatography analysis [96]. The NO reductive degradation activity by terminal


19

oxidases can be observed under strictly anaerobic conditions both spectroscopically (Figure 1.12) and amperometrically (Figure 1.13). Measurements have to be performed (virtually) in the absence of O2 (i.e., by degassing and N2 equilibrating the sample) and in the presence of an O2 scavenging system, such as glucose-glucose oxidase, in the presence of catalase to avoid accumulation of H2O2. The spectroscopic determination consists in measuring at different time intervals the residual concentration of NO, during anaerobic incubation with oxidase and excess reductants (Figure 1.12). In more detail, one driving syringe of the stopped flow apparatus is filled with such a solution of NO and oxidase, and mixed at the appropriate times, with deoxy-hemoglobin (or myoglobin) in excess over NO. In the presence of an enzymatic NO degradation activity, a timedependent decrease of free NO is observed, detected as a decrease in the yield of nitrosyl-hemoglobin [96] (Figure 1.12, right panel). Similar results can be obtained amperometrically by directly monitoring NO in solution using a selective electrode [96–98]; such an experiment is depicted in Figure 1.13, where the NO reductase activity of purified cbb3 oxidase from P. stutzeri is compared with the activity of bona fide NO-reductase purified from P. denitrificans.

[HbFe2+NO - HbFe2+] 60

Baseline

[ΝΟ] µΜ

∆ OD

0.2

0

aa3OX

40 20

ba3OX caa3OX

0.2 500

600 λ (nm)

700

0

0

10 20 Time (min)

30

FIGURE 1.12 Reductive degradation of NO by heme-copper oxidases: spectroscopic assay. Typically, one driving syringe of the photodiode-array stopped flow apparatus is filled with an oxidase solution (mammalian or bacterial) and incubated anaerobically with NO for a suitable period of time (minutes). At appropriate times, the enzyme-NO solution is mixed with deoxy hemoglobin in excess over NO; the reaction between NO and Hb is very rapid and is complete within 100 ms, yielding the difference spectra depicted in the left panel. Notice that, at longer incubation times, the extent of the difference spectra decreases due to NO disappearance from solution. Right panel: The very slow NO degradation observed in the presence of mammalian aa3 oxidase (5 µM) is identical to baseline, and is likely due to chemical reaction with reductants. In contrast, ba3 (1.5 µM) and caa3 (0.3 µM ) oxidases from T. thermophilus bacteria degrade NO at a significant rate. (Modified from Giuffrè et al., Proc. Natl. Acad. Sci. USA (1999) 96: 14718–23.)

20

Nitric Oxide, Cell Signaling, and Gene Expression NO-reductase

[NO] 5 µM

[NO] 10 µM

cbb3OX

5

1 Time (min)

FIGURE 1.13 Reductive degradation of NO by heme-copper oxidases: amperometric assay. The NO consumption by the cbb3 oxidase (0.1 µM) from P. stutzeri (left panel) is measured anaerobically in the presence of excess reductants, ascorbate and tetra-methylp-phenylenediamine (TMPD), and is compared with the activity of bona fide NO-reductase (3 nM) purified from P. denitrificans. (Modified from Forte et al., Eur. J. Biochem. (2001) 268: 6486–91.)

1.9 PERSISTENCE OF NO IN THE MITOCHONDRION The almost incredible number of physiological actions that NO appears to have suggests that, if out of control, its interaction with biological targets may become pathological. NO in cells and tissues is freely permeable through the membranes [36, 37]. It is worth recalling that both NO and O2 are more soluble in the phospholipid membranes than in water [5, 99, and references therein]. Thus, the reaction between NO and O2 is facilitated in the membrane (hydrophobic) compartments where a decreased lifetime of NO is expected. As pointed out by Shiva et al. [5] and relevant to pathophysiology, in the presence of O2 the lipid reach environment of atherosclerotic plaques, for instance, can be responsible for a lower NO bioavailability to the mitochondrial targets (CcOX) and the close-by endothelial sites. In addition, the enhanced intramembrane reaction between NO and O2 would yield an overproduction of N2O3, an efficient NO+ donor, leading to an increase of S-nitrosated derivatives [5]. According to the two-inhibition mechanisms presented earlier, when dealing with NO and CcOX, one would expect the production of nitrite at low electron flow levels through the respiratory chain, whereas at higher levels the release of NO is expected [3]. The production of nitrite by isolated mitochondria, attributed to a mitochondrial NOS, is well documented [21, and references therein]. It is still unclear whether inhibited CcOX releases NO in the environment as such, or


21

as peroxynitrite or oxidized to NO2− [77, 87]. This is not a trivial point because all these species, including the nitroxyl anion (NO−) as recently put forward by Shiva et al. [100], can participate in reactions relevant to mitochondrial metabolism and more, in general, to cell/tissue physiology or pathology [12]. Some apparently futile reactions, such as NO binding to and release from reduced CcOX, or the back production of NO from oxidized products, may lead to a stationary persistence of NO in the cell/mitochondrion environment. In addition, the reaction of NO with ubiquinol (QH2) recycles NO [5, 12, 101]. This reaction generates ubisemiquinone (Q.) and a nitroxyl anion (NO−), which is cytotoxic and elicits biological responses similar to those of NO and peroxynitrite [100]. Interestingly, the nitroxyl anion can also readily be converted back to NO by metalloproteins [12, 101, 102]. If released in excess and persistent in the mitochondrial environment, NO upon increasing of superoxide ions (O2−) would react forming ONOO−, a powerful oxidizing agent detrimental not only to mitochondrial complexes [103, 104], but also for membranes, many other proteins, and nucleic acids. The reactions of ONOO− with all putative targets and the role played by mitochondria is a matter of intensive investigation [102, 105, 106, and references therein], but it is outside the scope of this review. In addition, peroxynitrite can be back reduced to NO by ascorbate; whether the conversion of ONOO− to NO demands or not the presence of CcOX still needs to be clarified [107]. The presence of NO is, indeed, of physiological relevance in the proximity of the endothelium, where a higher NO tension is required to maintain the correct vessel tone [15]. Particularly under chronic hypoxic conditions such as those characteristic of neurodegeneration, however, a detrimental cycle could be triggered by NOSs activation because the NO released, by further inhibiting respiration, might worsen the hypoxic effects. As outlined by Moncada and Erusalimski [9], the persistent blockage of CxOX by NO may induce different responses of patho-physiological relevance; depending on the type of cell facing the NO pulse, whether able or unable to promptly activate glycolysis (e.g., astrocytes or neurons, respectively), a twostage time response has been proposed [9, 108, 109]. The initial event is the efficient inhibition of the respiratory chain by NO, and the accumulation of reducing equivalents leading to an overproduction of superoxide ions. In the first stage, the mitochondrial superoxide dismutase would generate hydrogen peroxide at a concentration level compatible with cell signaling and physiology; at this stage, a correct balance between the radical species NO and O2− is set, and production of highly reactive (toxic) peroxynitrite is minimal. In contrast, in the second stage, the persistence of CcOX inhibition should lead to the accumulation of peroxynitrite, thereby triggering a more stable modification (inactivation) of the respiratory chain components, including the S-nitrosation of complex I, and activating cell responses such as apoptosis and necrosis. From the bioenergetic point of view, one key event seems to be the prompt activation of glycolysis, with production of ATP used to maintain the mitochondrial membrane potential and prevent cell death [109]. Consequently, parallel experiments were performed exposing nervous cells to pulses of exogenous NO (DETA-NO), causing no

22


significant toxicity in astrocytes but irreversible damage of neurons [110]. Among the numerous issues still to be elucidated, the important question regarding the NO concentration level that needs to be maintained in the cell remains to be addressed, including the length of time required to shift from a physiological to a pathological type of cell response.

1.10 ACKNOWLEDGMENTS The authors thank the Ministero dell’Istruzione, dell’Università e della Ricerca (MIUR) of Italy (PRIN “Bioenergetica: genomica funzionale, meccanismi molecolari e aspetti fisiopatologici” and FIRB, RBAU01F2BJ_001) for support.

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103. Cooper CE, Davies NA, Psychoulis M, Canevari L, Bates TE, Dobbie MS, Casley CS, Sharpe MA. Nitric oxide and peroxynitrite cause irreversible increases in the K(m) for oxygen of mitochondrial cytochrome oxidase: in vitro and in vivo studies. Biochim. Biophys. Acta. (2003) 1607:27–34. 104. Cooper CE, Davies NA. Effects of nitric oxide and peroxynitrite on the cytochrome oxidase K(m) for oxygen: implications for mitochondrial pathology. Biochim. Biophys. Acta. (2000) 1459: 390–396. 105. Radi R, Cassina A, Hodara R, Quijano C, Castro L. Peroxynitrite reactions and formation in mitochondria. Free Radic. Biol. Med. (2002) 33:1451–1464. 106. Beckman JS. Protein tyrosine nitration and peroxynitrite. FASEB J. (2002) 16:1144. 107. Barone MC, Darley-Usmar VM, Brookes PS. Reversible inhibition of cytochrome c oxidase by peroxynitrite proceeds through ascorbate-dependent generation of nitric oxide. J. Biol. Chem. (2003) 278:27520–27524. 108. Gegg ME, Beltran B, Salas-Pino S, Bolanos JP, Clark JB, Moncada S, Heales SJ. Differential effect of nitric oxide on glutathione metabolism and mitochondrial function in astrocytes and neurones: implications for neuroprotection/neurodegeneration? J. Neurochem. (2003) 86:228–237. 109. Almeida A, Almeida J, Bolanos JP, Moncada S. Different responses of astrocytes and neurons to nitric oxide: the role of glycolytically generated ATP in astrocyte protection. Proc. Natl. Acad. Sci. USA (2001) 98: 15294–15299. 110. Beltran B, Mathur A, Duchen MR, Erusalimsky JD, Moncada S. The effect of nitric oxide on cell respiration: A key to understanding its role in cell survival or death. Proc. Natl. Acad. Sci. USA (2000) 97:14602–14607.

Nitric 2 Mitochondrial Oxide Signaling in Synaptic Plasticity and Cell Death Alberto Boveris, Silvia Lores-Arnaiz, Juanita Bustamante, and Analía Czerniczyniec University of Buenos Aires, Buenos Aires, Argentina

CONTENTS 2.1 2.2 2.3 2.4 2.5

Introduction ...............................................................................................29 The Mitochondrial Production of NO and the NO Effects in Mitochondria ....................................................................................30 NO and Neuronal Plasticity......................................................................34 NO and Apoptosis.....................................................................................36 Conclusion.................................................................................................38 References .................................................................................................39

2.1 INTRODUCTION Nitric oxide (NO) is a free radical, originally described as the endothelial relaxation factor, which is now considered as an intercellular messenger in physiological processes, such as vasodilation and regulation of blood flow, neurotransmission, plaquetary aggregation, and inmunological response, as well as an intracellular regulator of mitochondrial respiration. NO is synthesized from L-arginine, NADPH, and O2, by the isoenzymes of the NO synthases (NOS) family. Three isoforms have been cloned and characterized: nNOS (NOS1, neuronal NOS, originally identified as a constitutive form in neuronal tissue), iNOS (NOS2, inducible NOS, originally identified as a form inducible by cytokines in macrophages and hepatocytes), and eNOS (NOS3, endothelial NOS, originally identified as a constitutive form in vascular endothelial cells) (1, 2). A large series of cellular enzymatic activities is affected by NO concentrations in the range of the physiological levels. Considering the diffusion properties of NO, which equals a few cell diameters (3), it is difficult to set the limits between intracellular and 29

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intercellular NO effects. Guanilate cyclase and cytochrome oxidase, in both cases with NO binding to an iron of the enzyme active center, are two metabolically important enzymes that are physiologically regulated by NO. In the first case, enzyme activation affords the mechanism for intercellular communication, and in the second case, the respiratory inhibition by NO generated intracellularly or in a neighbor cell appear as equally likely. The recognition of a NO synthase (NOS) localized in the mitochondria (mitochondrial NOS [mtNOS]) prompted the concept of NO-mediated intraorganellar regulation of mitochondrial function. The intracellular localization of the NOS isoforms, with the already known bipolar distribution in mitochondria and cytosol (including in this latter the NOS isoforms of caveolae and plasma and endoplasmic reticulum membranes) as well as the cellular levels of substrates and Ca2+, forms the molecular basis of the NOdependent regulatory mechanisms. The cellular conditions in which NO diffuses from mitochondria to cytosol, as well as the conditions in which NO diffuses from cytosol to mitochondria, are the current questions for the complex processes of intracellular signaling that modulate the cell cycle and the cellular responses in almost all types of tissues. This chapter discusses the processes and molecular mechanisms associated with neuronal dynamic plasticity and the cell death pathway in Ca2+-induced thymocyte apoptosis, two processes in which mitochondrial NO exhibits a regulatory role.

2.2 THE MITOCHONDRIAL PRODUCTION OF NO AND THE NO EFFECTS IN MITOCHONDRIA In the early 1990s, the extraordinary research activity on biological NO production identified the three classic genomic NOS (nNOS, iNOS, and eNOS). Concerning the biologically important intracellular localization of these enzymes, immunoytochemical assays with antibodies against eNOS gave the first evidence of a NOS located in mitochondria (4–6), which was named mtNOS and advanced as a regulator of mitochondrial and cellular respiration (6). The determination of mtNOS enzymatic activity was elusive for a few years, until two independent groups, Ghafourifar and Richter (7) working in Zurich and Giulivi and co-workers located at Los Angeles and Buenos Aires (8), succeeded in demonstrating NO production by rat liver mitochondria and mitochondrial fragments. The original observations in liver mitochondria were later extended to brain (9,10), heart (11, 12), thymus (13), kidney (14), and diaphragm (15) mitochondria, and the skepticism on mtNOS as a contamination produced during cell fractionation was largely dissipated. Mitochondrial NO production is performed by mtNOS, a classic NOS in biochemical terms, that requires NADPH (KM = 15 µM), arginine (KM = 12 to 60 µM), O2 (KM = 37 to 73 µM), and Ca2+ for enzyme activity. Calmodulin, tetrahydrobiopterine, and thiols increase the enzymatic activity (16). The intramitochondrial concentrations of NADPH, arginine, O2, and Ca2+ are in excess or in the range needed for enzymatic activity. The methodology currently used to determine mtNOS activity

Mitochondrial Nitric Oxide Signaling in Synaptic Plasticity and Cell Death

31

has been recently evaluated (17); mtNOS activity provides rates of 0.30 to 0.90 nmol NO/min.mg protein in mitochondria and submitochondrial preparations. In a pivotal contribution, Giulivi and co-workers (18) sequenced the 1429 amino acids of rat liver mtNOS, and found it identical to nNOS splice variant, α, which was mirystoylated and phosphorylated in post-translational processes. Transcripts corresponding to nNOSα were found in liver, brain, heart, kidney, skeletal muscle, lung, testis, and spleen (18). It is then clear that mtNOS is a constitutive protein of the inner mitochondrial membrane with its substrates present in the mitochondrial matrix. Interestingly, a whole series of physiological situations were reported to affect the level and activity of mtNOS: They were upregulated during brain development (19) by cold exposure (20) and by chronic hypoxia (21), and down-regulated by thyroxine (22) and angiotensin II (23). Pharmacological treatments with enalapril, the converting-enzyme inhibitor, were effective in increasing mtNOS activity (12, 14, 23) and administration of haloperidol and chlorpromazine decreased brain mtNOS activity (9, 24). The intramitochondrial NO steady-state concentrations were initially calculated in the relatively low side of 50 nM NO from the rates of NO production by submitochondrial membranes (25). The recent recognition of different rates of mitochondrial NO release in states 4 and 3 (resting state without ADP and active state with ADP, respectively) and of the role of membrane potential in upholding a high mtNOS activity (26) led to the high side, at 200 nM, of the estimation for the intramitochondrial steady-state NO levels. The higher value emphasizes in the NO diffusion from mitochondria to cytosol as a physiological process. In the tissues under physiological conditions, with a steady-state oxygenation of 20 to 40 µM O2, an intramitochondrial steady-state concentration of 200 nM NO corresponds to an inhibition of cytochrome oxidase of 26 to 42% (27). Diffusion to the surrounding medium (equivalent to cytosol) of 29 nM NO was electrochemically determined after supplementation of a single mitochondrion with Ca2+ (11). NO and O2•− metabolism in the mitochondrial matrix are linked by the very fast and diffusion-limited reaction between NO and O2•− to produce peroxynitrite (ONOO−). The oxidative utilization of NO is the main (70 to 80%) pathway of NO metabolism, and at the same time, it provides about one half (50%) of intramitochondrial O2•− utilization. The reductive utilization of NO by ubiquinol and cytochrome oxidase provides a minor (20 to 30%) but effective pathway of NO catabolism (24). Figure 2.1 illustrates a scheme of the integrated mitochondrial metabolism of O2•− and NO, and remarks on the diffusion of NO and H2O2 from mitochondria to cytosol. The effects of NO on mitochondrial electron transfer were recognized in 1994 by two British research groups that reported the inhibition of brain and muscle cytochrome oxidase (complex IV) activity by low NO concentrations, in a reversible and O2-competitive biochemical process (28, 29). The observation was rapidly confirmed by other groups using NO-donors and pure NO in liver, heart, and brown fat mitochondria (30–33). NO levels of 0.05 to 0.10 µM decreased cytochrome oxidase activity and mitochondrial respiration to one-half. The inhibition

32


QBP NADH–DH

UQH mtNOS

Mn–SOD 10–30 µM

O2– 0.2–0.3 nM

NO 100–300 nM NO 0.8–1.3 nmol/min

H2O2 50–100 nM

ONOO– 5–10 nM

H2O2 0.1–1.0 nmol/min

FIGURE 2.1 Mitochondrial NO metabolism. The concentrations below the chemical symbols and abbreviations correspond to physiological steady-state concentrations and levels. The rates indicate that, below NO and H2O2, outside mitochondria correspond to the diffusion from mitochondria to cytosol, expressed in nmol/min.mg mitochondrial protein. (NADH-DH, NADH-dehydrogenase; UBP, ubiquinone binding protein.)

is reversible by dilution, hemoglobin addition, or exposure to O2•−. The O2competitive inhibition of cytochrome oxidase makes the inhibition more marked at low O2 concentrations. A detailed mathematical model advances a set of equations that explain the known facts of the NO mediated inhibition of cytochrome oxidase and cell respiration (34). Ubiquinol-cytochrome c reductase (complex III) is a second NO-sensitive point in the mitochondrial respiratory chain (31, 32); half inhibition of electron transfer between cytochromes b and c occurs at 0.1 to 0.2 µM NO, enhancing the production of O2•− and H2O2 in submitochondrial particles and in mitochondria (32). A third NO-sensitive point is located at NADH-dehydrogenase (complex I); in this case, ONOO− appears as the effective inhibitory agent (35–36). Figure 2.2 illustrates the direct and indirect effects of NO in the mitochondrial respiratory chain, in the latter case through ONOO− formation. The reversible effects of NO on cytochrome oxidase and ubiquinol-cytochrome c reductase are two parts of the regulation of mitochondrial respiration by NO: on one hand, the inhibition of cytochrome oxidase and on the other, the provision of O2•− to remove or decrease the respiratory inhibition because it is a physiological need with high steady-state concentrations of NO (32), as in ischemia-reperfusion and in inflammation. The irreversible effects of ONOO- on complexes I and III are related to situations in which sustained high levels of ONOO− lead to mitochondrial dysfunction.

Mitochondrial Nitric Oxide Signaling in Synaptic Plasticity and Cell Death ONOO– irreversible 50–100 nM

NO reversible 50–100 nM

c

I

UQH2

e–

e–

F

e–

F e– Cu–Fe

O2

IV

H2O

III

UQ II

Succinate NADH

33

NO irreversible 100–200 nM ONOO– irreversible 50–100 nM

FIGURE 2.2 Sites of action of NO in the mitochondrial respiratory chain. Direct effects of NO and indirect, through ONNO formation, are indicated. The concentrations below the statement of reversible or irreversible effect correspond to the effector level that produces half-maximal inhibitory effects.

Concerning the important question if mtNOS enzymatic activity regulates mitochondrial functions under physiological conditions, Giulivi and co-workers described the modulation of mitochondrial O2 uptake and H2O2 production by the activity of mtNOS (37–39) in isolated mitochondria, later confirmed by others (14, 23). The current view is that mtNOS does regulate mitochondrial respiration (40) and that mtNOS, cytochrome oxidase, and F1-ATPase are the three regulatory proteins of cellular O2 uptake and energy production. The biochemical activity of mtNOS (expressed as nmol NO/min.mg protein) is usually measured as the difference in NO production between the assay performed in the presence of the substrate arginine and in the presence of a competitive inhibitor of NOS as NMMA or NNA (17). The regulatory activity of mtNOS is determined in isolated mitochondria by the difference in the rates of O2 uptake or H2O2 production between the system supplemented with arginine and superoxide dismutase, which provides the highest NO levels, and the system in the presence of a NOS competitive inhibitor and hemoglobin, which provides the lowest NO level (14, 23). The regulatory activity of mtNOS is expressed as percentage of state 3 respiration (usually 15 to 40%) or as the NO-dependent inhibition of respiration (usually 25 to 65 ng-at O/min.mg protein). The regulatory capacity of NO, estimated as the ratio of the rates of regulated O2 uptake/NO production or of regulated H2O2 production/NO production, is approximately 70 for O2 uptake and approximately 0.4 in H2O2 production (14, 23). The activity of mtNOS in isolated mitochondria markedly increased the [O2 ]0.5 of mitochondrial

34


oxygen uptake from 1.2 to 1.6 µM O2 in the case of inactive mtNOS, to 2.8 to 3.4 µM O2 in the case of an active mtNOS (24, 37, 41). However, it is not clear to what extent these effects, observed with isolated mitochondria, occur under physiological conditions in myoglobin-containing tissues and in blood perfused organs, due to the high affinity of the two previously mentioned hemoproteins for NO (42).

2.3 NO AND NEURONAL PLASTICITY Plasticity is the ability of the brain to reorganize neural pathways based on new experiences. Changes in synaptic ultrastructure, dendritic ultrastructure, and neuronal gene expression may contribute to molecular mechanisms of synaptic plasticity (43). It has been described that brains of rats reared in enriched environments have a higher number of synapses by neuron, greater size of synaptic connection zones, more dendritic density, higher proportion of glial tissue by neuron, and higher production of neurotrophic factors than those of animals reared in impoverished environments (44). NO has been implicated in an increasing number of experimental models of plasticity in the formation of long-term memory (LTM) and in spatial learning in rats (45, 46). Brain mtNOS activity was observed in rat and mouse mitochondria (9, 10, 24) and analyzed by Western blot, which mainly consisted of a 147kDa nNOS, but also had a minor quantity of eNOS (47). Brain mtNOS activity is regulated by physiological and experimental conditions: We have demonstrated that mtNOS activity is inhibited by central nervous system (CNS) drug treatment such as the antipsychotics haloperidol (9) and chlorpromazine (24). A different and complementary sequential expression of mtNOS and nNOS in developing rat brain was reported (19). In the nervous system, glutamatergic neurotransmission through NMDA receptors increases intracellular Ca2+ concentrations activating nNOS and NO stimulates soluble guanylate cyclase and increases intracellular cGMP levels. (48). The NO-cGMP signaling pathway is of importance in the formation of longterm potentiation (LTP), a model of synaptic plasticity involving the persistent enhancement of excitatory neurotransmission (46), and also in the formation of long-term memory (LTM) and spatial learning in rats (45). NO has been proposed as a retrograde messenger during induction of LTP in the CA1 region of the hippocampus and is thought to be necessary for growth of nerve fibers because nNOS is enriched in regenerating axons (49). Deletion of the genes that encode nNOS and eNOS isoforms reduce the inducibility of LTP, and selective NOS inhibitors impair spatial learning (50). The aging process is characterized by an acceleration of the rate of loss of hippocampal neurons and dendritic branching (51), a decrease in hippocampal plasticity (52), and increased cognitive deficits, particularly spatial learning and working memory (WM) deficits. Because aging weakened WM function, we tested the hypothesis that an enriched environment can preserve animals from the memory impairment associated with aging. This protective effect exerted


35

by exposure of the animals to enriched environments may be mediated by an increased neuronal plasticity through NO-dependent mechanisms. Female rats were randomly assigned at weaning and for all their lives to one of two rearing conditions: an enriched environment or a standard wire cage. In the enriched environment condition, animals were housed in groups of 5 rats, in cages of 50 × 98 × 54 cm, furnished with a large cord pending for the top of the cage, inclined surfaces, and suspended bridges (44). In the standard cage condition, animals were housed in groups of 3 to 4 rats in cages of 29 × 21 × 34 cm. At 10 weeks of age, both groups were trained in spatial WM in an 8-arm radial maze with 8 extra-maze cues. Old rats of 27 months of age from an enriched environment performed better in spatial WM, with a significantly higher (66%) percentage of days in keeping the WM criterion than standard reared rats (Table 2.1). These results support the concept that the exposure of animals to enriched environments markedly attenuates the effects of aging on spatial WM performance. Animals exposed to enriched environments also had a NOS activity both in the cytosolic and mitochondrial compartments (155% and 73%, respectively), as compared with rats from the standard cages (Table 2.1). The observation associates NO levels with synaptic plasticity and with the prevention of spatial cognition impairment during aging. In addition, mtNOS protein expression was markedly higher in brain mitochondria from enriched-housed rats than in standard-cage animals, indicating an up-regulation of the mitochondrial enzyme by the increased neurological activity associated with the enriched environments (Figure 2.3). These results suggest that NO takes part in the mechanisms involved in spatial WM development and maintenance. Changes in mitochondrial gene expression have been reported as involved in developmental neuroplasticity; a subset of mitochondrial genes regulated by visual experience in the first month of cat life was identified. Several genes— ATPase 6, cytochrome b, and NADH dehydrogenase subunits 4 and 2—were elevated in normal cats at 5 weeks and in dark-reared cats at 20 weeks (plasticity genes); others, such as NADH dehydrogenase subunits 3 and 5, were the opposite (anti-plasticity genes) (53). Decreased electron transfer in mitochondrial complex I was reported in the substantia nigra of patients with Parkinson’s disease (54), and a decreased expression of the subunit 4 of NADH dehydrogenase was observed in the hippocampus, inferior parietal lobule, and cerebellum of Alzheimer’s disease patients (55). A reduction in complexes I and IV of the mitochondrial respiratory chain has been reported in association to the aging process, whereas complexes II and III were unaffected (56, 57). We have observed an increased complex I activity in association with neuronal plasticity (Table 2.1), a fact that supports the idea that the higher neurological activity associated with enriched environments prevents the loss of activity of mitochondrial respiratory complexes in aging and neurodegenerative diseases.

36


TABLE 2.1 Spatial Working Memory and Brain Enzyme Activities of Rats Kept in Enriched Environments (Enriched) and Standard Reared Rats (Control) Condition Spatial working memory (%) Cytosolic nNOS activity (nmol/min.mg) mtNOS activity (nmol/min.mg) Complex I activity (nmol/min.mg)

Control 41 ± 11 0.31 ± 0.06

Enriched 68 ± 7* (66 %) 0.79 ± 0.10** (155%)

0.30 ± 0.04 19 ± 2

0.52 ± 0.12* (73%) 34 ± 8 (79%)

In parentheses, the enriched-environment associated percentages of parameter increase. Spatial WM is expressed as the percentage of days that the animals kept WM in the 8-arm maze, with a maximum of one error in 5 days, over the total number of training days. Enzyme activities, mtNOS, and complex I, were determined as reported (47) and expressed in nmol substrate/min.mg protein. *p < 0.05; ** p < 0.01 (Student’s t test). Source: From Lores-Arnaiz et al., Mol. Aspects. Med. 25: 91–101 (2004).

Control

Enriched

Amino terminus

147 kDa

1

11

Carboxy terminus 147 kDa

1

47

FIGURE 2.3 Western blot analysis of the mtNOS of brain mitochondrial membranes from standard (control) or enriched-reared animals of 27 months of age. The anti-nNOS antibodies against amino and carboxy terminal groups were from Santa Cruz Laboratories, Santa Cruz, California.

2.4 NO AND APOPTOSIS NO has been frequently referred as an intracellular regulator of cell death programs in biological systems; however, both pro- and anti-apoptotic roles have


37

been assigned to NO (58, 59), likely by the different type of cells used and to the different NO steady-state concentrations to which cells were exposed. Thymocytes constitute one cell type in which ample experimental evidence indicates a fast execution of the apoptotic process (60–62). The specialized NOS of mitochondria, mtNOS, has been recognized as the first enzyme with activity that is markedly increased during Ca2+-mediated thymocyte apoptosis (13). In a recent study (63), we analyzed the kinetics of a series of biochemical and cellular events in thapsigargin-induced thymocyte apoptosis in terms of the times required to observe half-maximal effects (t½), which ordered from lower to higher indicate the sequence of occurrence in the execution of the mitochondria-dependent cell death program (Figure 2.4). The process is triggered by the sudden increase in cytosolic Ca2+ (t½ = 2 min) produced by thapsigargin addition. The increase in Ca2+ levels is associated with the activation of a series of enzymes, including the whole NOS family, and with the initiation of apoptotic and necrotic cell death programs (64, 65). The initial rise in Ca2+ levels was immediately followed by the almost parallel increases in the activity of mtNOS (t≡ = 15 min) and in the H2O2 cellular steady-state concentration (t≡ = 18 min). The similar kinetics suggests an effective inhibition of complex III activity with an increased mitochondrial production of O2•− and H2O2. The raised H2O2 cellular level provides a molecular mechanism for a Fenton-type increased hydroxyl radical formation and an accelerated lipoperoxidation process that was detected by the increased cellular levels of the stable products, ROOH (t½ = 27 min) and TBARS (t≡ = 30 min). The increased lipid oxidation was accompanied by membrane damage and mitochondrial dysfunction with loss of respiratory control, inner membrane depolarization, and cytochrome c release (t≡ = 101, 129, and 133 min, respectively). The described phenomena, which involved mitochondria and mitochondrial products with their diffusion and effects in the cytosol, extend from thapsigargin addition (time 0) to the period of cytochrome c release (133 min). Endoplasmic reticulum underwent processes that were initiated following the earlier described mitochondrial changes: a markedly increased NO production by the endoplasmic reticulum NOS (erNOS), which has been identified as an eNOS (13) (t≡ = 48 min), the transcription of UDP-GT mRNA (t≡ = 52 min), and the increased UDP-glucosyltransferase (UDP-GT) activity (t≡ = 187 min). The protein of this enzyme behaves as a chaperone in the quality control of protein folding and is associated with the response to various cellular situations. The UDP-GT response appears limited and overwhelmed by the massive oxidative and nitrosative stress initiated by a high cytosol Ca2+ level. It is apparent that an intracellular cross talk between mitochondria and endoplasmic reticulum, with Ca2+ and NO as the intracellular signals kept at high levels, constitute the essential feature of the execution of the mitochondriadependent cell death program, which is induced by thapsigargin. The combined contribution of mitochondria and endoplasmic reticulum keeps a high cellular NO level, detected as the DAF-2-NO adduct (63), and estimated as 350 to 500 nM NO in the period from 1 to 4 h after thapsigargin addition. The cross talk between mitochondria and endoplasmic reticulum is necessary for apoptosome

38

Nitric Oxide, Cell Signaling, and Gene Expression Respiratory control mtNOS

Membrane potential

Ca2+

10

NO 8

Cvt c content

erNOS

UDP-GT Caspase 3

6 ROOH 4

DNA cleavage

H2O2 TBARS UDP-GT mRNA

2

0 10

100 Time (min)

2

15 18

27 30

46 52 49

101 133 180 210 260 125

FIGURE 2.4 Time course of thapsigargin-induced thymocyte apoptosis in terms of the times for half-maximal effects (t≡), indicated in the lower panel (in minutes), and with the indication of the quantitative changes of the considered parameter (expressed as times that the 0 time parameter was found increased [dark gray bars] as indicated by the ordinate scale). The decreases in mitochondrial respiratory control, membrane potential, and cytochrome c content are indicated as percentages of time 0 values (empty bars indicate the 100% reference values; the internal gray bars indicate the decreased activity). The time scale is logarithmic.

assembly and for the activation of caspases and proteases, both processes being the key steps for the irreversible phase of apoptosis (62). The effective inhibition of thapsigargin-induced apoptosis by NOS inhibitors, observed as an inhibition of DNA fragmentation (63) indicates that mtNOS and endoplasmic reticulum eNOS are required sources of NO for triggering mitochondria-dependent apoptosis. Another simultaneous role of mitochondria during the first and reversible phase of early apoptosis is exerted through the release of H2O2 to activate cytosolic JNK that attached to mitochondria catalyzes phosphorylation of Bcl-2 and Bclxl as well as intramitochondrial proteins (66). The process inactivates Bcl-2 and Bcl-xl and counteracts their antiapoptotic action (67), likely exerted by keeping homeostatic ion fluxes through mitochondrial membranes.

2.5 CONCLUSION The early recognition that O2•−, H2O2, and NO are able to initiate reactions harmful to cell and tissues is now complemented by the concept that the three molecules


39

are carefully regulated metabolites capable of signaling the regulatory devices of the biochemical and genetic systems of the cell. At present, the three chemical species are considered to participate in integrated processes of intracellular regulation and in intercellular signaling, communication, and cytotoxicity. The fine regulation by H2O2 of cell function was advanced by Antunes and Cadenas (68) when they demonstrated that Jurkat T-cells submitted to H2O2 levels below 0.7 µM are in a proliferative state, whereas at 1.0 to 3.0 µM H2O2 cells develop programmed cell death; at levels higher than 3.0 µM H2O2, cells undergo necrosis. H2O2 and .NO share the physical properties of being uncharged and highly diffusible through biological membranes, and, consequently, the biological adequacy for cellular and intercellular signaling. Evidence exists that H2O2 and .NO are able to modulate mitogen-activated protein kinases (MAPKs), the widespread integral components of intracellular phosphorylation and dephosphorylation signaling cascades involved in cell survival, proliferation, differentiation, and death. The interactions are complex and seem to involve intracellular glutathione and ONOO−. Both H2O2 and .NO diffusing from mitochondria to the cytosol signal a high mitochondrial energy charge.

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Nitric Oxide, Cell Signaling, and Gene Expression 46. Kemenes I, Kemenes G, Andrew RJ, Benjamin PR, O’Shea M. Critical timewindow for NO-cGMP-dependent long-term memory formation after one-trial appetitive conditioning. J. Neurosci. 22: 1414–1425 (2002). 47. Lores-Arnaiz S, D’Amico G, Paglia N, Arismendi M, Basso N, Lores-Arnaiz MR. Enriched environment, nitric oxide production and synaptic plasticity prevent the aging-dependent impairment of spatial cognition. Mol. Aspects. Med. 25: 91–101 (2004). 48. Snyder SH, Jaffrey SR, Zakhary R. Nitric oxide and carbon monoxide: parallel roles as neural messengers. Brain Res. Review 26: 167–175 (1998). 49. Gonzalez-Hernandez T, Rustioni A. Expression of three forms of nitric oxide synthase in peripheral nerve regeneration. J. Neurosci. Res. 55: 198–207 (1999). 50. Holscher C. Nitric oxide, the enigmatic neuronal messenger: its role in synaptic plasticity. Trends Neurosci. 20: 298–303 (1997). 51. Sapolsky RM. Stress, the Aging Brain and the Mechanisms of Neuron Death. MIT Press, Cambridge, MA (1992). 52. Bodnoff SR, Humphrey AG, Lehmann JC, Diamond DM, Rose GM, Meaney M J. Enduring effects of chronic corticosterone treatment on spatial learning, synaptic plasticity and hippocampal neuropathology in young and mid-aged rats. J. Neurosci. 15: 61–69 (1994). 53. Yang C, Silver B, Ellis SR, Mower G. Bidirectional regulation of mitochondrial gene expression during developmental neuroplasticity of visual cortex. Biochem. Biophys. Res. Commun. 287: 1070–1074 (2001). 54. Mizuno Y, Ikebe S, Hattori N, Nakagawa-Hattori Y, Mochizuki H, Tanaka M, Ozawa T. Role of mitochondria in the etiology and pathogenesis of Parkinson’s disease. Biochim. Biophys. Acta 1271: 265–274 (1995). 55. Aksenov MY, Tucker HM, Nair P, Aksenova MV, Butterfield DA, Estus S, Markesbery WR, The expression of several mitochondrial and nuclear genes encoding the subunits of electron transport chain enzyme complexes, cytochrome c oxidase, and NADH dehydrogenase, in different brain regions in Alzheimer´s disease. Neurochem. Res. 24: 767–774 (1999). 56. Lenaz G, D’Aurelio M, Merlo PM, Genova ML, Ventura B, Bovina C, Formiggini G, Parenti CG. Mitochondrial bioenergetics in aging. Biochim. Biophys. Acta 1459: 397–404 (2000). 57. Navarro A, Sanchez del Pino MJ, Gomez C, Peralta JL, Boveris A. Behavioral dysfunction, brain oxidative stress, and impaired mitochondrial electron transfer in aging mice. Am. J. Physiol. Regul. Integr. Comp. Physiol. 282: R985–R992 (2002). 58. Jianrong L, Billiar TR. The anti-apoptotic actions of nitric oxide in hepatocytes. Cell Death Differ. 6: 952–955 (1999). 59. Shen YH, Wang XL, Wilcken DEL. Nitric oxide induces and inhibits apoptosis through different pathways. FEBS Lett. 433: 125–131 (1998). 60. McConkey DJ, Nicotera P, Hartzell P, Bellomo G, Wyllie AH, Orrenius S. Glucocorticoids activate a suicide process in thymocytes through an elevation of cytosolic Ca2+ concentration. Arch. Biochem. Biophys. 269: 365–370 (1989). 61. Zhivotovsky B, Orrenius S, Brustugus OT, Doskeland SO. Injected cytochrome c induces apoptosis. Nature 391: 449–450 (1998). 62. Kroemer G, Dallaporta B, Resche-Rigon M. The mitochondrial death/life regulators in apoptosis and necrosis. Annu Rev Physiol 60: 619–642 (1998).


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63. Bustamante J, Di Libero E, Monti N, Fernandez-Cobo M, Cadenas E, Boveris A. Kinetic analysis of thapsigargin-induced thymocyte apoptosis. Free Radic. Biol. Med. 37: 1490–1498 (2004). 64. Clapham DE. Calcium signaling. Cell 80: 259–268 (1995). 65. Nakamura K, Bossy-Wetzel E, Burns K, Fadel MP, Lozyk M, Goping IS, Opas M, Bleackley C, Green DR, Michalak M. Changes in endoplasmic reticulum luminal environment affect cell sensitivity to apoptosis. J. Cell. Biol. 150: 731–740 (2000). 66. Schroeter H, Boyd CS, Ahmed R, Spencer JP, Duncan RF, Rice-Evans C, Cadenas E. c-Jun N-terminal kinase (JNK)-mediated modulation of brain mitochondrial function: new target proteins for JNK signalling in mitochondrion-dependent apoptosis. Biochem. J. 372: 359–369 (2003). 67. Srivastava RK, Sollott SJ, Khan L, Hansford R, Lakatta EG, Longo DL. Bcl-2 and Bcl-xl block thapsigargin-induced nitric oxide generation, c-Jun NH2 -terminal kinase activity and apoptosis. Mol. Cell Biol. 19: 5659–5674 (1999). 68. Antunes F, Cadenas E. Cellular titration of apoptosis with steady-state concentrations of H2O2: submicromolar levels of H2O2 induce apoptosis through Fenton chemistry independent of the cellular thiol state. Free Radic. Biol. Med. 30: 1008–1018 (2001).

Nitric 3 Mitochondrial Oxide and Redox Signaling Modulation of Cell Behavior María Cecilia Carreras, Soledad Galli, Daniela P. Converso, and Juan José Poderoso University of Buenos Aires, Buenos Aires, Argentina

Enrique Cadenas University of Southern California, Los Angeles, California

CONTENTS 3.1 3.2 3.3 3.4 3.5 3.6 3.7 3.8 3.9 3.10 3.11 3.12 3.13 3.14

Pathways of NO Utilization: Mitochondrial Production of Oxygen Active Species ..........................................................................................46 The Interplay between NO and H2O2 Steady-State Concentration .........49 The Very Low Oxidative Stress Level: Proliferating Effects of H2O2 ....51 Cell Cycle Arrest by H2O2 .......................................................................53 The Low-Level Oxidative Stress: p38 MAPK Cascade .........................54 The High Oxidative Stress Level: JNK and NF-κB ...............................56 The Effects of H2O2 on Cyclins ...............................................................57 NO and the Modulation of Cell Proliferation .........................................57 NO Synthases and Tumors ......................................................................58 NO, H2O2, and Cell Apoptosis ................................................................59 mtNOS and Life Processes ......................................................................61 Bioenergetics, Mitochondrial Biogenesis, and Cell Events ....................62 Conclusions and Perspectives ..................................................................64 Acknowledgments ....................................................................................65 References .................................................................................................65

Mitochondria are the central organelles in cell bioenergetics. Most of the available oxygen is consumed in the electron transfer chain and is placed in the inner membrane of the two membranes that limit the differentiated mitochondrial 45

46


compartment. Electron transfer through mitochondrial complexes I–IV is joined to proton pumping across the inner membrane creating a proton electrochemical gradient between the intermembrane space and the matrix. This gradient (~0.15 V) is dissipated by the reentry of protons through ATPase channels that couple ATP synthesis to the electron transfer activity. From a classic perspective, it is accepted that the rate of this process is regulated by O2 and substrate availability as well as ADP/ATP ratio in response to cell demands. In the last few years, significant modulatory effects of nitric oxide (NO) resulted from its high-affinity binding to cytochrome oxidase, the final electron acceptor of electron transfer chain (1). In addition, mitochondria produce oxygen active species by auto-oxidation of ubisemiquinone, a transitional intermediary redox state of membrane ubiquinol. About 2 to 3% of utilized O2 undergoes one-electron reduction by ubisemiquinone, forming superoxide anion (O2−) (Reactions 3.1 and 3.2). Most of O2− is dismutated by mitochondrial superoxide dismutase (Mn-SOD) to H2O2, which is freely diffusible to cytosol (2); in addition, mitochondrial O2− can be driven to cytosol through voltage-dependent anion channels (3). UQH− + e− → UQ−. (Reaction 3.1) UQ−. + O2 O2− → + UQ (Reaction 3.2) The ubisemiquinone pool and O2- production rate are increased by utilization of specific compounds, such as antimycin, which blocks electron flow between cytochromes b and c (2). In addition to reversible inhibition of cytochrome oxidase, NO reportedly induced inhibitory effects on the b-c1 region at complex III, leading to direct ubiquinol oxidation (4, 5). Considering that NO metabolism involves regulatory aspects on O2 uptake and O2−/H2O2 production in mitochondria, and the effects of H2O2 on gene expression and cell signaling, it is surmised that mitochondrial NO has a significant role in the modulation of life processes.

3.1 PATHWAYS OF NO UTILIZATION: MITOCHONDRIAL PRODUCTION OF OXYGEN ACTIVE SPECIES The synthesis of NO from L-Arg and O2 is catalyzed by a family of enzymes named NO synthases (NOS) (6). Three classical cytosolic isoforms have been described: neuronal NOS (nNOS or NOS I), inducible NOS (iNOS or NOS II), and endothelial NOS (eNOS or NOS III). Novel mitochondrial variants of NOS (mtNOS) have been recently described in the inner membrane of rat liver (7, 8), thymus (9), and brain (10). The roles and properties of NOS enzymes are different. nNOS and eNOS (11, 12) are constitutively expressed in different tissues. These enzymes are

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

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activated by Ca2+ pulses following stimulation of cell surface receptors by bradykinin, acetylcholine (i.e., eNOS), or excitatory amino acids, such as glutamate (i.e., nNOS) (10), to facilitate or potentiate neural transmission or to induce vasodilation in response to changes in blood flow. In contrast, iNOS is not constitutive, and Ca2+-independent. Inflammatory mediators, such as cytokines, tumor necrosis factor alpha (TNF-α), interferon γ, and LPS, activate transcription factors, such as NF-κB or AP-1, which modulate iNOS gene expression (13). In accordance with the studied tissue, mtNOS has different kinetics and immunological properties. In rat tissues, Elfering et al. (14) reported that mtNOS is identical to nNOSα with post-translational modifications, similar to acylation at N-termini and phosphorylation in Akt sensitive Ser1412 residue. Furthermore, this enzyme is constitutively expressed, Ca2+-dependent, and subjected to modulation by drugs (15) and hormones (16), or during rat brain and liver development (10, 17). Cytosolic NO is able to diffuse into mitochondria. Diffusion coefficient of NO in aqueous solution is similar to that of O2, in about 4 × 10−6cm2.s−1 (18); activation of endothelium is compatible with a concentration in the arterial wall of about 2 µM NO. Most NO binds to cytosolic compounds such as myoglobin (19, 20). Therefore, mitochondrial NO coming from canonical cytosolic NOS results considerably lower, in about 30 to 100 nM (19, 21), although it increases by tenfold after iNOS induction in experimental endotoxemia (22). In this situation, increased NO and NO-derived O2− enhance intramitochondrial peroxynitrite formation (23). New evidence supports the notion that mitochondrial NO steady-state concentration is mainly sustained by constitutive activity of mtNOS (24). Changes in the expression and activity of NOS and particularly of intramitochondrial NOS will be followed by significant variations in NO steady-state level in the mitochondrial compartment. NO effects in mitochondria are exerted at different levels. As demonstrated by Cleeter et al. (25), Brown (26), and Poderoso et al. (4, 27), NO reversibly binds to Cu2+ B center of cytochrome oxidase and consequently inhibits O2 uptake of isolated mitochondria (4, 25, 27), isolated rat heart (19), and other organs (28), as well as the whole animal (29). NO-dependent inhibition of O2 uptake is achieved at very low physiologic NO concentration; 50 to 100 nM NO inhibits by a half the activity of cytochrome oxidase (4, 27). NO inhibits electron transfer between cytochromes b and c1 in the respiratory chain at 0.3 to 0.5 µM (4, 19); prolonged exposure to 0.5 to 1 µM NO causes further inhibition of mitochondrial complex I, a hallmark of Parkinson’s disease (30), in intact cells (31) and isolated mitochondria (32). Some authors reported that NO also inhibits complex II activity, which probably depends on the time of exposure and on the mitochondrial type (33, 34). Consequently, NO renders an increase in the reduction level of the mitochondrial components, which favors the reactions of NO with ubiquinol and complex I.

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NO oxidizes ubiquinol to the respective semiquinone with a second-order rate constant of 2.3 × 103 M−1 s−1 (5, 27) (Reaction 3.1): UQH + NO → NO−+ UQH−. (Reaction 3.3) Ubisemiquinone decays through Reaction 3.2 and finally, O2− can react with NO to form peroxynitrite anion or dismutate to H2O2, catalized by Mn-SOD (Reactions 3.4 and 3.5): O2− + NO → ONOO−

(Reaction 3.4)

Mn-SOD O2− + O2− + 2H+ → H2O2 + O2 (Reaction 3.5) Considering the rate constants of Reactions 3.4 and 3.5 (2 × 1010 M−1 s−1 and 2.3 × 109 M−1 s−1, respectively, [35, 36]), O2− participation in mitochondrial metabolism will depend on NO and Mn-SOD concentrations (Figure 3.1). In agreement, supplementation of sub-mitochondrial particles with complex I or II substrates increases NO utilization by 10- to 20-fold; in the opposite situation, addition of SOD decreases NO utilization, prolongs its mean life, and increases H2O2 (27). From the described reactions, it is surmised that: 1. Most of NO decays in mitochondria by Reactions 3.4 and 3.5. 2. Depending on its concentration and on Mn-SOD level, mitochondrial NO utilization elicits a sustained production of H2O2 (37). From this perspective, the regulation of the pathways of NO utilization and mitochondrial production of NO, superoxide anion, H2O2, and peroxynitrite (Figure 3.1) has a significant participation in life processes. In the last few years, cumulative evidence indicated that the production of H2O2 and the oxidative stress level play an important role in the activation of signaling molecules that control the complex machinery involved in cell proliferation, differentiation, apoptosis, and senescence. Redox status is related to the activity of growth factors and to cell transformation and cancer. The underlying idea is that the grading expression and activity of NOS isoforms modulates H2O2 concentration and oxidative stress level. As reported in cell transformation, concomitant changes in Mn-SOD have two effects: increasing cytosolic H2O2 and prolonging NO effects on mitochondria (38) (Reactions 3.4 and 3.5). The two actions may signal for different targets and physiological or pathological responses; this assumption is in line with the fact that some effects following changes in SOD expression does not depend exclusively on produced H2O2 (39).


49

Gpx +

[H2O2]SS _

–

ONOO NO mtNOS

O2–

H2O2

Catalase Thioredoxin reductase

MnSOD

+

FIGURE 3.1 Most cytosolic H2O2 comes from mitochondria. Modulation of H2O2 steadystate concentration ([H2O2]ss) depends on the mitochondrial production and utilization of nitric oxide and on the activity of SOD and catabolizing enzymes.

3.2 THE INTERPLAY BETWEEN NO AND H2O2 STEADY-STATE CONCENTRATION The assessment of steady state implies the assumption that rates of production and utilization of the compounds are equalized (+d[H2O2]/dt = −d[H2O2]/dt and −d[NO]/ dt = +d[NO]/ dt). The main cellular source of O2− and H2O2 is mitochondria (40) (Figure 3.1). The dismutation of O2− to H2O2 is strictly compartmentalized. In microsomes, the reaction is catalyzed by Cu/Zn-SOD whereas in mitochondria, it is by Mn-SOD; compartmentalization is valid as well for the enzymes that catabolize H2O2, catalase, and gluthatione peroxidase (GPx) (Reactions 3.6 and 3.7): Catalase H2O2 + H2O2 → 2H2O + O2 (Reaction 3.6) GPx H2O2 + 2GSH → 2H2O + GSSG (Reaction 3.7) The rates of Reactions 3.6 and 3.7 depend on the respective second-order rate constants (5 × 107 and 4.6 × 107 M−1s−1 for catalase and gluthatione peroxidase, respectively [40]) and on enzyme concentration in the different tissues and cell fractions. In cytosol, catalase concentration ranges between 1.2 × 10−6 M (liver) and 3.8 × 10−8 M (heart) (41). Mitochondrial concentration of catalase has been estimated at 7.2 × 10−7 M (heart). In liver, cytosolic and mitochondrial concentration of gluthatione peroxidase is about 10−7 M, (42); the mitochondrial concentration of Mn-SOD in liver is two orders of magnitude higher than GPx: 0.3 to 1.1 × 10−5 M (35). Thioperoxidase (Trx) has gained significance in the last few years; different isoforms are found in cytosol (TrxR1) and mitochondria (TrxR2). Cells induced with a TrxR2 dominant negative and co-stimulated with EGF produced more hydrogen peroxide than non-transfected cells, with increased progression from G1 to S phase; cell proliferation and protein tyrosine phosphorylation of many proteins, including ERK, is also enhanced (43).

50


At 1 to 2 µM, NO-dependent H2O2 production rate in isolated liver and heart mitochondria is about 0.15–0.2 nmol H2O2/min.mg mitochondrial protein (4, 27). Thus, considering a close GPx concentration in the different cell compartments (42), a similar mitochondrial and cytosolic [H2O2] ss* should be expected. In accordance with this expectation, [H2O2]ss has been calculated in about10-8 M in rat liver cytosol and mitochondria (44), stimulated perfused liver (42), or after diffusion in hepatocytes (45). In rat liver mitochondria, the physiological [NO]ss** level has been estimated in 0.5 to 1 × 10−7 M (19, 21); most likely, at physiological O2 concentration (~5 to 10 µM), the production rates of O2− and H2O2 by mitochondria are higher than expected (46). At 0.1 µM NO, the +d[H2O2]/dt is about 0.34 × 10−6 M.s−1 (27). Considering the reduction to H2O by GPx and catalase at respective concentrations of 2.72 × 10−6 M and 1.2 × 10−6 M, and the rate constants for Reactions 3.6 and 3.7, the NO-dependent [H2O2]ss could be calculated as follows (40): +d [ H 2 O 2 ] ⁄ dt [ H 2 O 2 ] ss = -------------------------------------------------------k [ GPx ] + k [catalase] –6

(3.1)

–1

0.34 × 10 Ms [ H 2 O 2 ] ss = -----------------------------------------------------------------------------------------------------------------------------------------------------------------7 –1 –1 –6 7 –1 –1 –6 [ 5 × 10 M s ( 2.72 × 10 M ) + 4.6 × 10 M s ( 1.2 × 10 M ) ] NO-dependent [H2O2]ss = 0.18 x 10−8 M Considering that in liver, cytosol [H2O2]ss is about 0.82 × 10−8 M (40), NO mitochondrial utilization accounts for 20 to 30% of steady-state concentration. This data emphasizes the importance of mitochondrial NO metabolism in the modulation of oxygen active species, particularly at close physiological NO concentrations. It is well known that bacteria activate the expression of antioxidant enzymes in response to oxidative stress. However, this reaction cannot be easily extrapolated to mammalian cells (47); constitutive antioxidant defenses emphasize the role of H2O2 variations as a key signaling mechanism. Supplementing cells with exogenous H2O2 requires taking into account the activity of constitutive antioxidant enzymes that will set the [H2O2]ss to a considerably lower value. Because NO and H2O2 act in a concerted manner, it is sometimes difficult to ascertain specific effects to one or to the other active compound (37).

* [H2O2] ss = steady-state concentration of H2O2. ** [NO]ss = steady-state concentration of NO.


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3.3 THE VERY LOW OXIDATIVE STRESS LEVEL: PROLIFERATING EFFECTS OF H2O2 Although high level of reactive oxygen species is often associated with cytotoxicity, H2O2 can mediate the transduction of intracellular signals involved in growth and transformation. Proliferating mammalian cells have a broad response to oxidative stress; up to 3 to 15 µM H2O2 in the media exerts a significant mitogenic response in fibroblasts (48). Tenfold higher H2O2 concentration induces temporary cell cycle arrest followed by an increase in the expression of genes encoding antioxidant enzymes. At higher concentrations, H2O2 induces an almost permanent growth-arrest without cell damage. This status allows the cells to function normally, but they will not divide (48). Grading effects of H2O2 have been reported by Antunes and Cadenas in Jurkat T-cells (49); 0.7 µM [H2O2]ss represented the edge between survival and apoptosis in these cells, a level that is two orders of magnitude higher than that measured in different cell lines and tissues in the physiological conditions (40). Li and Holbrook reported an opposite response to 5 to 30 µM H2O2 (proliferation and apoptosis) in liver cells (50). Likewise, cell proliferation depends on a low, very precise H2O2 concentration; [H2O2]ss of E17-P2 proliferating hepatocytes ranged in about 10−11 M and increased by 100-fold in quiescent P30-P90 differentiated cells. H2O2 scavenging by 1 mM N-acetylcysteine (NAC) further increased the proliferation rate of neonatal P2 hepatocytes, but a higher (i.e., 5 to 10 mM) NAC concentration arrested the cells (17). Exposing P2 hepatocytes to 50 µM H2O2 also induced cell arrest without apoptosis. Differential responses to low and high oxidative stress levels have also been reported in liver regeneration. In the regenerating liver, increased production of mitochondrial oxidants is linked to cytokine-dependent proliferation and local expression of TNF-α (51, 52). In opposition, LPS-massive induction of TNF-α and activation of NF-κB, followed by mitochondria-mediated high-grade oxidative stress, are associated with damage, and cell apoptosis or necrosis (53). According to determinants of [H2O2]ss (Equation 3.1), Chae et al. observed that H2O2-dependent proliferation was inhibited by catalase in osteoblasts (54). Brown et al. infected aortic smooth muscle cells with adenovirus containing complementary deoxyribonucleic acid (cDNA) for human catalase (55). After the reduction of H2O2 concentration, proliferation became clearly diminished and apoptotic rate increased, an effect possibly related to a COX-2-dependent mechanism. In the same way, Tsai et al. (56) reported that incubation of vascular smooth cells with antioxidants, such as NAC, results in a reduction of cell viability and increase of the apoptotic rate, suggesting that acute setting of cell endogenous reactive species takes part in cell survival. H2O2 proliferative effects depend on selective activation of signaling pathways. In an attempt to identify these mechanisms, Preston et al. studied the response of Rat-1 fibroblasts to different oxidative stress conditions (57) (i.e., addition of catalase abolished both proliferation and ERK1/2 [MAPK44/42] phosphorylation) (Figure 3.2). In contrast, JNK-1 activity increased following the

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addition of catalase or H2O2 (at two opposite H2O2 levels). Only “toxic” H2O2 levels induced phosphorylation of p38 MAPK. Thus, authors attributed the inhibition of Rat-1 proliferation to the differential activation of these signaling pathways by oxidative stress. Recently, we have detected that ERK1/2 are able to translocate into mitochondria during brain development (58) and that ERK1/2 activation depended on H2O2 yield. In vitro, ERK1/2 activation in mitochondria was maximal at 1 µM H2O2 concentration, but markedly decreased at 50 µM H2O2 or in the presence of antimycin. We reported as well ERK1/2 activation in embryonic hepatoblasts or in isolated P2 hepatocytes exposed to 1 µM H2O2 (17), which decreased in the quiescent adult cells concomitant with p38MAPK activation (Figure 3.2). Therefore, we suggested that in association with high cell proliferation rate, solely ERK is activated at very low oxidative stress level. Differential effects of H2O2 are also detected in transformed cells. Our group reported increased proliferation in tumoral lung LP07 and mammary LMM3 cell lines at 1 µM H2O2, whereas cells became arrested without apoptosis at 50 to 100 µM H2O2 (59); these effects were abolished by co-incubation with catalase inhibitor 3-amino-1,2,4-triazole (ATZ). Median H2O2 required in the media to solely activate ERK and to induce proliferation in cultured cells can be estimated in about 2.3 µM (1–5), in hepatocytes or LPO7 tumoral cells (17, 50, 59, 71).

[H2O2]ss Cell cycle arrest

Proliferation

0.3 × 10−11 M

10−8 M

0.5 × 10−10 M 10−9 M

P-ERK ½ P-p38MAPK 11.16

2

0.63

0.095

0.075

Median H2O2 Supplementation for MAPK activation Proliferation P-ERK ½ 2.3 µM (1–5) (17, 50, 59, 71)

Cell cycle arrest apoptosis P–p38MAPK 0.03 mM (0.02–0.05) (17, 50, 59, 77, 79)

Apoptosis P–JNK 0.3 mM (0.05–1) (50, 77, 79, 94, 95)

FIGURE 3.2 The interdependence between H2O2 steady-state concentration ([H2O2]ss) and cell behavior (upper, [17]) is related to P-ERK ½ /P-p38MAPK ratio (middle) and depends on the differential redox activation of MAPK, as confirmed by supplementing different cell types with exogenous H2O2 (bottom; median concentration, references in brackets).


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H2O2 and oxidative stress have been considered to have a major role in oncogenic transformation (60–62). In an elegant study, Arnold et al. (63) reported that transfection of oxidase Nox1 to NIH 3T3 cell line led to a fivefold increase in cellular H2O2 level, which resulted in a transformed aggressive tumoral phenotype; cells reverted to a normal phenotype by co-transfection with catalase. These effects indicate that the major role of H2O2 is to activate genes related to the proliferating cascade. In accordance with this finding, tumor promotion in a rat liver epithelial cell line was enhanced by H2O2 that preferentially induced the expression of c-fos, c-jun, and c-myc (64). Growth factors, such as PDGF, BFGF, and EGF, have been demonstrated to trigger H2O2 production (65, 66). H2O2 is able to increase EGFR phosphorylation contributing to enhance tumorigenic effects related to abnormal expression of growth receptors (67). Hamada et al. enhanced the malignant potential of weakly malignant cell line ER-1 by continuous EGF stimulation and the changes were irreversible after 1 month of treatment. The effects of EGF were related to increased intracellular peroxide levels and DNA oxidative damage as they were almost completely inhibited by co-incubation with NAC (68). Duration of MAPK activation determines whether stimuli lead cells to proliferation or differentiation (69). Activation of myeloid leukemia cell line TF-1a by granulocyte/macrophage-colony stimulating factor induced only a transient activation of MEK and ERK1/2 associated to 50% increase in cell proliferation, whereas prolonged stimulation with 10−8 to 10−6 M PMA rendered 91 to 98% cell differentiation, with no proliferation. It is highlighted that ERK1/2 are activated by reactive oxygen species, as generated by PMA in leukocytes (69). As H2O2, peroxynitrite (ONOO−) may also activate ERK and Raf-1. This effect was maximal at 100 µM ONOO− in cultured rat myofibroblasts and was inhibited by NAC or prevented by the concomitant utilization of MEK inhibitor PD98059 (70).

3.4 CELL CYCLE ARREST BY H2O2 As mentioned previously, H2O2 is able to inhibit cell progression into the cell cycle, leading to arrest, apoptosis, or necrosis (Figure 3.3). The differential activation of signaling cascades may depend on cell H2O2 steady state and SOD concentration. The hypothesis implies that active oxygen species effects develop through differential activation of specific kinase signaling pathways, as suggested by recent evidence. Kwon et al. (71) reported that low 10 to 30 µM H2O2 increased ERK1/2 activity and expression in adult rat ventricular myocytes, neither affecting survival nor activating JNK, p38MAP, or Akt kinases. Instead, higher H2O2 concentration (100 to 1000 µM) increased apoptotic rate or caused both apoptosis and necrosis. The apoptotic effect of 100 µM of H2O2 was inhibited by transfection with a JNK dominant-negative, and potentiated by ERK inhibitor U0126 or Akt inhibitor LY 294002. The H2O2-dependent transition from cell proliferation to cell cycle arrest and differentiation depends as well on other stimuli such as Ca2+ release; in prostate tumor spheroids, incubation with 0.1 to 1 µM H2O2 upregulated c-fos and enhanced tumor growth in connection with release of Ca2+

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Stem cell Low energy charge Few mitochondria ••

•

• Proliferation

Transformation Low energy charge Dysfunctional mitochondria

• • • • • • Cell cycle arrest • • • • • • • • •

Nitric oxide

• • • • • •

H2O2 Diﬀerentiation

High energy charge Many mitochondria • • • • • •

• • • • • • • • •

• • • • • • • • •

• • • Apoptosis

FIGURE 3.3 The different cell responses depend on a continuous grading of nitric oxide and derived H2O2, produced, utilized and released by mitochondria, and on the size of mitochondrial pool and its energy charge.

from cell stores, whereas 200 µM H2O2 arrested cell cycle and induced p27, concomitantly with prolonged Ca2+ influx (72). Furthermore, H2O2 may activate factors that negatively modulate the cell cycle; TGF-β inhibits fetal hepatocyte growth by arresting cells in G1; associated mitochondria-mediated apoptosis was attributed to oxygen active species and was blocked by antioxidants (73). In addition, complex effects of Myc to coordinate cell growth and progression through the cell cycle (74) may rely on redox status: c-myc-dependent apoptosis of hepatoma cell was reported to be associated to elevated intracellular H2O2 levels (75).

3.5 THE LOW-LEVEL OXIDATIVE STRESS: P38 MAPK CASCADE In addition to extracellular signal-related kinases (ERKs) and stress-activated protein kinase/c-Jun N-terminal kinase (SAPK/JNK), p38 MAPK are important biomolecules in cell proliferation, survival, and apoptosis induced by extracellular stimuli (76). Kurata (77) reported that low 20 µM H2O2 causes rapid p38 MAPK activation in lymphoid cells, which results in phosphorylation of MKK3/6 and p38 MAPK, and in activation of transcription factors ATF-1 (CREB) and ATF-2. In this study, H2O2 effects were associated with growth arrest for 24 h and inhibition of cell division in phase M, and were reverted by NAC or by specific p38 MAPK inhibitor


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SB203580; in this condition, apoptotic rate did not increase. Activation of p38 MAPK cascade and ATF-1 and 2 sustained the activation of antioxidant genes, such as SOD, thus balancing redox cell status. Injection of p38 MAPK promoted similar effects in Xenopus oocytes although transition was inhibited in S/M instead of in M phase, probably by lack of p53 in these cells. Early activation of p38 MAPK by oxidative stress should not depend on intracellular calcium movements. H2O2-dependent activation of ERKs and p38 MAPK may precede many steps before a significant [Ca2+] release from intracellular and mitochondrial stores is detected; likewise, progression of cell injury reflects the Ca2+-dependent tyrosine kinase-induced activation of SAPK/JNK (78). In different cell lines, median H2O2 concentration in the medium required to activate p38MAPK is estimated in about 34 (20 to 50) µM (17, 50, 59, 77, 79). In normal development, p38 MAPK is only phosphorylated at quiescent stages, but it can be activated in proliferating hepatocytes by supplementation with H2O2 (17, 80) (Figure 3.2). Consequently, D-cyclins and proliferation rate are negatively correlated to phospho-p38; an opposite correlation is obtained when cells are exposed to antioxidants or kinase inhibitors (17). In studies about mechanisms of disease, exposure of neuronally differentiated SK-N-BE cells to amiloyd protein induced early generation of oxidative stress, resulting in p38MAPK and c-Jun aminoterminal kinases (JNKs) activation and apoptosis (81). Similarly, low-density lipoproteins (LDL) signaling in smooth muscle cells involves IL-8 transcription, the generation of H2O2, the phosphorylation of p38 MAPK and activation of AP-1, and NF-κB (82). The extent of H2O2 stimuli and the complex interplay among NO, oxygen active species and the signaling cascades may explain a different modulation of progression through life processes (Figure 3.3). For instance, activation of p38 MAPK concomitant with cell cycle arrest may eventually undergo apoptosis in the presence of NO that also activates p38 MAPK pathway; in accordance with this finding, the p38 inhibitor SB203580 blocks pro-apoptotic effect of NO in SH-SY5Y neurons (83). These NO effects on p38 MAPK signaling probably result in Bax translocation to mitochondria, a well-known intermediary of programmed cell death (84). Concerning to the mechanisms of H2O2 or NO on signaling, both may react with thiol groups of cysteine, methionine, or other aminoacids to modify the activity of tyrosine kinases and tyrosine phosphatases (85, 86). Recent evidence indicates that phosphatase PP2A is a major regulator of MAPK phosphorylation in brain: Inhibition of PP2A activity via reversible oxidation of cysteine thiols modulates the activation of MAPKs in response to hydrogen peroxide and oxidative stress (87). Other mechanisms involve the activation of specific phosphorylation cascades that participate in the progression through the cell cycle; for instance, H2O2 activates p70s6k in mouse epidermal cells, which plays an important role in the transition from G0/G1 phase to S (88). It is noticeable that ERK1/2 remain phosphorylated at a high oxidative stress level. Thus, final effects on cell cycle activity shall depend on the relative activity of cascades that promote proliferation or apoptosis. Aguirre-Ghiso et al. reported

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that modulation of ERK/p38 activity ratio predicted the in vivo behavior in ~ 90% of the studied cell lines (89). Similarly, we demonstrated that high ERK/p38 activity ratio was representative of normal liver proliferating phenotypes, whereas low ERK/p38 activity ratio was associated to quiescent cells and correlated with [H2O2]ss (r:0.96) (17). The interplay among the different MAPK pathways contributes to the modulation of their own activity and their effects. A direct protein-protein interaction between ERK1/2 and p38 reduces ERK1/2 activity (90). A recent study demonstrated that p38 MAPK activation decreased H2O2induced ERK activation through a PP2A-dependent mechanism in cardiac ventricular myocytes (91). Masuda et al. suggested that phosphorylation of Ser446 of MKP-7 by ERK contributed to suppress p38MAPK and JNK activation (91); interaction of these two MAPK pathways define a novel cellular mechanism that allows fine modulation of apoptosis during oxidative stress (91, 92).

3.6 THE HIGH OXIDATIVE STRESS LEVEL: JNK AND NF-ΚB At a high oxidative stress level, cascades dependent on NF-κB and JNK activation are recruited, with 95% of cells entering into apoptosis (93). Median H2O2 required to activate JNK in cultured cells can be estimated in about 310 µM (50 to 1000) (50, 77, 79, 94, 95) (Figure 3.2). The activation of JNK is specifically linked to the leakage of mitochondrial oxidants. The effects of oxidants on JNK cascade may cooperate upstream of other cell signaling molecules, such as Ca2+; addition of an intracellular Ca2+ chelator inhibited H2O2-induced phosphorylation of SAPK/JNK in Chinese hamster V79 cells exposed to hydrogen peroxide (96). In this experimental model, p38 and ERKs were also activated by H2O2 but they did not require Ca2+ movements. Similar effects were described by Salh et al., who observed H2O2-induced cellular injury depended on JNK activation. (95). In addition, redox-dependent pro-apoptotic role of JNK in lung fibroblasts was recently related to cooperative activation of TNF receptor 1 (97). JNK-dependent phosphorylation is important for the stabilization of pro-apoptotic p53 protein, and for effects of Bcl-2/Bax (98, 99). Interestingly, the SAPK/JNK pathway may also be activated in transformed cells by markedly decreasing the cell H2O2 concentration (57). The activation of NF-κB may involve pro-proliferative or pro-apoptotic consequences. Concerning the latter, some chemoprotective agents, such as non-steroidal anti-inflammatory agents (NSAIDs), could act by stimulating both the production of oxygen active species and NF-κB (100). Similarly, oxidative stress elicited by complex I inhibitor MPP+ activates JNK and NF-κB in SH-SY5Y neuroblastoma cells (101). These effects imply activation of MEK and MEKK-1 and loss of Raf-1, which are likely caspase-mediated processes. In contrast, targeted Raf-1 to mitochondria improved Bcl-2-mediated resistance to apoptosis (102).


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3.7 THE EFFECTS OF H2O2 ON CYCLINS Mammalian cell cycle progression depends on the sequential activation of a family of serine-threonine kinases: the cyclin-dependent kinases (Cdks). These kinases form specific complexes with different types of cyclins −D, E, A, and B−, which regulate the movement of cells through the cell cycle from the G0-G1/S/G2/M phase.* The D-type cyclins are associated with early steps in the transition from G0-G1 to S. Thus, quiescent cells (G0) contain low levels of D-type cyclins. Growth factors induce synthesis of cyclin D1; its translocation to the nucleus requires activation of Ras, Raf1, MKK1/2, ERKs, and cETS-2. Mitogen withdrawal cancels cyclin D1 synthesis; the enzyme is ultimately ubiquinated and degraded. ERKs stimulate cell proliferation and induction of active cyclin D1 by enhancing AP-1 activity. In contrast, p38 MAPK and JNK transcriptionally down-regulate cyclin D1. Casanovas et al. reported that oxidative as well as osmotic stress decreases cyclin D1 stability, the former by activating p38 pathway (104). This effect is due to phosphorylation of cyclin D1 at Thr286, which leads to the ubiquitination of the protein. Awad and Grappuso (80) observed similar effects. They reported a temporal inverse correlation between activation of p38MAPK and cyclin D1 content during liver development or liver regeneration in the rat. Accordingly, our group recently reported that hepatocyte behavior specifically depends on cell ERK/p38MAPK activity ratio, which is high during proliferation and low at quiescence, and respectively associates to high and low D-cyclin expression (17). Direct effects of H2O2 include overexpression of Cyclindependent kinase inhibitor (CKI) p21(Waf-1) and lower expression of Cdk; RNA level, protein, and kinase activity of Cdk2 are decreased 72 h after H2O2 stress (104). Oxidative stress may also modulate transition through other cell cycle phases by affecting cyclins other than cyclin D, such as cyclin E (late G1/S phase), cyclin A (S phase), and cyclin B (G2/M) (105).

3.8 NO AND THE MODULATION OF CELL PROLIFERATION NO has been associated to cell proliferation and oncogenesis, as well as apoptosis or inhibition of apoptosis. Because H2O2 or peroxynitrite also participate in cell cycle modulation and drive to apoptosis, it is sometimes difficult to discern the true effector in mitochondria, where the species cohabit because of NO metabolism (19, 21). Increased expression of iNOS is observed during proliferative bursts of hepatocytes. In this way, NO participates in the process of liver regeneration that follows partial hepatectomy. A disruption of iNOS gene severely inhibits the proliferative response to liver resection in transgenic mice (106). This effect appears to be the consequence of NO inhibitory effects on TNF-α pro-apoptotic * M denotes mitosis and S DNA synthesis; G1 and G2 are the gap phases preceding DNA synthesis and mitosis, respectively.

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response. Thus, the balance between TNF-α proliferative and apoptotic effects could depend on the expression and activation of NF-κB (a well-known inducer of iNOS trancription) and redox mechanisms that activate iNOS gene transcription (107, 108). Moreover, NO may inhibit NF-κB in a negative feedback regulation loop. NO could participate in carcinogenesis as well. Payne et al. (110) observed that NO and expression of iNOS isoform in colonic mucosa is associated to production of oxygen active species, oxidative stress, apoptosis resistance, and increased proliferation and transformation. Moreover, although in smooth muscle cells NO is currently understood as anti-proliferative by antagonizing effects of ET1 and angiotensin, tumoral angiogenesis involves increased expression of eNOS and NO participates in the in vivo proliferative action of VEGF on endothelium (110). NO activation of MAPK and fibroblast growth factor (FGF-2) via cGMP could correlate with angiogenesis and tumor progression (111). In normal liver development, we reported a concerted progressive increase of mtNOS and H2O2 in the transition from high to low proliferation rate. The increase of mtNOS was preceded by a burst of modified nNOS increase in cytosol with higher nNOS gene transcriptional activity during the first days of life (17). The same effects were observed after birth in rat brain development, just in the transition from neuroblast proliferation to cell cycle arrest in the structural plasticity period (10). Authors reported anti-proliferative effects of NO on smooth muscle and tumoral cells. Pervin et al. (113) observed nitric-oxide-induced cytostasis and cell cycle arrest of the MDA-MB-231 human breast cancer cell line. NO suppressed the synthesis of cyclin D1, probably by an effect other than changing mRNA expression, although translational or post-translational control of cyclin expression was subsequently involved.

3.9 NO SYNTHASES AND TUMORS Although NO synthases are present in a variety of tumors, the role of NO in tumoral biology remains controversial. iNOS protein and mRNA are found in transitional cell carcinoma of the bladder but not in normal urothelium (113). Morcos et al. reported that both normal urothelial and bladder T24 and MTB-2 cancer cells have Ca2+-dependent NOS activity but only transformed cells have Ca2+-independent iNOS activity (115). After supplementation with L-Arginine, stimulation of urothelial cell growth was attributed to activation of Ca2+-dependent constitutive NOS, and an anti-proliferative effect was attributed to iNOS. In accord with this finding, the control of in situ carcinoma of the bladder by administration of intravesical BCG was suggested to be the result of induced iNOS, by releasing cytotoxic and antitumoral NO. In addition, Klotz et al. (116) observed by immunohistochemistry in human bladder cancer that malignant cells were highly iNOS positive, whereas surrounding cells were not; in addition, eNOS related to blood vessels was found only in the stroma of tumors. Studies performed in urinary bladder urothelium after the Chernobyl accident (116) reported high iNOS expression and 8-hydroxy2 deoxyguanosine (8-OhdG) content associated to high carcinoma incidence and


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increased tumor suppressor p53 protein. It is noticed that p53 may be inactivated either by radiation or by NO, the latter through nitration of tyrosine residues (117). Administration of L-Arg increased mammary tumor EMT-6 size and cell proliferation in vivo and in vitro; however, only in vitro effects could be blocked by NOS inhibitor aminoguanidine (118). As reported by Orucevic et al. (120) most authors believe eNOS-like activity related to angiogenesis to be linked to tumor invasiveness, whereas iNOS is probably related to anti-tumoral effects. In addition, Reveneau found a significant inverse correlation between NOS activity and tumor grade; moreover, iNOS positive tumors (most of human breast tumors) are associated to the presence of estrogen receptors (a known marker of prognosis) (121). The same inverse correlation was found by Tschugguel et al. (122) in grade I–III ductal carcinomas. Increased expression of iNOS was also observed in colorectal cancer by Ropponen et al.; the authors reported that iNOS intensity was higher in low grade tumors Dukes A and B than in malignant C and D stages; this encouraged them to attribute a protective role to the enzyme (123). Other investigators reported that iNOS facilitated proliferation and metastatic potential; in this way, expression of iNOS in breast cancer has been reported to be correlated with metastatic disease (123) and malignancy (124). Discrepancies in NO effects on tumorigenesis may arise from variable mitochondrial NO concentration, increasing either the production of H2O2 or the matrical oxidants such as peroxynitrite. It is conceivable that a low NO is required to form mitochondrial H2O2 and to stimulate proliferation. In the opposite situation, Shi et al. and Xie et al. proposed that defective iNOS with poor activity may stimulate tumor growth by restricting the host defense mechanism (126, 127). In addition, we found a very low mtNOS activity with poor response to Ca2+ and low mitochondrial H2O2 in mice tumors and tumoral cell lines associated to permanent proliferation and growth (59); tumoral cells were very sensitive to exogenous H2O2, and likewise, it is surmised that defective mtNOS and low mitochondrial NO contribute to uncontrolled cell division. Similarly, Renaudin et al. found loss of NOS1 expression in high-grade renal cell carcinoma, whereas it is currently expressed in normal renal cells or benign renal tumors (128).

3.10 NO, H2O2, AND CELL APOPTOSIS Apoptosis is a complex process that involves membrane receptors, effectors, regulatory enzymes, and transcription factors. Two major pathways induce apoptosis: one involves death receptors and is exemplified by Fas-mediated caspase8 activation; the other is the stress- or mitochondria-mediated caspase-9 activation pathway (128, 129). Both pathways converge on caspase-3 activation, resulting in nuclear degradation and cellular morphological change. Activation of caspase9 is followed by release of cytochrome c. Release of the solubilized pool of cytochrome c into the cytosol occurs by permeabilization of the outer mitochondrial membrane mediated by pro-apoptotic Bcl-2 family proteins, notably Bax and Bak, or by Ca2+-triggered mitochondrial permeability. Prevention of cytochrome c release by Bcl-2 is consistent with the anti-apoptotic effect of the latter.

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Recent investigations indicate that cytochrome c extrusion occurs by a two-step process, initiated by a disruption of the association of the hemoprotein with cardiolipin, the phospholipid that anchors cytochrome c to the outer surface of the inner mitochondrial membrane (129). Oxygen active species and NO are important regulators of apoptotic pathways and the regulation of redox mitochondrial functions is the central focus in apoptosis research. Cytochrome c release follows the peroxidation of cardiolipin (129). Increase of Ca2+ intracellular level consecutive to oxidative cell injury opens the mitochondrial transition pore, lowers the efficiency of oxidative phosphorylation, and favors cytochrome c release (130). Different agents that promote apoptosis also promote mitochondrial changes; for instance, pro-apoptotic p53 induces gene transcription of redox related genes encoding proteins that lead to oxidative stress (131). Moreover, oxidative stress induces p53 and kinases, including apoptosis signalregulating kinase 1 (ASK1), JNK, and p38 MAPK kinase; Trx inhibits apoptosis signaling not only by scavenging intracellular ROS in cooperation with the GSH system, but also by inhibiting the activity of ASK1 and p38 MAPK. Pro-apoptotic activity of NO has been demonstrated in different cells and tissues (132, 133). The mechanism underlying the process may include activation of SAPK/JNK and p38MAPK, inhibition of ERK and damage to DNA (134, 135), as well as increase of p53 expression and p21 activation, the contribution of caspases and inhibitory Bax effects (135). Other pro-apoptotic effects include NOdependent tyrosine nitration of cytochrome c that may facilitate its release from mitochondria (136). Many pro-apoptotic effects of NO may be mediated by peroxynitrite. Moreover, intercellular induction of apoptosis may require the fine biochemical interplay between oxygen and nitrogen species (137); this mechanism could be important in the control of transformation in cells (138). In particular, caspases are modulated by oxygen active species and NO. Caspases exist as zymogens, which contain specific motifs, with a catalytic site cysteine susceptible to modulation by either oxygen species or nitrosylation/denitrosylation; denitrosylation occurs during proteolytic activation of caspase-3. Hydrogen peroxide blocks processing of the proenzyme by a thiol oxidation mechanism (139). NO has anti-apoptotic effects as well. NO is able to induce or inhibit NF-κB expression, a well-known promoter of iNOS gene. NF-κB expression and activity correlates with NO concentration. The inhibition or stimulation of NF-κB are related to proliferative or apoptotic effects. Thus, both NF-κB and NO (likely depending on cell type and concentration) may be protective or induce apoptosis. It was suggested that increased iNOS expression and resulted NO act as a negative regulatory feedback modulator of NF-κB activity (140). This transcription factor is probably important during development. NF-κB is induced by pro-proliferative MAPK and inhibited by p38 MAPK pathway. Disruption of NF-κB gene is associated to embryo lethality; many anti-apoptotic pathways such as Bcl-2 are induced by NF-κB. Accordingly, new therapeutics of cancer include the inhibition of NF-κB pathway (141). Other anti-apoptotic NO-pathways include S-nitrosylation and inactivation of thiol of caspases and up-regulation of anti-apoptotic genes such as that of heme oxygenase.


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3.11 mtNOS AND LIFE PROCESSES The discovery of mtNOS in 1998 by two independent groups of investigators added a new perspective to the knowledge of modulation of mitochondrial functions (7, 8, 24). It is now clear that activation of mtNOS by L-Arg and/ or Ca2+ is followed by a decrease in mitochondrial O2 uptake and mitochondrial transmembrane potential (24, 142), an effect almost completely prevented by NOS inhibitors. Moreover, activation of rat liver mtNOS drives to increased production of oxygen active species, because of the oxidative metabolism of NO (Figure 3.1) (143). The regulatory properties of mtNOS on O2 uptake have been explored by our group in different experimental models. In hypothyroid rats, we reported an increased expression and activity of liver mtNOS, which was completely reverted by hormone replacement (16). The activity of mtNOS correlated with both decreased systemic and mitochondrial O2 uptake. It is known that thyroid hormone exerts proliferative effects, whereas hypothyroidism is associated to diminished proliferation rate and mitochondrial biogenesis (144); whether the hypothyroidinduced decrease in cell proliferation is related to mtNOS activity and the production of oxygen active species is not established yet. Similar changes were observed by our group in rats acclimated to a cold environment; after a week of exposure to 4°C, liver and muscle mtNOS increased allowing high energy intake, which favored the synthesis of fat insulation, to maintain the body temperature (145). Modulation of mtNOS is related to cell responses and to growth and development. NO and oxygen active species are important in synaptic plasticity. Rat brain and cerebellum mitochondria produce H2O2 in the presence of NO (10). In the perinatal period, a definite time-course of brain mtNOS is observed: Brain and cerebellum mtNOS expression and activity and H2O2 increase from the last days of gestation up to the first 10 postnatal days and then, it sharply decreases; this time-course is opposite to cytosolic nNOS, which is practically absent in fetal development and increases a few days after birth. Interestingly, mitochondrial Mn-SOD follows an identical time-course to that of mtNOS. A temporal induction of mtNOS and Mn-SOD may cooperatively generate the H2O2 required to arrest neuroblasts and to start the process of synaptic connection and elimination. A connection between mtNOS, mitochondrial H2O2 production rate, cell [H2O2]ss, D-cyclin expression, and the rate of proliferation was delineated in normal rat liver development. A progressive increase of mtNOS content elicits crescent mitochondrial H 2 O 2 production rate and cell [H 2 O 2 ] ss that decreases ERK/p38MAPK activity ratio and the expression of cyclins D1-3. These effects progressively decrease the rate of proliferation up to quiescence. Effects can be reverted in isolated P2 proliferating cells by reducing cell [H2O2]ss with antoxidants (17). Mitochondrial NOS is almost absent in embryonic hepatoblasts at E17-19 days leading to very low oxidative stress level, which is required for activation of proliferative cascades to proceed into the cell cycle. We reported the same instance

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in mice tumoral cells, which lack significant mtNOS activity (59); thus, “embryonic” redox status is shared by both developing and tumoral cells. The mtNOS content and activity have a considerable impact on cell death. Activation of mtNOS ends in cytochrome c release from mitochondria, a wellknown promoter of apoptosis, by activation of caspase 9 (146). The activition of mtNOS leads to increased mitochondrial efflux of Ca2+. Addition of pro-apoptotic agents, such as etoposide and methylprednisolone, to a thymocyte suspension increased NO production by mitochondria, a fact associated to mild uncoupling of oxidative phosphorylation and to a marked decrease in GSH and cytochrome c content. Accordingly, a partial inhibitory effect on apoptosis was achieved by thymocyte pretreatment with NOS inhibitors, L-NMMA or Nitro-L-arginine (8). An increase in mitochondrial NO has been associated to pro-apoptotic effects of MPTP, a compound that promotes Parkinsonism and apoptosis in neural cells (147).

3.12 BIOENERGETICS, MITOCHONDRIAL BIOGENESIS, AND CELL EVENTS Energy demand in proliferation and differentiation is a task in the understanding of mitochondrial signaling. Komarova et al. reported a transition from glycolytic anaerobic metabolism in cultured primary osteoblasts, to four- to fivefold increased aerobic metabolism in mature cells (149); differentiation was associated to high mitochondrial activity and membrane potential. Similarly, inhibition of mitochondrial protein translation by chloramphenicol blocks avian myoblast QM7 cell line differentiation (149). In this study, persistently diminished mitochondrial activity contrasted with high oxidative activity at the terminal myogenic differentiation stage and changes were similar to those produced by the uncoupler FCCP (150). Moreover, in myoblasts, destruction of mtDNA by ethidium bromide inhibits differentiation, but not cell proliferation. It is then surmised that promotion of proliferation entails a controlled inhibition of mitochondrial respiration. In support, the activity of most tumoral and normal proliferating mitochondria is uniformly maintained at 20 to 30% with respect to quiescent organelles (17, 59). Likewise, low electron transport-coupled ATP synthesis correlates with faster tumor growth (151) and high invasive behavior (152). Altered mitochondrial structure and function, the reduction of mtDNA copies, and reduced expression of genes regulating ATP synthesis have been recently reported in transformed human fibroblast cell lines (153), and in cells from renal carcinoma (154). The exception to a general rule about decreased oxidative capacity and increased cell proliferation rate is oncocytoma, a tumor, which exhibits high mtDNA and mitochondrial number (155). Promotion of proliferation is not the mere result of lowering O2 uptake and ATP synthesis, but a consequence of decreasing O2− and H2O2 production rate at complexes I and II–III of the electron transfer chain. At high H2O2 concentration, transfection of mitochondrial Mn-SOD increases maturation, promotes differentiation, and suppresses malignant behavior of several cell types. Over-


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expression of Mn-SOD and mitochondrial subunits is also observed in senescent cells, allowing the hypothesis that aging is a hyperdifferentiating state. The mitochondrial complex I inhibitor rotenone impairs energy metabolism, but increases by several times the H2O2 production rate at complex I; rotenone promotes cell differentiation, lowers the basal hepatocyte proliferation, and decreases the incidence of hepatocellular carcinoma (156). Excessive production of reactive oxygen species and resultant mitochondrial damage may end in cell death, however (157). In human neuroblastoma cells, energy charge of mitochondria and ATP level is decreased by exposure to H2O2, with a marked inhibition of proliferation and delayed cell death (158). A disproportionate increase of H2O2, as catalyzed by overexpressed SOD, may also damage nuclear DNA (159). In the same way, excitotoxicity induced by stimulation of NMDA receptors by glutamate increases mitochondrial Ca2+ uptake and the production of oxidants, leading to necrosis or apoptosis of primary neuronal cultures by interfering with energy production, thus lowering delta ϕ potential and ATP level (160). The pro-apoptotic effects of Ca2+ may be enhanced by NO because it is released by the same excitatory mechanism (161). Some authors hypothesize that changes in the mass and number of mitochondria control the number of cell divisions between determination and terminal differentiation (162). From this point of view, there could be a mitochondrial differentiation signal, which would counteract the nuclear preventing of differentiation signal. Analysis of the expression of oxidative phosphorylation genes in liver during development reveals two programs: short and long. The former (differentiating program) is controlled post-transcriptionally and promotes rapid mitochondrial phenotypic changes. The latter (proliferating pathway) is prolonged, controlled both at transcriptional and post-transcriptional levels, and is responsible for increasing mitochondrial mass (163). On these bases and considering that mitochondria are multiplied several times over the number of cell divisions, the ratio of mitochondria/nucleus will increase up to terminal differentiation (Figure 3.3). From this perspective, it appears that mitochondrial biogenesis is the cornerstone in the sequential proliferating-differentiating process. As elegantly noticed by von Wangenheim and Peterson (162), more than 1000 eukaryotic species devoid of mitochondria never became multicellular. Agents that increase mitochondrial biogenesis as etoposide and genistein allow cells to undergo differentiation. In contrast, the number of mitochondria in stem cells is low, which allows maximal proliferation rate and self-renewal. As recently reported by Nisoli et al., mitochondrial biogenesis is stimulated by NO by increasing peroxisome proliferator-activated receptor γ coactivator 1 α (PGC-1) (164). The increase of mitochondrial mass also results in the augment of NO and H2O2 yields, both contributing to the proliferative and differentiating programs (17). Low intramitochondrial NO yield and very low oxidative stress level associate to partial inhibition of electron transfer chain and cell proliferation (17). We already mentioned that high NO yield is linked to cell cycle arrest, and peroxynitrite is associated to mitochondrial dysfunction (19) and to a derangement of tissue function (18). Specifically, high matrical NO and O2− levels and ONOO− formation can inhibit

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mitochondrial complex I, a rotenone-like effect (165). This could limit proliferation in most tissues whereas in others, such as the substantia nigra of the brain, it renders the cell more sensitive to apoptosis (166, 167). In this way, oxidation of mtDNA by ONOO− or active O2 species could act as a dequalinium, restricting the cell proliferative phase (55); oxidative-nitrosative effects of ONOO- on mitochondrial membrane proteins and lipids could be partially prevented by reactions with membrane components, such as ubiquinol (168).

3.13 CONCLUSIONS AND PERSPECTIVES The balance among growing, maturating, and dying is still a mystery of living organisms. The number and characteristics of the participants in this process is so extensive that it is almost impossible to delineate common mechanisms for all organisms. We attempted to analyze the influence of reactive N2 and O2 species. In the last 20 years, mitochondria have turned out to be the main source of O2and its product of dismutation H2O2. Furthermore, in the last 5 years, NO was introduced as one of the most important modulators of O2 uptake and as a source of reactive oxygen species in mitochondria. In the presence of NO, mitochondria releases H2O2, or forms peroxynitrite in the matrix compartment (Figure 3.1). Activity of Mn-SOD and the relative NO steady-state concentration will determine the rate of these respective reactions (27). Thus, the activity of NOS, particularly mtNOS, has a definite role in the redox signaling in living cells. If NO matrix concentration sets H2O2 release, then the activity of many life processes will depend on this mechanism. Grading concentrations of H2O2 elicit the differential activation of signaling pathways, which drive cells from proliferation to cell cycle arrest, differentiation, or cell death (Figure 3.2). The basis for these effects involves coordinated activation of different MAPKs, cooperation of growth factors and hormones, as well as the inhibition or stimulation of apoptosis. Ultimately, transcriptional as well as post-transcriptional events will result in regulation of multiple genes, such as cyclin D one. Moreover, oxidative stress may contribute to and sustain cell transformation. In this setting, disruption of mitochondria, which leads to a decreased energy charge or an increased production of oxidants, could play a significant role in the control of cell cycle and in carcinogenesis. Eukaryotic cells are endosimbiotic products of prokaryotes and mitochondrial precursors. Ancient bacteria were able to form and metabolize NO in the nitrificating-denitrificating pathways. The arrival of oxygen and the presence of mitochondria gave strength to the process of proliferation and differentiation conducive to the evolution of species. The mitochondrial NO-mediated formation of O2 active species was likely a substantial part of the selective pressure applied to primitive organisms to evolve, to grow, and to be different.


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3.14 ACKNOWLEDGMENTS This research was supported by grants M026 and M027 from the University of Buenos Aires, Agencia Nacional de Promoción Científica y Tecnológica (PICT 08468), and Fundación Perez Companc in Buenos Aires, Argentina.

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of 4 Functions Mitochondrial Nitric Oxide Synthase Pedram Ghafourifar Marshall University, Huntington, West Virginia

Alfredo Saavedra-Molina Universidad Michoacana de San Nicolás de Hidalgo, Morelia, México

CONTENTS 4.1 4.2 4.3 4.4 4.5 4.6 4.7 4.8 4.9 4.10 4.11

Introduction ...............................................................................................77 NO and NO Synthases ............................................................................78 Functions of NO for Mitochondria .........................................................79 Mitochondrial Nitric Oxide Synthase (mtNOS) .....................................81 Ca2+-Dependence of mtNOS ...................................................................82 Effects of mtNOS on Mitochondrial Bioenergetics ................................83 Mitochondrial Ca2+ Homeostasis .............................................................84 Heart mtNOS and Its Ca2+-Dependence .................................................84 Ca2+ and Mitochondrial Apoptosis ..........................................................86 Mitochondria, mtNOS, Oxidative Stress, and Apoptosis .......................86 Does mtNOS Link Hypoxia/Reoxygenation, ONOO-, Apoptosis, and Release of Cytochrome c? .............................................88 4.12 Antioxidants Prevent Hypoxia/Reoxygenation-Induced Oxidative Injury .......................................................................................89 4.13 Does mtNOS Play a Role in Hypertension? ...........................................89 4.14 Acknowledgments .....................................................................................91 References .................................................................................................91

4.1 INTRODUCTION The important role of nitric oxide (NO) in biology has been the subject of many studies in the past two decades. Several biological properties of NO are mediated through increase in intracellular cyclic GMP levels; however, many key functions 77

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of NO are cyclic GMP-independent. NO readily reacts with hemoproteins, thiols, and superoxide (O2−). Mitochondria possess several hemoproteins, such as cytochrome c oxidase (COX); they contain cysteine thiol-containing proteins, such as caspases; and they remain one of the main cellular sources of O2−. Thus, mitochondria are one of the prime intracellular targets for NO. Physiologically relevant concentrations of NO reversibly react with COX in an oxygen concentration-dependent manner that resembles a pharmacological competitive antagonism between NO and O2. This reaction plays a critical role in regulating mitochondrial oxygen consumption in many cells, tissues, and organs. The interaction of NO with mitochondrial thiol-containing proteins, such as caspase-3, is also reversible but in a redox- and pH-sensitive manner. This reaction plays a role in regulating mitochondrial apoptosis machinery. The reaction of NO with O2− is extremely fast and the product, peroxynitrite (ONOO−), is a potent oxidative nitrogen species. Reactions of ONOO− with mitochondrial susceptible targets are irreversible and cause mitochondrial malfunctioning, oxidative injury, and apoptosis. Although unwanted apoptosis is involved in the pathogenesis of many diseases, apoptosis deficiency may serve as one of the crucial mechanisms underlying some forms of cancer. This viewpoint can turn the perception of peroxynitrite from the ugly nitrogen oxide congener into a biologically desired one. The discovery of a mitochondrial NO synthase (mtNOS) has opened new windows in the field of NO research. Most laboratories have observed that mtNOS is continuously active, generates NO in a Ca2+-sensitive manner, and that mtNOSderived NO regulates mitochondrial respiration, transmembrane electrochemical potential (∆ψ), transmembrane pH gradient (∆pH), Ca2+ homeostasis, and ATP synthesis. The mtNOS-derived intramitochondrial ONOO− formation has been also reported by several laboratories. mtNOS-derived ONOO− causes oxidative stress and the release of cytochrome c from mitochondria, and is important for apoptosis. Few apparently inconsistent reports exist in the mtNOS literature. This chapter attempts to summarize the present understanding of mtNOS and its functions in biology, and discusses the reasons for the apparent inconsistencies.

4.2 NO AND NO SYNTHASES The discovery of NO (Palmer et al., 1987) as the endothelium-derived relaxation factor (EDRF; Furchgott and Zawadski 1980) changed our perception of NO from a noxious gas to that of a beneficial molecule of utmost importance in biology (Koshland 1992). In biology, NO is synthesized by NO synthase isozymes (NOS; EC 1.14.13.39) in a two-step, five-electron oxidation of the terminal guanidino nitrogen of L-arginine with N-hydroxy-L-arginine as the intermediate. This reaction stoichiometrically consumes O2, requires 1.5 moles NADPH, and produces L-citrulline as the final co-product. Three distinct isoforms of NOS have been well characterized. These enzymes do not exert a tissue-specific pattern of expression; however, they are commonly referred as to endothelial (eNOS), neuronal (nNOS), and inducible NOS (iNOS) (Moncada et al., 1991). The nNOS and eNOS isozymes are constitutively expressed, whereas expression of iNOS

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generally occurs after cells are challenged with an immunological or inflammatory stimulus. Cytoplasmic Ca2+ closely regulates the activity of the constitutive isoforms and these isoforms demonstrate a typical interaction with calmodulin. The activity of iNOS does not increase when cytosolic Ca2+ rises, although calmodulin is necessary for its activity. iNOS forms a tight complex with calmodulin at very low Ca2+ concentrations and thus appears to be Ca2+-independent. NO is a colorless gas that dissolves in deoxygenated aqueous solutions up to 2 mM, depending on the water/buffer used (Hogg and Kalyanaraman 1998). Although NO can react with O2, this reaction occurs in a manner that is second order in NO and first order in O2 concentrations. Therefore, at physiological concentrations of NO and O2 simple autoxidation of NO is not the main route of NO degradation (Kharitonov et al., 1994); however, NO reacts with O2− with rate constant of 1.9 × 1010 M−1 s−1 (Kissner et al., 1997) to produce peroxynitrite (ONOO−). Conversion to ONOO−, therefore, can be the main fate of NO in many biological systems.

4.3 FUNCTIONS OF NO FOR MITOCHONDRIA Long before knowing that stimulated macrophages produce NO and that NO produced by these macrophages regulates mitochondrial functions of the neighboring cells, Brown et al. (1998) and Granger and Lehninger (1982) reported that cytotoxic macrophages inhibit mitochondrial respiratory complexes I and II. This report can be considered to be the first evidence for inhibition of mitochondrial respiratory complexes by biologically produced NO. NO has physicochemical properties very similar to that of O2. NO readily binds to the O2 binding site of the reduced COX and reversibly regulates the respiration (Brudvig et al., 1980; Torres et al., 1995). NO decreases the oxygen consumption of purified COX, sub-mitochondrial particles, and mitochondria, as well as many cells including hepatocytes, brain nerve terminals, and astrocytes (Brown and Cooper 1994; Brown et al., 1995; Carr and Ferguson, 1990; Cleeter et al., 1994; Richter et al., 1994; Schweizer and Richter 1994; Takehara et al., 1995, 1996). Concentrations of NO required to inhibit COX and mitochondrial respiration is within what has been measured in a several biological systems (Brown 1995). Collectively, inhibition of non-mitochondrial NO synthesis results in stimulation of respiration in numerous biological systems (Brown et al., 1995; Borutaite and Brown 1996; Hurst et al., 1996; Lizasoain et al., 1996; Szabo et al., 1996; Takehara et al., 1995; 1996; Xie and Wolin 1996). Electrons enter the mitochondrial respiratory chain from complex I or II and flow down to the complex IV to reduce O2 to water. The electron flow through the respiratory chain is coupled to extrusion of protons from mitochondrial matrix into the intermembrane space. The inner membrane is impermeable to most cations including protons and protons can reenter mitochondria through ATP synthase machinery to produce ATP from ADP. The chemiosmotic principle established by pioneering work of Mitchell in 1950s (reviewed by Mitchell and Vectorial 1977) postulates that this proton extrusion establishes an electrostatic

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gradient across the coupling membrane, the mitochondrial transmembrane potential (∆ψ) negative inside, and an electrochemical gradient (∆pH) alkaline inside. Although the ∆ψ varies in different cells, its is generally much higher than the cell membrane potential. In succinate-energized rat liver mitochondria, the ∆ψ is about −180 mV. The O2 binding site of COX is highly specialized for O2; however, NO exerts physicochemical properties very similar to O2 that allow NO to bind to this binding site and subsequently inhibit the O2 consumption. NO inhibits the O2 consumption at physiologically relevant concentrations of NO in a competitive, reversible, and dose-dependent manner resembling a pharmacological competitive antagonism of NO with O2 (Ghafourifar et al., 2000). The ∆ψ is the driving force for mitochondria to take up cations such as Ca2+. Mitochondria are one of the main cellular calcium stores and can accommodate relatively large quantities of calcium; however, intramitochondrial ionized calcium concentration ([Ca2+]m) is kept very low by different mechanisms. Mitochondria precipitate the [Ca2+]m to form matrix electron dense granules, or exchange the [Ca2+]m by other cations such as H+ or Na+. By competing with O2, NO inhibits COX activity and, therefore, decreases ∆ψ, ∆pH, and [Ca2+]m (Ghafourifar and Richter 1999) as well as ATP formation (Saavedra-Molina et al., 2003) (See Scheme 4.1). Na+ or H+ Ca2+

Inhibition

∆ψ

Ca2+

IM

NO Matrix

Ie− II

OM

Ca3(PO4)2 (Ca3(PO4)2)3 Ca(OH)2

NADH NAD+ FAD+

IMS

H2 O

O2 Q

FADH2

III e−

ADP

H+

ATP

IV

cyto c

V

H+ H+

∆pH H+

SCHEME 4.1 Mitochondrial structure and functions. Mitochondria consist of distinct suborganelle compartments: the outer membrane (OM), the inner membrane (IM), the matrix, and the intermembrane space (IMS). These compartments are different in composition, electrochemistry, and redox status.


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The respiratory chain complexes are embedded in IM. The chain consists of four complexes (I to IV), the coenzyme Q (ubiquinone, Q), and the ATP synthase that is often referred as to complex V. Cytochrome c (cyto c) is the only respiratory chain member that is not embedded in IM. These complexes are functionally arranged in an electrochemical hierarchy based on their redox potentials. The respiratory chain provides a unique broad spectrum of redox potentials varying from –280 mV (complex I) to +250 mV (complex IV). Electrons enter the chain from complex I or II by oxidation of NADH or FADH2, respectively, flow down the chain to complex IV, and reduce O2 to H2O. Coupled to the electron flow, protons are pumped from the matrix into the IMS. The proton extrusion establishes a transmembrane potential (∆ψ, negative inside) and an electrochemical gradient (∆pH, alkaline inside) across the coupling membrane. The IM is impermeable to H+ and protons can reenter the matrix through the ATP synthase machinery. The ∆ψ is the driving force for mitochondria to uptake and retain cations such as Ca2+. Although mitochondria can accommodate relatively large quantities of calcium, the intramitochondrial ionized calcium concentration ([Ca2+]m) is maintained very low by at least two mechanisms: 1. The [Ca2+]m is precipitated in the matrix to form electron dense granules. The chemical structure of these granules may vary in mitochondria of different cells; however, these granules generally consist of calcium phosphate and calcium apatite. 2. The [Ca2+]m is released from mitochondria in exchange with other cations such as H+ or Na+. NO competes with O2 for the O2 binding site of complex IV and inhibits the O2 consumption in a reversible manner. Inhibition by NO of O2 consumption decreases the ∆ψ, ∆pH, and ATP. Exogenously added NO produced by the NO donor S-nitroso-N-acetylpenicillamine (SNAP) decreases the ATP synthesis by rat heart mitochondria (Table 4.1). Thus, 1, 3.5, and 7 µM NO dose-dependently decrease the ATP synthase activity to 62%, 44%, and 23%, respectively. These concentrations of NO do not affect the ATPase activity per se (Table 4.1; Saavedra-Molina et al., 2003), indicating that NO decreases the ATP synthesis via inhibition of the mitochondrial respiratory chain, and not the ATPase. Similar results have been obtained with mitochondria of endothelial cells (Ramachandran et al., 2004).

4.4 MITOCHONDRIAL NITRIC OXIDE SYNTHASE (mtNOS) Between 1995 and 1996, several studies suggested possible existence of a mitochondrially located NOS. Immunohistochemical studies using NADPH diaphorase staining (Bates et al., 1995; Frandsen et al., 1996) and silver-enhanced gold-immunolabeling (Bates et al., 1996) provided evidence for the presence of NOS-like proteins within mitochondria of different organs. Co-localization of eNOS with succinate dehydrogenase as a mitochondrial marker (Kobzik et al., 1995), and cross-reaction of mitochondria with antibodies directed against eNOS (Bates et al., 1995; 1996) or nNOS were reported (Frandsen et al., 1996). One

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TABLE 4.1 Effect of NO on ATP Synthesis and ATPase Activity in Rat Heart Mitochondria NOa 0 1.0 3.5 7.0 Oligomycin

ATPb 142 ± 15 88 ± 4 63 ± 3 33 ± 9 —

ATPasec 0.28 ± 0.02 0.24 ± 0.04 0.25 ± 0.03 0.27 ± 0.02 0.03 ± 0.001d

a. Concentrations of NO (µM) was achieved by using SNAP. b. ATP (nmoles/mg.min) was measured using the method as described. (From Castrejón et al., 1997. Arch. Biochem. Biophys. 346:37–4.) c. ATPase activity (µmoles/mg/min) was measured using the method as described. (From Guerra et al., 1995. Arch. Biochem. Biophys. 321:101–7.) d. Oligomycin was used as a positive control and added at a final concentration of 0.05 mg/ml. (Reproduced with permission).

study (Kobzik et al., 1995) also reported a faint NOS activity in rat diaphragm muscle mitochondria; however, they did not rule out the presence of nonmitochondrial NOS or the influence of the urea cycle in citrulline formation, which was used as the only NOS activity assay. In 1997, the first report on the presence of a constitutively expressed and continuously active NOS in mitochondria (i.e., mtNOS), its association with mitochondrial inner membrane, and determination of its activity were published (Ghafourifar and Richter 1997; also see Richter et al., 1999). It was also reported that mtNOS is Ca2+-sensitive (i.e., mtNOS activity increases with elevated [Ca2+]m) and that mtNOS exerts substantial control over mitochondrial O2 consumption and ∆ψ. Subsequently, these findings were confirmed by us (Ghafourifar and Richter 1999; Ghafourifar et al., 1999; Bringold et al., 2000; reviewed in Ghafourifar and Richter 2001; Szibor et al., 2001) and by several other groups (Arnaiz et al., 1999; Lacza et al., 2001; Manzo-Ávalos et al., 2002; Liang et al., 2002; Kanai et al., 2001; Carreras et al., 2001; López-Figueroa et al., 2000); however, few reports have published controversial results (see below).

4.5 CA2+-DEPENDENCE OF mtNOS One group used a single ADP-affinity column purification and purified a protein from mitochondrial matrix fraction that cross-reacts with an iNOS antibody and generates L-citrulline in an L-arginine-dependent and Ca2+-independent manner (Tatoyan and Giulivi 1998). The same group reported on an iNOS antibody cross-reacting protein in swine heart mitochondria that produces very low levels of L-citrulline in an L-arginine-dependent and Ca2+-independent manner (French et al., 2001). It has to be noted that many buffers traditionally used


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to investigate mitochondria, including the buffers those investigators have used, contain high concentrations (≥ 1 mM) of Mg2+. It is well known that Mg2+ blocks mitochondrial Ca2+ uptake (McKean 1991; Tsuda et al., 1991; Votyakova et al., 1993), and it can inhibit NOS activity (Howard et al., 1995). We and other researchers (Ghafourifar et al., 1999; Kanai et al., 2001) have clearly demonstrated that the blockade of liver and heart mitochondrial Ca2+ uptake (e.g., by ruthenium red or by collapsing ∆ψ) drastically decreases mtNOS activity and in a recent report, Manzo-Ávalos et al. (2002) demonstrated that Mg2+ potently inhibits mtNOS activity. Therefore, using high concentrations of Mg2+ in mtNOS research appears unnecessary, and in the presence of Mg2+, mtNOS can appear Ca2+-insensitive (reviewed in Ghafourifar 2002).

4.6 EFFECTS OF mtNOS ON MITOCHONDRIAL BIOENERGETICS mtNOS plays a major role in regulating mitochondrial bioenergetics. Intramitochondrially produced NO inhibits COX and decreases O2 consumption, ∆ψ and mitochondrial matrix pH (Ghafourifar and Richter 1997; Ghafourifar and Richter, 1999; Ghafourifar et al., 1999). Inhibition of the basal mtNOS activity increases basal mitochondrial O2 consumption and ∆ψ (Ghafourifar and Richter 1997), causes mitochondrial matrix alkalinization, and provides a resistance to the sudden drop of ∆ψ induced by mitochondrial Ca2+ uptake (Ghafourifar and Richter, 1999; Ghafourifar et al., 1999). These findings indicate that mtNOS is continuously active and regulates mitochondrial respiration and respiration-dependent processes. mtNOS provides a feedback regulatory mechanism that protects mitochondria against Ca2+ overload. Increased [Ca2+]m stimulates mtNOS and increases intramitochondrial NO (Ghafourifar and Richter 1997, Ghafourifar et al., 1999; Kanai et al., 2001) which causes Ca2+ efflux from mitochondria via at least two mechanisms: 1. Passive Ca2+ efflux as a result of decreased ∆ψ 2. Active Ca2+ release through intramitochondrial formation of pro-oxidants, such as ONOO−, which stimulates specific mitochondrial Ca2+ release pathway with preserved ∆ψ. The latter mechanism involves oxidization of mitochondrial pyridine nucleotides (Bringold et al., 2000). mtNOS plays a critical role in regulating the respiratory control ratio (RCR) values. In the absence of mtNOS activity, exogenously added NO decreases the RCR in a typical dose-dependent manner (Figure 4.1A). When mtNOS is stimulated by loading mitochondria with 0.1 µM Ca2+, basal RCR exhibits a lower value; however, these mitochondria are protected against further decrease in RCR induced by exogenously added NO (Figure 4.1B) (Saavedra-Molina et al., 2003).

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Thus, low levels of mtNOS-derived NO protect mitochondria against a drastic loss of RCR induced by high concentrations of exogenous NO (Saavedra-Molina and Devlin 1997; Saavedra-Molina et al., 2003). When mtNOS is over-stimulated by further increasing [Ca2+]m, exogenously added NO inhibits the state 3 and increases the state 4 of respiration, and thus, decreases the RCR (Figure 4.1C and Figure 4.1D). Overstimulation of mtNOS causes intramitochondrial peroxynitrite formation that induces oxidative damage to mitochondrial susceptible targets including the respiratory chain complexes, and can affect the generation of certain reactive oxygen species, such as H2O2, without affecting the ATP synthesis (Brookes and Darley-Usmar 2002).

4.7 MITOCHONDRIAL CA2+ HOMEOSTASIS Mitochondria remain one of the main cellular components of cellular Ca2+ oscillation and they actively participate in physiological cellular Ca2+ turnover (Robb-Gaspers et al., 1998; Rizzuto et al., 1998; reviewed in Pozzan et al., 2000; Rizzuto et al., 2000). The ∆ψ that renders the inside of the mitochondrial inner membrane negatively charged is the driving force for mitochondria to take up relatively large amounts of Ca2+ very rapidly. Intramitochondrial free Ca2+ concentration ([Ca2+]m) is maintained very low by several mechanisms, however, including release and precipitation of the Ca2+ to form electron-dense granules (Miyata et al., 1991; Coll et al., 1982; Carafoli 1987; reviewed in Tyler 1992). The content of these granules may vary in different physiological and pathological conditions (Ashraf and Bloor 1976; Karcsu et al., 1983); however, they consist mainly of tricalcium phosphate and hydroxyapatite. Earlier reports have suggested that rat liver and heart mitochondria contain 1 to 2 nmol Ca2+ per mg mitochondrial protein. Considering 7.2 × 109 mitochondria in each mg of mitochondrial protein and assuming a volume of 7.1 µm3 for each mitochondrion (Loud 1968), the intramitochondrial ionized calcium concentration is about 2 to 4 µM. Recent studies have detected lower [Ca2+]m (e.g., ≤ 100 nM in heart mitochondria) (Sheu and Sharma 1999; Miyata et al., 1991). In fact, intramitochondrial ionized Ca2+ in situ is much smaller than that in the ER (reviewed in Pozzan et al., 1994).

4.8 HEART mtNOS AND ITS CA2+-DEPENDENCE Several groups have reported heart mtNOS (Kanai et al., 2001; French et al., 2001; Manzo-Ávalos et al., 2002; Liang et al., 2002). Kanai identified an nNOS in mouse heart mitochondria that produces NO in a typical Ca2+-dependent manner. Manzo-Ávalos (2002) and Liang (2002) also reported that heart mitochondria generate NO in a Ca2+-sensitive manner. Our laboratory has observed that rat heart mtNOS cross-reacts with nNOS antibodies and produces NO in a Ca2+-sensitive fashion (unpublished results); however, one group (French et al., 2001) did not observe that heart mtNOS is Ca2+-dependent. As discussed previously, many buffers traditionally used to investigate mitochondrial functions,

Functions of Mitochondrial Nitric Oxide Synthase 100

100

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80 ∗

∗

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∗

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RCR (%)

80

20 0

85

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80 ∗

∗

∗

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∗

20

RCR (%)

RCR (%)

∗

20

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∗

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(NO) (µM)

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∗

∗

20 0.0

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FIGURE 4.1 Effect of NO on the respiratory control ratio. Oxygen consumption of isolated heart mitochondria (0.3 mg/ml) was supported with 800 µmoles ADP. The state 3 and 4 respiration (nmoles O2/min.mg) in the absence of Ca2+ and exogenous NO were 79 ± 8 and 10 ± 3, respectively. Panels represent the following: (A) no Ca2+ added; (B) 0.1 µM Ca2+; (C) 1.0 µM Ca2+; (D) 2.0 µM Ca2+. The concentration of free Ca2+ was calculated using the software Winmaxc 2.0. Data represent mean ± SEM, n ≥ 3, *p < 0.05.

including the ones used in that study, contain high concentrations of Mg2+. This cation is a known mitochondrial Ca2+ uptake blocker (McKean 1991; Tsuda et al., 1991; Votyakova et al., 1993) and blockade of mitochondrial Ca2+ uptake drastically decreases mtNOS activity (Ghafourifar et al., 1999; Kanai et al., 2001). Additionally, Mg2+ can inhibit the activity of NOS (Howard et al., 1995). The direct inhibitory effect of Mg2+ on mtNOS activity has been recently reported (Manzo-Ávalos et al., 2002; discussed below). Magnesium is an important intracellular cation involved in several functions, such as a co-factor for the Na+/Ca2+ ATPase and regulating co-transport of Na+, K+, and Cl− (Flatman 1991), and mitochondria maintain a dynamic Mg2+ homeostasis. For example, addition of cyclic AMP to mitochondria induces a net efflux of matrix Mg2+ (Romani et al., 1991) and Mg2+ appears to have a regulatory effect on the oxidative phosphorylation in intact heart mitochondria (Rodríguez-Zavala and Moreno-Sánchez 1998). Intramitochondrial matrix free Mg2+ controls the

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rate of L-citrulline synthesis through a direct interaction with carbamyl-phosphate synthase I (ammonia) (Rodríguez-Zavala et al., 1997). Therefore, intramitochondrial Mg2+ affects mitochondrial L-citrulline synthesis via pathways other than mtNOS. A recent report elucidates the direct effect of heart mitochondrial matrix free Mg2+ on mtNOS activity (Manzo-Ávalos et al., 2002). Incubation of heart mitochondria with increasing concentrations of extramitochondrial Mg2+ causes a dose-dependent decrease in heart mtNOS activity from 24 to 59% for 0.2 to 3.2 Mg2+, respectively mM (Figure 4.2), indicating that physiological range of Mg2+ inhibits the mitochondrial synthesis of NO (Manzo-Ávalos et al., 2002). Thus, in the presence of high concentrations (mM range) of Mg2+, as used in some recent studies (Giulivi et al., 1998; French et al., 2001), mtNOS exhibits lower activity and can appear Ca2+-insensitive (reviewed in Ghafourifar 2002).

4.9 CA2+ AND MITOCHONDRIAL APOPTOSIS Prolonged elevated cytoplasmic Ca2+ induces apoptosis. This type of apoptosis requires mitochondrial Ca2+ uptake (Smaili et al., 2000; Pacher et al., 2001; Hajnoczky et al., 2000; Stout et al., 1998; Kruman and Mattson 1999) and is prevented when elevation of [Ca2+]m is prevented (Stout et al., 1998; Kruman and Mattson 1999; Baek et al., 1997; Kruman et al., 1998; reviewed in McConkey and Orrenius 1997). Elevated Ca2+-induced apoptosis occurs with increased NOS activity (Stout et al., 1998; Almeida et al., 1998) and is prevented by the function of mitochondrial MnSOD (Keller et al., 1998; González-Zulueta et al., 1998) or by scavenging ONOO− (Kruman et al., 1998). We have suggested that mtNOS mediates the elevated Ca2+-induced apoptosis by generating ONOO− intramitochondrially (discussed below).

4.10 MITOCHONDRIA, mtNOS, OXIDATIVE STRESS, AND APOPTOSIS Mounting evidence indicates the crucial role of mitochondria in apoptosis (Green and Reed 1998; Kroemer 1999; Ghafourifar et al., 2000; Gottlieb 2000). Interaction with mitochondria is one of the prime early events in apoptosis induced by many apoptogenic factors such as ceramide, Bax, or NO. Additionally, mitochondria possess crucial pro- and anti-apoptotic proteins, such as Bax, caspase, AIF, and Bcl-2. Moreover, release of proteins, such as cytochrome c, triggers apoptosis in many cells. Thus, mitochondria are called the “switchboard of apoptosis” (Ghafourifar and Richter 2001; Szibor et al., 2001). Although apoptosis is an evolutionarily conserved mechanism needed for normal cell and tissue homeostasis (Kerr et al., 1972), unwanted apoptosis is the underlying mechanism in numerous pathological conditions such as heart ischemia/reperfusion (Gottlieb and Engler 1999). Many studies report that during hypoxia/reoxygenation [Ca2+]m is increased, ONOO− is elevated, cytochrome c is released, and mitochondria are malfunctioning. Mitochondria possess a Ca2+-sensitive NOS that is stimulated


87

(NO)m (µM)

.5

.4

.3

.2

.1 0

0.2

0.4

0.8

1.6

3.2

(Mg2+)e (mM)

FIGURE 4.2 Effect of Mg2+ on heart mtNOS activity. Rat heart mitochondria (1 mg/ml) were incubated in the medium containing 120 mM KCl, 20 mM MOPS, 0.5 mM EGTA, pH 7.40 in the presence of increasing concentrations of Mg2+ and mtNOS activity was measured as described. (From Manzo-Ávalos et al., 2002. Amino Acids 22:381–9.) Each point represents the mean ± SEM of at n ≥ 5 in duplicate. *p < 0.05.

upon elevation of [Ca2+]m, stimulation of mtNOS generates ONOO−, releases cytochrome c induces oxidative stress and malfunctioning mitochondria. Thus, it is plausible that mtNOS is involved in hypoxia/reoxygenation-induced cytochrome c release and oxidative stress (see below). In this view, a substantial number of recent reports indicate that endogenously formed NO induces apoptosis (Keller et al., 1998; Stout et al., 1998; GonzálezZulueta et al., 1998; Estevez et al., 1998; Brune et al., 1997; Messmer et al., 1996; Ferrante et al., 1999; Umansky and Schirrmacher 2001; Borutaite et al., 1999; Ghafourifar 1999b; Ghafourifar et al., 1999) through mechanisms that involve formation of RNOS such as ONOO− (Keller et al., 1998; Ferrante et al., 1999; Leist et al., 1997). This form of apoptosis is accompanied by mitochondrial dysfunction (Almeida et al., 1998; Keller et al., 1998) and perturbed mitochondrial redox balance (Keller et al., 1998). As discussed previously, prolonged elevated cytosolic Ca2+ induces apoptosis (reviewed by McConkey and Orrenius 1997) through a mechanism that: 1. Requires elevation of [Ca2+]m (Stout et al., 1998; Almeida et al., 1998; Kruman and Mattson 1999). 2. Occurs with increased NOS activity (Stout et al., 1998; Almeida et al., 1998).

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3. Is prevented by lowering mitochondrial O2− (Keller et al., 1998; González-Zulueta et al., 1998) or by scavenging ONOO− (Kruman et al., 1998) Mitochondria produce NO in a Ca2+-dependent fashion (Ghafourifar and Richter 1997; Ghafourifar et al., 1999; Bringold et al., 2000; Kanai et al., 2001), and they are well-known sources of O2−. Because the reaction of NO and O2− with the rate constant of 1.9 × 1010 M−1 s−1 (Kissner et al., 1997) is extremely rapid, intramitochondrial Ca2+-dependent ONOO− formation is, therefore, very likely. We have demonstrated that elevation of [Ca2+]m stimulates mtNOS, causes the release of cytochrome c, and induces mitochondrial oxidative stress through intramitochondrial ONOO− formation (Ghafourifar et al., 1999).

4.11 DOES mtNOS LINK HYPOXIA/REOXYGENATION, ONOO-, APOPTOSIS, AND RELEASE OF CYTOCHROME C? Hypoxia/reoxygenation increases myocardial Ca2+ and decreases ∆ψ (Ylitalo et al., 2000; Kloner and Jennings 2001). Prevention of this increased Ca2+ attenuates the cardiac damage (Vander Heide et al., 1994). Interestingly, hypoxia/reoxygenation also increases [Ca2+]m (Toyo-oka et al., 1989; Weiss et al., 2003) and causes mitochondrial malfunctioning (Pepe 2000). Although hypoxia per se induces necrotic cell death in cardiomyocytes, hypoxia/reoxygenation induces apoptosis (Gottlieb and Engler 1999) predominantly through mitochondria (Kang et al., 2000). Hypoxia/reoxygenation ignites the apoptosis machinery of mitochondria (Weiss et al., 2003) and induces cytochrome c release (Vanden Hoek et al., 2003) and inhibition of cytochrome c release inhibits reoxygenation-induced apoptosis (Kang et al., 2000). Moreover, formation of RNOS (Xie and Wolin, 1996; Xie et al., 1998) and loss of mitochondrial oxidative phosphorylation (Steinmann and Storm 1997; Xie 1998) are caused by hypoxia/reperfusion, but not hypoxia per se. Additionally, NO and ONOO− are increased during hypoxia/reoxygenationinduced cardiac injury (Xie et al., 1998; Ma et al., 2000; Ronson et al., 1999; Nakamura et al., 2000) and attenuation of NO or ROS (Ihnken et al., 1997) or augmentation of GSH (Nakamura et al., 2000) improves the injury. Likewise, lessening levels of mitochondrial O2− by overexpressing MnSOD (Chen et al., 1998; Xie et al., 1998) or scavenging ONOO− (Xie et al., 1998) protects myocardial cells against hypoxia/reoxygenation-induced injury. Mitochondria possess a Ca2+-sensitive mtNOS and elevation of [Ca2+]m causes intramitochondrial formation of RNOS, such as ONOO−, induces oxidative stress and release of mitochondrial cytochrome c, and causes mitochondria malfunction. Thus far, it appears very likely that mtNOS plays an undetected role in oxidative stress and apoptosis during hypoxia/reoxygenation. A very recent study has ignited supportive evidence. It has been reported that heart mtNOS activity significantly decreases during the hypoxia: from


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0.36 ± 0.02 µM in the control to 0.24 ± 0.01 µM after a 5 min hypoxia, to 0.22 ± 0.01 µM after 15 min and to 0.17 ± 0.02 µM at 30 min after the onset of hypoxia (Figure 4.3A). Upon reoxygenation, mtNOS activity was increased and reached the normal value of 0.34 ± 0.01 µM after 5 min of reoxygenation (Figure 4.3A). This study also suggests that heart mtNOS-derived NO can be stored during hypoxia (Saavedra-Molina et al., 2003). Intact heart mitochondria energized with glutamate plus malate produces 1 µM NO at atmosphere condition (Figure 4.3B). When these mitochondria are incubated in hypoxia for 24 min (Figure 4.3B), a large amount of NO (7 µM; p < 0.05) is released from these mitochondria upon reoxygenation. The concentration of oxygen during the 24min hypoxia is depicted in the insert for Figure 4.3B.

4.12 ANTIOXIDANTS PREVENT HYPOXIA/REOXYGENATION-INDUCED OXIDATIVE INJURY Deleterious effects of oxidative stress and the beneficial roles of antioxidants in reducing the oxidative injury during aging or pathological conditions including hypoxia/reoxygenation have been broadly studied (Ronson et al., 1999; Ames et al., 1993; Das and Maulik 1994; Kloner et al., 1989; Cadenas and Davies 2000; Vanden Hoek et al., 2003). A significant loss in tissue hydrophilic antioxidants, such as ascorbate and GSH, has been observed during hypoxia/reoxygenationinduced injury in isolated rat heart (Haramaki et al., 1998). Glutathione reverses the deleterious effects of ONOO− (Nakamura et al., 2000) by converting the ONOO− produced during hypoxia/reoxygenation to nitrosothiols and related products (Ronson et al., 1999). Alpha-tocopherol acetate, a water-soluble vitamin E derivative, significantly suppresses the ischemia/reperfusion-induced increase in 8-hydroxydeoxyguanosine (8-OH-dG) levels, a marker of oxidative damage (Yang et al., 1999). Raxofelast, a hydrophilic vitamin E analogue, prevents lipid peroxidation (LPO) elevation and GSH depletion and recovers the deteriorated functions of rat heart induced by ischemia/reperfusion (Campo et al., 1998). Ascorbic acid and Trolox, a potent ONOO− scavenger hydrophilic analogue of alpha-tocopherol, reduce oxidative injury induced by hypoxia/reoxygenation in isolated perfused rat hearts (Molyneux et al., 2002). Trolox also inhibits apoptosis induced by exogenously added NO (Umansky and Schirrmacher 2001; Takabayashi et al., 2000) or by prolonged increased cytosolic Ca2+ (Vergun et al., 2001), and prevents LPO of heart mitochondrial membranes (Santos and Moreno 2001).

4.13 DOES mtNOS PLAY A ROLE IN HYPERTENSION? NO, one of the most powerful vasodilators, is involved in the pathology of diseases such as hypertension (Moncada et al., 1991; Ignarro et al., 1999). For example, NOS activity is decreased in hypertension (Shesely et al., 1996; Miyamoto et al., 1998) and NOS inhibitors increase the blood pressure (Qiu et al., 1998). Possible

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Hypoxia Reoxygenation Normoxia

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FIGURE 4.3 Effect of hypoxia-reoxygenation on heart mtNOS activity. Heart mitochondria (1 mg/ml) were incubated in the medium containing 120 mM KCl, 20 mM MOPS, 0.5 mM EGTA, pH 7.40 plus 5 mM glutamate-malate, 10 mM NaCl, 5 mM NaH2PO4. Hypoxia was obtained by saturating the medium with 95% N2/5% CO2 and reoxygenation with 95% O2/5% CO2. The gas exchange did not cause any pH changes (not shown). Panel A: mtNOS activity was measured as described. (From Manzo-Ávalos et al., 2002. Amino Acids 22:381–9.) Panel B: Heart mitochondria (0.3 mg/ml) incubated in hypoxia followed by reoxygenation. Data represent the mean ± SEM of n ≥ 3 in duplicates. *p < 0.05.


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involvement of mtNOS of heart and kidney, two organs highly involved in the pathology of hypertension, was recently studied (Aguilera-Aguirre et al., 2002). This study also reported that the basal [Ca2+]m is significantly lower in both organs from spontaneously hypertensive rats (SHR) compared with normotensive genetic control WKY. This study also reported that inhibition of heart and kidney mtNOS inhibits the Ca2+-induced permeability transition pore (PTP) opening in normotensive (WKY), but not in hypertensive SHR. This finding suggests that heart and kidney mitochondria from SHR would have lower basal NO production than that of normotensive controls (Aguilera-Aguirre et al., 2002). In a supportive finding, this study demonstrates that basal heart and kidney mitochondrial matrix pH (pHi) of SHR is lower than in WKY ones. It worth reminding that NO added exogenously or produced by mtNOS decreases the pHi and inhibition of mtNOS increases the pHi (Ghafourifar and Richter 1999). These data suggest that mtNOS may play a role in physiological regulation of blood pressure.

4.14 ACKNOWLEDGMENTS This work was supported by the National Institute on Aging (AG023264-02; to P.G.), National Center for Research Resources, Marshall University Center of Biomedical Research Excellence (1 P20 RR020180; to P.G.), and the Mexican grants of CONACYT (43705; to A.S-M.) and CIC-UMSNH (2.16; to A.S-M.).

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A Mediator 5 Peroxynitrite: of Nitric Oxide-Dependent Mitochondrial Dysfunction in Pathology Celia Quijano, Adriana Cassina, Laura Castro, Marianela Rodriguez, and Rafael Radi Universidad de la República, Montevideo, Uruguay

CONTENTS 5.1 5.2 5.3

Introduction ............................................................................................100 Peroxynitrite Formation in Mitochondria .............................................101 Peroxynitrite Reactions and Their Role in Mitochondrial Dysfunction and Apoptosis ...........................................102 5.3.1 Peroxynitrite Reactivity ..............................................................102 5.3.2 Peroxynitrite-Dependent and Independent Pathways Involved in Tyrosine Nitration in Mitochondria ........................103 5.3.2.1 Peroxynitrite-Mediated Nitration of Tyrosines ...........103 5.3.2.2 Peroxidase, Peroxide, and Nitrite-Mediated Nitration of Tyrosine ...................................................105 5.3.2.3 Fenton Chemistry and Tyrosine Nitration ...................106 5.3.2.4 ·NO Reaction with Tyrosyl Radicals Leads to Tyrosine Nitration ........................................................106 5.3.2.5 Tyrosine Nitration Is a Radical Termination Reaction That Evidences ·NO Formation ...................107 5.3.3 Peroxynitrite Reactions with Components of the Energy Metabolism .................................................................................107 5.3.3.1 Electron Transport Chain Components and ATP Synthase .......................................................................107 5.3.3.2 Intermembrane Components ........................................108 5.3.3.3 Matrix Components .....................................................109 5.3.4 Peroxynitrite Reactions with Components of the Apoptotic Machinery .................................................................109

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5.4

5.5 5.6 5.7


5.3.5 Peroxynitrite Reactions with Mitochondrial Antioxidants ........110 Peroxynitrite Mediates Mitochondrial Dysfunction in Pathology ........115 5.4.1 Diabetes ......................................................................................115 5.4.1.1 Peroxynitrite Formation in Diabetes ...........................115 5.4.1.2 Evidence of Peroxynitrite-Mediated Mitochondrial Dysfunction in Diabetes ..............................................116 5.4.2 Sepsis ..........................................................................................117 5.4.2.1 Peroxynitrite Formation in Sepsis ...............................117 5.4.2.2 Evidence of Peroxynitrite-Mediated Mitochondrial Dysfunction in Sepsis ..................................................118 5.4.3 Neurodegenerative Diseases .......................................................119 5.4.3.1 Peroxynitrite Formation and Mitochondrial Dysfunction in Neurodegenerative Diseases ...............119 Mitochondrial-Targeted Pharmacology .................................................121 Conclusions ............................................................................................122 Acknowledgments ..................................................................................122 References ...............................................................................................123

5.1 INTRODUCTION Mitochondria, due to their ability to couple the energy released during electron transfer to ATP synthesis, constitute the major site of bioenergetic control in most mammalian cells. In addition, because numerous mitochondrial proteins participate in apoptosis, they are now acknowledged as key sites of cellular death control. Mitochondria are also the major intracellular source of superoxide (O2.−) and hydrogen peroxide (H2O2), and a primary locus for the reactions of reactive species with target molecules. In this context, peroxynitrite (ONOO−), a potent oxidant and nitrating species, product of the fast reaction between nitric oxide (.NO) and superoxide, can be formed within mitochondria and impact in its homeostasis. Mitochondrial dysfunction is a relevant factor in many human diseases, including neurodegenerative diseases, ischemia-reperfusion injury, aging, inflammatory damage, and diabetic complications. Reactive oxygen and nitrogen species have also been implicated as causative or contributory agents in many of these pathologic processes. In particular, peroxynitrite-mediated damage is being increasingly screened in different disease states, and is hypothesized to be involved in the onset and progression of different pathologies. In this scenario, the role of peroxynitrite in mediating mitochondrial dysfunction and/or apoptotic signaling arises as an active and promising area of research, directed to unravel the molecular basis of many diseases and in the rational design of drugs.

Peroxynitrite

101

5.2 PEROXYNITRITE FORMATION IN MITOCHONDRIA As peroxynitrite anion (ONOO−) is formed in the diffusion-controlled reaction (k ~ 1 × 1010 M−1s−1) of ·NO with O2•− [1–4], its formation is closely related to the variation in the formation and decomposition rates, as well as to the diffusion of these radicals. Within mitochondria, superoxide is formed in the outer mitochondrial membrane, in the matrix, and on both sides of the inner mitochondrial membrane (recently reviewed by Turrens [5]); complex I and III of the electron transfer chain is responsible for most of the superoxide produced under normal and pathological conditions [6–9]. The steady-state concentration of superoxide ([O2.−]ss) in the mitochondrial matrix can be calculated considering the rate of superoxide production by mitochondria (Rp) (estimated value, Rp = 0.57 × 10−6 M s−1 [10]), and its decomposition rate by the reaction catalyzed by Mn-SOD (kSOD = 2 × 109 M−1 s−1 [11], [SOD] 1 × 10−5 M), rendering values of 10−11 M. This value is in agreement with the [O2.−]ss measured in E. coli [12]. In the same way, a mitochondrial intermembrane superoxide steady-state concentration results from superoxide production toward this compartment [13] and from the activity of intermembrane Cu,Zn-SOD [14–16]. Mitochondrial superoxide production is largely increased during different pathological conditions [17–22], and its physiological steady-state concentration is about five- to tenfold higher than in the cytosol and nucleus [10]. ·NO is formed by ·NO synthases (NOS), which catalyze the N-oxygenation and oxidative cleavage of the guanidino group of arginine. The constitutive forms, namely endothelial NOS (eNOS) and neuronal NOS (nNOS) produce shortlasting, small quantities of ·NO, whereas the inducible NOS (iNOS) has a longlasting release of higher quantities [23]. ·NO formation rates and steady-state concentrations of ·NO ([·NO]ss) also differ with the physiological or pathological situation, increasing during diseased states [24–30]. More recently, a mitochondrial NOS (mtNOS) has been described [31–35]. This enzyme is a variant of the nNOS isoform and has been found in liver, heart, and bladder [33, 36, 37]. Mitochondria isolated from the liver and heart reportedly produce ·NO [36], which modulates oxygen consumption, ATP [38] and superoxide production [39], as well as apoptosis [31]. Mitochondrial ·NO production in pathology is not well defined. However, roles in bladder radiation damage [37], cardiomyocyte protection during calcium overload [33], and sepsis [24] have been reported, but this area still requires extensive further study. Although NOS enzymes are the major source of ·NO, it has been proposed that complex III of the electron transport chain can reduce nitrite to ·NO [40], resulting in an alternative source of mitochondrial ·NO. An increase in either superoxide or ·NO formation, due to mitochondrial or non-mitochondrial sources, will result in an increase in peroxynitrite formation [40a]. The anionic character of superoxide (pKa = 4.7) limits its diffusion across membranes [41, 42] and although superoxide has been described to cross the membranes through anion channels [43, 44], its reactivity is mostly confined to

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the compartment where it is formed. ·NO, on the contrary, is a hydrophobic and diffusible molecule that readily crosses cellular membranes [45–47]. ·NO diffusion to mitochondria is confirmed by all the studies that describe NO actions on the electron transport chain, such as regulation of cell respiration [48, 49]. Once inside the mitochondria, ·NO will face many targets; in physiological conditions superoxide, due to its low concentrations, will probably be out-competed by other targets, such as metalloproteins [50], particularly cytochrome oxidase [51–55]. Nevertheless, increases in superoxide concentrations, such as those observed in pathological situations, will shift the reactivity of ·NO toward superoxide because the reaction becomes kinetically favored. In fact, peroxynitrite formation is reported to occur during sustained mitochondrial ·NO production and many nitrated mitochondrial proteins identified under these conditions [56]. In vitro and in vivo evidence for peroxynitrite formation in mitochondria has been thoroughly revised in a previous review [57] and new evidence is presented throughout this text. An alternative route of peroxynitrite formation in mitochondria is the reaction of nitroxyl anion (NO−) with molecular oxygen (O2) [58, 59]. Nitroxyl anion can originate from the reaction of ·NO with ubiquinol [60] or cytochrome c [61].

5.3 PEROXYNITRITE REACTIONS AND THEIR ROLE IN MITOCHONDRIAL DYSFUNCTION AND APOPTOSIS 5.3.1 PEROXYNITRITE REACTIVITY Peroxynitrite anion (ONOO−) and its conjugated acid peroxynitrous acid (ONOOH, pKa = 6.8) quickly react with many target molecules present in the mitochondria and their reactions can be roughly divided in two groups [1]: 1. Direct reactions leading to one or two-electron oxidation of targets, such as ferrous cytochrome c or thiols. 2. Indirect reactions that involve the rupture of peroxynitrite peroxo-bond (O-O) and lead to the formation of the secondary radicals, carbonate radical (CO3.−), nitrogen dioxide (·NO2) and hydroxyl radical (.OH). These radicals mediate one-electron oxidation of target molecules, nitration, and hydroxylation reactions. In mitochondria, carbon dioxide, metalloproteins, and thiols constitute preferential targets for direct reaction with peroxynitrite, due to their abundance and high kinetic constants. Carbon dioxide (CO2) is mainly produced in the decarboxylation reactions of pyruvate dehydrogenase and the citric acid cycle, so its concentration is particularly high in the mitochondrial matrix (~1.5 to 2 mM). Carbon dioxide reacts with peroxynitrite (k = 4.6 × 104 M−1s−1 at pH 7.4 and 37ºC) rendering nitrogen dioxide and carbonate radical [62–64], which will, in turn, oxidize and nitrate target molecules. The peroxynitrite/carbon dioxide path-

Peroxynitrite

103

way is highly relevant in mitochondria, and contributes to the shorter half-life of peroxynitrite in mitochondria (t1/2 ~ 3 to 5 ms), in comparison with the extracellular media (t1/2 ~ 10 ms) [57]. Although many of the reactions of peroxynitrite in mitochondria will be, in fact, mediated by carbonate radical and nitrogen dioxide, the direct reaction of peroxynitrite with metalloproteins is highly relevant because it produces radicals in the proximity of the metal center that lead to site-specific oxidation or nitration of amino acids near to the active site of the protein, where the metal is usually found. These reactions can have profound effects in activity, most notably the nitration of tyrosine-34 in the active site of Mn-SOD [65, 66] leads to enzyme inactivation via a manganese-dependent mechanism [67].

5.3.2 PEROXYNITRITE-DEPENDENT AND INDEPENDENT PATHWAYS INVOLVED IN TYROSINE NITRATION IN MITOCHONDRIA In summary, peroxynitrite and its derived radicals mediate oxidation and nitration reactions and their reactivity has been recently described in terms of nitroxidative stress [68]. Oxidation processes mediated by peroxynitrite occur with higher yields than nitration [69–71] and can well be responsible for changes in cell or mitochondrial homeostasis (e.g., protein and non-protein thiol oxidation [72–75]). However, although oxidation reactions can be mediated by different reactive oxygen and nitrogen species, nitration is a more specific biomarker denoting ·NOderived species involvement during mitochondrial oxidative damage. Peroxynitrite causes the nitration of both tyrosine and tryptophan residues forming 3-nitrotyrosine [76], 5- and 6-nitrotryptophan [71, 77], respectively, and 3-nitrotyrosine was initially considered a “footprint” of peroxynitrite formation in vivo. Nevertheless, other nitrating agents have been described; therefore, nitration per se might not be considered conclusive evidence of peroxynitrite formation. We will now describe in detail the biochemical pathways involved in nitration reactions, due to their relevance in the assessment of peroxynitrite formation in pathology, as well as the alternative nitration pathways that may occur in mitochondria. 5.3.2.1 Peroxynitrite-Mediated Nitration of Tyrosines Peroxynitrite nitration reactions proceed through its direct reaction with a Lewis acid (LA) that promotes the homolysis of the O-O bond with the concomitant formation of an oxidizing intermediate (LA-O.−) and nitrogen dioxide [1]. In general, the reaction can be represented as ONOO− + LA → LA-O.− + ·NO2 In mitochondria carbon dioxide, transition metal centers (Men) and in a minor degree protons (H+) are the Lewis acids involved in this process, whereas carbonate radical (CO3.−), oxo-metals (Men+1 = O) and hydroxyl radical (.OH), are

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the oxidizing intermediates, respectively [78–82]. These oxidizing species abstract one electron from the tyrosine aromatic ring, forming a tyrosyl radical that then recombines with nitrogen dioxide, forming 3-nitrotyrosine (see Figure 5.1, central part of the drawing).

e

.NO Tyr H2O2 P-Fe

NO−2

e−

I

f

O2

. Tyr

O2.−

Tyr

TyrNO2

cyt c

III

Mn

IV

V

.NO

ONOO− CO2

a

Tyr

TyrNO2

2

Q

II

P-Fe3+

.NO

e−

Tyr

O

− . Tyr e . 4+ P-Fe O

Tyr . P-Fe4+ O

3+

Tyr

b

CO3.−

Mn+1 .NO

.NO

2

2

. Tyr Mn

TyrNO2

HCO3− . Tyr

TyrNO2

Mn

c

NO2−

.NO

2

H2O2 2+

ATP - Fe

4+

3+

ATP - Fe = O Tyr

ATP - Fe d

. Tyr

TyrNO2

Peroxynitrite

105

Peroxynitrite nitration yields are highly increased by carbon dioxide and transition metals, such as iron, copper, and manganese [70, 80–82], which are found in mitochondria bound to proteins or low molecular weight chelators. The metal centers of Mn-SOD (matrix), Cu,Zn-SOD (intermembrane space), and catalase (matrix of heart mitochondria) quickly react with peroxynitrite [67, 83] and can promote the nitration of tyrosines [67, 70, 84, 85]. Iron bound to low molecular weight chelators is also capable of increasing peroxynitrite nitration yields [70, 76]. Due to the abundance of iron chelators in mitochondria, such as adenine nucleotides, tri- and di-carboxylic acids, and phosphate, this nitrationpathway could also become relevant when iron or other transition metals are mobilized from their storage proteins [86]. 5.3.2.2 Peroxidase, Peroxide, and Nitrite-Mediated Nitration of Tyrosine In addition to the peroxynitrite-mediated pathway, 3-nitrotyrosine is formed, both in vitro and in vivo, by heme peroxidases, hydrogen peroxide, and nitrite [87–91]. Heme peroxidases can catalyze the oxidation of nitrite to nitrogen dioxide by hydrogen peroxide [92]. Nitrogen dioxide can, in turn, promote the nitration of free and protein tyrosine, oxidizing tyrosine to tyrosyl radical (k = 3.2 × 105 M−1s−1 at pH 7.5 and 20°C), which then recombines with a second molecule of nitrogen dioxide (k ~ 3 × 109 M−1s−1 at pH 7.5 and 20°C) [93]. Nitrogen dioxide alone is an inefficient nitrating agent, first, because two nitrogen dioxide molecules are needed to nitrate one tyrosine, and second, because nitrogen dioxide oxidation of tyrosine is slow in comparison with other reactions, such as the oxidation of thiols (2–5 × 107 M−1s−1 at pH 7.4 and 22°C [94]) [86]. FIGURE 5.1 Biochemical mechanisms of mitochondrial protein tyrosine nitration. The drawing illustrates potential mechanisms of protein tyrosine nitration in different mitocondrial compartments. The central part of the drawing depicts the peroxynitritedependent pathways of mitochondrial nitration. Peroxynitrite formed from the diffusioncontrolled reaction between nitric oxide and superoxide can react with transition metalcontaining proteins (e.g., Mn-SOD) to form oxo-metal complexes plus nitrogen dioxide (a); the oxo-metal complexes can attack vicinal tyrosines, yielding tyrosyl radical that recombines with nitrogen dioxide to yield protein 3-nitrotyrosine. Peroxynitrite can also react with carbon dioxide to yield carbonate and nitrogen dioxide radicals that can also yield 3-nitrotyrosine (b). The lower and upper parts of the cartoon depict peroxynitriteindependent mechanisms of nitration. In the lower part, low molecular weight chelates of transition metals (e.g., ATP- Fe2+) react with hydrogen peroxide to form an oxo-metal complex that can alternatively attack nitrite (c) or tyrosine (d) to form the corresponding radicals, which then form 3-nitrotyrosine. In the upper part, a hemeprotein (e.g., cytochrome c reacts with hydrogen peroxide to form an oxo-ferryl iron, which can attack nitrite to form nitrogen dioxide (f), and a tyrosyl radical. Nitrogen dioxide then reacts with tyrosyl radicals forming 3-nitrotyrosine. An alternative mechanism of nitration has been postulated that involves the transient formation of nitrosotyrosine (e), which can evolve to 3nitrotyrosine, presumably through the intermediate formation of iminoxyl radical.

106


Mitochondrial nitration by the peroxidase pathway would require mitochondrial peroxidases. In this sense, cytochrome c is a mitochondrial protein with a weak peroxidase activity that can promote hydrogen peroxide-mediated oxidation of different substrates [95–100]. Moreover, the peroxidatic activity of cytochrome c is augmented when oxidatively damaged [100] and was recently reported to catalyze nitration of low molecular weight aromatics and protein tyrosine residues, including self-nitration [101] (see Figure 5.1, upper part of the drawing). Though depletion of cytochrome c significantly reduced nitration by hydrogen peroxide and nitrite in mitoplasts [101], its role mediating mitochondrial nitration in vivo is yet to be established. 5.3.2.3 Fenton Chemistry and Tyrosine Nitration Fenton chemistry involves the oxidation of organic molecules by ferrous ion (Fe+2) and hydrogen peroxide; this reaction, which has been known for over 100 years [102], is still a matter of debate and appears to be involved in new radical pathways such as tyrosine nitration. Free and chelated ferrous ion (Fe2+) react with hydrogen peroxide producing strong oxidizing species, whose nature is still under debate, being hydroxyl radical and hypervalent iron (IV) compounds, such as an oxo-ferryl (Fe4+ = O) species, alternative candidates [103]. Protein tyrosine nitration occurs in the presence of hydrogen peroxide, nitrite, and both heme and free transition metals; the oxoferryl species is considered responsible for nitrite and tyrosine oxidation to nitrogen dioxide and tyrosyl radical, respectively, leading to 3-nitrotyrosine formation (see Figure 5.1, lower part of the drawing) [104, 105]. This nitration pathway heavily relies on iron or copper availability and as we know these metals are usually found bound to proteins. Superoxide can mobilize iron from isolated aconitase [106]; and in cells, increased intramitochondrial formation of superoxide inactivates mitochondrial aconitase [107] and probably results in iron release from its active site. In addition, ·NO decreases ferritin levels and the activity of complexes I, II, and IV of the respiratory chain, with a concomitant increase in the labile iron pool [108]. Increases in superoxide and ·NO will be followed by increases in hydrogen peroxide and nitrite concentrations; therefore, the same scenario that will promote peroxynitrite formation can lead to nitration through this “Fenton-like” pathway [104, 105]. 5.3.2.4 NO Reaction with Tyrosyl Radicals Leads to Tyrosine Nitration An alternative pathway that also leads to tyrosine nitration involves the reaction of a tyrosyl radical with ·NO to form 3-nitrosotyrosine that is then monovalently oxidized to iminoxyl radical, and then to 3-nitrotyrosine [109]. In particular, cytochrome c tyrosine nitration can be mediated by hydrogen peroxide, which generates the tyrosyl radical and ·NO (see Figure 5.1e) [110]. We must note that although free nitrosotyrosine can reversibly generate tyrosyl radical and ·NO

Peroxynitrite

107

[111], in certain heme-proteins, such as cytochrome c, this adduct is oxidized by the heme rendering the stable 3-nitrotyrosine residue. 5.3.2.5 Tyrosine Nitration Is a Radical Termination Reaction That Evidences ·NO Formation All the nitration pathways described herein are radical processes in which a tyrosyl radical, formed by a highly oxidant species, recombines with either nitrogen dioxide or ·NO, forming the same end product, 3-nitrotyrosine. As recently stated in reviews by Turko and Murad [112] and Radi [86], the different nitration pathways (peroxynitrite-, peroxidase-, Fenton-, and nitric-oxidedependent nitration pathways) probably overlap in vivo and specific conditions will favor one over the other. Still, we must note that most of the available evidence on mitochondrial nitration points toward the involvement of peroxynitrite-dependent instead of independent pathways, as will be substantiated in further sections of this chapter.

5.3.3 PEROXYNITRITE REACTIONS ENERGY METABOLISM

WITH

COMPONENTS

OF THE

Mitochondrial energy metabolism is impaired in many pathologic conditions in which peroxynitrite is reportly formed and detailed examples will be presented later in this chapter. Moreover, proteomic analysis of mitochondria obtained from septic and diabetic animals reveal that several proteins, components of the energy metabolism, are nitrated in vivo, including ATP synthase and enzymes of the citric acid cycle [113, 114]. This section will discuss in vitro studies with isolated mitochondria or purified enzymes, which have helped to clarify the role of peroxynitrite and its mechanism of action in mitochondrial dysfunction. 5.3.3.1 Electron Transport Chain Components and ATP Synthase Experiments using isolated mitochondria demonstrate that although ·NO competes with oxygen for the binding to cytochrome c oxidase (complex IV) and modulates the respiratory rate [51–54], peroxynitrite reactions with complex I, II, and V produce their inactivation [53, 115–119]. A proteomic approach combined with a selective labeling of mitochondrial thiols confirms the presence of free thiol residues on the matrix surface of respiratory complexes I, II, and IV [74]; and inactivation of complex I, in isolated mitochondria exposed to peroxynitrite, depends on thiol oxidation and formation of S-nitrosothiol-derivatives [49, 116]. In addition, positive immunostaining for 3-nitrotyrosine in complex I was seen in dopaminergic cells exposed to peroxynitrite fluxes [120] and nitration of specific tyrosine residues, namely Tyr122 of subunit B14 and Try46, Tyr50, Tyr51 of subunit B15, was confirmed when enriched complex I preparations were exposed to peroxynitrite [119]. In fact, complex I

108


inactivation is observed in Parkinson’s disease [9], sepsis [121], and diabetes [122]. Therefore, studies concerning its interactions with peroxynitrite in vivo appear as a relevant issue. Inactivation of complex II may also involve the oxidation of the critical thiol of succinate dehydrogenase, present in the dicarboxylate binding site (Cys252 of subunit A), as observed during Trypanosoma cruzi exposure to peroxynitrite [123, 124]. Nitrated ATP synthase is also found in septic animals [125] and in isolated mitochondria. ATP synthase is inactivated when exposed to peroxynitrite [53, 115, 119] and nitrated during excess ·NO production [56]. A critical tyrosine (Tyr368 of beta subunit of F1) or a thiol residue involved in F1F0 interaction [126] are the candidate targets responsible for peroxynitrite-mediated enzyme inactivation; nevertheless this remains to be established. Incubation of isolated cytochrome c with peroxynitrite results in the direct one-electron oxidation of the ferrous heme (P-Fe2+) (k = 2.3 × 105 M−1s−1) forming ferric cytochrome c (P-Fe3+) [127], which is in turn nitrated by peroxynitritederived radicals [100, 100a]. Cytochrome c is also nitrated in vivo, during chronic allograft nephropathy [128], renal, and brain ischemia reperfusion [129, 130]. Nitration of cytochrome c not only impairs its function as electron carrier in mitochondria but also results in a gain of peroxidatic activity [100, 100a, 101], and may be involved in its migration to the cytosol [130]. Cytochrome c is present in high concentrations in the intermembrane space (>1 mM) [96], making it a preferential target for peroxynitrite reactions. Although isolated complex IV is inactivated by peroxynitrite [131, 132], in intact mitochondria, complex IV-dependent respiration is not affected by peroxynitrite [53, 115]. In fact, the reduced form of cytochrome c oxidase may serve to catalytically decompose peroxynitrite by a two-electron reduction to nitrite [133]. 5.3.3.2 Intermembrane Components Mitochondrial creatine kinase (Mt-CK) was found nitrated in vivo in animal models of inflammatory challenge, and diabetes [113, 114] and its activity is impaired during diabetic cardiomyopathy [134], heart ischemia [135], and in an animal model of amyotrophic lateral sclerosis [136]. Inactivation of Mt-CK by peroxynitrite has been observed after the exposure of intact mitochondria to peroxynitrite, at concentrations under which respiration is not affected [137]; thus, it appears as an extremely sensitive target. Studies with the isolated enzyme reveal that the enzyme is inactivated solely through the oxidation of the active site residue, Cys278 [137, 138], which reacts with peroxynitrite with a high rate constant (k = 9 × 105 M−1s−1)[139]. Tyrosine nitration and Trp223 oxidation are also observed during the incubation of the enzyme with peroxynitrite but are not the primary trigger of Mt-CK inactivation [137]. Peroxynitrite also promotes the dissociation of Mt-CK octamers to dimers through the oxidation of Met267 and the nitration of Trp264 or Trp268, located at the dimer/dimer interface [138]. The octameric structure of Mt-CK is crucial for the transport of phosphocreatine and

Peroxynitrite

109

creatine between the mitochondria and the cytosol, and controls the mitochondrial permeability transition [140]. Therefore, peroxynitrite-mediated oxidation of the enzyme will probably have a high impact on mitochondrial and cellular physiology. Carnitine palmitoyltransferase I activity is decreased and nitrated in the hearts of septic suckling rats [141, 142], and exposure of mitochondria to a peroxynitritegenerating system composed of superoxide and ·NO fluxes results in enzyme inactivation [141]. 5.3.3.3 Matrix Components Mitochondrial aconitase is nitrated in septic animals [113]. In vitro studies revealed that its iron sulfur cluster (4Fe-4S) is rapidly oxidized by peroxynitrite in a direct reaction (k = 1.4 × 105 M−1s−1) that disrupts the cluster to the 3Fe-4S form, releases free iron, and leads to enzyme inactivation [106, 143]. Therefore, although reaction of peroxynitrite-derived radicals with aconitase tyrosines occurs, cluster oxidation determines aconitase inactivation. Aconitase inactivation has also been observed in cells in which peroxynitrite was being formed by increased mitochondrial superoxide production and exposure to a ·NO donor [107]. Aconitase is also inactivated by superoxide (k ~ 107 M−1s−1) [106] and may even be inactivated by ·NO and S-nitrosothiols in certain conditions [107, 144]. Nevertheless, in the presence of both ·NO and superoxide, peroxynitrite formation will probably prevail, the latter being the oxidant responsible for aconitase inactivation. Isocitrate dehydrogenase was found nitrated in isolated mitochondria after mtNOS activity was stimulated [56]. In vitro studies reveal that in the isolated enzyme peroxynitrite induces S-nitrosylation of Cys305 and Cys387, inactivating the enzyme, along with nitration of Tyr280 [145]. Peroxynitrite also nitrates and inactivates isolated α-ketoglutarate dehydrogenase [146]. The activity of this enzyme is severely reduced in many neurodegenerative diseases, including Alzheimer’s disease [147] and Parkinson’s disease [148]. Other key enzymes for mitochondrial energy production that were identified as nitrated in vivo include succinyl-CoA:3-oxoacid CoA-transferase (SCOT), which is also inactivated [114, 149, 150], glutamate dehydrogenase [113], and several enzymes of the β-oxidation of fatty acids [113].

5.3.4 PEROXYNITRITE REACTIONS APOPTOTIC MACHINERY

WITH

COMPONENTS

OF THE

The mitochondrial apoptotic pathway is activated by the release of cytochrome c and other pro-apoptotic factors, including caspase-9 and AIF, from the mitochondria that promote the assembly of the apoptosome, the activation of downstream caspases and DNA cleavage. Cytochrome c release occurs through the mitochondrial permeability transition pore (PTP), the main components of which are

110


believed to include the voltage-dependent anion channel (VDAC), the adenine nucleotide translocase (ANT), and cyclophylin D (CyP-D). Other proteins, including cytosolic kinases, mitochondrial creatine kinase, and Bcl/Bax family proteins, are associated with the pore and have a modulatory action. ANT is an inner membrane integral protein that forms a complex with the outer membrane VDAC, and oxidation of ANT vicinal thiols leads to the recruitment of CyP-D, present in the matrix, forming the basic unit of the pore. Calcium overload and oxidative stress promote pore opening whereas adenine nucleotides exert an inhibitory action [151, 152]. Currently, widespread evidence indicates that reactive species are important mediators of apoptosis. Exposure of cells to reactive oxygen species leads to cytochrome c release from the mitochondria and apoptosis, whereas antioxidants and overexpression of antioxidant enzymes, such as Mn-SOD, Cu,Zn-SOD, phospholipid hydroperoxide glutathione peroxidase, and thioredoxin reductase, inhibit caspase activation and apoptosis (thoroughly reviewed by Iverson and Orrenius [153]). In particular, challenge of cells with peroxynitrite results in apoptosis [120, 154–156], and different apoptotic stimuli increase peroxynitrite formation in neurons [130, 157, 158], macrophages [159, 160], and endothelial cells [161]. Studies with isolated mitochondria have demonstrated that peroxynitrite can induce PTP opening through the oxidation of thiols in ANT, promoting cytochrome c release [162–164], part of which might be nitrated [130], and that calcium strongly influences peroxynitrite-mediated permeability transition [165]. Peroxynitrite oxidation of critical thiols also promotes increased hydrolysis of pyridine nucleotides that result in an increase in calcium release through the PTP [166]. Proteomic studies reveal that VDAC is nitrated in mitochondria of diabetic and septic rats [113, 150] and in cells where iNOS is activated [125]; nevertheless, the functional consequences of this posttranslational modification are yet to be established.

5.3.5 PEROXYNITRITE REACTIONS ANTIOXIDANTS

WITH

MITOCHONDRIAL

Mn-SOD is a key antioxidant enzyme that catalyzes the dismutation of superoxide to oxygen and hydrogen peroxide. Peroxynitrite reacts with the metal center (Mn3+) of the enzyme (1 × 105 M−1s−1) [67] and inactivates the enzyme through the nitration of the critical residue Tyr34 [65, 66], in a metal dependent process [67]. Mn-SOD nitration has been reported in numerous pathological states both in humans and animal models (Table 5.1) [113, 114, 128, 129, 167–172a]. Peroxynitrite inactivation of Mn-SOD will probably lead to an increase in superoxide steady-state concentration, favoring peroxynitrite formation and resulting in a positive feedback process that promotes mitochondrial damage. Indeed, in studies of protein nitration, Mn-SOD appears as an early target [125, 128], even preceding organ dysfunction [128].

Peroxynitrite

111

TABLE 5.1 Mitochondrial Proteins Nitrated in Pathologic Conditions Disease Condition Diabetic mouse hearta

Nitrated Protein SCOT Creatine kinase Peroxiredoxin 3 Trifunctional protein VDAC-1

Observation SCOT decrease in activity

Ref. [114]

Diabetic rat heartb

SCOT

SCOT decrease in activity

[150]

Cerebral cortex of rats subjected to brain ischemia/reperfusionc

Cytochrome c

Cytochrome c release to the cytosol Swollen mitochondria

[130]

Cerebrospinal fluids of patients with ALS, Alzheimer’s disease, and Parkinson’s diseaseb

Mn-SOD

[172]

Spinal cord from FALS mouse

Aconitase ATP synthase Creatine kinase Mn-SOD

[172a]

Aged rat aortad, e

Mitochondrial proteins Mn-SOD

Lung from a LPStreated rat (Inflammatory disease model)a

Aconitase Creatine kinase VDAC Malate dehydrogenase

Reduced .NO biodisponibility Enhanced O2.− formation

[168]

[113]

112


TABLE 5.1 Mitochondrial Proteins Nitrated in Pathologic Conditions (continued) Disease Condition Liver from a LPStreated rata

Nitrated Protein Aconitase ATP synthase VDAC Glutamate-oxaloacetate transaminase 2 Mn-SOD Hydroxymethylglutaryl-CoA synthase 4-Trimethylamino butyrataldehyde dehydrogenase Glutamate dehydrogenase Short chain 3-OH acyl CoA dehydrogenase D-β-OH butyrate dehydrogenase 3-Ketoacyl CoA thiolase Rhodanese

Observation

Ref. [113]

Kidney from a LPStreated ratf

SCOT

SCOT decrease in activity

[149]

Heart from a LPStrated suckling ratb

Carnitine palmitoyltransferase I (CPT I)

CPT I decrease in activity

[142]

Diaphragm from a LPS-treated ratg

Mitochondrial proteins

Uncoupling of oxidative phosphorylation Decrease in diaphragmatic force

[27, 244]

Eye from a LPStreated rat (endotoxin-induced uveitis model)b

Mn-SOD

Mn-SOD increase in activity and in expression

[169]

Chronic allograft rejection of human kidneyb, h

Mn-SOD

Mn-SOD decrease in activity, increase in expression and aggregate formation Kidney rejection

[171]

Chronic allograft nephropathy of rat kidneyb

Mn-SOD Cytochrome c

Mn-SOD decrease in activity Renal dysfunction

[128]

Peroxynitrite

113

TABLE 5.1 Mitochondrial Proteins Nitrated in Pathologic Conditions (continued) Disease Condition Ischemia/reperfusion of rat kidneyb

Nitrated Protein Mn-SOD Cytochrome c

Observation Mn-SOD decrease in activity Cytochrome c and Mn-SOD release to the cytoplasm Decrease in ATP levels and apoptosis Renal dysfunction

Ref. [129]

Kidney from angiotensin IIinfused rat (Hypertension model)e, i

Mn-SOD

Mn-SOD decrease in activity

[167]

Heart from apoE-/mouse (Hypercholesterolemia model)e

Mn-SOD

MtDNA damage

[170]

Heart from mouse exposed to cigarette smokee

Mn-SOD

Mn-SOD and ANT decrease in activity MtDNA damage

[170]

Techniques used for the identification of nitrated proteins: a

Two-dimensional electrophoresis, Western blot with anti-TyrNO2, and matrix-assisted laser desorption ionization/time-of-flight mass spectrometry (MALDI-TOF/MS) of tryptic fragments. bImmunoprecipitation with anti -TyrNO and Western blot with anti-SCOT, anti-MnSOD, anti-Cyt 2 c, or anti-CPT I. cConfocal inmmunohistochemistry with anti-TyrNO and anti-Cyt c (not conclusive). 2 dImmunohistochemistry with a transmission electron microscope and anti-TyrNO 2. eImmunoprecipitation with anti-MnSOD and Western blot with anti-TyrNO 2. fWestern blot with anti-TyrNO , purification of nitrated SCOT, and microsequence analysis. 2 gMitochondrial purification and Western blot with anti-TyrNO . 2 hImmunoprecipitation with anti-TyrNO , Western blot with anti-TyrNO , and microsequence anal2 2 ysis. iImmunoprecipitation with anti-MnSOD, protein hydrolysis, and HPLC detection of TyrNO . 2

Glutathione (GSH) is a relevant mitochondrial antioxidant that directly scavenges reactive species, and participates with GSH peroxidase in the detoxification of peroxides. The efficiency of GSH as an antioxidant resides in the existence of GSH reductase that catalytically reduces glutathione disulfide to GSH at the expense of NADPH [173]. Glutathione is found in high concentrations in the mitochondria (5–10 mM), and reacts with peroxynitrite

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in a two-electron oxidation process leading to the formation of glutathione disulfide (GSSG) [174] (k = 1350 M−1s−1 [175]). Glutathione also reacts with peroxynitrite-derived radicals, such as carbonate radical, hydroxyl radical, and nitrogen dioxide, forming the one electron oxidation product, glutathionyl radical (GS.) that either recombines to glutathione disulfide [176] or reacts with glutathione forming the disulfide radical anion (GSSG.−)[177]. GSH peroxidase (GPx) has a selenocysteine residue that undergoes a fast two-electron oxidation (k = 8 × 106 M−1s−1) reducing peroxynitrite to nitrite. Because the oxidized selenium in the enzyme can be reduced back to the selenol by glutathione, the enzyme was initially considered a peroxynitrite reductase [178]. Although GSH peroxidase protects different molecules from peroxynitritemediated oxidation and nitration in vitro and in cell lysates [178], it does not appear to do it in intact cells. Hepatocytes from GPx knockout mice were more resistant to peroxynitrite-mediated cytotoxicity and nitration than their wild-type counterpart [179]. Therefore, the role of GPx as a peroxynitrite reductase in vivo requires further study. Tocopherols and ascorbate constitute a relevant antioxidant system that is mainly in charge of protecting membrane lipids from oxidative stress. α-Tocopherol and γ- tocopherol inhibit peroxynitrite-induced lipid peroxidation in liposomes of unsaturated fatty acids [180], and their protective actions are due in part to their reactions with peroxynitrite-derived radicals [181–183]. Tocopherols can also reduce the lipid peroxyl radicals (LOO.) (k = 106 M−1s−1) [184], generated during peroxynitrite reaction with lipids [185], thus terminating free radical chain reactions. The tocopheroxyl radical generated in this reaction can then be reduced back to tocopherol by ascorbate, accounting for ascorbate antioxidant actions in membrane [186]. Aside from its synergistic actions with tocopherol, ascorbate can directly react both with peroxynitrite (k = 236 M−1s−1) [187] and with peroxynitrite-derived radicals [188]. Ascorbate can also repair peroxynitrite oxidation targets, such as tyrosyl radicals, thus regenerating tyrosine [189]. Exposure of isolated mitochondria to peroxynitrite results in a decrease in both ascorbate and α-tocopherol levels, with a concomitant increase in α-tocopherolquinone [190, 191], and the main oxidation product of α-tocopherol reaction with peroxynitrite [183, 192]. Moreover, ascorbate depletion precedes and α-tocopherol decreases parallels, the impairment of oxidative phosphorylation, and reduction in respiratory control observed in mitochondrias incubated with peroxynitrite [190]. Ubiquinone could also play a role as a mitochondrial antioxidant because peroxynitrite-derived radicals promote the one electron oxidation of ubiquinone to ubisemiquinone, which could, in turn, be reduced back to ubiquinone by complex I and II of the respiratory chain. Moreover, ubiquinone supplementation protects mitochondria exposed to peroxynitrite from nitration and loss of respiratory control [193].

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5.4 PEROXYNITRITE MEDIATES MITOCHONDRIAL DYSFUNCTION IN PATHOLOGY Protein tyrosine nitration, due to its relative stability, has become a marker of NO-derived species involvement in pathogenic processes. As presented in Table 5.1, nitration of mitochondrial proteins has been observed in a wide variety of pathological conditions, and in many cases, it accompanies mitochondrial or organ dysfunction. Tyrosine nitration has been assessed by a number of techniques, including analytical and immunochemical detection of 3-nitrotyrosine (see Table 5.1); however, the detection of nitrated proteins does not always imply that a relevant percentage of a given target has been modified, or that the activity of a protein has been affected. Very few reports have dealt with this issue [167], and in fact, biological nitration yields are low. Under inflammatory conditions, 1 to 5 3-nitrotyrosine residues per 10,000 tyrosine residues are detected [90]. Mn-SOD, SCOT, and carnitine palmitoyltransferase I are the only mitochondrial proteins reported to be both nitrated and inactivated in vivo (Table 5.1) and of these only Mn-SOD inactivation is known to be solely due to tyrosine nitration [65]. Alternatively, nitrated proteins reportedly undergo an enhanced turnover [194], and recent data indicate that denitration processes can take place in mitochondria [125, 195]. Therefore, nitration events might be somehow underestimated when protein 3-nitrotyrosine is measured. We will now describe the following in further detail: peroxynitrite formation, mitochondrial protein nitration, and protein/mitochondrial/cell dysfunction in diabetes, sepsis, and neurodegenerative diseases.

5.4.1 DIABETES 5.4.1.1 Peroxynitrite Formation in Diabetes Diabetes causes development of vascular complications and epidemiological evidence strongly suggests a correlation between hyperglycemia and the appearance of these complications [196, 197]. Strikingly, the four main pathogenic molecular mechanisms of hyperglycemia (increased polyol pathway flux, increased advanced glycation end product [AGE] formation, activation of protein kinase C, and increased hexosamine pathway flux [198]) are linked to an increase in mitochondrial superoxide formation [199, 200]. Increased mitochondrial superoxide production has been described in hyperglycemic bovine aortic endothelial cells [199], islets of Langherhans [19], retinal cells, and retinal endothelial cells [18]. Moreover, islets of Langherhans and retinas of diabetic mice produce more superoxide than their non-diabetic counterparts [18, 19]. Other reported sources of superoxide in diabetic animals or cultured cells exposed to hyperglycemic conditions are uncoupling of ·NO synthase [201, 202], aldose reductase [202], NAD(P)H oxidase [203], and increased xanthine oxidase levels in liver, aorta, and plasma of diabetic rats [204]. Nevertheless, mitochondrial superoxide production appears to precede these events and could even affect eNOS activity as

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well as promote NAD(P)H oxidase activation through PKC activation and the hexosamine pathway [198, 205]. ·NO metabolism is also altered in diabetes. ·NO-dependent vasodilation is impaired in animal models [206] and humans with type 1 and type 2 diabetes [207–211], and hyperglycemia produces endothelial dysfunction in healthy subjects [212]. The fact that antioxidants revert this impairment [208, 209, 212, 213] and plasma 3-nitrotyrosine levels are increased in diabetic patients [214], during hyperglycemic clamp to healthy subjects [215] and postprandial hyperglycemia [216], has led to the proposition that increased superoxide formation is responsible for the decrease in ·NO availability in these situations. Alternative mechanisms, such as an increase in asymmetric dimethyl arginine (an endogenous NOS inhibitor [217]), inhibition of eNOS by posttranslational modification [205], and eNOS uncoupling [201, 218], probably contribute as well. Experiments with aortic endothelial cells exposed to hyperglycemic conditions further support peroxynitrite formation, because increased tyrosine nitration [161, 201, 219], along with reduced ·NO levels and cyclic GMP production (·NO availability) [161], despite an increase in ·NO synthase expression [219], are observed. In these reports, enhanced superoxide is reported to arise from NADH oxidase activation [219] or from eNOS uncoupling [201]. Nevertheless, mitochondrial superoxide formation also increases in aortic endothelial cells exposed to hyperglycemia [199]. Therefore, though not reported yet, peroxynitrite could also be formed inside the mitochondria. In this sense, a recent report indicates that eNOS is associated to mitochondria in endothelial cells, that ·NO accumulates in mitochondria and that hyperglycemia decreases mitochondrial ·NO levels with the concomitant formation of higher oxidant species [220]. Further studies are required to define if peroxynitrite formation occurs in mitochondria of diabetic endothelial cells and if it has a role in endothelial dysfunction and cell apoptosis that lead to diabetic vascular disease. 5.4.1.2 Evidence of Peroxynitrite-Mediated Mitochondrial Dysfunction in Diabetes Development of cardiac myopathy is a complication usually observed in diabetic patients. Myocardial biopsies obtained from diabetic patients, a mouse model of diabetes, and rat hearts perfused with a hyperglycemic solution clearly indicate an association between apoptotic and necrotic death of myocytes, endothelial cells and fibroblasts, as well as protein tyrosine nitration [221–223]. Moreover, the voltage-dependent anion channel (VDAC) is also found nitrated in diabetic heart mitochondria [114] where the permeability transition is increased [224]. Furthermore, mitochondrial energy metabolism is impaired in the hearts of diabetic rats [122, 134, 225, 226] where many mitochondrial proteins are nitrated (see Table 5.1) [114, 150]. In particular, certain mitochondrial enzymes involved in energy homeostasis, such as creatine kinase and complex I of the respiratory chain, are nitrated [114], and their activities decreased in diabetic heart [122, 134]; they are also nitrated and inactivated by peroxynitrite in vitro [53, 115, 137,

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138, 227]. Moreover, SCOT is also found both nitrated and inactivated in the diabetic heart [150]. Retinopathy is a microvascular complication usually developed by diabetic patients. In animal models of diabetes, an increase in tyrosine nitration in the retina was observed associated with retinopathy [228] and accompanying the breakdown of the blood retina barrier [229]. Apoptosis is also increased in both patients with diabetic retinopathy [230], and in the retinas of diabetic rats, where cytochrome c migrates to the cytosol, Bax migrates to the mitochondria, and caspase activity augments [231]. Manganese (III) tetrakis (4-benzoic acid) porphyrin (MnTBAP) and uric acid, scavengers of peroxynitrite and peroxynitritederived radicals, respectively, decreased apoptosis and nitration of hyperglycemic retinal endothelial cells [229, 231]. According to this data, peroxynitrite clearly appears to be involved in mitochondrial dysfunction in the diabetic heart and to participate in the development of diabetic retinopathy and endothelial dysfunction, although the role of mitochondrial impairment in the latter processes requires further study.

5.4.2 SEPSIS 5.4.2.1 Peroxynitrite Formation in Sepsis An important increase in ·NO production is observed during the septic inflammatory response [24]. Nitrite and nitrate concentrations in plasma are augmented ~20-fold in septic rats [25], and in rat diaphragm and heart, a twofold increase in ·NO concentration was measured [25–27]. In humans, plasma and tissue nitrite/nitrate are also elevated (approximately twofold in muscle [121]), though not so much as in rodents [232]. The source of most of the ·NO is iNOS, whose expression increases [233, 234], although eNOS and nNOS expression are also enhanced, but in a minor degree, in skeletal muscle from septic rats [235]. Mitochondrial ·NO production by mtNOS is also augmented during lipopolysaccharide (LPS)-induced septic shock in rat diaphragm, liver [24], and lung [236], and appears to inhibit complex I and IV of the mitochondrial respiratory chain. The use of NOS inhibitors, such as NG-methyl-L-arginine (L-NMMA) and NG-nitroL-arginine methyl ester (L-NAME), has unambiguously indicated that augmented ·NO is involved in the pathophysiology of endothelial and contractile dysfunction in sepsis [26, 27, 232, 237, 238]. Increased superoxide formation is also observed in the diaphragms of septic rats [239], and studies with polyethylene glycol-SOD (PEG-SOD) demonstrated its participation in the development of diaphragm muscle dysfunction [240, 241]. Interestingly, mitochondria extracted from endotoxemic rats indicated an increase in hydrogen peroxide formation that probably reflects increased mitochondrial superoxide production [27, 242]. Xanthine oxidase and NADH oxidase have been also reported to increase their superoxide production during sepsis [25, 243]. Mitochondrial protein nitration is reported to occur in multiple tissues during endotoxemia, including rat diaphragm [27, 244], eye [169], kidney, heart [149],

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lung, and liver [113], and specific mitochondrial proteins prone to nitration have been identified (see Table 5.1). Although, in sepsis, alternative nitration routes such as myeloperoxidase-catalized oxidation of nitrite may become relevant, reports of SOD-mediated inhibition of mitochondrial nitration point toward peroxynitrite as the main nitrating agent in mitochondria [241]. 5.4.2.2 Evidence of Peroxynitrite-Mediated Mitochondrial Dysfunction in Sepsis An association between ·NO overproduction, antioxidant depletion, mitochondrial dysfunction, and patient outcome has been established in septic shock patients, implicating bioenergetic failure in the pathophysiological mechanisms of multiorgan failure [121]. Evidence for oxidant-mediated mitochondrial dysfunction involvement in the pathogenic mechanism of skeletal muscle, heart, and liver failure in septic shock will be subsequently exposed. Sepsis induces a severe and persistent alteration of skeletal muscle characterized by an increase in muscle catabolism that produces tissue wasting [245], decreased muscular force [27, 240] and impaired tissue oxygen extraction [246], all due to an impairment in mitochondrial energy metabolism. Loss of skeletal muscle contractile function during endotoxemia occurs concomitantly with an increase in ·NO, superoxide formation, and mitochondrial protein nitration [27, 240, 247]. Furthermore, isolated mitochondria from septic diaphragm tissues demonstrate decreased oxygen consumption during state-3 respiration, along with uncoupling, and these alterations are prevented by the administration of PEGSOD, SOD-mimetics, or NOS inhibitors to the animals during endotoxin exposure [27, 241, 242, 244]. In cardiac tissue, intense 3-nitrotyrosine immunoreactivity is found in biopsies from septic patients with myocarditis [248]. Experiments with endotoxemic rats have demonstrated that enhanced generation of superoxide and ·NO accompany the impairment of cardiac work and efficiency, as well as the decrease in oxygen consumption and ATP levels [25, 249]. Although the reduction in the number of mitochondria is probably involved in the bioenergetic failure of the tissue [249], the nitration and inactivation of a relevant mitochondrial protein, carnitine palmitoyltransferase I, evidences peroxynitrite impact in cardiac energy homeostasis [141, 142]. Liver failure and eventual poor outcome of septic rats is associated with an increase in ·NO production, complex I inactivation and ATP depletion [250]. Isolated mitochondrias from this tissue exhibit swelling, increased iNOS expression [251], and nitrated mitochondrial proteins, including aconitase, ATP synthase, and enzymes of beta-oxidation [113]. All these observations strongly suggest that peroxynitrite mediates mitochondrial dysfunction, which gives rise to contractile impairment of skeletal muscle and heart, as well as liver failure, and open a wide area of research.

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5.4.3 NEURODEGENERATIVE DISEASES 5.4.3.1 Peroxynitrite Formation and Mitochondrial Dysfunction in Neurodegenerative Diseases NO production in enhanced neurodegenerative diseases [28–30] and experiments with NOS inhibitors and NOS knockout mice confirm its neurotoxicity [28, 252–255]. Different NO sources have been described; on the one hand, excess release of the excitatory neurotransmitter glutamate acting via the NMDA receptors is considered responsible for increased nNOS activation and neuronal cytotoxicity, known as excitotoxicity, in stroke [28], ALS [256], Parkinson’s disease [257], and Alzheimer’s disease [258]. Alternatively, cytokine-stimulated activation of iNOS in astrocytes or microglia is reported to mediate neurotoxicity in Parkinson’s disease [254], Alzheimer’s disease [259, 260], stroke [261], and multiple sclerosis [262]. Mitochondrial superoxide appears to play a relevant role in the pathogenesis of neurodegenerative diseases as well. The sensitivity of the nervous system to increases in mitochondrial superoxide is evidenced by the high degree of neuronal degeneration and impaired neurological phenotype of SOD2 knockout mice [263, 264], as well as by the fact that Mn-SOD expression correlates with the degree of neurodegeneration in ischemia/reperfusion [265], ALS [266], and Parkinson’s disease [267]. Furthermore, superoxide levels in mitochondria are increased during excitotoxic and ischemic challenge to cortical neurons [20–22] and correlate with cell death [22, 268]. An increase in protein tyrosine nitration has been described in all the mentioned neurodegenerative diseases, both in human patients [172, 262, 269–272] and animal models [28, 130, 172a, 267, 273, 274], but very few reports are available about the specific nitration of mitochondrial proteins [130, 172]. The expression levels of MnSOD and a mitochondrial peroxiredoxin are related to the resistance of neurons to ·NO-mediated toxicity and nitration [273, 275, 276]; however, supporting peroxynitrite involvement in mitochondrial damage. Neurodegenerative diseases, such as Parkinson’s disease, Alzheimer’s disease, multiple sclerosis, amyotrophic lateral sclerosis (ALS), and stroke, are characterized by alteration of mitochondrial function, which is suggested to be involved in their pathogenesis [29, 257]. Current evidence of mitochondrial peroxynitrite formation and its impact in mitochondrial energy metabolism and apoptosis, in an acute (stroke) and a chronic (Parkinson’s disease) neurodegenerative process, will now be discussed. 5.4.3.1.1 Stroke Stroke is characterized by a period of ischemia followed by reperfusion of the tissue. In this period neurons undergo apoptosis, calcium homeostasis is altered, and mitochondrial energy metabolism impaired [277], implicating mitochondrial damage in the pathogenesis of the disease. ·NO, mitochondrial superoxide, and peroxynitrite formation are increased in the brains of animals undergoing cerebral ischemia and reperfusion, and are clearly involved in neurotoxicity [28, 261, 265,

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278, 279]. Moreover, highly oxidant species have been detected inside the mitochondria from rats undergoing brain ischemia [280] and Mn-SOD overexpression decreases protein nitration along with infarct volume [157], suggesting that specific intramitochondrial formation of peroxynitrite could be involved in ischemic cell death. Moreover, mitochondrial nitration [130] and Mn-SOD expression levels [281] correlate with cytochrome c release and DNA fragmentation in mice with cerebral ischemia, implying peroxynitrite in mitochondrial-mediated apoptosis. In what refers to the mitochondrial energy metabolism, in the brains of animals undergoing brain stroke electron transport, respiratory control ratio, ATP synthesis, and mitochondrial polarization are impaired [277]; but despite being sensitive peroxynitrite targets, no direct evidence linking peroxynitrite with their dysfunction in vivo has been provided yet. 5.4.3.1.2 Parkinson’s Disease Parkinson’s disease is characterized by the loss of dopaminergic neurons in the substantia nigra and by the presence of Lewy bodies in remaining nerve cells. 1-Methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) causes a Parkinson’s-like syndrome in humans and animals and has been extensively used in animal models of the disease [282]. Increased ·NO production [252–255, 257] and mitochondrial superoxide [267, 283] are involved in the development of Parkinson’s disease, although alternative superoxide sources, such as dopamine metabolism, microglial NADPH oxidase, and cyclooxygenase-2 (COX-2), are also considered important [282, 284]. 3-Nitrotyrosine has been observed in the Lewy bodies of Parkinson’s patients [272], and Mn-SOD overexpression decreases tyrosine nitration and neurotoxicity in animals treated with MPTP [267]. Furthermore, nitrated Mn-SOD was found in the cerebrospinal fluid of Parkinson’s disease patients [172]; thus, peroxynitrite could be formed inside the mitochondria and eventually diffuse to the cytoplasm and promote the already reported nitration of α-synuclein in Lewy bodies and tyrosine hydroxylase [285–287]. The molecular mechanisms responsible for neuronal degeneration and death in Parkinson’s disease are not fully understood, but one theory maintains that mitochondrial dysfunction and excess oxidant formation can trigger cell death. Electron transport chain complex I activity is reduced in the substantia nigra and platelets of Parkinson’s disease patients [9, 282, 288], and in fact, in vitro experiments demonstrate that neurotoxins that induce Parkinson’s disease, such as MPTP, rotenone, and zinc, inhibit complex I of the electron transport chain and α-ketoglutarate dehydrogenase, promoting an increase ROS formation [9]. Apoptosis is enhanced in the substantia nigra of Parkinson’s disease patients and MPTP-treated mice. [289–291]. In isolated mitochondria, MPTP promotes the opening of the of the mitochondrial transition pore inducing cytochrome c release in a Bax and oxidant-dependent process [292, 293]; therefore, the mitochondrial pathway and oxidant formation are indeed involved in apoptosis.

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5.5 MITOCHONDRIAL-TARGETED PHARMACOLOGY Selective increase in the antioxidant capacity of mitochondria could be a useful therapy in the diseases where mitochondrial dysfunction due to oxidative damage is implied, and in fact, overexpression of antioxidant enzymes, such as Mn-SOD, has, as a result, been protective in a wide range of processes [267, 275, 294–296]. Nevertheless, gene therapy is still far from clinical application, and SOD protein delivery to tissue is difficult due to protein inability to cross membranes, low half-life in blood, and immunogenicity [297]. Therefore, much hope is posed on low molecular weight antioxidants. In this sense, pioneer work by Murphy and colleagues has recently reported that attaching a lipophilic cation to an antioxidant molecule highly increases its accumulation in the mitochondrial matrix, due to the large potential across the organelle inner membrane [298, 299]. Following this strategy, different antioxidant molecules have been successfully delivered to the mitochondria and have protected mitochondria from oxidative challenge and cells from oxidant-induced apoptotic death [298–301]. In this scenario, manganese porphyrins appear as promising antioxidants to target to mitochondria. Manganese porphyrins were first described as superoxide dismutase mimics [302], but were later acknowledged to react with peroxynitrite with rate constants ranging from 105 to 107 M−1s−1 and to serve as peroxynitrite decomposition catalysts in the presence of reductants [303]. Incubation of neurons with neuroprotective concentrations (200 µM) of MnTBAP or manganese (III) meso-tetrakis ([N-ethyl] pyridinium-2-yl) porphyrin (MnTEPyP) resulted in the accumulation of 1.7 and 2.5 ng per ml of mitochondrial protein, respectively [21], yielding intramitochondrial concentrations of 10 and 16 µM, respectively. Kinetic considerations clearly indicate that MnTEPyP (k = 3.4 × 107 M−1s−1 [304]) would outcompete both carbon dioxide and cytochrome c in the reaction for peroxynitrite. MnTBAP and MnTEPyP have been successfully used in different pathologic situations involving peroxynitrite formation and mitochondrial dysfunction [231, 244, 268, 305–307]. Very few reports, however, have dealt with the improvement of their cellular targeting [308], and improvement of their targeting to mitochondria (only 10% and 20% MnTBAP and MnTEPyP are distributed into the mitochondria, respectively [21]), along with its pharmacological implications, is still to be done. Thiols with low pKas could also be good peroxynitrite scavengers due to their high reaction constants with peroxynitrite [309]. In fact, a recent report indicates that in vivo gene transfer of peroxiredoxin-3 to rat brain decreases nitration and cell death associated with excitotoxicity [273]. Peroxiredoxin-3 is a mitochondrial antioxidant protein that belongs to the peroxiredoxin family. These enzymes detoxify peroxynitrite, as well as hydrogen peroxide, through the fast reaction with a thiol present in the active site (k = 106 to 107 M−1s−1) [310, 311]. This fast reaction is probably due, at least in part, to the fact that the reactive thiol has a pKa < 5 [310]. Though the pKas of low molecular thiols are not as low as those

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found in the active sites of enzymes, catalytic rate constants as high as 7 × 103 M–1s–1 have been reported [309], and the relation between thiol structure and reactivity remains a promising area of research, including the generation of thiol containing peptides. Another area of pharmacological development could be related to the generation of mitochondrial-targeted compounds that are able to react with peroxynitritederived radicals, namely carbonate radical, oxo-metals, and nitrogen dioxide. In this sense, tyrosine-containing peptides appear to be an interesting alternative.

5.6 CONCLUSIONS From the data presented herein, mitochondria appear as a cellular locus for peroxynitrite formation in pathology. We have addressed the intramitochondrial formation of peroxynitrite, the biochemistry of mitochondrial protein nitration, and the role of peroxynitrite in promoting mitochondrial energetic dysfunction and apoptosis. At a pathophysiological level, peroxynitrite-mediated mitochondrial damage is related to organ dysfunction in the diabetic heart and septic skeletal muscle. Current evidence also suggests peroxynitrite involvement in the impairment of other septic and diabetic tissues as well as in brain function in neurodegenerative diseases. Much of the information of peroxynitrite reactions in vivo comes from the identification of 3-nitrotyrosine residues in proteins of human biopsies or tissues from animal models of these diseases; however, it is important to appreciate that peroxynitrite-dependent oxidation reactions are probably more abundant than nitration and also relevant in the development of cellular and tissue dysfunction. Furthermore, tyrosine nitration is not always related to the functional alteration of the target proteins. The comparison of in vitro and in vivo data reveals that of all the nitrated mitochondrial proteins identified in vivo (see Table 5.1) only MnSOD, creatine kinase, ATP synthase, cytochrome c, and aconitase have been demonstrated to be, in fact, inactivated by peroxynitrite in vitro. In addition, cytochrome c nitration results in a potentially toxic “gain-of-function” through the acquirement of peroxidatic activity [86, 100–101]. Of these, only Mn-SOD has been found both nitrated and inactivated in vivo along with SCOT and carnitine palmitoyl transferase I. Finally, we have assessed current knowledge on mitochondrial targeting of antioxidants that could eventually give rise to efficient pharmacological treatments in pathologies where peroxynitrite mediates mitochondrial dysfunction.

5.7 ACKNOWLEDGMENTS This research was supported by grants from Fondo Clemente Estable (Uruguay) to Larua Castro, Fogarty-National Institutes of Health, the Howard Hughes Medical Institute, and the Guggenheim Foundation to Rafael Radi.

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of Glucose 6 Modulation Metabolism by Nitric Oxide in Astrocytes and Neurons Juan P. Bolaños, María Delgado-Esteban, Pilar Cidad, Paula García-Nogales, and Angeles Almeida Universidad de Salamanca, Salamanca, Spain

CONTENTS 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8 6.9 6.10 6.11 6.12 6.13 6.14

Introduction ............................................................................................146 NO formation in Neural Cells ...............................................................146 Neurotoxic vs. Neuroprotective Roles for NO .....................................147 Role of Astrocytes in Glucose Homeostasis in the Brain ....................148 Main Glucose Carriers in Neural Cells .................................................148 NO Up-Regulates Glucose Uptake by Astrocytes ................................149 Glucose Utilization through the Pentose–Phosphate Pathway: The Role of NO and Peroxynitrite ........................................................152 Low Doses of Peroxynitrite May Cause Neuroprotection: The Role of Glutathione ........................................................................153 Glucose Metabolism and Glutamate-Induced Neurotoxicity ...............154 Nitric Oxide and Glycolysis: What Is the Role of Glyceraldehyde-3-Phosphate Dehydrogenase? .....................................155 NO Triggers Glycolytic Activation and Protects Astrocytes from Cell Death .....................................................................................156 On the Mechanism Whereby NO Stimulates Glycolysis in Astrocytes ...................................................................................... 157 Concluding Remarks .............................................................................159 Acknowledgments ..................................................................................159 References ...............................................................................................159

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6.1 INTRODUCTION It is well known that the brain is a vulnerable organ that depends on an efficient supply of oxygen and energetic substrates from the blood. Thus, a transient lapse in such supply may lead to unconsciousness and, after only a few minutes, to irreversible alterations possibly causing neuronal death [1]. In fact, neurotransmission is one of highest energy-demanding processes in mammals, and a continuous supply of metabolic substrates is essential for keeping the axonal membrane potential active [2]. To aid neurons in this process, astrocytes surround both capillary blood vessels [3–5] and synaptic spaces [6, 7]. This allows glial cells to remove synaptic neurotransmitters [8] and to shuttle metabolic substrates between intracerebral vessels and neurons [9, 10]. The blood–brain barrier represents a restriction of energy availability for the central nervous system by reducing the range of metabolic substrates to those having an efficient carrier system. In the mammalian brain, glucose represents the main source of energy because after mitochondrial oxidative catabolism it generates ATP [1]. The endothelial cells of the blood–brain barrier express a suitable glucose transport system, the energy-independent glucose transporter GLUT1, which has a sufficiently high kM for glucose that prevents any failure in substrate supply under normoglycemic conditions [11, 12]. Accordingly, astrocytes are the first neural cell type to take up glucose in the central nervous system and thus knowledge of the factors that regulate glucose uptake and metabolism may be relevant for the understanding of neuronal energy metabolism. This chapter focuses on recent findings suggesting that nitric oxide (NO) would be a signalling molecule that physiologically modulates glucose metabolism in astrocytes and neurons. Given the importance that has been given to NO as a both a neurotransmitter and a neurotoxin, the metabolic signalling pathways modulated by NO may be important for understanding the apparent controversy as regards the roles of NO in neuronal cell death/survival decisions.

6.2 NO FORMATION IN NEURAL CELLS NO is a neural messenger [13] that is synthesized by the NO synthase (NOS)catalyzed reaction from L-arginine [14–16]. NO activates soluble guanylate cyclase (sGC) and thus participates in the transduction pathway involving cyclic GMP-dependent kinases [17, 18]. In addition, NO inhibits mitochondrial cytochrome c oxidase (complex IV) and thus leads to the as yet not fully elucidated consequences of the modulation of cellular respiration [19–25]. The formation of NO in neurons occurs following the interaction of synaptic glutamate with the NMDA (N-methyl-D-aspartate) receptor, an ion channel that allows the entry of extracellular Ca2+ to form a complex with calmodulin, which activates constitutive neuronal NOS (nNOS or NOS1) [16]. In glial cells (astrocytes, microglia, and oligodendrocytes), NO is mainly formed after the transcriptional induction of a calcium-independent inducible NOS isoform (iNOS or NOS2). Such induction takes place along inflammatory-like situations, such

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 147

as the interaction of the endotoxin lipopolysaccharide (LPS) and certain cytokines, such as interferon-γ, tumor necrosis factor-α or interleukin-1β, with their respective plasma membrane receptors (for reviews, see [26–29]). iNOSdependent NO might serve as a signalling defense against external insults, but it also diffuses to neighboring neurons, where it may cause either cellular damage or the activation of survival pathways. NO is also formed in the endothelial cells by the constitutive Ca2+-dependent activity of endothelial NOS (eNOS or NOS3) [16]. Regardless of these major sites of NO production, most brain cell types are known to be able to express several NOS isoforms. Thus, astrocytes also produce NO via nNOS activity [30–33], and both endothelial cells [34] and neurons [35] can express iNOS after LPS plus cytokine treatment. NO reacts with superoxide (O2²-) to form peroxynitrite anion (ONOO–) [36]. Peroxynitrite is a compound that is only stable in alkaline solutions, having a pKa of ~6.8, and is thus rapidly protonated at physiological pH values to form peroxynitrous acid (ONOOH) [37]. The half-life of peroxynitrous acid is ~1 sec, giving rise to chemical species with hydroxyl radical (²OH)-like reactivity and nitrogen dioxide (²NO2). The latter two free radicals would then spontaneously form the more stable compound nitric acid (NO3-) at physiological pH [37]. The occurrence of such radical-mediated reactions confers peroxynitrite pro-oxidant properties that are thought to be responsible for the execution of the neurotoxic NO-mediated responses [37–40]; however, this controversial issue has been revisited recently [41].

6.3 NEUROTOXIC VS. NEUROPROTECTIVE ROLES FOR NO Overproduction of NO by nNOS activity may be neurotoxic [42–44] and may play a role during neurodegeneration and ischaemia (e.g., see reviews [40, 45–48]). Certain pro-inflammatory conditions, such as sepsis or brain ischaemia, occur along cytokine-mediated iNOS gene overexpression in glial cells, which strongly release NO, causing damage to the neighboring neurons [28, 49]. When formed in excess, NO-derived ONOO– causes damage to DNA and triggers the activation of the DNA repairing system poly(ADP-ribose) synthetase (PARS), followed by depletion of ATP stores [50–54]. In addition, ONOO- inhibits aconitase [55, 56] and interferes with the mitochondrial respiratory chain [57–59]. In contrast with neuronal-derived NO, the role of glial-derived NO during cerebral damage following ischaemia is controversial. Thus, whereas cytokines and growth factors mediate ischaemia-mediated iNOS induction in glial cells following neurotoxicity [60, 61], other authors have suggested that these factors would be neuroprotective [62, 63]. For instance, hypoxia followed by re-oxygenation episodes releases interleukin-1 in cultured rat astrocytes [64], and NOmediated neuronal ischaemic insult can be prevented by pretreatment with certain growth factors (fibroblast growth factor, epidermal growth factor, insulin-like growth factor, glial-derived neurotrophic factor or transforming growth factor-β1) both in vivo and in vitro [65-68]. The role of ischaemia-mediated iNOS induction in glial cells has been reviewed elsewhere [69–73].

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6.4 ROLE OF ASTROCYTES IN GLUCOSE HOMEOSTASIS IN THE BRAIN Regardless of the critical importance of glucose as the main source of precursors for mitochondrial ATP biosynthesis [1], this substrate is also the main precursor for the biosynthesis of fatty acids, cholesterol, neurotransmitters, amino acids, glycerol-3-phosphate, and, within the brain, in astrocytes [74], for the formation of glycogen. Furthermore, an important fraction of glucose entering the brain is metabolized through the pentose–phosphate pathway, which is the main cellular source of NADPH and ribose-5-phosphate. Finally, under long-term starvation or during the suckling period, the brain can be adapted to use alternative substrates such as lactate or ketone bodies [75–80]. Astrocytes play a critical role in brain energy homeostasis, in part because they are strategically localized within the brain cellular network [3, 81]. Thus, they channel metabolic substrates between the blood and neurons, and through their end-feet processes, they surround blood vessels [4, 5] and can easily take up the glucose that arrives through endothelial cells. Alternatively, the astrocyte processes that surround synapses [6, 7] modulate glucose uptake as a function of synaptic activity. Finally, once glucose has been taken up by astrocytes, these rapidly and efficiently distribute this metabolite (in the form of glucose-6-phosphate) to the most inaccessible zones of the brain through gap junctions [82, 83]. As mentioned previously, astrocytes are the only neural cell type that accumulates glucose in the form of glycogen [84]. Accordingly, astrocytes represent an important source of energy under stressful conditions, such as those involving a reduction in the supply of blood and oxygen (ischaemia) to the brain, and under physiological conditions, such as neurotransmission. In fact, synaptic activity stimulates the degradation of glycogen to glucose, which can be converted to lactate [85–87]. The lactate thus formed is released to the interstitial space and is then taken up by neurons, which can use this substrate as an alternative source of energy [76]. A large body of evidence is now reinforcing this hypothesis, and it has been suggested that lactate can preserve neuronal activity during hypoglycaemic episodes, and can even exert a neuroprotective role under certain pathophysiological conditions [88, 89] (reviewed in [80]). Because the ability of astrocytes to store glycogen is very limited [85, 86], however, this polymer would only transiently support such neuroprotection. Thus, astrocytes must express suitable systems to take up glucose so efficiently that, even under stressful conditions, they can account for the well-known ability of these cells to support neuronal energy metabolism.

6.5 MAIN GLUCOSE CARRIERS IN NEURAL CELLS In mammalian cells, glucose is taken up through both the sodium-dependent (SGLT) and the sodium-independent (GLUT) families of glucose transporters. The former group contains at least six members (SGLT1-6) that are expressed at the apical membrane of kidney and intestine epithelial cells, but not in brain cells.


The SGLT take up glucose as a function of the glucose concentration gradient and therefore indirectly require energy in the form of ATP [90]. The second family (GLUT) is expressed in all cells, including neural cells, which take up glucose through a facilitative, energy-independent process favored by the glucose concentration gradient [91]. Up to 14 GLUT members have been identified in humans [92–95], but in the brain the major glucose transporters expressed are GLUT1 and GLUT3, although GLUT4, GLUT5, GLUT2, and GLUT8 are also expressed to a lesser extent. Possibly, all brain cell types express GLUT1 [96], although its expression in cultured neurons is believed to be an adaptive consequence to the in vitro conditions [97]. The expression of GLUT3 is confined to neurons [98, 99], and in light of the kinetics parameters of the glucose uptake process, it appears that GLUT3 transporter activity is responsible for the high affinity of cerebellar neurons for glucose [97, 100, 101]. This observation has led to the notion that GLUT3 activity in neurons would confer protection to these cells against hypoglycaemic episodes [97]. Furthermore, brain GLUT4 immunoreactivity has been found to be restricted to GLUT3-expressing neurons [102], suggesting that both transporters could cooperate under conditions of a lack of glucose supply.

6.6 NO UP-REGULATES GLUCOSE UPTAKE BY ASTROCYTES In vitro activation by incubation of cells with LPS and/or cytokines is known to stimulate glucose utilization in a NO-dependent manner in a broad range of cell types such as macrophages, epithelial cells, smooth muscle cells, endothelial cells, fibroblasts, or pancreatic islets [23, 103–108]. Such a response to NO was first interpreted to be a suicide-like action of the cells in the presence of large amounts of the radical [103]; however, further reports have now established that increased glucose utilization might serve to protect cells from the toxic actions of excess NO. Thus, upon activation, not only astrocytes [109] but also macrophages [106] or epithelial cells [108] respond to an increased consumption of glucose both through glycolytic and pentose–phosphate pathways in an apparent attempt to compensate for the loss of ATP and glutathione oxidation generated by NO and/or other pro-oxidant derivatives. The effects and regulation of these metabolic pathways by NO and peroxynitrite in astrocytes and neurons will be discussed in the following sections. Here, we shall first focus on the hypothesis that any increase in brain oxidative glucose metabolism must be preceded by the specific stimulation of the uptake process in astrocytes. Indeed, the rate of 2-deoxy-D-[U-14C]glucose uptake increases significantly in astrocytes previously activated with LPS [110] (Figure 6.1). 2-Deoxy-Dglucose is a glucose analogue that is taken up by the cells and phosphorylated by hexokinase(s), but the product (2-deoxy-D-glucose-6-phosphate) is not further metabolized and is therefore entrapped by the cells. The measurement of radioactive cells thus represents a good index of the uptake process, at least when this

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(a) Astrocytes

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UT GL

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FIGURE 6.1 Nitric oxide up-regulates glucose uptake. (a) Upon injurious stimuli, astrocytes respond producing NO from iNOS activity, leading to a cyclic GMP-independent activation of glucose uptake. The mechanism responsible for such an effect involves the decrease in the cellular energy charge caused by the interference of NO with cytochrome c oxidase, followed by phosphorylation of AMP-activated protein kinase (AMPK) [158]. (b) In skeletal muscle, NO also promotes glucose uptake. In these cells, muscle contraction triggers AMPK activation, which phosphorylates (and activates) eNOS. The NO so-formed would induce GLUT4 protein translocation through a mechanism involving cyclic GMP.

process is measured at relatively low extracellular glucose concentrations. In fact, due to the relatively low kM of hexokinase(s) (~100 µM), the activity of this enzyme, if saturated, might limit the glucose uptake process itself. Glucose taken up by activated astrocytes is most efficient at extracellular glucose concentrations ranging below 0.25 mM [110], but is not evident at higher glucose concentrations, strongly suggesting an increased affinity of activated astrocytes for glucose. The mechanism(s) responsible for such an increased affinity of astrocytes for glucose appears to be both transcriptional and posttranslational. In this context, the high-affinity GLUT3 glucose transporter is undetectable in astrocytes, but constitutively expressed in neurons [97, 110, 111]. Shortly (as from ~4 h) after incubation of astrocytes with LPS (1 µg/ml), with a low-oxygen atmosphere (~3% O2) or with low-glucose medium, however, a dramatic increase occurs in the abundance of GLUT3 mRNA and protein [110], possibly reflecting a transcriptional effect triggered by LPS. In fact, this increased mRNA abundance can be abolished by coincubating the cells with a NF-κΒinhibitor [110]. Moreover, because NF-κΒ is also


involved in LPS-mediated iNOS expression [112], it can be envisaged that both NO production and the increased affinity for glucose may be coordinated in astrocytes upon the development of an unforeseen stressful condition. Additionally, blockade of iNOS activity in LPS-stimulated astrocytes partially prevents the increased 2-deoxyglucose from being taken up, which strongly suggests that endogenously formed NO may be involved in the stimulating effect. This is further supported by the observation that exogenously applied NO triggers an increase in the rate of 2-deoxyglucose taken up in control (LPS-untreated) astrocytes [110]. The mechanism responsible for the NO-dependent increase in 2-deoxyglucose uptake in astrocytes has not yet been elucidated but, in view of the fact that in these cells NO blocks mitochondrial function [23], it may be speculated that the increase in glucose uptake would be a compensatory bioenergetic effect (Figure 6.1a). In addition, the up-regulation of the GLUT3 transporter by LPS and cytokines would also be advantageous for the brain under those (pathophysiological) circumstances in which the supply of glucose to the tissue decreases and NO increases, such as in brain ischaemia. Regarding the mechanism whereby NO would be involved in direct stimulation of glucose uptake, others have reported—in skeletal muscle cells—that NO activates glucose uptake in a cyclic GMP-dependent fashion [113, 114] by promoting GLUT4 translocation to the plasma membrane [115] (Figure 6.1b). Moreover, this pathway can be triggered by 5′-AMP-activated protein kinase (AMPK), a cell energy sensor that is activated in response to high AMP:ATP ratios [116] and that triggers the phosphorylation of metabolic substrates to maintain the energy balance [117]. The endothelial NOS isoform (eNOS) can also be activated by AMPK during the energy loss associated with ischaemia in the rat heart [118]. Activation of AMPK by the AMP analogue 5-aminoimidazole-4-carboxamide ribonucleoside (AICAR) triggers eNOS-mediated NO production, which is responsible for the cyclic GMP-dependent glucose uptake observed in skeletal muscle cells [115]. In cells that express mainly GLUT1 but not GLUT4, GLUT2, or GLUT3, such as rat liver epithelial clone 9 cells [119], 3T3-L1 pre-adipocytes, or myoblasts [120], AMPK also stimulates glucose uptake [120, 121]. In contrast to GLUT4-expressing muscle cells, however, the stimulation of GLUT1 by AMPK does not involve translocation, but, instead, activation of the transporter at the plasma membrane [120, 121]. Furthermore, the underlying mechanism is NO- and cyclic GMP-independent [121]. In keeping with these results, we have recently found evidence consistent with the notion that GLUT3-mediated increased glucose uptake by NO occurs via AMPK through a cyclic GMPindependent mechanism (P. Cidad, A. Almeida, and J.P. Bolaños, personal communication). Accordingly, the mechanism through which NO and AMPK stimulate glucose uptake appears to depend on the cell type and on the major glucose carrier expressed in that cell type. In astrocytes, the modulation of mitochondrial function by NO may thus represent an indirect mechanism for the regulation of glucose metabolism.

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6.7 GLUCOSE UTILIZATION THROUGH THE PENTOSE–PHOSPHATE PATHWAY: THE ROLE OF NO AND PEROXYNITRITE It is well known that the intracellular content of glutathione (GSH) plays a key role in dictating neuronal vulnerability against NO and peroxynitrite. Thus, both species potently oxidize sulfhydryls, including GSH [122, 123]. Accordingly, neuronal GSH oxidation has been proposed to be a contributing factor leading to the mitochondrial damage and neurotoxicity associated with nitrosative stress [49, 124, 125]. Unlike neurons, astrocytes efficiently maintain GSH in its reduced redox status, even under conditions of excessive endogenous NO formation [23, 58]. Among the factors possibly involved in this different cellular susceptibility, glucose oxidation through the pentose–phosphate pathway appears to play a key role. Indeed, this metabolic route is considered essential during GSH regeneration from oxidized glutathione (GSSG). For instance, hepatocytes are prone to hydrogen peroxide (H2O2)-mediated activation of glucose-6-phosphate dehydrogenase (G6PD), the enzyme that catalyzes the first rate-limiting step in the oxidative branch of the pentose–phosphate pathway [126, 127]. Furthermore, stimulation of this pathway in neurons [128] and astrocytes [129] has been proposed to elicit a protective action against H2O2 toxicity through the pentose–phosphate pathway activity-mediated production of NADPH, a cofactor necessary for GSH regeneration from GSSG [130, 131]. Consistent with those studies, glucose utilization through the pentose–phosphate pathway has been found to be strongly stimulated in astrocytes by LPS (1 µg/ml, 18 h) treatment (Figure 6.2). Moreover, such activation has been reported to be responsible for the self-protection of astrocytes against endogenous NO-mediated GSH oxidation [109]. Because LPS-stimulated astrocytes synthesize O2•− through iNOS-dependent activity [132], it was next hypothesized that peroxynitrite might, at least partially, be responsible for the observed activation of the pentose–phosphate pathway in these cells. Thus, it was found that treatment of astrocytes with peroxynitrite (10 boli of 50 µM each during 5 min) triggered a rapid activation of the pentose–phosphate pathway, together with the accumulation of NADPH concentrations [133]. Investigation of the mechanism responsible for such a rapid effect revealed that peroxynitrite-stimulated G6PD activity in intact cells, as demonstrated by an increase in the ratio of 6-phosphogluconate to glucose-6-phosphate (i.e., the product and substrate of G6PD, respectively) [133]. Furthermore, the expression of a plasmid construct encoding G6PD led PC12 cells to demonstrate enhanced pentose–phosphate pathway activity, NADPH accumulation, protection against GSH oxidation, and resistance to apoptotic cell death. Conversely, expression of a plasmid coding for a G6PD antisense mRNA decreased glucose oxidation through this pathway, lowered NADPH concentrations, raised GSH oxidation, and increased the vulnerability of the cells [133]. Taken together, these results strongly suggest that glucose utilization through the pentose–phosphate pathway may play a key neuroprotective role against nitrosative stress (Figure 6.2).

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 153 Astrocytes

Neurons

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Glutamate

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−

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+

FIGURE 6.2 Nitric oxide formation is associated with glucose-6-phosphate dehydrogenase activation and glutathione regeneration. Upon injurious stimuli (inflammation, ischemia, spesis in astrocytes, or glutamate receptor overstimulation in neurons), iNOS (in astrocytes), or nNOS (in neurons) activities forms NO and oxidizes glutathione; however, defense mechanisms appear to occur in both cell types focused to prevent further glutathione oxidation. Thus, in astrocytes a co-induction of glucose-6-phosphate dehydrogenase (G6PD) exists (i.e., the rate-limiting enzyme of the pentose–phosphate pathway and, in neurons as well as in astrocytes), NO-derived peroxynitrite triggers G6PD translocation and activation. In both cases, active G6PD generates NADPH, which is a necessary cofactor for the regeneration of glutathione, thus promoting cytoprotection.

6.8 LOW DOSES OF PEROXYNITRITE MAY CAUSE NEUROPROTECTION: THE ROLE OF GLUTATHIONE As mentioned previously, neurons are vulnerable cells that upon exposure to either endogenous [39, 134] or exogenous [135] excess NO rapidly (within 1 h) undergo apoptotic death. Strikingly, pretreatment of neurons with peroxynitrite (either 10 boli of 50 µM for 5 min or continuously from 1 mM SIN-1) fully prevents apoptosis shortly (1 h) after NO treatment, although this protection decreases progressively thereafter (to ~60% of protection after 4 h, and to ~25% after 8 h) [133]. Furthermore, these changes in peroxynitrite-mediated neuroprotection indicated a good time-course correlation with the observed changes in the redox glutathione status. Because glutathione oxidation has been implicated in

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NO-mediated neuronal apoptosis [49, 134, 136, 137], it is tempting to speculate that the neuroprotection exerted by peroxynitrite would be associated with its ability to activate pentose–phosphate pathway activity and to generate NADPH. In fact, peroxynitrite triggers a rapid (within 5 min) trans-compartmentalization of G6PD in both astrocytes and neurons [133], a phenomenon previously observed for the activation of G6PD by growth factors in renal cells [138]. If so, maintenance of the reduced status of GSH after peroxynitrite treatment would occur at the expense of increased NADPH availability to serve as the co-factor for glutathione reductase activity (Figure 6.2). The transient neuroprotective role for peroxynitrite [133] is in apparent contradiction with the widely held assumption that peroxynitrite would be the NOderived neurotoxic effector molecule [37–39, 58]. Apparently, the relationship between peroxynitrite-mediated interference with key energy metabolic targets and the time-course of the observed neurotoxicity is an issue that should be revisited. Furthermore, an increasing body of evidence now suggests that NO might play a protective role against O2•−-mediated neurotoxicity [139–142] and H2O2-mediated cytotoxicity [143, 144]. In view of the spontaneous formation of peroxynitrite through the reaction of NO with O2•− [37] and in the light of our own results, it is tempting to speculate that peroxynitrite would mediate these NO-derived protective responses. Taken together, our results [133] may provide a clue for understanding the existing controversy concerning the role of NO formation in cell death/survival decisions.

6.9 GLUCOSE METABOLISM AND GLUTAMATEINDUCED NEUROTOXICITY Exogenously added glucose appears to contribute to cellular protection against glutamate-induced neurotoxicity in primary cortical neurons. Thus, the presence of D-glucose (20 mM) has been found to completely abolish glutamate (100 µM for 5 min) mediated increases in neuronal apoptotic death, an issue that has also been associated with neuronal protection against ATP depletion [145]. This phenomenon cannot be ascribed to a putative nonspecific effect due to the presence of high glucose concentrations in the extracellular medium, because the inactive enantiomer L-glucose, which is not recognized by hexose transporters, is not able to prevent such glutamate-mediated neurotoxicity or ATP depletion. Interestingly, D-glucose is able to restore ATP status only 24 h after challenge by glutamate, but not after 3 h, suggesting that the long-term neuroprotective effect of the sugar would be due to its intracellular uptake and further intracellular metabolism. To gain further insight into the metabolic pathway responsible for such neuroprotection, the rate of lactate release was measured in the culture medium to determine the possible existence of a putative role for the glycolytic pathway in ATP compensation and recovery from glutamate-mediated cell death. In contrast with this hypothesis, lactate concentrations were unchanged by glutamate exposure. This rules out:


1. The possibility that glutamate-mediated ATP depletion might be due to glycolytic inhibition 2. The notion that the neuroprotective effect of D-glucose would be due to a compensative increased glycolytic rate [145] In contrast to the glycolytic pathway, most intracellular glucose taken up during or after glutamate challenge appears to be oxidized through the pentose–phosphate pathway, because D-glucose (but not L-glucose) 1. Abolishes the glutamate-mediated increase in GSH oxidation to GSSG 2. Prevents the loss of NADPH caused by glutamate-receptor stimulation [145] Because NADPH is a necessary co-factor of glutathione reductase during GSH regeneration from GSSG, our results are consistent with the idea that the neuroprotective effect of D-glucose would be due to its metabolism through the pentose phosphate pathway [145]. Whether such a pathway might exert neuroprotection in vivo is unknown, but it is interesting to note that post-mortem brain samples from Alzheimer’s disease patients exhibit decreased glucose transporters GLUT1 and GLUT3 [97], which could point to a limitation in glucose utilization in degenerating neurons.

6.10 NITRIC OXIDE AND GLYCOLYSIS: WHAT IS THE ROLE OF GLYCERALDEHYDE-3-PHOSPHATE DEHYDROGENASE? In cellular extracts obtained from a range of cellular systems treated with either endogenous or exogenous NO an inhibition occurs in the activity of glyceraldehyde-3-phosphate dehydrogenase (G3PD) [146, 147], a non-limiting rate NAD+requiring glycolytic enzyme. GADP together with phosphoglycerate kinase (PGK), is a glycolytic enzyme responsible for the substrate level phosphorylation of ADP to form ATP during glycolysis [148]. The observation that NO inhibits maximal G3PD activity is intriguing [149] because, under identical conditions, NO also stimulates the flux of glucose consumption through glycolysis as measured by the rate of lactate formation [150]. One possible explanation for this apparent paradox is that the degree of G3PD inhibition than can be caused by NO may not be sufficient to limit glycolytic flux [151] because G3PD catalyses a non-rate-limiting step in this pathway; however, such a possibility does not explain why the glycolytic flux is increased. A recent report has revealed that NO donors trigger G3PD release to the cytosol from membrane protein band 3, leading to an enhancement of enzyme activity in red blood cells [152]. Such a mechanism would explain why maximal G3PD activity has consistently been found to be unaltered or inhibited in extracts obtained from NO-treated cells despite having an increased glycolytic rate. However, previous studies addressing

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different kinds of interactions between NO (and ONOO−) and G3PD might provide novel insight into the relationship between G3PD activity and the glycolytic pathway. Initially, it was reported that NO triggers S-nitrosylation of the critical Cys149 residue of G3PD [146, 147], and such protein modification reversibly inhibits G3PD in vitro activity. Later studies revealed that this modification favors the subsequent linking of NADH to G3PD [153], leading to an irreversible inhibition of the enzyme activity. Whether NADH or NAD+ is the co-factor that actually links to G3PD is controversial [153]. In this context, Wu et al. [154] have suggested that NAD+, instead of NADH, would be the co-factor that actually interacts with the protein, such that the NAD-G3PD complex would bind to actin to remain inactive. Furthermore, the latter authors reported that the increased glycolytic flux to lactate observed in cells treated with NO might provide the necessary NAD+ to facilitate the binding of this co-factor to G3PD protein, thus further stimulating its inactivation. Most cell systems tend to decrease their ATP concentrations upon exposure to NO, possibly through a mechanism involving the inhibition of mitochondrial ATP synthesis (see previous sections). Certain cell systems do not exhibit such an effect, however, and, instead, decrease their ATP stores in correlation with G3PD inhibition. This applies to the observation that NO-mediated inhibition of G3PD dehydrogenase activity may occur in parallel with an increase in G3PD acyl phosphatase activity [155]. This observation led to the suggestion that NO could trigger the uncoupling of the glycolytic flux from the substrate level phosphorylation, finally leading to ATP depletion. The latter observation suggests that NO, despite promoting increased flux through glycolysis (see the proposed mechanism below), abolishes ATP generation in this pathway. Nevertheless, the facts reveal that NO stimulates glycolytic pathway activity in astrocytes, where the enhanced glycolytically generated ATP contributes to maintaining the cell energy status [135]. In view of the relevance and recent findings regarding the mechanism through which NO triggers glycolytic flux enhancement, this issue is discussed in the following section.

6.11 NO TRIGGERS GLYCOLYTIC ACTIVATION AND PROTECTS ASTROCYTES FROM CELL DEATH Astrocytes are considered glycolytic cells because mitochondrial toxins, such as antimycin, strongly stimulate glucose metabolism through the glycolytic pathway, whereas neurons are non-glycolytic cells [156, 157]. As with other exogenous mitochondrial toxins, incubation of astrocytes and neurons with NO (1.4 µM from 0.5 mM DETA-NO) strongly inhibits (by ~85%) cellular respiration [135]. The inhibition of mitochondrial respiration occurs simultaneous to an enhanced glycolytic rate in astrocytes, but not in neurons, as assessed by analyses of lactate concentrations in intact treated cells (Figure 6.3). Furthermore, despite an initial slight decay in ATP concentrations, astrocytes are able to prevent themselves from further ATP depletion and cell death, unless glucose is removed from the


media [135]; however, neuronal ATP stores indicate a progressive decrease, ending in depletion and rapid cell death. These correlations strongly suggest that the NO-mediated inhibition of cellular respiration triggers an up-regulation of glycolytic flux rate in certain cell types, such as astrocytes (but not in neurons), thereby preventing them from ATP depletion [135]. Nevertheless, how NO activates the glycolytic pathway in astrocytes or why astrocytes and neurons respond differentially to NO by up-regulating glycolysis are questions that remain to be elucidated (Figure 6.3).

6.12 ON THE MECHANISM WHEREBY NO STIMULATES GLYCOLYSIS IN ASTROCYTES Besides hexokinase and pyruvate kinase, 6-phosphofructo-1-kinase (PFK1) is the key rate-limiting step in the glycolytic pathway [148]. In fact, PFK1 activity increases in astrocytes upon inhibition of mitochondrial ATP synthesis with potassium cyanide, oligomycin, or NO [158]. It should be noted that the increase in PFK1 activity brought about by NO treatment can only be seen in measurements of fructose-6-phosphate (F6P) and fructose-1,6-bisphosphate (F1,6P2) concentrations (i.e., the substrate and product of PFK1, respectively) in neutralized-perchloric cell extracts, but not in in vitro assays of PFK1 enzyme activity. Accordingly, the NO-mediated increase in PFK1 activity would be allosteric, a notion reinforced by the observation that intracellular levels of fructose-2,6bisphosphate (F2,6P2) (i.e., the most powerful PFK1 allosteric activator [159]) can be rapidly (~5 min) and time-dependently (up to 60 min) accumulated in astrocytes, but not in neurons [158] (Figure 6.3). RT-PCR analysis of the F2,6P2forming enzyme, 6-phosphofructo-2-kinase (PFK2), has demonstrated that astrocytes mainly express isoform PFK2.3 (i.e., the one with the highest kinasebisphosphatase activity ratio) [160]. Western blotting against such isoenzyme reveals higher protein contents in astrocytes than in neurons. Furthermore, when PFK2.3 protein expression is silenced by the RNA interference approach, astrocytes are unable to increase F2,6P2, PFK1 activity, and lactate accumulation upon exposure to NO, strongly suggesting that PFK2.3 would be an essential step for NO-mediated allosteric PFK1 activation and glycolysis flux stimulation [158]. Investigation of the molecular mechanism whereby NO triggers PFK2 and PFK1 activation has indicated that it is a cyclic GMP-independent mechanism requiring prior inhibition of mitochondrial respiration [158] (Figure 6.3). Thus, the inhibition of astrocyte respiration by NO lowers ATP concentrations by ~25% [135] and may lead to subsequent elevations in AMP concentrations. In the heart, a response to in vivo ischaemia consists of the stimulation of AMP-activated protein kinase (AMPK)-triggered enhancement of PFK2.2 isoform activity [161]. In astrocytes treated with NO, AMPK is also activated, as judged by the phosphorylation of the Thr172 residue in the α1 subunit of AMPK [158]. Moreover, the abolition of AMPK-α1 subunit protein by the RNA interference approach renders astrocytes unable to form F2,6P2 and to activate PFK1 and the glycolysis

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Nitric Oxide, Cell Signaling, and Gene Expression Infammation, Sepsis, Hypoxia, Ischemia, Neuronal necrosis, ...

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FIGURE 6.3 Proposed signalling pathway through which NO rapidly activates glycolysis in astrocytes and leads to cytoprotection. Upon injurious stimuli, astrocytes synthesize NO, which down-regulates mitochondrial function at the level of cytochrome c oxidase. The subsequent decrease in the cellular energy charge phosphorylates and activates AMPK, which activates PFK2 to synthesize F2,6P2 (i.e., the most potent PFK1 allosteric effector). This cascade results in the rapid activation of the glycolytic pathway, thus conferring protection to astrocytes. In contrast, such a mechanism is not fully developed in neurons, in which the appropriate PFK2 isoform expression is not detectable (see Reference 158).

flux upon NO treatment [158]. Together, these observations are compatible with the notion that the mechanism through which NO activates glycolysis involves an inhibition of mitochondrial respiration that leads to AMP enhancement. AMP, when transiently increased, would activate AMPK, which phosphorylates and activates PFK2 to form F2,6P2. Thus, elevations in F2,6P2 would stimulate PFK1 activity allosterically, leading to a rapid increase in the glycolytic flux [158] (Figure 6.3).


What can this signaling pathway teach us about the role of NO in cell death and survival pathways? The Western blotting and biochemical data obtained by Almeida et al. [158] indicate that neurons virtually lack operative PFK2 activity. Neurons rapidly die upon treatment with doses of NO that do not kill astrocytes, and the abolition of either AMPK or PFK2.3 significantly increases apoptotic cell death in NO-treated astrocytes. These data indicate that the ability of NO to activate glycolysis affords protection to cells with the necessary biochemical machinery, such as PFK2, to rapidly respond to the mitochondrial inhibition and AMP elevations that follow exposure to NO [158]. Thus, the synthesis of ATP by substrate level phosphorylation at glycolysis may represent a transient response of some cells to provide glycolytic intermediates for other metabolic pathways and/or to compensate for the energy failure that would otherwise lead to cell death.

6.13 CONCLUDING REMARKS Following inflammatory brain stimuli, astrocytes actively synthesize nitric oxide and peroxynitrite. These nitrogen-derived species trigger a cascade of biochemical effects, including alteration of mitochondrial function and of the glutathione redox status in both astrocytes and neighboring neurons. Furthermore, under such nitrosative stress, astrocytes demonstrate remarkable resistance, despite having their mitochondria impaired, whereas neighboring neurons demonstrate vulnerability. In this review, we have discussed recent evidence that strongly suggests that nitrogen-derived species modulate key regulatory steps in glucose metabolism that would be responsible for the cellular resistance. These involve up-regulation of the high-affinity glucose transporter, the stimulation of glycolysis at the level of 6-phosphofructo-1-kinase, and activation of the pentose–phosphate pathway at the level of glucose-6-phosphate dehydrogenase. We conclude that the orchestrated stimulation of glucose-metabolizing pathways by nitric oxide would be a transient attempt of certain neural cells to compensate for an impaired energy status and oxidized glutathione, and thus emerge from an otherwise potentially neuropathological outcome.

6.14 ACKNOWLEDGMENTS J.P.B. was funded by the M.C.Y.T. (SAF2001-1961) and J.C.yL. (SA081/04). A.A.P. was funded by the FIS (03/1055) and J.C.yL. (SA020/02). P.G.-N. is the recipient of a Postdoctoral Marie Curie Fellowship.

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Oxide Cell 7 Nitric Signaling Mediated by cGMP Emil Martin, Iraida Sharina, Aurora Rachel Seminara, Joshua Krumenacker, and Ferid Murad University of Texas Health Science Center, Houston, Texas

CONTENTS 7.1 7.2

Introduction ............................................................................................168 The NO/cGMP Pathway ........................................................................169 7.2.1 NO Synthases—Upstream Element of the NO/cGMP Signaling ..................................................................169 7.2.1.1 The Three Musketeers of NO Synthesis .....................169 7.2.1.2 Ca2+-Regulation of NOSs ............................................169 7.2.1.3 Sub-Cellular Localization of NOS and Its Dynamics 170 7.2.2 Soluble Guanylyl Cyclase—Switching from NO to Second Messenger cGMP .......................................................................171 7.2.2.1 Structure-Functional Organization of sGC— Receptor for NO ..........................................................171 7.2.2.2 Role of Heme in sGC Function ..................................172 7.2.2.3 Dynamics of Sub-Cellular Localization of sGC .........173 7.2.3 Cyclic Nucleotide-Gated Channels—cGMP Effector Molecule .......................................................................174 7.2.3.1 CNG Channel Expression and Composition ...............174 7.2.3.2 Structure and Regulation of CNG Channels ...............174 7.2.3.3 CNG Channel Function ...............................................175 7.2.3.4 Pathologies Related to CNG Channels .......................176 7.2.3.5 CNG Channel Regulation by NO/cGMP Signaling ...176 7.2.4 Phosphodiesterases and NO/cGMP—Keeping the Cyclic Nucleotides in Check .................................................................177 7.2.4.1 Characteristics of Different PDE Families Affecting the Levels of Intracellular cGMP ...............178 7.2.5 cGMP-Dependent Protein Kinases—Divergence of the NO/cGMP Pathway ....................................................................182 167

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7.3

NO-cGMP Signaling in Selected Various Cellular Processes ..............183 7.3.1 NO/cGMP Signaling and Vascular Relaxation .........................183 7.3.1.1 Reduction of Intracellular Ca2+ ...................................183 7.3.1.2 Ca2+ Desensitization of the Contractile System ..........184 7.3.1.3 Regulation of Thin Filament Function ........................185 7.3.2 NO/cGMP Signaling and Platelet Aggregation .........................185 7.3.2.1 Ca2+ Homeostasis .........................................................186 7.3.2.2 Surface Receptors ........................................................186 7.3.2.3 Cytoskeletal-Associated Proteins ................................187 7.3.3 NO/cGMP Signaling and Neurotransmission ............................187 7.3.3.1 Synaptic Plasticity .......................................................188 7.3.3.2 Perception of Pain ........................................................188 7.3.3.3 NO in Neurotransmission ............................................189 7.3.4 NO/cGMP Signaling and Kidney Function ...............................189 7.3.5 NO/cGMP in Reproduction .......................................................190 7.3.6 NO/cGMP in Bone Homeostasis ...............................................190 7.3.7 NO/cGMP Signaling and Gene Regulation ...............................191 7.3.8 NO/cGMP Signaling and Apoptosis ..........................................191 7.3.9 NO/cGMP and Intestinal Motility .............................................192 7.3.10 Self-Regulatory Interactions in NO/cGMP Cascade .................192 Closing Remarks ....................................................................................193 References ...............................................................................................193

7.4

7.1 INTRODUCTION Over the last two decades our understanding of nitric oxide’s (NO) biology evolved from the perception of it as a hazardous atmospheric pollutant [1] to a key secondary messenger molecule mediating intra- and intercellular signals, only to return later as one of the culprits in various pathologic processes and illnesses. Since the postulation [2, 3] and subsequent demonstration [4, 5] of the NO’s function as endothelium-derived relaxation factor (EDRF), NO was reportedly involved in a plethora of physiologic processes that include, but are not limited to, smooth muscle relaxation, inhibition of platelet and leukocyte aggregation, attenuation of vascular smooth muscle cells proliferation, neurotransmission, and immune defense. NO has also been implicated in the pathology of many inflammatory diseases, including arthritis, myocarditis, colitis, and nephritis, as well as a large number of pathological conditions such as amyotrophic lateral sclerosis (ALS), cancer, diabetes, and neurodegenerative diseases [6–15]. The variety of physiological and pathophysiological processes with which NO is associated is a direct reflection of the concentration dichotomy of NO-dependent effects. On one hand, pathophysiological processes and NO-dependent protein modifications, such as nitrosylation and nitration, require large concentrations of NO (>1 µM) or accumulation of NO metabolites [16]. On the other hand, NO-dependent smooth muscle relaxation, vasodilation, platelet, and leukocyte aggregation and neurotransmission require only nanomolar concentrations of NO.

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Several processes have been documented to occur at low NO levels. Nanomolar concentrations of NO inhibit the terminal complex IV (cytochrome oxidase) from the mitochondrial respiratory chain [17–19] with a half-inhibitory concentration in the range of 60 to 270 nM [17]. A more detailed review of this process is presented in the accompanying chapter of this book. Another early recognized effect of low concentration of NO is the increase in intracellular levels of 3′–5′ cyclic guanosine monophosphate (cGMP) upon exposure to various NO donors and endogenous NO production [2, 20]. The goal of this chapter is to discuss the NO-cGMP signaling pathway, the key players, and their interaction, as well as to review some of the cellular and physiological processes regulated or affected by this pathway.

7.2 THE NO/CGMP PATHWAY 7.2.1 NO SYNTHASES—UPSTREAM ELEMENT NO/CGMP SIGNALING

OF THE

7.2.1.1 The Three Musketeers of NO Synthesis Physiological synthesis of NO is performed by nitric oxide synthase (NOS), a specialized enzyme that catalyzes the O2-dependent oxidation of the guanidino nitrogen of L-arginine resulting in the formation of NO and L-citrulline. The NOS enzyme is the first element of the NO/cGMP pathway. Three independent isoforms of NOS termed NOSI (first identified in neuronal tissue and also termed nNOS), NOSII (first detected in macrophages and also termed iNOS), and NOSIII (first detected in endothelium and also termed eNOS) have been characterized. All isoforms are homodimers that share homology in their C-terminal half, termed the “reductase domain,” to the cytochrome P450 reductase. The N-terminal half of the enzyme termed the “heme or oxygenase domain” contains the binding site for the heme group and substrate L-arginine. In addition to oxygen, L-arginine, and the heme prosthetic group, catalysis of NO requires the reduced form of nicotinamide adenine dinucleotide phosphate (NADPH), flavine adenine dinucleotide (FAD), and mononucleotide (FMN) as well as tetrahydrobiopterin, all of which are bound to the enzyme. Various L-arginine analogs are widely used as competitive and irreversible inhibitors of NOS enzymes. Their administration is used as one of the key tests to determine the involvement of NO-dependent pathways in investigated processes. The flow of electrons from the reductase to oxygenase domains is essential for the NO catalysis and is the subject of an enzyme’s in vivo regulation. 7.2.1.2 Ca2+-Regulation of NOSs All NOSs contain a consensus sequence between the reductase and oxygenase domain capable of binding calmodulin. The transfer of electrons between the reductase and oxygenase domain is regulated by the binding of calmodulin. NOS1

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and NOS3 isoforms bind calmodulin only after an increase in intracellular [Ca2+]i (> 500 nm). Under normal conditions, many agonist-receptor interactions result in transient transmembrane Ca2+ fluxes or release of Ca2+ from intracellular storage compartments. The transient nature of such fluxes dictates the transient production of NO by NOSI and NOSIII and is necessary for physiological signaling through the NO/cGMP pathway. Interaction of calmodulin and the NOSII isoform, however, can occur at resting [Ca2+]i (< 100 nm), which makes the NOSII isoform independent of changes in intracellular Ca2+. The NOSII enzyme is fully active and capable of generating large fluxes of NO soon after its synthesis. These large and prolonged fluxes determine the anti-microbial and anti-parasitic function of the NOSII isoforms [21]; however, such prolonged output of NO results in a significant increase of intracellular cGMP levels, which could be detrimental as demonstrated by the precipitous drop in blood pressure in cases of septic shock. Large concentrations of NO generated by the NOSII enzyme under inflammatory conditions are the basis of deleterious nitrosative stress. Various aspects of this nitrosative stress are covered in detail in the accompanying chapters of this book. This enzymatic property of NOSII dictates a tight regulation of the amount of NOSII isoform expressed [21], which occurs mainly at the transcriptional level. 7.2.1.3 Sub-Cellular Localization of NOS and Its Dynamics NOS enzymes display multiple sub-cellular localizations. Originally described in mainly the cytosolic compartment, the NOSI enzyme was later detected in a membrane-bound form as well [22]. NO synthesis in the nervous system is predominantly regulated by the influx of Ca2+ through receptor-dependent channels, in particular following the stimulation of the NMDA receptor by glutamate [23]. The NOSI enzyme possesses a PDZ domain, which interacts with PSD 93 and 95 proteins [24, 25]. PSD 95 also binds directly to the NMDA receptor [26], thus acting like a scaffolding protein allowing NOSI to be exposed to the Ca2+ entering the ion channel of the activated NMDA receptor. In skeletal muscle, the NOSI enzyme is targeted to the membrane structures due to the association with α1-syntropin [25], which shares homology with PSD95 and contains a PDZ domain. NOSI is also found to bind to the protein inhibitor of NOS (PIN) protein [27], which may act as an NOS inhibitor [27] and as an axonal transport protein for NOSI [28]. NOSI can also be inhibited by the interaction with caveolin-1 and caveolin-3, which scaffold the NOSI enzyme to the membrane [29, 30] and interfere with calmodulin binding to the enzyme. NOSIII was initially described in both soluble and particulate fractions from the bovine aorta endothelial cells [31]. Interaction of the NOS enzyme with caveolin proteins was originally described for the NOSIII enzyme [32, 33] shortly after the discovery of NOSIII association with plasmalemmal caveolae [32]. NOSIII is distinct from other NOS isoforms by its acylation, such as palmetoylation and myristoylation [34], which is required for enzyme sequestration to the membrane [35]. Caveolin-1 and caveolin-3 were co-precipitated with NOSIII in


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endothelial cells and cardiac myocytes, respectively [33]. It appears that a counterbalancing modulation of NOSIII activity by caveolin and calmodulin exists, which is also regulated by the level of NOSIII acylation [36]. Compartmentalization of various signaling molecules, such as growth factor and hormonal receptors, G proteins, and protein kinases in caveolae, may facilitate the coupling between agonist stimulation and NOSII activation. The NOSII isoform was found in both membrane and cytosolic fractions [37, 38] and detected in macrophages in association with intracellular vesicles [39]. The molecular mechanism of this association remains to be determined. NO produced by the NOS enzyme is an easily diffusible and membrane permeable messenger. The distance of NO diffusion (up to 300 µm in diameter) implies that synthesized NO can influence the function of proteins not necessarily located in the same cell. This property of the NO molecule is the basis for the paracrine effects of NO because the main receptor of NO is often located in the cytosol of the adjacent cell.

7.2.2 SOLUBLE GUANYLYL CYCLASE—SWITCHING FROM NO TO SECOND MESSENGER CGMP As the intracellular calcium fluxes are transient, so is, under normal conditions, the production of NO by the Ca2+-dependent NOSI and III. To preserve and amplify the original signal(s) the NO messenger recruits the next player in the NO/cGMP cascade—soluble guanylyl cyclase (sGC). 7.2.2.1 Structure-Functional Organization of sGC— Receptor for NO sGC is a member of a large family of receptor proteins, which upon activation with various ligands enhance their catalytic ability to synthesize cGMP from GTP [40]. Soluble guanylyl cyclase was originally identified as a guanylyl cyclase activity in the cytosolic fraction of cellular or tissue lysates. This cytosolic activity differed from the activity of membrane-bound GC activity by its response to Ca2+, ATP, detergents, and kinetics of cGMP synthesis [41]; however, the main distinction of cytosolic GC activity was its activation by sodium nitroprusside, nitroglycerine, azide, hydroxylamine, and NO [42]. These findings made it possible to postulate [2], and later confirm in several systems [43, 44], that NO is the physiological activator of sGC. This NO-dependent activation of sGC and the coupled switch of the message from NO to cGMP is the key step in the NO/cGMP pathway. Exposure of sGC enzyme to NO donors transforms a weak and inefficient enzyme with low cGMP output (specific activity 0.01 to 0.1 µmole cGMP/min/mg sGC) [45] into a robust high-output sGC with a specific activity several hundred-fold higher (10 to 35 µmole cGMP/min/mg sGC) [45–47] sGC is a heterodimer composed of one α and one β subunit [48]. To date, two isoforms for each sGC subunit have been identified (α1, α2, β1 and β2) in mammals [49, 50]. The most common sGC isoform α1β1 is detected throughout

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the body, although lungs, nervous system, and liver exhibit the highest level of expression. The α2β1 heterodimer was found in the uterus, placenta, and brain [51]. Despite detection of β2 mRNA in the kidney and liver [52], no sGC heterodimers containing β2 subunit have been identified so far in vivo. sGC heterodimers were indicated to be the active form of sGC [53] because expressed α1/α1 and β1/β1 homodimers were inactive [54]. Interestingly, recombinant β2/β2 homodimer also had enzymatic activity [55], although only in the presence of non-physiologic concentrations of Mn2+ ion. The physiological meaning of this characteristic remains to be determined. The C-terminal portions of both α and β subunits are highly homologous to the C-terminal portions of all guanylyl cyclases and to the catalytic region of the adenylyl cyclases [56]. This portion of enzyme contains the cyclase homology domain carrying the catalytic function [56]. The catalytic domain of GC enzymes has been modeled according to the tertiary structure determined for adenylyl cyclases [57] and residues responsible for the substrate specificity predicted. Site-directed mutagenesis confirmed this prediction by conversion of NO-activated GC into an NO-activated AC [58]. Deletion probing of the N-terminal portion of both the α and β subunit clearly demonstrated their importance in the NO-dependent up-regulation of sGC activity [59, 60]. The segment between the regulatory and catalytic domains of sGC subunits is regarded as the region necessary for dimerization [59]. 7.2.2.2 Role of Heme in sGC Function sGC is a hemoprotein containing one protoporphyrin type IX heme moiety per heterodimer [61]. It is the heme moiety of the enzyme that is the target of NO. Extensive analysis of the purified sGC by UV-Vis [45, 62], EPR [63], resonance Raman spectroscopy [64, 65] demonstrated that heme is in a ferrous fivecoordinated, high-spin histidyl complex similar to deoxyhemoglobin and deoxymyoglobin; however, in contrast to deoxyhemoglobin and deoxymyoglobin, sGC is stable in aerobic conditions and does not bind oxygen. This resistance to oxygen is the key property of sGC that allows the enzyme to be constantly available for binding of NO. Direct kinetic measurements of the NO binding [66] indicated that NO binds extremely fast to form a hexacoordinated intermediate with a kon of 1.4 × 108 M−1s−1. This intermediate, however, in which the heme iron retains coordination bonds with both the NO ligand and proximal histidine residue, is transient and is rapidly transformed (k = 2.4 × 105 M−1s−1) into a more stable pentacoordinated NO-heme complex [66] with a half-life ranging from few seconds to several minutes [67]. These binding kinetics indicate that sGC acts as an extremely fast and specific receptor for NO. Kinetic comparison of the transformation dynamics of the NO-heme adduct and of the catalytic properties of sGC clearly demonstrate that formation of pentacoordinated heme due to the cleavage of the iron-histidine bond coincides with the transition of sGC into a high-output mode [66]. These conclusions correlate well with the finding that heme-deficient sGC can be efficiently activated by the protoporphyrin IX alone, which does not make any coordinating bonds without NO [68].


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However, the cleavage of the proximal coordination does not appear to be the only requirement for activation of the enzyme. Site-specific mutagenesis of sGC identified residue histidine 105 of the β1 subunit as the proximal ligand of heme [69, 70]. The heme deficient sGC mutant carrying phenylalanine substitutions in position 105 of β subunit did not demonstrate increased activity even when reconstituted with heme or protoporphyrin IX, despite the absence of the heme-coordinating bond. Interestingly, when His105 was substituted with a cysteine residue, the mutant enzyme displayed a high-output activity when assayed in thiol-free conditions [46]. This high-output activity was significantly inhibited by thiols. We suggested that the thiol-sensitive function of the cysteine 105 residue of the mutant is to stabilize the structure of the heme pocket. In the wild type enzyme, the protoporphyrin moiety of the heme group could have such a function. Thus, the sGC heme moiety appears to play a dual role: 1. Heme acts as a negative regulator of the regulatory domain through the coordination bond with His105. 2. The heme prosthetic group supports the structure of the regulatory domain permitting the activating function of the regulatory domain. Binding of NO to the heme abolishes the inhibitory function of the heme, but preserves its positive effect on the regulatory domain. Despite the in-depth analysis of the changes that occur in the heme-surrounding area as well as a good understanding of the catalytic domain provided by the tertiary structure of the related adenylyl cyclase [57, 71], the mechanism of the intramolecular signal transduction is still unknown. It is possible that only after determination of the tertiary structure of full-length sGC with and without the NO ligand will we gain insights into the precise mechanism that provides the switch from NO-mediated to cGMP-mediated message in the NO/cGMP pathway. 7.2.2.3 Dynamics of Sub-Cellular Localization of sGC sGC is largely a cytosolic protein; however, several recent reports suggest that, at least in certain cases, dynamics occur in the sub-cellular localization of sGC. The α2/β1 heterodimer was found to be co-precipitated with PSD95 in synaptosomes [72]. This interaction appears to be dependent upon the PDZ domain of PSD 95 and the C-terminal portion of the α2 subunit of sGC. Such localization assembles NOSI and sGC in close proximity to the NMDA receptor, suggesting that it is the α2/β1 isoform in neuronal tissues that could be the main receptor for the NOSI-derived NO. Studies performed in platelets and lung endothelial cells suggested that sGC translocates to membrane of platelets and caveolae of endothelial cells in a manner dependent on increased intracellular Ca2+ [73]. It is speculated that such translocation of sGC to membrane compartments sensitizes it to the effects of NO [73] due to conglomeration of the upstream players of the NO/cGMP pathway into one location.

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7.2.3 CYCLIC NUCLEOTIDE-GATED CHANNELS—CGMP EFFECTOR MOLECULE Cyclic nucleotide-gated (CNG) channels constitute a special class of downstream effectors of cGMP. CNG channels are directly activated by cyclic nucleotides through a binding site within the channel protein. It appears that all CNG channels respond, at some level, to both cAMP and cGMP. As discussed below, however, some CNG channels are able to distinguish between the two ligands. First recognized for their ability to control light sensitive conductance in rod photoreceptors [74], CNG channels have also been found in cone photoreceptors [75], olfactory sensory neurons [76], pineal gland [77], kidney, testis, and heart [78]. Following cyclic nucleotide binding, CNG channels nonselectively allow the passage of divalent cations through the plasma membrane and into the cell to mediate, among other processes, photo, and olfactory reception. cGMP production in the context of CNG channel regulation mainly refers to that produced by membrane-associated (particulate) guanylyl cyclase; however, an increasing amount of evidence indicates that NO-stimulated cGMP production through the soluble guanylyl cyclase (sGC) isoforms could also be important for CNG channel modulation. 7.2.3.1 CNG Channel Expression and Composition To date, the majority of information defining CNG channel expression has primarily focused on retinal photoreceptors and olfactory sensory neurons. Six CNG channel members have been identified in mammals and can be divided into two subtypes, the α subunits (CNGA1-4) and the β subunits (CNGB1 and 3) [79]. Although CNGA1-3 can all exhibit activity as a homomeric channel when expressed in heterologous systems, the CNGA4 and CNGB subunits cannot; however, co-expression of CNGA1 and CNGB1 subunits results in a channel with permeation, cyclic nucleotide specificity, and regulation similar to native channels [80, 81]. Native CNG channels form a heterotetrameric structure composed of three subunits and one β subunit [82–84]. In rod photoreceptors, the heterotetramer is composed of three CNGA1 and one CNGB1 subunits [85–87]. The CNG channels of cone photoreceptors consist of CNGA3 and CNGB3 subunits [85–89], whereas CNG channels from olfactory neurons are composed of CNGA2, CNGA4, as well as an alternatively spliced form of CNGB1 [90–95]. The stoichiometry of CNG channel formation in olfactory neurons is yet unknown. In addition to retina and olfactory sensory neurons, expression of CNG channels was also found in the hippocampus, heart, testes, kidney, pancreas, adrenal gland, and colon [96–102]. The function of CNG channels in these latter tissues is not completely understood. 7.2.3.2 Structure and Regulation of CNG Channels Each subunit of the CNG channels contains six transmembrane segments (S1–S6), a P-loop, and intracellular amino and carboxy-terminal regions [83, 103–106].


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The four subunits of each heterotetrameric CNG channel are organized to form a central pore through which the ions pass [82–84]. This is structurally similar to other P-loop containing ion channels such as the bacterial potassium channel KcsA [107]. The P-loop consists of an α helix (C-helix or pore-helix), which is thought to undergo a conformational change upon opening. The intracellular carboxy-terminal region of the CNG channel contains the cyclic nucleotide binding domain (CNBD) and a region termed the C-linker. The CNBD shares sequence similarity with domain regions of other proteins that bind cGMP, such as cGMP and cAMP-dependent protein kinases and the E. coli protein known as catabolite activator protein (CAP) [108]. Photoreceptor CNG channels do exhibit a high degree of specificity for cyclic nucleotides. For example, the free energy of opening for CNGA1 channels in the presence of cGMP is lower than with cAMP because the opening of the channel by cGMP occurs up to three orders of magnitude more readily [94, 108–111]; however, both cAMP and cGMP can fully stimulate channel activity when using saturating conditions. Olfactory channels, on the other hand, appear to have similar affinities and specificity for cAMP and cGMP [93, 95, 112, 113]. The ligand specificity for the CNGA1 rod channels have been demonstrated to rely on a threonine (T560) and aspartic acid residues in the C-helix (D604). Also important is a region within the C-linker that serves as an allosteric regulatory site by transition metals, such as Ni+2, Zn+2, Cd+2, Co+2, and Mn+2 [84, 110, 114, 115], which enhance responses to the ligand. Besides the binding of cyclic nucleotides, modulation of CNG channels has been demonstrated to occur by various additional mechanisms. For example, tyrosine phosphorylation of the CNGA1 ligand-binding domain decreased its sensitivity to cyclic nucleotides [116, 117]. Serine/threonine phosphorylation of rod channels or specific PKC phosphorylation of CNGA3 channels were also demonstrated to decrease ligand sensitivity [118, 119]. In CNGA2 channels, PKC phosphorylation increased nucleotide sensitivity [120]. CNG channels have also been demonstrated to be glycosylated, which may affect their function [92, 106, 121]. Lipids, such as diacylglycerol (DAG), can also affect channel activity [122, 123]. There appears to be an endogenous Ca+2 binding protein other than calmodulin, which regulates cGMP-dependent modulation of cone CNG channels [110, 124]. Finally, the sensitivity of CNG channels was affected by circadian rhythms [125]. 7.2.3.3 CNG Channel Function The mechanism of CNG channel function is best characterized in retinal CNG channels, which were first described in the plasma membrane of the outer segment of rod photoreceptors. In the absence of light, cGMP levels are relatively high in cells containing rod photoreceptors. This results in the opening of CNG channels after the binding of cyclic nucleotides to CNBD. The inward flow of Ca+2 and Na+ ions depolarizes the membrane in the outer segments of the rods. When the retina is exposed to light, a phototransduction cascade is activated following rhodopsin activation and GTP hydrolysis by the G protein transducin [126]. GDP

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bound transducin then activates phosphodiesterase activity and cGMP hydrolysis. Decreased cGMP levels directly affect the CNG channels and inhibiting cation passage, resulting in hyperpolarization and decreased glutamate release from presynaptic terminals [127]. Decreased channel activity means decreased cytoplasmic Ca+2 levels [128] and subsequent stimulation of cGMP synthesis through particulate guanylyl cyclase [129, 130]. It was demonstrated that increased cGMP levels actually increase the affinity of the CNG channels for cGMP [131], and decrease the activity of rhodopsin [132, 133]. Therefore, although Ca+2 is permeable through the CNG channel, it also acts as a negative feedback regulatory mechanism to block the permeability of monovalent cations [134, 135]. In rod and olfactory channels, Ca+2 passage into the cell activates calmodulin, then binds to the CNGA2 amino terminal domain and down-regulates channel activity. This is proposed to be the mechanism for olfactory adaptation [136–139]. Ca+2 entry into the cell through the CNG channels is balanced by the Ca+2 exit through a Na+/Ca+2-K+ exchange [140, 141]. 7.2.3.4 Pathologies Related to CNG Channels The importance of CNG channel activity in physiology has been confirmed with the identification of related pathologies thought to be caused by CNG malfunction. For example, retinitis pigmentosa, a heterogeneous group of diseases that result in blindness caused by the degeneration of rod and cone photoreceptors, is reportedly caused by mutations in the gene encoding the CNGA1 subunit [142]. Furthermore, mutations in either the CNGA3 or CNGB3 genes, which are expressed in cone photoreceptors, cause total color blindness [143–145]. The disruption of CNG channels in gene knockout models (in particular CNGA2, CNGA3 and CNGA4 channels) has also been established, and has shed some light into their physiological function. In this regard, animals deficient in CNGA2 gene do not exhibit cAMP-sensitive olfactory channel activation and suffer from anosmia [146]. Animals with disruption of this gene also appeared to display biochemical and morphological changes in the olfactory sensory and surrounding periglomerular and atypical neurons [147]. CNGA2 knockout mice may also exhibit attenuated long-term potentiation in the hippocampus [148]. Deletion of the CNGA3 subunit resulted in mice that did not display conemediated photoresponses, whereas the rod responses were completely functional [149]. CNGA4 channel deletion results in a defect in odorant-dependent adaptation [150]. 7.2.3.5 CNG Channel Regulation by NO/cGMP Signaling Although the majority of information gained about CNG channel function and regulation has been done through exogenous expression of CNG or activation of particulate soluble guanylyl cyclase, increasing evidence suggests that NO may also play a role in the stimulation of CNG channels. Retinal cells express NOSI and produce NO that is released into cone terminals, which activates soluble


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guanylyl cyclase [151–154]. In olfactory neurons, as mentioned earlier, cAMP plays a critical role in their functionality. NO may also play a critical role in the regulation of the olfactory channel. Stimulation of olfactory sensory neurons by an NO donor occured concomitantly to an increase in cGMP levels [155]. Whether or not this activation occurs through a cGMP-dependent mechanism is questionable; however, mutation of a specific cysteine residue to serine within the channel abolished the NO-induced activity. It has been proposed that nitrosylation of cysteine’s thiol by NO is responsible for channel activation [155, 156]. Interestingly, nitrosylation-dependent activation of CNG channels may be specific to olfactory sensory neurons because rod and cone photoreceptor channels in a C. elegans model were not activated in a similar manner [154, 157, 158]. A more detailed analysis of direct effect of NO on the CNG channels is reviewed in an accompanying chapter of this book. Evidence of NO-stimulated channel activation through cGMP production also exists. For example, odorant-stimulated cGMP production in rat olfactory neurons could be blocked by NOS inhibition and NO scavenging [159]. In Xenopus olfactory neurons, exogenous NO or cGMP induced similar currents, suggesting that NO activation occurs through cGMP [160, 161]. In addition, rat retinal ganglion cells were found to respond to NO stimulation by CNG channel activation [162]. It is important to note, however, that developing and regenerating olfactory sensory neurons and not mature olfactory sensory neurons (OSN) have been demonstrated to exhibit NOS activity [163–166]. With this in mind, carbon monoxide has also been suggested as an activator of the soluble guanylyl cyclases in OSNs [167]. Presently, the role of NO in CNG channel modulation has not been fully clarified and requires further investigation.

7.2.4 PHOSPHODIESTERASES AND NO/CGMP—KEEPING CYCLIC NUCLEOTIDES IN CHECK

THE

Cyclic nucleotide phosphodiesterases (PDEs) regulated by cGMP binding and performing cGMP hydrolysis have critical roles in NO/cGMP signal transduction. PDEs catalyze the breakdown of cGMP produced by soluble guanylyl cyclase in response to NO and play an important part in determining the actual tissue cGMP levels by balancing and modulating the production-decomposition process. PDEs cleave the cyclic nucleotide phosphodiesther bond between the phosphorous and oxygen atoms at the 3′-position [168]. The catalytic mechanism involves a nucleophilic attack by the OH− of an ionized molecule of water, production of which is facilitated by divalent metal cations bound in the conserved metal binding motifs within PDEs. The catalytic domains of all known mammalian PDEs contain two Zn2+ binding sequences (HX3HXn(E/D)) arranged in a tandem [169]. PDEs were first discovered by Sutherland and co-workers more than five decades ago [170, 171]. Presently, more than 50 different mammalian PDEs exist, which are divided into 11 enzyme families based on coding sequence, domain

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structure and regulation [172]. Some of these families contain several isoforms coded by independent genes, many of which have several alternative splicing forms resulting in proteins with different aminoacid sequences. This chapter concentrates on the PDEs that hydrolyze cGMP or are regulated upon binding of cGMP and therefore have the potential to participate in NO/cGMP signal transduction. Modulation of PDE activity introduces significant changes in tissue cGMP levels, which in turn alter the physiological response initiated by NO/cGMP signaling. Therefore, it is not surprising that all PDEs are highly regulated enzymes. General modes of PDE regulation include: 1. Regulation by substrate availability, which is accomplished by changes in concentration of cyclic nucleotides or competition by another cyclic nucleotide as in the case of dual specificity PDE1, 2, and 3 2. Regulation by extracellular signals inducing various intracellular signaling mechanisms such as phosphorylation (PDE3); Ca2+/calmodulin association (PDE1); and stimulation of PDE3 activity by insulin or PDE6 by photons through transducin system 3. Feedback regulation by allosteric binding of cGMP, which promotes changes in activity (PDE2, 5, 6, and10) 4. Cellular compartmentalization and changes in expression levels in response to chronic exposure to high levels of cyclic nucleotides (PDE3) [173] TABLE 7.1 Enzyme PDE1 PDE2 PDE3 PDE5 PDE6 PDE9 PDE10 PDE11

Isoforms A, B, C A A, B A A, B, C A A A

Substrate Specificity cGMP/cAMP cGMP/cAMP cGMP/cAMP cGMP cGMP cGMP cGMP/cAMP cAMP/cGMP

Regulation by cNMP None cGMP-stimulated cGMP-inhibited cGMP-stimulated cGMP-inhibited cAMP-inhibited cAMP-inhibited cAMP-inhibited

7.2.4.1 Characteristics of Different PDE Families Affecting the Levels of Intracellular cGMP 7.2.4.1.1 PDE1 All PDE1 enzymes are activated by Ca2+/calmodulin binding [174]. Three PDE1 variants are encoded by three different genes (PDE1A, PDE1B, and PDE1C) and demonstrate different catalytic properties and selectivity toward substrates. PDE1A and PDE1B genes each encode two splice variants (A1,2 and B1,2), and the PDE1C gene has five different amino-terminal and carboxy-terminal splice


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variants (C1-5). Although PDE1A and PDE1B enzymes hydrolyze cGMP with similar high affinity, their affinity for cAMP is relatively low, although PDE1B hydrolyzes cAMP more efficiently than PDE1A. All PDE1C variants hydrolyze both cAMP and cGMP with high efficiency [175], but structural variations of amino-termini of different isoforms introduce a difference in the ability of calmodulin to activate enzymes [176]. Levels of PDE1 proteins are modulated on transcriptional and posttranscriptional levels, whereas their activity varies depending on cAMP and Ca2+-dependent signaling [172]. PDE1 enzyme activity is important for the regulation of vascular cGMP levels and reactivity. Ca2+/CaM-dependent PDE activity was reportedly responsible for the hydrolysis of cGMP in rabbit aorta and inhibited norepinephrine-induced contractions [177]. PDE1A1 and PDE1B2 variants were identified in aortic vascular smooth muscle cells of several species including humans [178]. Selective expression of PDE1C in synthetic/activated human VSMC in contrast with absence of expression in contractile/quiescent VSMC suggested that induction of PDE1C expression could represent a useful marker of the phenotypic switch between these cells [179]. PDE1C has also been demonstrated to take part in olfactory fatigue [179] and insulin secretion [180]. Two PDE1 gene products (1A and 1C) are expressed in cardiac tissues from several species, mostly in a non-myocyte fraction instead of in cardiomyocytes [181]. Presently, data available on PDE1 expression in cardiomyocytes are incomplete and no changes in PDE1 expression in heart tissue have been reported accompanying cardiovascular stresses; however, the hypothesis of PDE1 involvement in modulation of cGMP levels during cardiomyocyte contractions requires further investigation. Although PDE1 protein and mRNA have not been described in vascular endothelial cells, PDE1 activity has been detected in lysates from bovine and human aortic vascular endothelial cells (VEC) [182], suggesting that some isoforms of PDE1 should be expressed in VEC. 7.2.4.1.2 PDE2 cGMP-stimulated PDE2 was one of the first cyclic nucleotide phosphodiesterase species characterized in rat liver extracts in the early 1970s. PDE2-dependent hydrolysis of cAMP and cGMP is stimulated up to tenfold with the submicromolar concentrations of cGMP by binding to allosteric regulatory sites known as GAF domains which are located in the N-terminal part of the enzyme [183]. The catalytic domain resides in the carboxy-terminal part of the enzyme similar to all other PDEs. A single PDE2 gene encodes three different splice variants (PDE2A1, A2, A3) expressed in different tissues. Cyclic GMP signaling through activation of PDE2 was described in adrenal granulose cells where it is involved in olfactory signal transduction pathway by inhibiting cAMP stimulated secretion of aldosterone [184]. The PDE2-A2 variant was expressed in the cardiomyocyte fraction of cardiac tissues in rat and human. PDE2 A2 activity inhibited accumulation of cAMP in

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cardiac myocytes, thereby decreasing L-type Ca2+ currents and contractile force of the heart; however, the magnitude of this effect was species-specific [190]. PDE2 expression was detected in human, bovine, and porcine vascular endothelial cells [185]. PDE2 plays a role in the increase of blood platelets aggregation by cAMP hydrolysis [186]. 7.2.4.1.3 PDE3 PDE3 enzymes have similar high affinity for cAMP and cGMP but the Vmax for cAMP is up to 10 times higher for cAMP than for cGMP. PDE3 enzymes do not contain allosteric GAF domains, such as PDE2 and PDE5, and therefore cGMP inhibits PDE3 activity through direct competition with cAMP for catalytic sites. For this reason, the PDE3 family was formerly referred to as cGMP-inhibited cAMP PDEs. The PDE3 family contains two genes, PDE3A and PDE3B, different in their tissue-specific expression. PDE3 enzymes contain two membrane association regions (NHR1 and 2) at the N-terminus. Differential start codon usage in PDE3A produces three splice variants (A1–3), predicted to have different numbers of NHR sites [187]. Subcelllular partitioning is different for all PDE3A variants. PDE3A1 is entirely a particulate protein, PDE3A2 is both soluble and particulate, and PDE3A3 is cytosolic [187]. PDE3A has been found in blood vessels, heart, megakaryocytes, and oocytes, whereas PDE3B is abundant in adipocytes, hepatocytes, brain, renal epithelium, and spermatocytes [188]. In rat and human vascular smooth muscle, both PDE3A and 3B are expressed, but have different sub-cellular localization. PDE3 is thought to mediate mostly cAMP-regulated processes, which include cardiac contractility, platelet aggregation, smooth muscle relaxation, and hormonal regulation [189]. Direct pharmacological inhibition of PDE3 activity increases L-type Ca2+ currents in isolated cardiomyocytes and contributes to the positive ionotropic effects of these inhibitors [190]. PDE3 inhibitors also relax isolated arterial and venous tissue, dilate blood vessels in vivo, inhibit VSMC proliferation in vitro and limit accumulation of neointima VSMC in arteries after vascular damage [172]. Less is known about PDE3 involvement in cGMP signaling; however, some studies in human atrial myocytes, renal vasculature and platelets have suggested that partial effects of NO-donors were due to the elevation of cGMP levels, which inhibited cAMP-hydrolyzing PDE3 activity [172]. 7.2.4.1.4 PDE5 PDE5 is a specific cGMP-hydrolyzing enzyme. A single PDE5 gene encodes three different isoforms of PDE5 (A1, A2, A3) which are dissimilar at their Nterminal ends [191]. Presently, the information about the functional significance of the differential tissue targeting of PDE5 isoforms is incomplete. PDE5 is a homodimer with each subunit containing one catalytic site, two allosteric GAF domains, and one PKA/PKG phosphorylation site [192]. All of the components required for PDE5 catalytic activity are contained within a single monomeric catalytic domain. The activity of the PDE5 enzyme is stimulated by


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PKG and PKA phosphorylation and is facilitated by cGMP binding to GAF domains. Occupation of the allosteric cGMP-binding sites is required for specific phosphorylation of Ser-92 by PKG or PKA. It is believed that cGMP binding at the catalytic site enhances cGMP binding at allosteric sites. This binding promotes PDE5 phosphorylation and, thus, an even further increase in the catalytic activity of the enzyme. This pathway was not directly proven in lysates, but existing experimental results in vivo are consistent with this model [192]. This process represents a negative feedback regulation of rising cGMP levels in cells, which makes PDE5 an attractive therapeutic target for maintenance of elevated intracellular cGMP levels. Several PDE5 inhibitors were developed and are currently used clinically. Sildenafil, the active component of the drug Viagra, is presently widely used in treatment of male erectile dysfunction. PDE5A1 and A2 are expressed in rat, bovine and human contractile/quiescent VSMC where it was recognized to be a valid therapeutic target not only in erectile dysfunction, but also in the treatment of pulmonary hypertension, a disorder with very poor prognosis [193]. PDE5 is an important regulator of NO/cGMP signaling in platelets. Typically, purified sGC enzyme is capable of generating cGMP as long as NO is present; however, measurements of cGMP accumulation in platelets under a continuous flux of NO generated by NO-donors resulted in a sharp increase of cGMP followed by a rapid decline of intracellular cGMP. This decline was not the result of decreased catalytic properties of sGC, but correlated with a change in phosphorylation and activation of PDE5 [194, 195]. This NO-induced activation and phosphorylation of PDE5 is a probable mechanism for a long-lasting negative feedback loop, which controls the extent of the cGMP response in human platelets upon exposure to NO. 7.2.4.1.5 PDE6 PDE6 is the primary effector of phototransduction in vertebrate photoreceptors. PDE6 is a heterotetramer in its inactive state, which is activated upon illumination through the transducin system in retinal rods. Hydrolysis of cGMP by the catalytically active heterodimeric PDE6 core promotes closure of cGMP-gated channels in cellular membranes [196]. Despite a well-described role of cGMP signaling in retinal cells, co-expression of PDE6 and sGC was never reported. As discussed previously, only a limited amount of evidence indicates that the NO/cGMP pathway is directly involved in phototransduction. 7.2.4.1.5PDE 9, 10, and 11 Several new PDE gene family members were recently identified [197, 198]. The PDE9 enzyme has a high affinity for cGMP. Four different splice variants of PDE9 have been identified in different tissues, but their functional significance is presently unclear [199]. PDE10 hydrolyzes cAMP with much higher efficiency than cGMP. Subsequently, cGMP hydrolysis by PDE10 is potently inhibited by cAMP in vitro, suggesting that cGMP-directed PDE10 activity can be regulated

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by cAMP concentrations in vivo [200, 201]. PDE10 contains two amino-terminal GAF domains, but their regulatory function has not yet been demonstrated. A number of tissue-specific splice variants of PDE11A have been identified, although in most cases their enzyme activities in the cell has not been confirmed [202-204] and their regulation by cyclic nucleotides is undetermined [204]. PDE11A may contain a unique structural feature. Truncated GAF domains in Nterminus were identified in several splice variants. Genomic studies have demonstrated the existence of separate promoters for two different splice variants that contain incomplete GAF domains [203]. The functional role of these incomplete GAF domains remains to be determined.

7.2.5

CGMP-DEPENDENT THE

PROTEIN KINASES—DIVERGENCE NO/CGMP PATHWAY

OF

cGMP-dependent protein kinases (PKG or cGK), homodimeric serine/threonine kinases, are considered the main effector molecule in the NO/cGMP pathway. Three different isoforms of mammalian PKG, which are termed PKGIα, PKGIβ, and PKGII [205], exist. PKGIα and PKGIβ are products of alternative splicing and differ only in approximately the first 100 amino acids [206, 207]. PKG II is a separate gene product with no similarity in the amino terminus to that of PKGI isoforms [208]. PKGI is a 75kDa cytosolic protein, whereas 85kDa PKGII is membrane-bound [209–211]. The PKG subunit is comprised of several domains present in all three isoforms. The carboxyl-terminal region of each polypeptide chain is conserved between both PKGI and PKGII and contains the catalytic domain that includes the ATP and protein substrate-binding sites. Immediately amino-terminal to the catalytic domain are two tandem allosteric cGMP-binding sites. The binding of cGMP causes a conformational change that is associated with activation of the kinase [212, 213]. Differences in the first 100 N-terminal residues affect the response of the enzyme to cGMP. The EC50 for cGMP is 10 times higher for PKGIβ than for PKGIα. An autoinhibitory/autophosphorylation domain, with autophosphorylation sites located within and near this domain [214–216], is located immediately amino-terminal to the cGMP-binding sites. The autoinhibitory domain of each of the PKGs contains a pseudosubstrate sequence that interacts with the catalytic site to block substrate access and thus maintains the kinase in an inactive state [217]. Autoinhibition is relieved by cGMP binding and autophosphorylation in PKGI [215, 217, 218] and cGMP binding alone in PKG II [212, 219]. Both PKGIα and PKGIβ undergo autophosphorylation, which increases the basal activity of the enzyme and the affinity to cyclic nucleotides [215]. The extreme amino terminus of the PKG protein contains the dimerization domain. Proteolytic cleavage of this domain produces a fully functional but monomeric PKG [220, 221]. Recombinant PKG lacking the dimerization and autoinhibitory domain (PKGcat), exhibits increased basal activity [222]. PKGIα and PKGIβ are found in platelets, smooth muscle, fibroblasts, leukocytes, glomerular mesangial cells, cardiomyocytes, endothelial, and neuronal


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cells [211]. Although their expression is, in many cases, very similar, some differences occur in their distribution. For example, the neurons express either PKGIα or PKGIβ isoforms, whereas platelets contain predominantly PKGIβ. The PKGII isoform is found in renal cells, lung, intestinal mucosa, pancreas, chondrocytes, and neurons [211]. Recruitment of the PKG enzymes by the NO/cGMP pathway provides a wide diversity in protein targets and cellular processes, which are affected by the activation of the pathway. It is at the step of PKG activation that the divergence of NO-dependent effects occurs.

7.3 NO-CGMP SIGNALING IN SELECTED VARIOUS CELLULAR PROCESSES 7.3.1 NO/CGMP SIGNALING

AND

VASCULAR RELAXATION

Because of the appreciation of the paracrine regulation of vascular smooth muscle cell (SMC) relaxation by EDRF, significant progress has been made in defining the cellular processes underlying the relaxation of smooth muscles under the influence of NO derived from endothelium or therapeutic NO-donors such as nitroglycerine. It is generally accepted that NO-dependent activation of sGC and the associated increase in intracellular cGMP is the principal event initiating SMC relaxation. Different agents, such as endothelin [223], acetylcholine [224], insulin [225, 226], estrogen [227, 228], and corticotrophin-releasing hormone [229], stimulate the endothelial cells to produce NO, which activates the cGMP production in nearby SMC. It is generally accepted that the subsequent PKG activation affects several processes, which results in relaxation of SMC. This chain of events is largely confirmed by studies of transgenic animals. Mice lacking the NOSIII gene, although viable, display a 30% increased mean arterial blood pressure [230]. PKGI−/− mice are viable, but 50% die before they reach 6 weeks of age [210] and also experience systemic hypertension, impaired calcium regulation, and vasorelaxation [210, 231, 232]. To date, reduction of cytosolic free Ca2+, Ca2+ desensitization of the contractile system and regulation of thin filament function are recognized as major processes affected by NO/cGMP pathway during SMC relaxation. 7.3.1.1 Reduction of Intracellular Ca2+ It is widely accepted that a decrease in the concentration of myoplasmic Ca2+ ([Ca2+]I) is a prerequisite for the relaxation of smooth muscle cells. Extrusion of Ca2+ across the plasma membrane was one of the earliest mechanisms of NO/cGMP-dependent relaxation of SMC [233]. NO-donors and cGMP-analogs significantly increase Na+-dependent Ca2+-efflux through activation of the Ca2+/ATPase pump [233]. The exact mechanism for this process is not finalized, but several possibilities have been postulated. PKG phosphorylates and activates phosphatidyl inositol kinase (PI-kinase) [234], which generates phosphatidyl

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inositol-4 phosphate capable of activating the Ca2+/ATPase pump. A 240-kDa protein associated with the purified Ca2+/ATPase pump was suggested to be necessary for pump activation after phosphorylation by PKG [235]. The Ca2+/ATPase pump is also responsible for the uptake of Ca2+ into the sarcoplasmic reticulum (SR), thus decreasing [Ca2+]I. This function is regulated by phospholamban, a demonstrated target of PKG [236]. Phosphorylation of phospholamban, which in SMCs is co-localized with PKG, increases Ca2+/ATPase pump activity. NO-donors inhibit voltage-gated Ca2+-channels [237–239], resulting in a decreased influx of Ca2+. Decreased availability of these L-type Ca2+ channels is mediated through a cGMP- [237, 238] and PKG-dependent mechanism [238], and direct phosphorylation of the channel by PKG has been suggested. Another proposed mechanism of NO/cGMP/PKG inhibition of L-type Ca2+ channels suggests an important role in this process for Ca2+-activated K+-channels (KCa). The NO-cGMP-PKG pathway increases the activity of KCa channels in vascular and tracheal SMCs by direct phosphorylation of the channel or of its regulator [240–243]. This increase of K+ efflux results in hyperpolarization of the plasma membrane leading to inhibition of the voltage-gated Ca2+ channel and Ca2+ influx. The release of Ca2+ from intracellular storage compartments, such as SR, also appears to be affected by PKG in SMCs. The inositol1,4,5 triphosphate (IP3) receptor located in the SR is a well demonstrated substrate of PKG [244, 245]. PKG phosphorylation of the IP3 receptor at serine 1755 reduces the channel activity of the IP3 receptor, thereby reducing the [Ca2+]I. Targeting of the PKG to the SR is facilitated by the IRAG protein (IP3 receptor-associated PKG substrate) [246], which is crucial for PKG-mediated inhibition of the IP3 receptor. Synthesis of IP3, the activator of IP3 receptor, also appears to be affected by the NO/cGMP pathway. It has been proposed that the activity of phospholipase C (PLC), which is responsible for the synthesis of IP3, is inhibited by cGMP either through a direct mechanism or through interference with the G protein-coupled receptor stimulation of PLC [247]. As mentioned previously, PKGI-deficient mice have an impaired regulation of intracellular Ca2+. Transfection of the PKGI−/− knockout, with PKGIα, but not PKGIβ, restores calcium regulation in smooth muscles cells [248]. 7.3.1.2 Ca2+ Desensitization of the Contractile System Contractions of smooth muscle cells occur after a certain threshold for [Ca2+]I is achieved. cGMP decreases the sensitivity of the contractile system to Ca2+ rendering it inactive, even at [Ca2+]I exceeding threshold values [249]. The balance between the activity of the myosin light chain kinase (MLCK) and myosin light chain phosphatase (MLCP) regulates the phosphorylation of myosin light chain at serine 19. This phosphorylation is necessary for the actin activation of myosin ATPase and subsequent cross-bridge cycling. The cGMP-PKG pathway stimulates the activity of the MLCP [250, 251]. PKG is targeted to the myosinbinding subunit (MBS) of the MLCP through a leucine zipper interaction, leading to the phosphorylation of the MBS and concomitant activation of the MLCP’s


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catalytic subunit [252]. PKG-dependent phosphorylation of telokin, a myosin binding protein, also enhanced MLCP activity [253]. In addition, PKG-dependent phosphorylation of MLCK decreases the affinity of this enzyme for calmodulin and MLCK activity [254]. The resulting net decrease of MLC phosphorylation reduces the sensitivity of the contractile system to Ca2+ and contributes to the relaxation of SMC. Phosphorylation of MLC is also regulated by NO/cGMP pathway through small GTPase RhoA. PKG-dependent phosphorylation of RhoA results in decreased activity of Rho-kinase [255], which normally down-regulates MLCP activity. SNP or 8-(2-chlorophenylthio)-cGMP also induces an increase in RhoA mRNA and protein expression, which was inhibited by PKG inhibitor (Rp)-8bromo-β-phenyl-1,N2-ethenoguanosine 3′:5′-phosphorothioate [256]. These data suggest that inhibition of RhoA-induced Ca2+ sensitization and actin cytoskeleton organization contribute to the vasodilator action of NO. 7.3.1.3 Regulation of Thin Filament Function Proteins that bind to thin filaments are logical targets for regulation of SMC contraction. Vasodilatory-stimulated phosphoprotein (VASP) was originally identified and characterized in platelets [257] and later indicated in all studied cells as a protein binding to actin filament and stress fibers. PKG phosphorylates VASP and decreases the number of focal adhesions in various cells [258–260], including vascular SM endothelial cells [259]; however, studies on VASP-deficient mice demonstrated that VASP is dispensable for vascular relaxation [261]. A different actin-associated protein, heat shock protein 20 (HSP20), may be an important modulator of SMC relaxation. Administration of cyclic nucleotides increased phosphorylation of HSP20 [262] at serine 16 by both PKA and PKG [263] and affected its association with actin filaments [264]. Actin-associated protein alpha-actinin appears to be the interacting partner for HSP20 [265]. Interestingly, cGMP does not affect HSP20 phosphorylation in umbilical SMC [266, 267], which is refractory to cGMP-dependent relaxation, supporting the role of HSP20 in cGMP-induced relaxation of SMC. Further studies are needed to identify the mechanism of HSP20-dependent modulation of SMC relaxation.

7.3.2 NO/CGMP SIGNALING

AND

PLATELET AGGREGATION

Under normal conditions, several endothelial products inhibit platelet adhesion and aggregation. Prostacyclin and 13-hydroxyoctadecanoic acid, which are products of cyclooxygenase and lypooxygenase pathways, as well as ectonucleotidase ADP hydrolase and NO, are regarded as the main endothelium-derived antithrombotic factors. Exposure of platelets to NO decreases or prevents their adhesion and aggregation [268, 269] and could even induce disaggregation of previously aggregated platelets [270]. Ample evidence demonstrates that this NO-dependent inhibition of platelet function relies on the cGMP pathway. 1H(1,2,4)oxadiazolo (4,3)-quinoxalin-1-one (ODQ), an inhibitor of sGC, blocks

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NO-dependent effect on platelets [271], whereas NO-independent activators of sGC mimic the NO-effects [384,385]. Administration of M+B22984, a PDE inhibitor, potentiated the anti-platelet effect of L-arginine [272]. To date, it is clear that platelets contain all the major players of the NO/cGMP pathway: cGMPproducing sGC [273], cGMP-dependent PKG [211], as well as several phosphodiesterases [274], including cGMP-regulated PDE5 and PDE2. Transgenic PKGIdeficient mice exhibit increased platelet activation during ischemia/reperfusion [232], demonstrating the key role of PKGI in the inhibition of platelet aggregation. Although the important role for the NO-cGMP pathway in the function of platelets is well documented, the mechanism of platelet inhibition is very complex and continues to be unraveled. 7.3.2.1 Ca2+ Homeostasis sGC-mediated increase in intracellular cGMP in platelets results in the reduction of intracellular Ca2+ through mechanisms similar to those in SMCs. The resulting decrease of intracellular Ca2+ impairs the activation of Ca2+-dependent PKC, Ca2+calmodulin dependent MLCK, and cytosolic phospholipase A2 among many other Ca+-dependent processes. Inhibition of Ca2+ release from intracellular stores [275, 276], increased Ca2+ extrusion [277] and decreased Ca2+ influx [278], as well as acceleration of Ca2+-ATPase-dependent refilling of Ca2+ stores [279], are some of the processes affected by the NO/cGMP pathway. Most platelet agonists activate IP3-mediated Ca2+ release from intracellular storage compartments. cGMP inhibits the function of the IP3-receptor via IP3R phosphorylation [280]. Although the PKG-dependent inhibition of IP3-receptors in SMC is dependent on the IRAG protein, it is not clear whether a similar mechanism is taking place in platelets. Protein kinase C (PKC) may also be inhibited indirectly through an undetermined mechanism of PLC inhibition. Both cAMP and cGMP inhibit phospholipase C through an undefined mechanism, possibly through the decrease of phosphoinositide turnover [281]. This results in the decreased synthesis of DAG, which in addition to Ca2+, is required for PKC activation. 7.3.2.2 Surface Receptors NO decreases the surface expression of P-selectin [282], a mediator of platelet aggregation needed at the initial attachment of platelets to endothelium. This process is at least partially dependent on PKG-dependent inhibition of PKC [282]. NO decreases, in a cGMP-dependent manner, surface expression of the CD63 glycoprotein and the fibrinogen receptor GPIIb/IIIa [283]. Fibrinogen, which forms cross-bridges between activated platelets, binds less efficiently to platelets due to an NO/cGMP-dependent decrease in the affinity of the GPIIb/IIIa receptor to its ligand [284]. The signaling mechanism mediated through the thromboxane TXA2 receptor, crucial for the recruitment of platelets, is also blocked by NO due to a PKGdependent phosphorylation of the carboxy-terminus of the receptor [285].


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The NO/cGMP/PKG pathway inhibits the platelet ADP receptor P2Y12 [286]; however, the mechanism of this inhibition remains to be elucidated. 7.3.2.3 Cytoskeletal-Associated Proteins Phosphorylation of the VASP protein correlates very well with platelet inhibition caused by cGMP- and cAMP-elevating agents [257]. In VASP-deficient mice cGMP- and cAMP-mediated inhibition of platelets was significantly diminished [261], through a mechanism that does not affect cytosolic Ca2+ concentration. VASP enhances actin polymerization and actin filament bundling [287], whereas phosphorylated VASP suppresses its association with actin exerting negative effect on actin dynamics [136]. VASP is not the only cytoskeleton-associated protein phosphorylated by PKG in platelets. Heat shock protein 27 [288] and LASP (LIM and SH3 domain protein) [289] have been identified as PKG substrates. PKG-dependent phosphorylation of these proteins may regulate actin microfilament and cytoskeletal reorganization in platelets. Interestingly, HSP27 is also a substrate for the MAPKAPK2, which is regulated in platelets by p38 MAPK [290]. cGMP-elevating agents inhibit agonist-induced activation of p38 MAPK [291], potentially decreasing the level of HSP27 phosphorylation. These opposite effects of cGMP-mediated phosphorylation of HSP27 probably reflect the presence of a finely tuned back loop regulation, which remains to be determined. Inhibition of collagen-stimulated platelet aggregation by NO-donors correlates in a dose-dependent manner with phosphorylation of the small GTP-ase rap1b [292] through PKG. Serine 179 residue of the protein has been identified as the position of PKG-dependent phosphorylation [293]. The role of rap1b in platelets is not completely understood; however, in leukocytes activation of rap1b induces cell adhesion, whereas its inactivation inhibits adhesion. Similar function could be envisioned for rap1b in platelets. In summary, the NO/cGMP pathway inhibits a broad range of platelet function, including adhesion, aggregation, degranulation, and disaggregation. High potency of anti-platelet function of NO is the result of cGMP/PKG interference with the platelet activation-signaling cascade at multiple sites.


AND

NEUROTRANSMISSION

NO was first characterized in the central nervous system as an intercellular messenger that increases cGMP levels following the activation of glutamate receptors [294]. Currently, ample evidence indicates that increased cGMP levels mediate a large number of NO actions in neuronal tissues. Although all three isoforms of NOS are found in the brain, it is believed that NOSI is the major isoform responsible for the neuronal signaling, neurotoxicity, synaptic plasticity, learning, and pain. Induction of the NOSII enzyme in glial cells occurs usually as an unspecific immune response and is associated with pathological conditions [295]. NOSIII function in the CNS is believed to be mainly involved in the

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regulation of vascular function, although it is also found in some neurons [296] and glial cells [297]. 7.3.3.1 Synaptic Plasticity An important role of NO in the modulation of synaptic plasticity was suggested following in vitro studies in which inhibition of NO prevented the development of long-term potentiation (LTP) [298–300]. sGC appears to be the main mediator of NO-dependent induction of LTP [298, 301]. Interestingly, LTP is only slightly reduced in NOSI or NOSIII null mice [302], though a substantially decreased LTP is present in animals deficient in both the NOSI and NOSIII genes [303]. The role of PKG in LTP as a presynaptic effector was proposed when PKG activators facilitated LTP in response to tetanic stimuli, and PKG inhibitors were able to block the effect [304]. High concentrations of PKG are found in many neuronal cells and areas of the brain [305]. Pyramidal neurons of the hippocampus contain high levels of PKGI, whereas neuronal cells in other hippocampal regions highly express PKGII [306]. Though PKGI- and II-deficient mice exhibit normal hippocampal histology and no significant difference in LTP, the inhibition of NOS by L-nitro-arginine attenuated LTP; however, cGMP analogs had no significant effect on LTP indicating that NO may affect LTP independently of sGC/cGMP/PKG [306]. It is likely that both cGMP/PKG-dependent and independent mechanisms exist. ADP-ribosylation and activation of Ca2+/CaM-dependent kinases are regarded as alternative mechanisms of NO-dependent regulation LTP. The effects of NO/cGMP/PKG on long-term depression (LTD) are conflicting. In Purkinje cells, evidence exists both for and against NO/cGMP-dependent LTD [307–310]. Purkinje cell-specific ablation of PKGI caused impairment in LTD and cerebellar learning [311]. LTD decreased when neuronal cells were treated with sGC antagonist ODQ [312], and increased in response to cGMP agonists [309] and the PDE5 inhibitor zaprinast [313]. Yet, these same reagents had no effect on both PKGI- and II-deficient mice [306], indicating the involvement of PKG. 7.3.3.2 Perception of Pain NO has been implicated at various levels of the nociceptive neural pathway; however, the role of NO changes according to the type of pain stimuli. Nociceptors or sensory neurons were demonstrated to exhibit NO/cGMP-dependent spinal sensitization, which was blocked by NOS inhibitors and sGC antagonists [314]. In addition, PKG inhibitors block glutamate and aspartate release from spinal nocioceptive neurons upon excitatory stimulation [315]. Inhibition of NO has anti-nocicpetive effects in models of thermal hyperalgesia or visceral pain [316, 317]; however, inhibition of NO synthesis exacerbates pain in models of mechanical hyperalgesia [318]. The exact mechanisms of pro- or anti-nocicpetive effects of the NO/cGMP pathway are not entirely clear.


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7.3.3.3 NO in Neurotransmission The role of NO as an inhibiting neurotransmitter of non-adrenergic, non-cholinergic (NANC) neurons has long been recognized [319]. A special term of “nitrergic nerves” has been adopted for nerves with functions that depend on release of NO [320]. Several lines of evidence support the role of the NO/cGMP pathway in neuronal firing and neurotransmitter release. PKG phosphorylated the synaptic vesicle protein rabphilin3A [321], indicating a possible role for PKG in exocytosis and vesicle transport. The NO/cGMP modulates the spontaneous firing of Purkinje neurons in cerebellar slices in the absence of excitatory stimuli. Activation of the NO-cGMP signaling pathway sustained an increase in spontaneous firing rate, which was blocked by inhibitors of sGC and PKG [322]. It has been demonstrated that NO causes synaptic suppression in the neuromuscular junction [323] via PKG-dependent phosphorylation of nicotinic acetylcholine receptors or dystrophin, respectively [324, 325], which are involved in targeting skeletal nNOS to the sarcolemma. In addition, NO acts presynaptically, in a cGMP/PKG-dependent fashion, to elicit a synaptic potentiation, facilitating neurotransmitter release [326, 327]. A wide network of peripheral nitrergic nerves innervates various organs throughout the body, with smooth muscles in gastrointestinal, vascular, respiratory, and urogenital systems being the most widely recognized target of their action. Several examples of the functional role of nitrergic nerves will be discussed below.


AND

KIDNEY FUNCTION

NO plays an important role in various physiological processes in the kidney, including salt and fluid reabsorption [328], tubuloglomerular feedback [329, 330], renin secretion [331], and renal hemodynamics [332]. All three isoforms of NOS, sGC, and both PKGI and PKGII are found in various part of the kidney (macula densa cells, juxtaglomerular apparatus, proximal tubes, collecting ducts, ascending and descending arterioles). In vivo experiments performed in animals demonstrated that infusion of substances like acetylcholine, which causes NO release, into the renal artery results in increased urinary volume and sodium excretion [333, 334]. Conversely, decreased water and sodium excretion was observed when inhibitors of endogenous NO production were infused into the kidney [333, 335, 336]. Administration of sGC and PKG inhibitors augmented tubuloglomerular feedback, which can be reversed by administration of cGMP analogs [329]. Renin is produced predominantly in renal juxtaglomerular granular (JGG) cells. JGG cells are encircled by cells with a high capacity for NO formation; however, analysis of NO effect on renin secretion is full of contradictory observations, which is echoed by similar confusing observation of the effects of cGMP analogs [331]. It has been suggested that inhibition of renin secretion is related to the direct effect of cGMP through activation of PKG [331]. Activation of rennin secretion by the NO/cGMP pathway, on the other hand, is related to activation

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of PKA, which occurs after cGMP-dependent inhibition of PDE3 and subsequent increase in cAMP [331]. Interestingly, this conclusion is supported by PKGII−/− mice, which exhibit decreased cGMP-mediated inhibition of renin expression, and decreased secretion in renal juxtaglomerular cells [337]. In addition, these mice exhibit hypertension associated with sodium and water retention. Although the pool of evidence demonstrating that NO/cGMP-dependent signaling regulates many aspects of kidney physiology, only a limited understanding and consensus exists about the molecular mechanism involved in this process.

7.3.5 NO/CGMP

IN

REPRODUCTION

NO mediates erectile function. NO, released from nitrergic nerves innervating the corpus cavernosum, activates sGC and increases cytosolic cGMP. This increase in cGMP modulates [Ca2+]i and regulates smooth muscle contractility and erectile function. The PDE5 specific inhibitor sildenafil increases cGMP, potentiating corpus cavernosum smooth muscle relaxation and penile erection [338]. Erectile responses were also elicited by intracavernous injection of cGMP and cAMP analogs, though the response was significantly more potent with cGMP analogs [339]. It should be noted that administration of PDE5 inhibitors does not aid erection in patients with complete loss of sacral nerve activity or absence of sexual arousal [340] emphasizing the crucial role of functional nitrergic nerves and pre-activation of NO/cGMP signaling in this process. Moreover, PKGIdeficient mice have a very low ability to reproduce and their corpora cavernosa fails to relax upon activation of the NO/cGMP signaling cascade [341]. In females, NO plays a role in keeping the uterus from contracting before term [342, 343], by increasing cellular cGMP levels and thus inhibiting myometrial contractility during pregnancy. A decrease in responsiveness to NO would appear to be involved in initiation of labor [344].

7.3.6 NO/CGMP

IN

BONE HOMEOSTASIS

NO generated from NOSII has been implicated as a potential mediator of cartilage pathophysiology in arthritis [345], which disassembles complexes formed at focal adhesion sites. PKG also was found to mediate the disassembly of focal adhesions triggered by thrombospondin or tenascin [259]. NO/cGMP was further implicated in the down-regulation of osteoclastic activity and bone resorption [346]. Several mechanisms may account for this effect. One mechanism is that by which cGMP inhibits the differentiation stage of osteoclast formation in bone marrow cultures [347]. Another mechanism involves PKG-dependent inhibition of ATP-dependent acid transport in reconstituted osteoclast membrane vesicles [348]. These studies suggest an inhibitory role of cGMP in bone resorption. NO also supports bone formation by inhibiting proliferation and by stimulating differentiation of osteoclastic cells [349]. Therefore, the NO/cGMP pathway appears to control osteoclastic, bone degrading, and bone producing activities.



AND

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GENE REGULATION

Over the last decade, an increasing amount of experimental evidence indicates that the NO/cGMP pathway regulates multiple genes through a variety of mechanisms. For example, expression of inflammatory mediators, such as TNF-α and COX-2 [350–352], are stimulated by NO-donors and cGMP analogs. On the contrary, NO donors and cGMP analogs decreased mRNA levels of plasminogen activator inhibitor 1 (PAI-1) [353]. It has been suggested that PKGI has a nuclear location sequence (NLS) for docking at the nuclear complex [354]. This NLS resides in the ATP binding domain of PKG and initiates a conformational change to regulatory domain [213], allowing it to translocate to the nucleus after cGMP binding [354]. Although in many cases, the exact mechanism of this regulation is not determined, evidence of PKG-dependent phosphorylation of various transcription factors exists. CREB, a transcription factor crucial to cell proliferation and survival is directly phosphorylated by PKG in a cGMP-dependent fashion [355]. ATF-1, a member of the same family of transcription factors as CREB, is also phosphorylated by cGMP-activated PKG and mediates transcriptional activation of RhoA [256, 356]. NO donors and cGMP elevating agents increase mRNA expression of c-fos and JunB, which collectively make up the AP-1 transcription factor [357]. PKG regulates promoters of c-fos, JunB, and Ca2+/calmodulin kinase [357–359] through cAMP response elements (CRE), AP-1 binding sites, and SRE serum response elements [360]. TFII-I, an important transcriptional regulator that interacts with a myriad of transcription factors, is directly phosphorylated by PKGIβ [361]. TFII-I, assisted by PKG phosphorylation, is able to transactivate the fos promoter, among others. Egr-1, a factor involved in cell survival and apoptosis, demonstrated increased mRNA and protein expression in response to NO donors and cGMP analogs [362], possibly through activation of PKG. The function of several transcription factors is also regulated by PKG through indirect mechanisms. NFκB activity can be either increased or decreased in response to NO, though regulation via the cGMP/PKG pathway is usually stimulatory. PKG directly phosphorylates IκBα, the inhibitory subunit of the NFκB complex [350]. Phosphorylation of IκB-α results in its degradation, followed by release and subsequent activation of NFκB. Evidence also indicates that PKG can directly phosphorylate the p60 and p65 subunits of NFκB [363], activating the critical stimulator of inflammatory mediators. Calcineurin, a calcium-dependent phosphatase, is inhibited by PKG phosphorylation, preventing nuclear translocation of NFAT [364]. PKG inhibits serum response factor (SRF)-dependent transcription by interfering with RhoA signaling in cardiomyocytes and VSMC [356]. The ultimate effect of this inhibition will depend on the differentiation state of the cells.


AND

APOPTOSIS

The outcome effect of the NO/cGMP pathway on cell death largely depends on the concentration of produced NO, cell type, and mechanism of apoptosis

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induction. NO production in SMCs, cardiomyocytes, and endothelial cells exhibits a PKG-dependent increase in apoptosis, due to the down-regulation of the Bcl-2 homolog Mcl-1 [365]. Alternatively, NO can inhibit caspase signaling by direct nitrosation of key cysteine residues [366, 367]. In addition to direct inhibition of caspases NO affects the apoptotic process through the NO/cGMP pathway as well. Apoptosis of serum deprived neuroblastomal cell cultures was prevented through an NO/sGC/PKG-dependent mechanism [368, 369]. An NOmediated hepatoprotective effect also appears to depend, at least partially, on cGMP synthesis. In an in vivo model of fulminant hepatic failure of mice, in which ActD/LPS induced hepatocyte cell death, apoptosis was inhibited by administration of NO donors or cGMP analogs. These protective effects of NO were blocked by the sGC inhibitor LY83583 [370]. NO-dependent prevention of apoptosis in rat cultured hepatocytes treated with TNFα/ActD was also in part dependent on the sGC activity because it was blunted by sGC inhibitor ODQ [371].

7.3.9 NO/CGMP

AND INTESTINAL

MOTILITY

NANC neurons that reside in the intestinal wall of the gastrointestinal tract contain NOS and produce NO [372, 373]. NO from NANC produces an increase in cGMP, causing smooth muscle relaxation. PKGI is highly expressed in the smooth muscle of the intestine [374]. PKGI deficient mice have distention of their gastrointestinal tract, pyloric stenosis, and muscle hypertrophy. Loss of peristalsis, retarded passage of intestinal content, and progressive stenosis are consequences of PKGI deficiency [210]. NOSI-deficient mice had a similar phenotype [375], indicating that PKGI activation occurs through NO-sGC signaling.

7.3.10 SELF-REGULATORY INTERACTIONS CASCADE

IN

NO/CGMP

The diversity of the NO/cGMP-dependent processes implies that multiple internal control mechanisms should be imposed at multiple sites in the signaling cascade. In addition to degradation of cGMP due to activation of cGMP-dependent phosphodiesterases, several other possible mechanisms of self-regulation were described. The NOS enzyme is subject to autoinhibition. Both exogenously added NO [376] and endogenously produced NO by NOS [377] significantly inhibits the output of produced NO. The most probable mechanism of this inhibition is the binding of NO to the catalytically crucial heme moiety of the enzyme [378], displaying a classical mechanism of enzyme inhibition by the reaction product. In vitro studies on purified sGC enzyme [379] also demonstrated some level of inhibition of sGC by reaction products pyrophosphate and cGMP [379]; however, this inhibition was achieved only at concentrations of the products significantly higher than expected under normal physiological conditions. Nevertheless, this type of sGC inhibition may still occur under pathophysiologic conditions (e.g., during acute inflammation or septic shock).


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Prolonged exposure of VSMC and vascular endothelial cells to NO donors, or activation of endogenous NOSII upon stimulation with inflammatory cytokines result in decreased levels of sGC mRNA, protein, and activity [380, 381]. Destabilization of sGC mRNA is believed to be responsible for these effects, although decreased transcription or translation of sGC was not ruled out. Similarly, continuous exposure to NO produced by NOSII or NO-donors down-regulated both PKGI protein and mRNA levels [382]. As described previously, PKG-dependent phosphorylation of PDE5 is a critical step in activation of the PDE5 enzyme [192]. Recently, a connection between the activity of sGC enzyme and PKG activity was reported [383]. Phosphorylation of the β subunit decreased NO-stimulated activity of sGC. This inhibition could be prevented by PKG-dependent activation of protein phosphatase activity, suggesting that tonic activity of PKG or its stimulation may regulate sGC activity through dephosphorylation of the β subunit.

7.4 CLOSING REMARKS Soluble guanylyl cyclase stimulated by NOS-derived NO produced in the same or neighboring cells increases intracellular cGMP levels. Through recruitment of several cGMP-effector molecules, such as CNGs, PDEs, and PKG, a myriad of processes are affected by the activation of the NO/cGMP pathway. Autocrine and paracrine effects of the NO/cGMP pathway have a wide range of physiological and pathophysiological consequences. The large spectrum of NO-induced effects makes it impossible to embrace all these processes in one comprehensive review. We focused only on a small section of this spectrum with the main goal of demonstrating the diversity of processes and complexity of mechanisms associated with NO-dependent synthesis of cGMP. A great number of excellent specific reviews of the role of NO in various systems have been published over the years, including the accompanying sections of this book. The readers might find them helpful in their search for the desired answer.

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of Cell 8 Regulation Signaling by Protein Nitrosylation/ Denitrosylation Joan B. Mannick University of Massachusetts Medical School, Worcester, Massachusetts

CONTENTS 8.1 8.2 8.3 8.4 8.5 8.6 8.7 8.8

Introduction ............................................................................................217 Specificity of Nitrosylation ...................................................................218 Denitrosylation .......................................................................................219 Fas-Induced Apoptosis ..........................................................................219 N-Methyl-D-Aspartate Receptor Signaling ...........................................221 Ryanodine Receptor S-Nitrosylation .....................................................222 Thioredoxin Signaling ...........................................................................224 Conclusion .............................................................................................226 References ...............................................................................................226

8.1 INTRODUCTION Accumulating data suggests that nitric oxide (NO) regulates a diverse array of biological processes via nitrosylation/denitrosylation of proteins (1). Nitrosylation is the binding of a NO group and denitrosylation is the removal of a NO group from a protein or peptide. Proteins have two main nitrosylation targets: reactive metal centers (metal-nitrosylation) and cysteine residues (S-nitrosylation). Nitrosylation of critical cysteines or metals regulates protein function. In addition, nitrosylation is specifically targeted and rapidly reversible, allowing cells to dynamically modify signaling in response to physiologic stimuli; however, unlike more traditional posttranslational modifications, such as phosphorylation, nitrosylation of proteins is, at least in part, non-enzymatically mediated. Instead, complex and incompletely understood intracellular redox chemistry results in the addition or deletion of NO groups from proteins. Thus, nitrosylation directly 217

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translates shifts in the cellular redox environment to changes in protein function. Although the activity of many proteins is modified by nitrosylation in cell free systems, it has been much more difficult to identify proteins that are endogenously nitrosylated in cells. Identification of endogenously nitrosylated proteins is technically challenging because the intracellular levels of nitrosylated proteins are at the limits of detection of currently available assays; however, strong evidence indicates that several cell-signaling pathways are regulated by endogenous nitrosylation. This chapter discusses how specificity and reversibility of nitrosylation reactions are achieved and reviews some of the signaling pathways that are regulated by nitrosylation/denitrosylation intracellularly.

8.2 SPECIFICITY OF NITROSYLATION Despite the fact that virtually all proteins contain cysteine residues, many proteins contain transition metals, and most if not all cells produce NO, only a precisely defined subset of targets is nitrosylated intracellularly. For instance, the skeletal muscle calcium release channel/ryanodine receptor has 50 free cysteine residues per subunit, but only 1 of the 50 thiols is S-nitrosylated under physiologic conditions (2, 3). The specificity of nitrosylation reactions is conferred in part by sub-cellular co-localization of nitrosylation targets with nitric oxide synthase (NOS), the intracellular enzyme that produces NO. An increasing number of S-nitrosylated proteins have been reportedly associated intracellularly with NOS (4–10). Because NO is a relatively reactive molecule, discrete co-localization of NOS with specific protein targets may ensure that NO (or related species) reacts preferentially with NOS-associated proteins. In addition to co-localization of nitrosylation targets with NOS, specificity of S-nitrosylation reactions is also conferred by consensus motifs flanking cysteine residues (11). Two known S-nitrosylation consensus motifs exist: the acid/base motif and the hydrophobic core motif. In the acid/base motif, the target cysteine is located between an acidic and basic amino acid either in the primary or tertiary structure of the protein (11–13). The importance of the acid/base motif for targeting S-nitrosylation reactions has been demonstrated using the protein methionine adenosyltransferase (MAT). MAT is the enzyme that synthesizes S-adenosylmethionine, a methyl donor for many biological methylation reactions. Each subunit of MAT contains 10 free cysteines, but only a single cysteine is targeted for S-nitrosylation, leading to an inhibition of MAT activity (14, 15). Examination of the tertiary structure of MAT reveals that the single S-nitrosylated cysteine is located between an acidic and basic amino acid. When the flanking acidic or basic amino acid is mutated, MAT is not longer S-nitrosylated and becomes unresponsive to NO (15). Hydrophobic cores in proteins provide a second consensus motif for S-nitrosylation reactions. NO and O2 are concentrated within hydrophobic regions leading to an increase in the rate of formation of S-nitrosylating species such as NO2 and N2O3 (16). The single cysteine residue that is targeted for S-nitrosylation in the ryanodine receptor is located within a hydrophobic pocket. Moreover,

Regulation of Cell Signaling by Protein Nitrosylation/ Denitrosylation

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formation of the hydrophobic pocket may be dependent on redox-driven conformational changes in the ryanodine receptor (2, 3).

8.3 DENITROSYLATION Nitrosylation is reversed by denitrosylation. Denitrosylation can be accomplished non-enzymatically due to the redox-sensitive nature of S-NO and metal-NO bonds. A variety of factors including reducing agents, thiols, transition metals, and UV light can break S-NO or metal-NO bonds resulting in protein denitrosylation (11, 17). Therefore, denitrosylation may simply require altering the redox environment of a protein. In addition, denitrosylation may be enzymatically mediated. Specifically, formaldehyde dehydrogenase selectively denitrosylates the S-nitrosylated peptide S-nitrosoglutathione (GSNO) (18). GSNO serves as a reservoir of NO in cells and donates its NO group to proteins in trans-nitrosation reactions. Therefore, mice or yeast with a targeted deletion of formaldehyde dehydrogenase have increased levels of not only GSNO but also S-nitrosylated proteins (18). Formaldehyde dehydrogenase knockout mice accumulate increased levels of S-nitrosylated proteins and have increased tissue damage and decreased survival after endotoxic or bacterial challenge (19). Moreover, the survival of the formaldehyde dehydrogenase knockout mice is significantly increased during endotoxic shock if the accumulation of S-nitrosylated proteins is abrogated by NOS inhibitor treatment. Thus, formaldehyde dehydrogenase may protect against the hazardous accumulation of S-nitrosylated proteins during nitrosative stress (19). In addition, the data indicate that intracellular levels of S-nitrosylated proteins and peptides are regulated not only by their rate of formation (NOS) but also by their rate of degradation (formaldehyde dehydrogenase). In summary, the specificity of nitrosylation is conferred both by co-localization of NOS with nitrosylation targets and by nitrosylation consensus motifs. Reversibility of nitrosylation is achieved by enzymatically and non-enzymatically mediated denitrosylation. The specificity and reversibility of nitrosylation enables cells to precisely and dynamically modify protein function in response to alterations (particularly redox alterations) in their environment. The following sections review some of the pathways that are regulated by endogenous S-nitrosylation/denitrosylation in cells.

8.4 FAS-INDUCED APOPTOSIS Apoptosis is a form of cell death that removes excess or unwanted cells from organisms. Apoptosis must be tightly regulated to avoid the accumulation of unwanted cells (as occurs in cancer) or the removal of desirable cells (as occurs in neurodegeneration). Apoptosis is triggered by exogenous stimuli or by cell surface receptors such as Fas. Fas stimulation leads to the activation of a family of cysteine proteases called caspases (20–22). Caspases are expressed as relatively inactive zymogens that are cleaved to form active tetrameric enzymes. Upstream

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caspases, such as caspase-8 and caspase-9, cleave and activate downstream caspases such as caspase-3. Downstream caspases cleave specific cellular proteins resulting in apoptotic cell death. Mitochondria are key sites of apoptosis regulation in cells. When mitochondria receive an apoptotic stimulus, they release a variety of pro-apoptotic molecules from their intermembrane space into the cytoplasm including cytochrome c. In the cytoplasm, cytochrome c forms a multi-protein complex with an adaptor protein called Apaf-1 and caspase-9 called an apoptosome. The apoptosome also cleaves and activates downstream caspases, amplifying apoptotic cascades (23–27). All caspases contain a critical catalytic site cysteine residue. S-nitrosylation of the catalytic site cysteine inhibits caspase activity in cell free systems (28). To determine if S-nitrosylation of the catalytic cysteine regulates caspase activity in cells, caspase-3 S-nitrosylation was examined in purified form from human lymphocyte lines at various time points before and during Fas-induced apoptosis (29). Before Fas stimulation, a subset of caspase-3 zymogens is S-nitrosylated on their catalytic site cysteine in resting cells. During Fas-induced apoptosis, caspase-3 zymogens are denitrosylated, allowing the catalytic site to function (29). Thus, S-nitrosylation/denitrosylation may serve as an on/off switch for caspase-3 activity during apoptosis. Of interest, it is predominantly the mitochondrial subpopulation of caspase-3 zymogens that is S-nitrosylated in resting human lymphocyte cell lines. Not only caspase-3 but also caspase-9 zymogens are S-nitrosylated predominantly within mitochondria (30) (Figure 8.1). Mitochondrial caspases reside within the mitochondrial intermembrane space and may be important for initiating mitochondrially regulated forms of apoptosis. It is unclear why mitochondrial caspase zymogens are specifically S-nitrosylated in resting cells; however, caspase zymogens autoactivate when they are brought into close proximity (31). It is possible that caspase activity must be inhibited by S-nitrosylation when caspase zymogens are relatively concentrated in the small mitochondrial intermembrane space to prevent aberrant autoactivation and induction of apoptosis. A modified yeast two-hybrid screen has been used to identify proteins that associate with S-nitrosylated but not denitrosylated caspase-3. NOS was found to bind to S-nitrosylated caspase-3 within mitochondria (10). Thus, co-localization of mitochondrial caspase-3 with NOS may maintain caspase-3 in an S-nitrosylated state. The finding that caspase zymogens are S-nitrosylated predominantly within mitochondria raises the possibility that mitochondria are a key site of protein nitrosylation in cells. In support of this hypothesis, another protein residing within the mitochondrial intermembrane space, cytochrome c, is also regulated by nitrosylation. Cytochrome c is nitrosylated on its heme iron in mitochondria during Fas-induced apoptosis and then is rapidly released into the cytoplasm (Figure 8.1). Heme nitrosylation of cytochrome c may enhance its pro-apoptotic activity (32). Thus, Fas-induced apoptosis is regulated both by nitrosylation of caspase zymogens and cytochrome c. Denitrosylation of the catalytic site cysteine


221

A. Mitochondria Cytc

Fas

Fe B.

C9

C3

Mitochondria

S–NO

S–NO

Cytc

FeNO

Cytoplasm

C9

C3 S

S

Cytoplasm

C. Mitochondria

S Apaf Cytc

C9 FeNO

Apoptosis C3 C3S

FIGURE 8.1 Regulation of Fas-induced apoptosis by protein nitrosylation. (A) In resting cells, mitochondrial caspases (C3 and C9) are inhibited by S-nitrosylation of their catalytic site cysteine. (B) After Fas stimulation, mitochondrial caspases are denitrosylated, allowing their catalytic site to function. Concurrently, cytochrome c (Cytc) is nitrosylated on its heme iron. Both mitochondrial caspases and cytochrome c are released into the cytoplasm. (C) In the cytoplasm, nitrosylated cytochrome c forms an apoptosome with Apaf-1 and denitrosylated caspase-9 (C9). The combination of cytochrome c nitrosylation and caspase denitrosylation results in enhanced caspase-3 (C3) cleavage and activation as well as enhanced apoptosis.

of mitochondrial caspase zymogens concurrently with heme nitrosylation of cytochrome c enhances caspase activation and promotes Fas-induced apoptosis (Figure 8.1).

8.5 N-METHYL-D-ASPARTATE RECEPTOR SIGNALING The N-methyl-D-aspartate (NMDA) class of glutamate receptor (NMDAR) regulates development, learning, and memory in the central nervous system (33, 34). The activity of the NMDAR must be tightly regulated because overstimulation of the receptor results in excitotoxic cell death of neurons and may be a contributing factor to a variety of neurodegenerative diseases including amyotrophic lateral sclerosis, Parkinson’s disease, Huntington’s disease and Alzheimer’s disease (35). The neuronal isoform of NOS (nNOS) is anchored to the NMDAR by

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the scaffolding protein PSD-95 (4, 7). Activation of the NMDAR by glutamate leads to an influx of calcium into neurons that activates receptor-associated nNOS (36). Local production of NO by nNOS results in specific S-nitrosylation of 3 proteins within the NMDAR complex (Figure 8.2). One of the 3 S-nitrosylated proteins is the NR2A subunit of the NMDAR (Figure 8.2). S-nitrosylation of cysteine 399 on the NR2A subunit downregulates NMDAR activity (37, 38). A second protein S-nitrosylated by NMDAR-associated nNOS activity is Dexras, a brain-enriched member of the Ras family of small monomeric G proteins (8). Ras family members are guanine nucleotide-binding proteins that cycle between inactive GDP-bound and active GTP-bound states to regulate a diverse array of cellular processes. Dexras and nNOS co-associate within the NMDAR complex by binding to the adaptor protein CAPON (Figure 8.2). Activation of nNOS by NMDAR stimulation leads to selective activation of Dexras (8). The mechanism by which NO activates Dexras is likely to be S-nitrosylation of a critical cysteine residue because NO activates another Ras family member, H-Ras, via S-nitrosylation (39–41). In addition, in vitro S-nitrosylation of Dexras by NO donors is associated with enhanced guanine-nucleotide exchange and increased concentrations of the active GTP-bound form of Dexras (8). Of interest, recent data suggests that the actual chemical process of S-nitrosylation instead of the product of S-nitrosylation interferes with guanine nucleotide substrate binding (42). Specifically, formation of a Ras radical intermediate, such as Ras-S., during the process of S-nitrosylation may reduce the affinity of Ras for its guanine nucleotide substrate, leading to enhanced guanine nucleotide dissociation (43). Finally, calcium-stimulated nNOS activity within the NMDAR complex leads to specific nitrosylation of guanylate cyclase (GC) (Figure 8.2). GC catalyzes the conversion of GTP to cGMP. GC co-associates with nNOS within the NMDAR complex via the scaffolding protein PSD-95 (6). When nNOS is activated after NMDAR stimulation, NO binds to the heme prosthetic group of GC. Heme nitrosylation of GC causes an up to 200-fold increase in activation of the enzyme (44, 45). The subsequent increase in cGMP levels modulates the activity of a variety of downstream proteins such as cGMP-activated protein kinases, cGMPregulated phosphodiesterases, and cGMP-gated ion channels (6). In summary, just as adaptor proteins co-localize kinases with phosphorylation targets and thereby contribute to the specificity and rate of kinase signaling reactions (46), similarly adaptor proteins within the NMDAR complex co-localize nNOS with three nitrosylation targets: NR2A, Dexras, and GC. Activation of nNOS within the receptor complex leads to selective nitrosylation of the receptorassociated proteins and modulation of downstream signaling events.

8.6 RYANODINE RECEPTOR S-NITROSYLATION Ryanodine receptors are a class of calcium release channel in the sarcoplasmic reticulum of muscles. In response to an action potential, ryanodine receptors release calcium from the sarcoplasmic reticulum leading to muscle contraction. The cardiac isoform of the ryanodine receptor (RyR2) co-localizes with nNOS


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A. Physiologic pO2

nNOS C3635-SNO RyR Ca2+ RyR

Muscle Contraction

B. Oxidative Stress

C3635-Sox

Sarcoplasmic reticulum

nNOS RyR Ca2+ RyR

Muscle Contraction

Sarcoplasmic reticulum

FIGURE 8.2 Regulation of muscle contraction by ryanodine receptor nitrosylation. (A) The ryanodine receptor (RyR) co-localizes with nNOS in the sarcoplasmic reticulum of muscle. At physiologic pO2, local production of NO by nNOS results in selective Snitrosylation of single cysteine residue (C3635) on the ryanodine receptor. S-nitrosylation of C3635 (C3635-SNO) activates the receptor leading to a release of calcium from the sarcoplasmic reticulum and muscle contraction. (B) At higher pO2 levels or under oxidative stress, C3635 may be oxidized (C3635-Sox) and is not nitrosylated. Consequently, NO does not activate the ryanodine receptor, calcium is not released from the sarcoplasmic reticulum, and muscle contraction is attenuated.

in the sarcoplasmic reticulum (9). As is the case with the NMDAR, co-localization of NOS with the RyR targets the RyR for S-nitrosylation leading to reversible activation of receptor function (2, 47). The chemistry underlying RyR S-nitrosylation has been most extensively studied using the skeletal muscle isoform (RyR1). Remarkably, RyR1 has approximately 50 free cysteine residues per subunit, but only a single cysteine residue (C3635) is S-nitrosylated by physiologic nanomolar levels of NO (2, 3). S-nitrosylation of C3635 leads to receptor activation and a NO-mediated enhancement of skeletal muscle contraction (48) (Figure 8.3). Moreover, S-nitrosylation of the RyR is dependent on pO2. A small subset of thiols on each subunit of RyR1 serves as a redox sensor. At tissue-level pO2 (10 mM Hg), the 6-8 thiols are reduced, thereby producing a NO-responsive conformation in the channel. NO may directly react with one of the reduced thiols, or reduction of the thiols may change the structure of the protein and expose a NO-reactive cysteine; however at higher levels of pO2 associated with oxidative stress in peripheral tissues (150 mM Hg), the 6-8 thiols are oxidized thereby preventing RyR S-nitrosylation and attenuating muscle contraction (2) (Figure 8.3). Thus, NO responsiveness of the RyR is tuned by the pO2 (2).

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Nitric Oxide, Cell Signaling, and Gene Expression Ca2+ NMDA receptor

G cyclase

S NO

PSD 95

Fe nNOS SH CAPON

Dexras-GDP

G cyclase FeNO

S NO Dexras-GTP

cGMP

FIGURE 8.3 Regulation of NMDA receptor signaling by protein nitrosylation. The NMDA receptor is a multiprotein complex in which the scaffolding proteins PSD-95 and CAPON bring nNOS into close proximity with three nitrosylation targets: the NR2A subunit of the NMDA receptor, the G protein Dexras, and guanylate cyclase (G cyclase). Stimulation of the NMDA receptor leads to an influx of calcium that activates nNOS. Local production of NO by nNOS leads to selective nitrosylation of a critical cysteine residue on NR2A leading to down regulation of NMDA receptor function. In addition local production of NO nitrosylates Dexras, leading to increased levels of the active GTP bound form of the enzyme. Finally, local NO production nitrosylates the heme iron of guanylate cyclase, leading to enzyme activation and increased cGMP production.

Oxidation of the redox sensor thiols during conditions of oxidative stress, such as muscle fatigue, may be a mechanism to limit muscle contraction (48). These studies also emphasize the importance of studying the regulation of cellular functions by NO under redox conditions (including pO2) that closely mimic the physiologic conditions in tissues.

8.7 THIOREDOXIN SIGNALING The thioredoxin system consists of two antioxidant oxidoreductase enzymes: thioredoxin (Trx) and thioredoxin reductase (TrxR) (49). TrxR catalyzes the reduction of the active site disulfide in Trx. Reduced Trx is a general protein disulfide reductant and scavenges reactive oxygen species (ROS). More recently, thioredoxin reportedly regulates intracellular S-nitrosothiol (SNO) levels and is regulated by S-nitrosylation (50). When thioredoxin is overexpressed, the SNO content of endothelial cells increases. Conversely reducing


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the level of endogenous thioredoxin in cells with antisense oligonucleotides reduces the intracellular levels of SNOs (50). Thus, thioredoxin may be required to maintain intracellular SNO concentrations. Moreover, thioredoxin itself is endogenously S-nitrosylated on cysteine 69 (C69). S-nitrosylation of C69 contributes to the redox regulatory activity of Trx because inhibition of C69 S-nitrosylation decreases the ROS scavenging function of Trx (50). S-nitrosylation of Trx also regulates tumor necrosis factor-α (TNF-α) signaling in endothelial cells. TNF-α-stimulation leads to a generalized reduction in intracellular protein S-nitrosylation and an increase in apoptosis in endothelial cells (51). Overexpression of Trx inhibits both the TNF-α-induced decrease in protein S-nitrosylation and increase in apoptosis (Figure 8.4). The inhibitory effects of Trx on TNF-α signaling are partially dependent on C69 S-nitrosylation. These data suggest that Trx can exert its complete redox regulatory and anti-apoptotic functions during endothelial cell signaling only when C69 is S-nitrosylated (50). TNF-α

TNF-α receptor

+

ROS

+

SNO

−

+

Apoptosis −

Thioredoxin

−

—C69-SNO

FIGURE 8.4 Regulation of TNF-α signaling by S-nitrosylation of thioredoxin. Stimulation of the TNF-α receptor on endothelial cells leads to an increase in reactive oxygen species (ROS) (52), a decrease in the level of intracellular S-nitrosylated proteins (SNOs), and an increase in apoptosis. S-nitrosylation of a critical cysteine on thioredoxin (C69-SNO) inhibits the effects of TNF-α on ROS production, protein S-nitrosylation, and apoptosis.

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8.8 CONCLUSION Nitrosylation is a specific, rapidly reversible posttranslational modification that regulates the function of an increasing number of intracellular proteins; however, the nitrosylation field is still in its infancy and it remains to be determined if nitrosylation is as ubiquitous a regulator or protein function as more traditional posttranslational modifications such as phosphorylation. Technical advances that allow more sensitive detection of endogenously nitrosylated proteins will help elucidate the role of protein nitrosylation in cell signaling. Moreover, although attention in the signal transduction field has been focused primarily on enzymemediated posttranslational modifications, such as phosphorylation, it is likely that nitrosylation is just one of multiple redox-mediated posttranslational modifications, such as sulfenic acid and mixed disulfide formation (13), that have important but as yet undiscovered roles in the regulation of cell signaling.

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Oxide and Caspase 9 Nitric Activation Judith Haendeler and Stefanie Dimmeler University of Frankfurt, Frankfurt, Germany

CONTENTS 9.1 9.2 9.3 9.4 9.5 9.6 9.7 9.8

Introduction ............................................................................................231 NO and Redox Homeostasis .................................................................232 Modulation of Caspases by NO in the Process of Apoptosis ..............234 Modulation of GTPases and Kinases by NO in the Process of Apoptosis .............................................................................235 The Pro-Apoptotic Effects of NO .........................................................236 A Potential Link between NO-Bioavailability, Caspase Activity, Apoptosis, and Aging ............................................................................237 Conclusion .............................................................................................238 Acknowledgment ...................................................................................239 References ...............................................................................................239

9.1 INTRODUCTION NO is a short-lived free radical gas with multiple biological effects. It is synthesized from L-arginine by three different isoforms of NO synthases and is a key molecule in regulating diverse biological processes, including neurotransmission, immune function, and vasoreactivity (Ignarro, 1989). Endogenous NO is synthesized at various rates by the three different NOS enzymes (Nathan and Xie, 1994). The constitutive enzymes, endothelial NOS (eNOS) and neuronal NOS (nNOS), produce low levels of NO (pmolar range) that can be rapidly and transiently increased during intracellular Ca2+ oscillations or upon phosphorylation (Andrew and Mayer, 1999; Dimmeler et al., 1999; Fulton et al., 1999). Inducible NOS (iNOS) is a Ca2+-independent enzyme, which releases nmolar amounts of NO. iNOS is regulated by transcriptional activation of protein expression in response to cytokines or endotoxins in response to inflammation or infection (Kroncke et al., 2000). The classic signalling pathways of NO are attributed to the direct activation of the soluble guanylate cyclase with concomitant cGMP production, which results in cGMP-dependent cellular signalling responses including activation of protein kinases, phosphodiesterases, and phosphatases (Andreopoulos and 231

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Papapetropoulos, 2000). Increasing evidence suggests that cGMP-independent NO effects contribute to cellular signalling, possibly by S-nitros(yl)ation of SHgroups. In the past few years, over 100 target proteins for S-nitros(yl)ation have been identified as recently reviewed by Stamler et al. (for a review, see Stamler et al., 2001). Among them are proteins of different classes like ion-channels, kinases, transcription factors, signalling molecules, and enzymes. Furthermore, NO reportedly plays an important role in apoptotic processes. It has been described as either promoting or preventing apoptosis in response to multiple stimuli in a cGMP-dependent or -independent manner. Apoptosis or programmed cell death is an energy-dependent process. This process required activation of different intracellular pathways dependent on the apoptosis-inducing stimulus used. Important contributors to the apoptosis induction program are the caspases, the mitochondrial amplification loop, and different protein kinases. The importance of caspases for apoptosis signalling was further supported by the use of the novel pancaspase inhibitor IDN-6556 in a first clinical trial in normal volunteers and patients with hepatitic dysfunction. The drug was well tolerated and the effects in hepatic impaired patients appear to be consistent with the administration of an effective hepatoprotective drug that delays cell death in hepatocytes (Valentino et al., 2003). This chapter summarizes: 1. The importance of the intracellular redox status to determine the fate of NO 2. The importance of S-nitros(yl)ation of caspases and protein kinases in inhibiting apoptosis 3. The pro-apoptotic effects of NO 4. The potential link between NO bioavailability, caspase activation, aging, and apoptosis

9.2 NO AND REDOX HOMEOSTASIS NO is a free radical. In contrast to the oxygen-centered radicals, it lacks the reactivity, which is normally inherent with radicals; however, reactive oxygen species can react with NO and this may result in more reactive molecules than NO itself. Reactions of NO include those with oxygen in its various redox forms, other nitrogen-centered molecules, and transition metals (Stamler et al., 1992; Yoo and Fukuto, 1995). The best-studied reaction of NO is to bind and react with transition metals or metal-containing proteins. The classic example is the guanylate cyclase. Guanylate cyclase belongs to the family of heme containing proteins and catalyzes the formation of cGMP. NO binds to the heme moiety of guanylate cyclase, which resulted in a conformational change and thereby activation of the enzyme (Ignarro, 1990). Its product, cGMP, modulates as a known second messenger the function of protein kinases, ion channels, phosphodiesterases, and other important targets (Andreopoulos and Papapetropoulos, 2000). One important example for cGMP

Nitric Oxide and Caspase Activation

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is the regulation of vascular smooth muscle relaxation. Vascular smooth muscle relaxation is mediated by a cGMP-dependent protein kinase that activates a calcium-sensitive potassium channel (Archer et al., 1994). Aside from the important role for NO to increase cGMP, various studies support the concept that NO directly influenced the redox homeostasis of the cell. Important contributors to the redox homeostasis are the mitochondria. Several studies demonstrated that NO controls the cell respiration by reacting with several respiratory chain complexes (Poderoso et al., 1999; Welter et al., 1996). The most sensitive target for NO to inhibit cell respiration is the terminal enzyme of the electron transport chain, cytochrome c oxidase. This has been documented from cells (Sarti et al., 1999; Stumpe et al., 2001) and tissues (Wolin et al., 1999), up to in vivo (Hare et al., 1995; Loke et al., 1999), demonstrating a possible role for NO to regulate the respiratory chain. Therefore, it is tempting to speculate that the interaction of NO with components of the respiratory chain has physiological roles. Indeed, it has been demonstrated that the inhibition of the cytochrome c oxidase by NO is characterized as being reversible and oxygen dependent (Brookes and Darley-Usmar, 2002; Shiva et al., 2004). Thus, the maintenance of the redox homeostasis involves regulatory mechanisms that are capable of sensing NO. This is further underscored by the finding that NO reacts with the heme-containing NOS enzyme, leading to feedback inhibition of NO production by NO (Adak et al., 1999). With respect to iron containing proteins, it has to be noted that the reaction of NO with Fe3+ of the heme moiety is reversible and leads to a reduction of the iron as described for the components of the respiratory chain. In contrast, NO also forms an irreversible, stable nitrosyl complex with Fe2+ in competition with O2. Another important reaction of NO is the reaction with nucleophilic centers of proteins through bimolecular nitrosation reactions, which results in nitrosamines and nitrosothiols. In the past few years, over 100 proteins have been identified that are targets for NO (for a review, see Stamler et al., 2001). Among them are proteins of different classes like ion-channels, kinases, transcription factors, signalling molecules, and enzymes. Several studies supported the concept that S-nitrosothiols are involved in the redox balance of the cell, possibly by regulating the blood flow, allowing targeted delivery of O2 and NO (Lipton et al., 2001; Pawloski et al., 2001). For hemoglobin, it was reported that the interaction of hemoglobin with the anion exchanger 1 promoted the deoxygenated structure of S-nitros(yl)ated hemoglobin, which leads to NO transfer to the membrane (Pawloski et al., 2001). The formation of an S-nitrosothiol at the ryanodine receptor/calcium release channel is restricted to physiological O2 concentration (Eu et al., 2000; Sun et al., 2001). Moreover, thioredoxin has been identified as a target for direct NO modification (Haendeler et al., 2002). Thioredoxin is an oxidoreductase, which directly reduces H2O2 in cells. Direct reaction with NO increased the activity of the enzyme, further supporting the concept that NO can influence the redox-status of a cell. Further, it is of particular interest that cell life cycle is associated with a progressive increase in the oxidative potential from the proliferative state to apoptosis. This can be demonstrated by the steady-state

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increase of intracellular H2O2 in vivo (Boveris and Cadenas, 2000). In turn, these changes in H2O2 may be directly attributed to the availability of bio-active NO in the cell. Thus, the formation of nitrosating species and the process of S-nitros(yl)ation indicate a sensitive dependence on the O2 tension, redox state, and transition metal content of the local redox homeostasis. It has further to be noted that the cellular microenvironment fundamentally contributes to the different effects of NO with particular respect to apoptotic processes.

9.3 MODULATION OF CASPASES BY NO IN THE PROCESS OF APOPTOSIS Caspases are a family of cysteine proteases. This family includes as of now 13 known mammalian members. They are ubiquitously expressed and prominent among the death proteases (Nicholson, 1999). They can be separated into initiator and executor caspases of the apoptotic pathways: the caspase-8 like initiator subfamily, associated with receptor-mediated apoptosis and the caspase-3 like subfamily, known as the molecular executors of apoptosis (Nicholson, 1999). Typically, caspases are present in the cell as zymogens. Upon activation, they cleave targets containing a specific recognition sequence. Among these targets are different signaling molecules leading to degradation of proteins and subsequently to DNA fragmentation. Numerous studies have demonstrated that caspases are molecular targets for direct interaction with NO. Most of the 13 known mammalian caspases can be modified by NO. The first evidence that NO could modulate caspases was reported in cell-free systems. Upon treatment with NO-releasing donors, recombinant human caspases were S-nitrosated at an essential cysteine in the active center of the molecules (Dimmeler et al., 1997b; Haendeler et al., 1997; Li et al., 1997). The formation of S-nitrosothiols led to complete inhibition of caspase activation. Moreover, the modulation by NO was reversible in the presence of an excess of free SH-groups, e.g., the presence of dithiothreitol. Several reports further pointed out that caspase-3 can be poly-S-nitrosated at up to three cysteines (Zech et al., 1999). These modulations by NO could be reversed by glutathione, leading to glutathionylation of caspase-3. Physiological significance to the process of caspase inhibition by NO was given in different cell culture models. Overexpression of caspase-3 increased apoptosis in endothelial cells (Rössig et al., 1999). This process was completely inhibited by exogenous given NO donors due to S-nitrosation of caspase-3 at the active site at cysteine-163. Moreover, Mannick et al. reported that pro-apoptotic stimuli reduced S-nitrosation of caspase-3 in MCF-7 cells, demonstrating a possible denitrosation process (Mannick et al., 1999). This process was reversible by increasing the endogenous steady-state level of NO (Mannick et al., 1999). These data implicate that caspases are potential direct targets for modulation by NO; however, it has to be noted that cGMP/protein kinase G-dependent mecha-


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nisms may also contribute to the suppression of caspase activation by NO (Kim et al., 1999; Kim et al., 1997). Further implications for the role of NO and caspase activation were given by studies involving endogenous NO production by NOS. Activation of all three isoforms of NOS inhibits caspase activation. In venous and pulmonary artery endothelial cells, caspase activation and apoptosis induction was dependent on inhibition of the eNOS (Ceneviva et al., 1998; Dimmeler et al., 1997b; Haendeler et al., 1997). NO derived from nNOS can increase S-nitrosation of caspase-3 which can suppress neuronal apoptosis induction (Park et al., 2002); however, other studies demonstrated that glutamate-induced activation of the nNOS leads to apoptosis induction (Rameau et al., 2003). Therefore, further studies are needed to clarify the physiological significance of inhibition of caspase activation by nNOS. In hepatocytes, activation of the iNOS inhibits apoptosis induction by modulation of the initiator and executor caspases through NO (Kim et al., 2000). Of note, S-nitrosation of caspases by NO-aspirin (NCX-4016) was detected in a concanavalin A-induced hepatitis mouse model. Thereby, NO-aspirin had similar effects than pancaspase inhibitors suggesting an important role for modulation of caspases by NO even in in vivo models (Fiorucci et al., 2002). Taken together these studies clearly support the idea that modulation of caspases by NO is a central mechanism for the anti-apoptotic properties of NO.

9.4 MODULATION OF GTPASES AND KINASES BY NO IN THE PROCESS OF APOPTOSIS Aside from the caspases, mitogen-activated kinases and their upstream signals (e.g., Ras GTPase) play crucial roles in signalling events leading to cell differentiation, proliferation, survival and apoptosis. Several studies now supported the idea that NO can also directly modulate kinases and Ras GTPase by S-nitrosation. Specifically, NO induces conformational changes in c-Ras through S-nitrosation of a critical redox-sensitive cysteine, which in turn leads to increased guanine nucleotide (GDP/GTP) exchange (Lander et al., 1997; Lander et al., 1995). c-Ras is an important activator of the ERK1/2 promoting cell proliferation and survival suggesting that an increase in c-Ras activity will induce cell survival. This hypothesis is supported by recent studies, which demonstrated that the physiological most important stimulus for the activation of eNOS and the protection against apoptosis of endothelial cells, the laminar flow in the blood vessel (named shear stress), increased the S-nitrosation and activity of c-Ras in human endothelial cells (Hoffmann et al., 2003). Given the fact that NO modulates a redox-sensitive cysteine residue in c-Ras, one may speculate that activation of c-Ras by NO is an intracellular answer of cells to respond to changes in the redox-status. Further evidence supports redox-sensitive cysteines, which are modulated by NO in other proteins (e.g., the c-Jun N-terminal kinases [JNKs]). JNKs can be activated by H2O2 and are sensitive to GSH underscoring its redox-dependent regulation (Adler et al., 1999). With respect to modulation

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by NO, recombinant JNK1 and JNK2 can be S-nitrosated after incubation with NO donors, which subsequently led to an inhibition of JNK activity. Furthermore, these findings could also be demonstrated in different cell types. Upon stimulation of iNOS, JNK was S-nitrosated and its enzymatic activity was abolished (Park et al., 2000). Of note, activation of JNK can lead to activation of caspases and JNK can be activated by caspase-3 pointing toward a possible amplification loop for apoptosis induction (Dietrich et al., 2004; Sabourin et al., 2000). Taking into account that NO can inhibit caspase-3 and JNK, respectively, by direct interaction would further support the importance of NO and its bioavailability in protecting against apoptotic processes in vivo.

9.5 THE PRO-APOPTOTIC EFFECTS OF NO NO can act in cytotoxic and cytostatic ways. High fluxes of NO produced by the iNOS or by mM concentrations of NO donors are cytotoxic for cells. The cytotoxicity of NO can be beneficial by killing bacteria and parasites; however, NO can induce tissue destruction. On a cellular basis, different mechanisms describing how NO exerts its pro-apoptotic effects are presented. Under physiological pH, NO can be oxidized by molecular oxygen to N2O3. N2O3 is a powerful nitrosating agent and its formation was demonstrated in different cell types (Espey et al., 2002; Jourd’heuil et al., 2003). N2O3 can directly deaminate DNA. Several studies in mammalian cells exposed to NO have suggested that nitrosative deamination of nucleobases contributes to “spontaneous” deamination in vivo and thereby induces direct DNA damage (Burney et al., 1999; Wink et al., 1991). Similar to the formation of N2O3 in cells NO can also react with superoxide to form ONOO-. ONOO- directly damages DNA, lipids, and proteins by oxidation (Estevez and Jordan, 2002; Virag et al., 2003). The direct DNA damage results in concomitant up-regulation of the p53 tumor suppressor (Messmer et al., 1994), which leads to an increase in the pro-apoptotic protein Bax and to induction of apoptosis (Knudson and Korsmeyer, 1997; Miyashita et al., 1994). Furthermore, NO has a direct effect on the cytochrome c oxidase as described in Section 9.2, which in turn leads to the inhibition of the respiratory chain and to an increase in superoxide production from the mitochondria. Therefore, several studies have documented that the direct effect of NO on proteins in the mitochondria will initiate the loss of the mitochondrial membrane potential and, thereby, induce cytochrome c release (for a review, see Cadenas, 2004). The release of cytochrome c leads to the formation of the apoptosome complex, which results in cleavage and activation of the death effector caspase-9 and finally induction of apoptosis (Hengartner, 2000). It is still unclear, however, why high doses of NO increase caspase activation and do not lead to an inhibition of caspases by oxidation or nitrosation of the active cysteine. It has to be noted that the inhibition of caspases by oxidation of the reactive cysteine has not been documented in vivo. Moreover, the increase in superoxide production induced by inhibition of the respiratory chain by high doses of NO could in turn activate the effector caspases. Thus, high doses of NO may nitrosate caspases, but the burst


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in endogenous superoxide may induce a reactivation of the caspases by an unknown mechanism. Moreover, the enhanced production of superoxide from the mitochondria leads to an increase in ONOO- formation in close proximity to the mitochondria. ONOO- has been demonstrated to irreversibly oxidize the mitochondrial complexes I, II, IV, and V as well as the mitochondrial membrane lipids (Brown and Borutaite, 2002; Keller et al., 1998). Thus, these oxidations lead to alterations in the permeability transition and, thereby, to the release of cytochrome c and subsequently to the activation of caspases and the induction of apoptosis. It has to be noted that increase in ONOO- will also inactivate other metabolic enzymes, such as aconitase, which contains Fe-S centers (Castro et al., 1994), and will activate DNA repair enzymes such as poly(ADP-ribose)synthetase (Zhang et al., 1994). Inhibition of ATP production from the mitochondria and activation of ATP consuming enzymes will in turn result in ATP depletion; however, apoptosis is an ATP-dependent process, therefore, the NO/ONOO−-induced cell death should be implicated in necrotic processes.

9.6 A POTENTIAL LINK BETWEEN NO-BIOAVAILABILITY, CASPASE ACTIVITY, APOPTOSIS, AND AGING NO exerts many protective functions in the vasculature including inhibition of platelet aggregation, neutrophil adhesion, endothelial cell apoptosis, and smooth muscle cell proliferation. These important anti-atherosclerotic functions of NO were evidenced by numerous experimental and clinical studies. For example, eNOS deficient mice develop hypertension, exhibit a reduced growth factorinduced angiogenesis, and accelerated atherosclerotic lesion formation (Huang et al., 1995; Moroi et al., 1998; Murohara et al., 1998). Moreover, an impairment of endothelial NO-synthesis predicts a worse outcome in patients with coronary artery disease (Schachinger et al., 2000). Classical risk factors, which are known to promote endothelial dysfunction in vivo, can induce endothelial cell apoptosis in vitro. Pro-inflammatory cytokines and the peptide hormone angiotensin II induced apoptosis in endothelial cells (Dimmeler et al., 1997c; Fujita et al., 2000). Likewise, oxidized low-density lipoprotein (LDL) and high concentrations of reactive oxygen species triggered apoptosis of endothelial cells (Dimmeler et al., 1997a; Harada-Shiba et al., 1998; Kontush et al., 2003; Sudoh et al., 2001). All of the mentioned pro-apoptotic stimuli triggered apoptosis induction in a caspase-3 dependent manner. Interestingly, aging of endothelial cells led to enhanced apoptosis sensitivity toward TNFα and oxidized LDL which was accompanied by an increase in caspase-3 activity and a reduction in NO bioavailability in aged endothelial cells (Hoffmann et al., 2001), suggesting a potential link between NO bioavailability, caspase activity, aging, and apoptosis. Recent studies, in animals and humans, further suggest that a link exists between aging and NO-bioavailability. In rats, the NO-bioavailability in aortas as well as the endotheliumdependent relaxation declined with age (Tschudi et al., 1996). Aging of humans

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could also be correlated with reduced NO-bioavailability (Zeiher et al., 1993). Moreover, endothelial cell apoptosis is enhanced in old monkeys, suggesting a link between aging and apoptotic cell death (Asai et al., 2000). Thus, these studies support the importance of NO for two processes, namely apoptosis and aging. Moreover, on a molecular level aging and apoptosis are connected.

9.7 CONCLUSION NO exhibits a double-edged role in apoptosis induction. On a molecular level, the different effects of NO on apoptosis may be explained by the reaction of NO with different molecular targets (Figure 9.1). Thus, the activation of several enzymes, which contribute to apoptosis, can be modulated by NO with either pro- or anti-apoptotic effects (Figure 9.1). NO can influence the redox homeostasis of the cell by reacting with oxygen in its various redox forms, other nitrogen-centered molecules, proteins, and transition metals. NO can alter the homeostatic balance by regulating the activation of caspases, which are the central mediators of apoptosis in physiological and pathological conditions. NO can reversibly inhibit caspases by forming S-nitrosothiols. In other cell types, NO can oxidize proteins in the mitochondria and thereby initiates the loss of the mitochondrial membrane potential and induces apoptosis. Because the NO-bioavailability and caspase-dependent cell death is central in numerous diseases, these studies support the concept that modulation of caspase activation and delivery of appropriate amounts of NO may offer promising therapies in human diseases.

NO

Initiator caspases

NO

RasGTP

Mito

caspase cascade

NO

cGMP

Respiratory chain Executor caspases

JNK1/2

O2− .

DNA damage

NO

N2O3 ONOO −

Apoptosis

FIGURE 9.1 NO and apoptosis.


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9.8 ACKNOWLEDGMENT The authors acknowledge the many important and relevant articles in this field that may not have been cited in this chapter due to space limitations.

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Effects of 10 Signaling Peroxynitrite in Mammalian Cells Lars-Oliver Klotz Heinrich-Heine-Universität Düsseldorf, Düsseldorf, Germany

CONTENTS 10.1 Introduction ............................................................................................245 10.2 General Considerations ..........................................................................246 10.2.1 Membrane Permeability and Intracellular Targets of Peroxynitrite ...........................................................................246 10.2.2 How to Prove the Involvement of Peroxynitrite in an Observed Biological Effect ...................................................247 10.3 Mechanisms of Peroxynitrite Signaling ................................................248 10.3.1 Tyrosine Nitration vs. Phosphorylation .....................................249 10.3.2 Role of Tyrosine Phosphatases in Peroxynitrite Signaling .......250 10.4 Cellular Signaling Effects of Peroxynitrite ...........................................251 10.4.1 Adaptive Responses to Exposure to Peroxynitrite ....................252 10.4.2 Growth Arrest and Cell Death ...................................................254 10.5 Acknowledgments ..................................................................................255 References ...............................................................................................256

10.1 INTRODUCTION The signaling effects of nitric oxide (NO) are, at least in part, also defined by NO reaction products with reactivities of their own that are discernible from those of the “mother compound”. One such product is peroxynitrite, which is generated in the nearly diffusion-controlled reaction of NO with superoxide (k2 ≈ 1010 M−1s−1). Thus, peroxynitrite may cause cellular responses that are at the intersection of the effects elicited by NO and by the action of superoxide. Whether or not peroxynitrite significantly contributes to these NO and superoxide effects in vivo is a matter of intense debate. It is, however, without question that the exposure of cells to peroxynitrite, either by application of authentic peroxynitrite or by generating NO and superoxide in parallel, such

245

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as by employing 3-morpholinosydnonimine (SIN-1), does affect cellular signaling pathways in several in vitro systems, including mammalian cell culture. The outcome of such treatment is distinguishable from both NO or superoxideelicited signaling. At first glance, it is not very surprising that a strong oxidant affects cellular signaling processes, especially those involved in the cellular stress response. What makes the study of oxidant-induced signaling, and of peroxynitrite effects in particular, interesting is the fact that “oxidative stress” is not the same independent of how it is brought about because oxidants differ in their biochemical reactivities, their lifetimes, and their sites of generation. In addition, a multitude of potential reaction types exist in the case of peroxynitrite that compose the final signaling pattern on target cells, including oxidation, nitration, and nitrosation reactions. It is not the aim of this chapter to provide an overview on peroxynitrite biochemistry, which has been done extensively elsewhere [1] and in this book (see Chapter 5). Instead, it is attempted to give an overview on peroxynitriteinduced signaling with a focus on general mechanisms of activation of signaling events by an oxidizing and nitrating species, and on the outcome of the modulation of signaling processes by peroxynitrite in terms of cellular proliferation.

10.2 GENERAL CONSIDERATIONS 10.2.1 MEMBRANE PERMEABILITY AND INTRACELLULAR TARGETS OF PEROXYNITRITE Although with a half-life of about 1 sec at pH 7.4 and 37°C [2] peroxynitrite is rather short-lived as compared to nitric oxide or hydrogen peroxide, it diffuses far enough to act on and in cells, even if applied from outside the cell. It readily crosses phospholipid membranes, as was demonstrated employing liposomal systems [3] as well as erythrocytes [4, 5]. In the presence of CO2 and other biological targets (see below), the half-life of peroxynitrite is significantly shortened and on the order of a few milliseconds, thus limiting the extent of membrane crossing. With a pKa of 6.8 at both 25°C and 37°C [2, 6], about 80% of the peroxynitrous acid/peroxynitrite couple will be in the anionic form at pH 7.4; accordingly, anion transporters appear to mediate its entry into cells, such as the band 3 anion exchanger in erythrocytes. The free permeability of liposomal membranes to peroxynitrite may be explained by diffusion through the lipid bilayer of the fraction of peroxynitrous acid present at physiological pH, although the formation of electrically neutral complexes of peroxynitrite with cations such as Na+ has also been hypothesized [5]. It can be concluded from these data that cellular targets for extracellularly formed peroxynitrite are not only found in the cell membrane but also intracellularly. Carbon dioxide, which is present at about 1 mM in plasma, and low-molecular weight thiols, such as glutathione (GSH), which is present in cells in millimolar concentrations, are major targets of peroxynitrite [7]. In addition, depending on the microenvironment, certain proteins were demonstrated to be preferred targets

Signaling Effects of Peroxynitrite in Mammalian Cells

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of peroxynitrite, such as myeloperoxidase in neutrophils or hemoglobin in erythrocytes. The selenocysteine-containing glutathione peroxidase also efficiently reacts with peroxynitrite, acting as a peroxynitrite reductase at the expense of glutathione [8, 9]; in contrast to other proteins targeted by peroxynitrite, glutathione peroxidase is reversibly oxidized by peroxynitrite, and this glutathione peroxidase/glutathione system is a likely candidate for a physiological defense mechanism against peroxynitrite.

10.2.2 HOW TO PROVE THE INVOLVEMENT AN OBSERVED BIOLOGICAL EFFECT

OF

PEROXYNITRITE

IN

Because no such enzyme specifically scavenges peroxynitrite as can be found for hydrogen peroxide, the dismutation of which is specifically catalyzed by the enzyme catalase, it is difficult to prove the involvement of peroxynitrite in an observed biological effect. The nitration of protein-bound tyrosine has been frequently used as a footprint for the action of peroxynitrite [10]; however, formation of 3-nitro-tyrosine is also achieved in systems with hypochlorite/nitrite [11, 12] as well as peroxidase/nitrite [13, 14]. Thus, to involve peroxynitrite in a biological effect, this will additionally have to be experimentally attenuated by use of more or less specific peroxynitrite scavengers, such as Mn- and Feporphyrins, selenocompounds, and others [7]. The search for peroxynitritespecific reactions that may be exploited analytically as markers for the action of peroxynitrite, is ongoing [15]. Not only is demonstration of peroxynitrite being the source of a biological effect a demanding task, but it is also difficult to assess the relative contributions of tyrosine nitration and oxidation reactions to observed peroxynitrite effects. Schroeder et al. [16] described the inhibitory effect of the flavanol (−)-epicatechin (Figure 10.1) on tyrosine nitration and various oxidation reactions by peroxynitrite and found that this compound, which is present in chocolate or green tea, inhibits tyrosine nitration by peroxynitrite several orders of magnitude more efficiently than it inhibits oxidation reactions by peroxynitrite. The oxidation reactions examined were the oxidation of thiols, the oxidative inactivation of glyceraldehyde-3-phosphate dehydrogenase or of purified soybean lipoxygenase-1 as well as the oxidation of 2′,7′-dichlorodihydrofluorescein in cells. Based on an efficiency of (−)-epicatechin in preventing the peroxynitrite-induced dimerization of tyrosine equal to that of preventing its nitration, it was hypothesized that the apparent selectivity of (−)epicatechin with regard to interfering with tyrosine nitration but not oxidation reactions is due to interference with tyrosyl radicals instead of to a direct interaction with peroxynitrite [17] (Figure 10.2). This is in line with data from Tibi and Koppenol [18] demonstrating that neither catechol nor a model flavonoid are scavengers of peroxynitrite (i.e., the rate of peroxynitrite decomposition is not changed by these compounds, both in the presence and absence of carbon dioxide). Urate (Figure 10.1) turned out to be another efficient inhibitor of tyrosine nitration [15, 19], also not by directly scavenging peroxynitrite but instead by interference with radical intermediates responsible for tyrosine nitration.

248

Nitric Oxide, Cell Signaling, and Gene Expression O

O

OH HO

NH

HN

OH

O

O

O

O

N N H H Uric acid

OH

O

N H

OH

pK = 5.4

N

HN

(−)-Epicatechin

N H

H+

O

OH

N

HN

N H

N H

O−

Urate

FIGURE 10.1 Structures of inhibitors of peroxynitrite-induced tyrosine nitration, epicatechin, and urate.

• OH Tyrosine

H2O

O •

O Tyrosyl radical Epicatechin

•OH

ONOO−

ONOOH H

+

•

×2

NO2

NO2 OH

OH

3-Nitrotyrosine

OH

Dityrosine

FIGURE 10.2 Hypothetical mechanism of epicatechin inhibition of peroxynitrite-induced tyrosine nitration: interaction of epicatechin with tyrosyl radicals [17].

10.3 MECHANISMS OF PEROXYNITRITE SIGNALING The oxidation and nitration of various biomolecules leads to generation of products that, in turn, may give rise to signaling events, such as the lipid peroxidation products 4-hydroxynonenal [20, 21], malondialdehyde [22], or acrolein [23] that are known to activate a variety of stress-responsive signaling cascades or DNA damage that triggers the cellular DNA damage response [24–26]. Dysregulation of Ca2+ concentrations with its obvious consequences for the activities of calciumdependent enzymes is a common phenomenon in cells exposed to oxidants, and


249

this may also be, in part, caused by oxidation of (membrane) lipids or proteins that regulate the sub-cellular distribution of calcium ions. As with most other oxidants, peroxynitrite may cause cell death by initiating these processes, leading to dysregulation and loss of cellular integrity. It appears that only a restricted and defined extent of (peroxynitriteinduced) damage may foster signaling events and regulatory processes in terms of a pro-survival stress response. This is nicely reflected even at the level of interaction of peroxynitrite with isolated proteins, for example isolated glyceraldehyde 3-phosphate dehydrogenase (GAPDH). GAPDH activity, which relies on the intactness of a crucial cysteine at the enzyme’s active site, very rapidly declined even at lowest peroxynitrite concentrations, followed, at higher peroxynitrite levels, by protein dimer and oligomer formation, probably due in part to tyrosine dimerization, and by tyrosine nitration [27]. Interestingly, GAPDH became highly susceptible for degradation by isolated 20S proteasome after exposure to peroxynitrite at concentrations well below those leading to significant inactivation, dimerization, and nitration. With higher peroxynitrite concentrations used, proteasomal degradation of the exposed GAPDH decreased, probably due to significant changes of the protein structure [27]. Hence, at low oxidant concentrations, a modified protein may be rapidly degraded, whereas at high concentrations the oxidative damage to the protein may become irreversible. Such an effect of peroxynitrite on proteasomal degradation was also seen in cultured cells exposed to authentic peroxynitrite or SIN-1, again with a bellshaped pattern: Increasing the oxidant dose enhances protein degradation until a maximum protein turnover is reached. Even higher peroxynitrite concentrations result in a decrease in protein turnover without significant loss in cell viability or proteasome activity [28]. These studies nicely demonstrate that adaptation in terms of the enhanced degradation of protein before significant protein deterioration may occur only within a defined range of oxidant concentrations beyond which damage is too extensive to be held transient and be exploited as a regulatory switch.

10.3.1 TYROSINE NITRATION

VS.

PHOSPHORYLATION

Not long after it was first demonstrated that peroxynitrite leads to the nitration of tyrosyl residues and the formation of 3-nitrotyrosine in vivo [29], it was demonstrated that this affects phosphotyrosine-dependent signaling in cell-free systems [30] and in cell culture [31–33] in that nitrated tyrosines cannot be phosphorylated (Figure 10.3). A recent example for this mutual exclusiveness of tyrosine nitration/phosphorylation is the peroxynitrite-induced tyrosine nitration of CD95 in rat hepatocytes, which prevents its tyrosine phosphorylation and the induction of apoptosis by CD95-L; vice versa, prior tyrosine phosphorylation of CD95 does not allow for significant nitration by exposure to peroxynitrite [33]. It was later reported that signal transduction pathways that rely on tyrosine phosphorylation, such as those described in detail below, are not blocked but are instead activated by peroxynitrite. This can mean that either tyrosine nitrations

250

Nitric Oxide, Cell Signaling, and Gene Expression ONOO− no dephosphorylation ATP NO2

(3)

OH

(1)

OH

O S P O− O−

PTK S−

ADP

no phosphorylation

(2)

O–PO 2−3

PTPase

H2O H2PO−4 ONOO−

SO(1-3)H inactive PTPase

FIGURE 10.3 Modulation of tyrosine (1) phosphorylation/dephosphorylation-based signaling effects by peroxynitrite: nitration of tyrosyl residues (3) prevents their phosphorylation by protein tyrosine kinases (PTK, left), whereas oxidatively inactivated protein tyrosine phosphatases (PTPases) leave phosphotyrosine residues (2) unaffected [89].

by peroxynitrite are preferably at sites not crucial for signaling or that some nitrations promote instead of block tyrosine phosphorylation-dependent signaling. Both possibilities apply: An example for the former is given by Schroeder et al. [16] who demonstrate that selective inhibition of tyrosine nitration by peroxynitrite employing (−)-epicatechin does not diminish activation of MAP kinases and the PI3K/Akt pathway by peroxynitrite, implying that activation is not due to, or negatively influenced by, tyrosine nitration. On the other hand, MacMillan-Crow et al. [34] suggest that c-Src activation by peroxynitrite might be due to tyrosine nitration, keeping the protein in an unfolded, active state. Similar to this hypothesis of tyrosine nitration of Src leading to its activation, Mallozzi et al. [35] demonstrated that nitrotyrosine-containing peptides can interfere with the intramolecular phosphotyrosine/SH2-interaction in Src-family kinases that keeps them in an inactive state, leading to activation, implying and assuming that nitrotyrosine and phosphotyrosine may compete for binding sites under certain circumstances.

10.3.2 ROLE OF TYROSINE PHOSPHATASES SIGNALING

IN

PEROXYNITRITE

All protein tyrosine phosphatases (PTPases) known so far rely on the presence of a cysteine in their active site which serves as a nucleophile accepting the phosphate moiety of the phosphatase substrate forming an intermediate phosphocysteine (Figure 10.3) [36, 37]. An oxidation of this cysteine residue (e.g., by peroxynitrite) or its alkylation (e.g., by naphthoquinone derivatives) [38, 39], would lead to the inactivation of the phosphatase concomitantly allowing kinase activities to become predominant (see Figure 10.3). Peroxynitrite is known to oxidize thiols to preferably form the respective sulfenic acids (RSOH) or disulfides (RSSR) and, at more acidic pH, thiyl radicals [40]. It was further demonstrated to efficiently inhibit both cellular protein tyrosine phosphatase


251

activity [32] and various isolated tyrosine phosphatases [41]. This inhibition is not reversible with DTT, indicating that oxidation is beyond the disulfide or sulfenic acid state, and probably sulfinic (RSO2H) or sulfonic acid (RSO3H) is formed [41]. Recently, another reversible oxidative modification of the active site of PTPases exposed to oxidants was described, the formation of a sulfenyl-amide [42, 43]. Hence, a mechanism for activation of signaling pathways by peroxynitrite may be envisaged that is based upon the oxidative inactivation of tyrosine phosphatases. Yet this cannot account for all oxidant-induced signaling; for example, the peroxynitrite-induced activation of Akt (protein kinase B) does not solely rely on the oxidative inactivation of a phosphatase: Peroxynitrite treatment renders cells refractory to subsequent Akt activation by growth factors, indicating that phosphatase inhibition cannot be the sole mechanism responsible for activation of the kinase [44]. If that were the case, phosphorylation would not be expected to be inhibited by peroxynitrite pretreatment, but instead enhanced due to the loss of negative regulation. Similarly, the activation of Src-family kinases in erythrocytes was demonstrated to not strictly depend on the inactivation of a tyrosine phosphatase [45]: In the presence of carbon dioxide, the inhibition of membranebound PTPases elicited by treatment of erythrocytes with peroxynitrite was attenuated; nevertheless, activation of src kinases did not appear to be affected.

10.4 CELLULAR SIGNALING EFFECTS OF PEROXYNITRITE Cellular reactions to stressful stimuli, such as the exposure to oxidants, are depicted in Figure 10.4. Although low oxidant concentrations may elicit proliferative effects, adaptive responses are induced at higher concentrations, followed by the need to stop proliferation and arrest growth at concentrations that cause considerable damage that requires time for repair or that leads to the induction of apoptosis. This has been demonstrated for hydrogen peroxide and Jurkat T-cells, with discrete ranges of steady-state concentrations of the oxidant allowing for cell proliferation (< 0.7 µM) or causing apoptosis (1 to 3 µM) and necrosis (>>3 µM) [46]. Similar cellular reactions can be expected for peroxynitrite. concentration Peroxynitrite

Proliferation

Adaptation

Growth arrest

Repair

Cell death

Apoptosis

FIGURE 10.4 Cellular stress responses to increasing concentrations of peroxynitrite.

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The mitogen-activated protein kinase (MAPK) pathways regulate cellular proliferation, differentiation, and cell death and appear to be activated by a great variety of stressful stimuli, including oxidative stress. The major MAPKs (for reviews, see [47–50]) are activated by upstream-kinases (the MAPK kinases, MKKs) by dual (Thr- and Tyr-) phosphorylation of a Thr-XTyr-motif and are “proline-directed” Ser/Thr-kinases that, in turn, phosphorylate their substrates on Ser- or Thr-residues at positions defined by the direct C-terminal attachment of proline. In addition to the classical mitogenactivated protein kinases (the extracellular-signal-regulated kinases [ERK] 1 and 2), which are activated by mitogenic stimuli such as growth factors, the MAPK group comprises stress-activated kinases: one subgroup, termed p38, with four members of apparent molecular masses between 38 and 43 kDa [51], and one, the c-Jun-N-terminal kinases (JNK), which consists of 10 isoforms with molecular masses between 46 and 57 kDa, which are derived from three genes, JNK-1 through JNK-3, and generated by alternative splicing [52]. These three MAPK-subfamilies are activated by more or less specific MKKs, which, in turn, are phosphorylated and activated by MKK kinases (MKKKs). Small GTP-binding proteins, such as Ras, are often involved in activation of these three-kinase-modules. Among the substrates of activated MAPKs are transcription factors such as Elk1, Sap, c-Jun, or ATF2. Upon phosphorylation by MAPKs, these factors are activated and lead to the expression of proteins constituting the leucine-zipper transcription factor AP-1 (e.g., c-Jun and c-Fos). Following activation of AP-1, the expression of genes with functional AP-1 elements in their promoter regions may be induced.

10.4.1 ADAPTIVE RESPONSES

TO

EXPOSURE

TO

PEROXYNITRITE

Peroxynitrite is known to activate all three previously mentioned MAP kinase family members, ERK 1/2, p38 and JNK, in a wide variety of cell types, including rat liver epithelial cells [53], rat fibroblasts [54], bovine endothelial cells [55], human neuroblastoma cells [56] and human neutrophils [57, 58]. Indeed, the induced expression of stress genes such as c-fos [59], heme oxygenase-1 [60], or the growth arrest and DNA damage-inducible (Gadd) proteins 34, 45, 153 [61], upon exposure of cells to peroxynitrite was described, although the role of MAPK in these effects is not yet fully elucidated. What leads to the activation of ERKs upon exposure to peroxynitrite, and what are the consequences of ERK activation? The consensus as to how ERKs are activated by peroxynitrite appears to be limited: It is known that the epidermal growth factor (EGF) induces the activation of ERKs by activating its receptor (EGFR), a receptor tyrosine kinase, followed by activation of downstream molecules such as Ras, Raf, and MKK 1/2 (the MKKs directly upstream of ERKs; also termed MEK 1 and 2, for MAPK/ERK kinase). Hydrogen peroxide was demonstrated to activate ERKs along the same pathway [62]. Regarding the effect of peroxynitrite, however, reports as diverse as the cell types used for the studies are available. According to the literature, activation of ERKs occurs:


253

1. Via EGFR and downstream targets [63] 2. Independent from EGFR and Raf, but dependent on activation of MKK1/2 [62] 3. Activation-independent from EGFR and even MKK 1/2, but partly coming from a Ca2+-dependent PKC-isoform [54] Employing (−)-epicatechin to selectively inhibit tyrosine nitration by peroxynitrite did not diminish activation of ERK- and p38-MAP kinases as well as the PI3K/Akt pathway by peroxynitrite in murine aortic endothelial cells, implying that activation is not due to, or negatively influenced by, tyrosine nitration [16]. Regarding the EGFR, tyrosine phosphorylation of the receptor was enhanced in human skin fibroblasts upon treatment with SIN-1 [44]; in A431 epidermoid carcinoma cells, however, EGF receptor molecules dimerized covalently upon treatment with peroxynitrite, presumably due to the formation of dityrosines, yet no increased activity in terms of autophosphorylation resulted from that dimerization. Instead, the activation by EGF of a downstream molecule, phospholipase C-γ1, was attenuated [64]. ERK activation by peroxynitrite in human neutrophils leads to induced expression of CD11b/CD18 and enhanced adhesion of such peroxynitritetreated leukocytes to lipopolysaccharide-treated endothelial cells [58]. Although here the role of ERK activation appears to be that of the induction of gene expression and protein synthesis, effects that are based upon substrates for ERK other than transcription factors also occur. Neutrophilic NADPH oxidase is activated via ERKs under certain circumstances [65]; phorbol esterstimulated activation of NADPH oxidase was demonstrated to be enhanced by low micromolar concentrations of peroxynitrite via MKK1/2. Another example for a possible biological effect of peroxynitrite via ERKs is gap-junctional intercellular communication (GJC): It is known that ERKs phosphorylate proteins regulating GJC, such as the gap-junctional protein connexin43—with phosphorylation blocking gap junctional cell–cell communication [66]. As would be expected, exposure of WB-F344 rat liver epithelial cells to peroxynitrite diminishes intercellular communication [67]; however, no proof exists regarding the involvement of ERK 1/2 yet. Another pathway emanating from growth factor receptors and involved in antiapoptotic and proliferative responses to growth factors, such as platelet-derived growth factor (PDGF) and EGF, is the phosphoinositide 3-kinase (PI3K)/Akt pathway. This pathway is also activated in response to stressful stimuli such as heat shock or hydrogen peroxide [68]. PI3K is upstream of Akt (protein kinase B) and regulates Akt activity by generating 3′-phosphorylated phosphoinositides that reside in the cell membrane, and leads to binding of the pleckstrin homology domain of Akt, concomitant with a translocation of Akt to the cell membrane. This relocalization of Akt renders it accessible to regulating kinases that phosphorylate Akt on Thr-308 and Ser-473 (for a review, see [69]). Exposure of human skin fibroblasts to both authentic peroxynitrite and SIN-1 leads to activation of the PI3K/Akt pathway, followed by phosphorylation of an established in vivo-substrate of Akt,

254


glycogen synthase kinase-3 (GSK-3). Both the EGFR and PDGFR A/B were activated by peroxynitrite, but only PDGFR A/B appeared to be responsible for activation of PI3K/Akt [44]. The non-receptor tyrosine kinase, c-Src, is involved in EGFR activation in some cases (for a review, see [70]). In addition, phosphorylation of the EGFR in endothelial cells exposed to H2O2 was indeed demonstrated to be inhibitable by addition of PP2, a Src tyrosine kinase inhibitor [71], which also blocked the activation of Akt by H2O2—known to rely upon EGFR activation [72]—in HeLa cells [73]. Peroxynitrite activates Src and Src-family kinases in various human cell types [34, 45, 74]. The role of Src in receptor tyrosine kinase activations by peroxynitrite, however, remains to be resolved. Src kinases do play an important role in peroxynitrite activation of AMPactivated kinase (AMPK) via PI3K [75, 76]. AMPK activation leads to stimulation of processes mediating replenishment of cellular ATP, such as fatty acid oxidation or glycolysis [77]. Interestingly, AMPK activation by peroxynitrite occurs without affecting cellular AMP concentrations [76]. Another adaptive response, the significance of which is immediately evident, is observed with cells exposed to peroxynitrite: A two- to sixfold up-regulation of cellular glutathione levels was described for bovine aortic endothelial cells and smooth muscle cells 16 to 18 h after exposure to SIN-1, which was partially prevented by SOD [78]. Peroxynitrite thus appears to enable the cell to better cope with a peroxynitrite stress.

10.4.2 GROWTH ARREST

AND

CELL DEATH

According to Figure 10.4, growth arrest is one possible cellular response to concentrations of peroxynitrite that do not allow for signaling processes in terms of an adaptive response to occur (e.g., because oxidative damage is significant). Growth arrest gives the cell time for repair of damage before proliferation resumes or to initiate apoptosis and regulated cell death if damage exceeds the cellular repair capacity. For example, the tumor suppressor gene product p53 is activated upon recognition of DNA damage and induces cell cycle arrest (e.g., by acting as a transcription factor that stimulates the production of the inhibitor of cyclindependent kinases, p21waf). Indeed, peroxynitrite is known to block cell proliferation and induces growth arrest at according concentrations [79]. In human keratinocytes, peroxynitrite-induced growth arrest was followed by the expression of keratin patterns that characterize terminal differentiation [79]. Interestingly, p53 binding to DNA target sequences was impaired instead of induced by peroxynitrite [80, 81]. This may be due to oxidation of the zinc-finger zinc-thiolate clusters of p53. It was similarly demonstrated that DNA binding of retinoid X-receptor / vitamin D-receptor heterodimeric zinc-finger transcription factors is abolished after exposure to peroxynitrite, which was not due to DNA damage [82]. The same effect was seen with NO, but NO effects were reversible in the presence of DTT whereas peroxynitrite-effects were not [82].


255

Peroxynitrite can induce apoptosis [31, 83–85]. Apoptosis has been linked with MAPK activation because a crucial role of p38 and JNK as pro-apoptotic stimuli in PC12 cells was proposed, whereas activation of ERK seemed to be anti-apoptotic [86]. Although the importance of JNK and p38 in mediating apoptosis following stress has been well documented in a variety of model systems, clear exceptions to this generality exist. In fact, cases exist in which JNK and p38 activations appear to play a role in protecting cells against apoptosis [87] as well as cases in which activation of ERKs is a prerequisite for apoptosis [88]. A treatise of the mechanisms of peroxynitrite-induced apoptosis and the role of mitochondria is beyond the scope of this article. The signaling effects of peroxynitrite outlined previously are summarized in Figure 10.5. ONOOH/ONOO− RTK

Akt

GSK-3

e

an

AMPK

br

P13K

em

JNK, p38

m

MKK1

ll Ce

c-Src

Raf-1

ERK1/2

AP-1

N uc leu

s GJC

Transcription DNA Neighboring cell

FIGURE 10.5 A summary of peroxynitrite-induced cellular signaling events (see text for details). Abbreviations: AMPK, AMP-activated kinase; ERK, extracellular signal-regulated kinase; GJC, gap junctional intercellular communication; JNK, c-Jun N-terminal kinase; PI3K, phosphoinositide 3-kinase; RTK, receptor tyrosine kinase (such as the EGF receptor or the PDGF receptor).

10.5 ACKNOWLEDGMENTS Research in the author’s laboratory is financially supported by Deutsche Forschungsgemeinschaft (Bonn, Germany; grants SFB 503/B1, SFB 575/B4, GRK 320).

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Oxide and Cell 11 Nitric Signaling: Redox Regulation of Ras Superfamily GTPases Jongyun Heo and Sharon L. Campbell University of North Carolina, Chapel Hill, North Carolina

CONTENTS 11.1 11.2 11.3 11.4

Introduction ............................................................................................264 Formation of Free Radical Species .......................................................265 Redox Potentials and Protein Target Sites of Free Radicals ................265 Ras Superfamily Redox-Sensitive GTPases ..........................................265 11.4.1 Redox-Sensitive Ras Superfamily GTPases ..............................266 11.4.2 Redox-Sensitive Rho and Rab Family GTPases .......................267 11.5 Mechanism of Redox-Active GTPase S-Nitrosylation .........................268 11.6 Characterization of NO-Mediated Guanine Nucleotide Exchange for Redox-Active GTPases ....................................................................273 11.7 Structural and Mechanistic Basis of NO-Mediated Guanine Nucleotide Exchange on Redox-Active GTPases in the Presence of O2 ................275 11.8 Superoxide Anion Radical Mediates Guanine Nucleotide Exchange on NKCD-Containing GTPases ...........................................279 11.9 Hydroxyl Radical Mediates Guanine Nucleotide Exchange on NKSD-Containing GTPases ..................................................................280 11.10 Structural and Mechanistic Basis for •NO2/O2• −- and OH•-Mediated Guanine Nucleotide Exchange on Rab and Rho GTPases ...................281 11.11 Role and Fate of Free Radical-Mediated GNE of Redox-Active GTPases .................................................................................................283 11.12 Redox Signaling and GEFs ...................................................................284 11.13 Concluding Remarks .............................................................................284 References ...............................................................................................285

263

264


11.1 INTRODUCTION The Ras superfamily consists of a number of small monomeric GTPases including Ras, Rho, and Rab subfamilies (1–4). These GTPases cycle between active GTP- and inactive GDP-bound states to regulate a diverse array of biological processes including cell proliferation, cell death, cell migration, and vesicular and nuclear transport (5–8). Hence, the GDP and GTP-bound states of Ras superfamily GTPases are highly regulated by multiple cellular factors given their critical cellular roles. Guanine nucleotide exchange factors (GEFs) (9–12) as well as free radicals facilitate exchange of GDP with GTP to produce the active GTP-bound form of the GTPase (13, 14), whereas GTPase-activating proteins (GAPs) enhance the slow intrinsic rate of GTP hydrolysis to produce the inactive GDP-bound form of the GTPase (11, 12, 15). In addition, guanine nucleotide inhibitors (GDIs) down-regulate the activity of a subset of GTPases (e.g., Rho, Rab subfamilies) by preventing membrane association as well as inhibiting guanine nucleotide dissociation (9, 12, 16, 17). Although the basic mechanisms by which GEFs, GAPs, and GDIs regulate their respective GTPase substrates have been revealed through numerous studies, the mechanism(s) of free radical-mediated GTPase activation is poorly understood and more complex than previously believed. A number of GTPases including H-, K-, and N-Ras contain redox-active residue(s) that are sensitive to free radical species, such as the nitric oxide (NO) derivative, nitrogen dioxide (•NO2), superoxide anion radical (O2• −), and hydroxyl radical (OH•). We have recently proposed a mechanism by which Ras is activated by NO (18). According to this mechanism, treatment of Ras with NO in the presence of O2 to produce •NO2, produces a Ras-thiyl radical intermediate. Consistent with this mechanism (18), previous studies have reported that the reaction of •NO2 with glutathione (GSH) and cysteine (CysSH) produces a glutathionyl radical (GS•) and cysteinyl radical (Cys-S•), respectively (19–21). Moreover, •NO2-mediated S-nitrosylation of GSH and CysSH proceeds through formation of a thiyl radical (i.e., GS• and Cys-S•), and this process is dominant over N2O3- or NO-mediated S-nitrosylation (19–21). A radical conversion process that generates a Ras-GDP guanine cation radical has been further implicated in facilitating release of Ras bound GDP (22). Although other mechanisms of NO-mediated Ras guanine nucleotide exchange (GNE), such as a non-radical-mediated transient structural change in the Ras GDP-binding site during the NO modification process, have not been ruled out, our data is most consistent with the proposed thiyl radical-mediated Ras GNE process. Hence, this chapter describes this radicalbased mechanism in detail. Furthermore, we postulate that although some of the free radical species may target distinct redox-active GTPase residues, the mechanisms of redox-mediated GNE are similar. In this context, the mechanism of free radical-mediated GTPase activation and the target specificity of the free radical toward the redox-active GTPases are reviewed.

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 265

11.2 FORMATION OF FREE RADICAL SPECIES Nitric oxide (NO) is produced by nitric oxide synthases (NOSs) (23), whereas reactive oxygen species (ROS) are produced by a variety of cellular metabolic and enzymatic processes (13, 14). In particular, NADPH oxidase and xanthine oxidase produce the ROS, superoxide radical (O2• −). O2• − can be converted into hydrogen peroxide (H2O2) by superoxide dismutase (SOD), which can be further converted to hydroxyl radical (OH•) by the transition metal (i.e., Fe)-catalyzed Harber–Weiss reaction (24). Moreover, various NO-derived reactive radical species (RNS), such as •NO2, can be formed by the reaction of NO with O2 as well as ROS (18). Figure 11.1 summarizes mechanisms of RNS and ROS formation. Both RNS and ROS contribute to a plethora of cellular and pathological processes including DNA damage and oxidative inactivation of certain proteins (13, 14).

A. NOS

NO

B. NADPH oxidase

O2

NO NO2

N2O3

SOD H2O2 O2 −

O2

−

+Η+ OH + O2 + H2O

FIGURE 11.1 Formation of RNS and ROS.

11.3 REDOX POTENTIALS AND PROTEIN TARGET SITES OF FREE RADICALS Although various redox-sensitive sites have been identified in cellular proteins (25), protein residues containing a thiol (R-SH) and hydroxyl (R-OH) group (e.g., cysteine and serine) are of interest in this chapter because these residues can form radical species upon reaction with RNS and ROS. By considering the redox potentials (oxidant/reductant) of •NO2/NO2−, O2• −/H2O2, R-O•/R-OH, R-O•/R-O, R-S•/RS−, and R-S•/R-SH at pH 7.0 (26), the target specificity of these RNS and ROS can be categorized as follows: •NO2 and O2• − can react with the thiolate form (R-S−) of the redox-active R-SH group to produce a protein-thiyl radical (R-S•), whereas OH• may react with the hydroxylate state (R-O−) of the R-OH group to produce a protein-hydroxyl radical (R-O•). Generation of protein radicals can alter the protein’s function and consequently its biological activity (vide infra).

11.4 RAS SUPERFAMILY REDOX-SENSITIVE GTPASES Small GTPases are present in eukaryotes, from yeast to human, and constitute a Ras superfamily consisting of more than 200 members (1). This superfamily is structurally classified into at least five sub-families: the Ras, Rab, Rho, Sar1/Arf, and Ran families (4–8). The Ras superfamily GTPases cycle between inactive GDP-bound and active GTP-bound states (8, 12). Although the Arf and Ran subclass of GTPases play critical roles in transport processes (27–30), this chapter

266


focuses on redox-sensitive GTPases that we have characterized, which include members of the Ras, Rab, and Rho subclass of GTPases. The guanine nucleotide state of Ras GTPases, is regulated by protein modulatory agents (8, 12). In particular, guanine nucleotide exchange factors (GEFs) facilitate exchange of GDP with GTP to promote GTPase activation (9, 10), whereas GTPase-activating proteins (GAPs) deactivate the GTPase protein by stimulating hydrolysis of bound GTP to GDP (11, 12). For other Ras superfamily GTPases, such as the Rho and Rab subclass, a distinct class of protein regulatory factors, guanine nucleotide inhibitors (GDIs), have been identified and characterized (12, 16, 17, 30). GDIs inhibit both membrane association and dissociation of GDP from their respective GTPases substrates to down-regulate GTPase activity. In addition to protein modulatory agents, such as GEFs, small molecule redox active agents play a role in regulating the GTPase activity of Ras. For example, biologically active free radicals (RNS and ROS) regulate Ras GTPase activity by stimulating GNE (13, 14, 31–34); however, the target specificities of RNS and ROS free radicals for Ras superfamily GTPases, and the mechanisms by which free radicals mediate activation of redox-active GTPases is not clear. Based on recent results obtained from our laboratory, two major classes of redox-sensitive GTPases, •NO2/O2• −- and OH•-sensitive GTPases, have been proposed. These free radicals are somewhat target specific as described below, in that a redoxactive R-SH group corresponding to a particular GTPase may be sensitive to •NO2 and O2• −, whereas both the redox-active R-SH and R-OH group in distinct GTPases are sensitive to OH•. Activation of Ras superfamily GTPases by targetspecific free radicals, in addition to GEFs, may be an important path to regulate a diverse array of cellular processes.

11.4.1 REDOX-SENSITIVE RAS SUPERFAMILY GTPASES Nearly all of Ras superfamily GTPases contain a guanine nucleotide-binding motif (NKXD motif) that specifically interacts with the guanine nucleotide base of GDP or GTP. Although the residue X in the NKXD motif is not well conserved in Ras superfamily members, a number of Ras and Rab GTPase family members contain a redox-active cysteine at this position, which we designate as the NKCD motif. We have recently proposed that NO gas (in the presence of O2) and xanthine oxidase (in the presence of its substrate xanthine and O2) form •NO2 and O2• −, respectively, and these radical species react with the Ras NKCD-containing cysteine to promote guanine nucleotide exchange on Ras. Within the Ras subfamily GTPases, Ras (H-, N-, and K-Ras), and Rap1 GTPases (both Rap1A and Rap1B) contain a redox-sensitive NKCD motif. Both Ras and Rap proteins has been implicated in a wide range of biological processes, from cell proliferation and differentiation to cell adhesion (4, 35, 36). For example, Ras targets directly or indirectly multiple downstream effectors such as phosphatidylinositol 3-kinase (PI3-K) and Raf kinase (37). Although the GTPase Rap is best characterized as a critical regulator of integrin-mediated cell adhesion and can influence the properties of other cell-surface receptors, its mechanism of action is poorly


understood (4, 35). Although Ras and Rap proteins can regulate distinct cellular effectors, they commonly regulate activation of the MAP kinase cascade in some cell types (38). Rab GTPases represent the largest subfamily, with over 60 distinct human Rab GTPases identified (39). These GTPases are best characterized for their role in regulation of vesicle trafficking (16, 39–41). A number of Rab GTPases including Rab1A and Rab3 (i.e., Rab3A, Rab3B, Rab3C, and Rab3D) contain a •NO2/O2• −-sensitive NKCD motif. However, some of the Rab GTPases, such as Rab2 (both Rab2A and Rab2B), contain a serine at position X in the NKXD motif, herein referred to as the NKSD motif, and may render the GTPase sensitive to OH•. Intriguingly, several yeast Rho GTPases, such as Sec4, also contain a NKSD motif that may be sensitive to OH•. The yeast Rho GTPases (i.e., Sec4, Rho1, Rho3, and Cdc42) play a key role in exocytosis (42). Although it has been previously demonstrated that Ras can be activated by NO both in vitro and in situ (endogenous) (31, 34, 43–47), a potential role for reactive free radicals, •NO2, O2• −, and OH•, in modulating redox-sensitive GTPase activity has not yet been clearly demonstrated. We have recently explored the mechanism by reactive radical species, such as •NO2, O2• −, and OH•, modulate the activity of select redox-active Ras superfamily GTPases in vitro, as described in this chapter. Although NO, generated endogenously from nNOS in neuronal cultures, reportedly enhances Ras activity (31), further studies will be required to determine how various redox agents directly alter redox sensitive Ras superfamily GTPases to regulate downstream signaling pathways, cellular processes, pathophysiological states, and stress responses. Such studies will also aid in understanding the complexities of free radical-mediated redox signaling in cells.

11.4.2 REDOX-SENSITIVE RHO

AND

RAB FAMILY GTPASES

A different class of redox-active GTPases, found mainly in the mammalian Rho and Rab family GTPases, contain a conserved redox-active cysteine, Cys18 (Rac1 numbering). Similar to the redox-sensitive NKCD motif found in Ras and Rap1A, Cys18 is sensitive to both •NO2 and O2• −. The cysteine is located at the end of the P-loop GxxxxGK(S/T) motif (residues 10 to 17, Ras numbering) and is conserved in nearly 50% of all Rho family GTPases. This P-loop-containing cysteine will be referred to herein as the GxxxxGK(S/T)C motif. Similar to Ras GTPases, mammalian Rho family GTPases are involved in regulating cell growth, differentiation, and cell death (5, 7–9, 48–51); however, these Rho GTPases are distinct in that they also regulate oxidant production as well as cell morphology through actin cytoskeletal rearrangements (51–56). RhoA regulates stress fiber formation, whereas Cdc42 and Rac1 are primarily involved in the formation of protrusive structures (49, 51, 55–57). Recently, it has been reported that Rac2 stabilizes the assembly of several proteins, including NADPH oxidase in phagocytes, which in turn triggers generation of bactericidal O2• − from NADPH oxidase (52). An isoform of Rac2, Rac1, may function to regulate the activity of NADPH oxidase in other cells (51–54). Because Rac regulates NADPH oxidase activity, and thus should be exposed to higher localized concentrations of O2• −, Rac may

268


be sensitive to regulation by O2• − via the redox active Cys18 residue. It is intriguing to speculate that O2• − may be involved regulating Rac activity through a feedback mechanism. Although many Rab GTPases contain either the •NO2/O2• −- or OH•-sensitive NKCD and NKSD motifs, respectively, an additional redox-active motif may also exist in several Rab GTPases. A putative •NO2/O2• −-sensitive redox-active GxxxxGK(S/T)C motif or OH•-sensitive redox-active GxxxxGK(S/T)S motif is also found where the GxxxxGK(S/T)S motif contains a serine in place of the redox-active cysteine in the GxxxxGK(S/T)S motif. Further studies will be necessary to understand whether free radicals mediate redox signaling of Rho and Rab GTPases in cells.

11.5 MECHANISM OF REDOX-ACTIVE GTPASE S-NITROSYLATION NO-mediated redox regulation is best characterized for the Ras GTPase (18), and we have recently proposed a mechanism, described in Figure 11.2 and Figure 11.3, for NO-mediated Ras GNE. According to this mechanism, NO reacts with O2 to produce •NO2, and •NO2 can further react with NO to produce dinitrogen trioxide (N2O3). The reaction is reversible, such that N2O3 can be degraded into and equilibrated with NO and •NO2. Therefore, NO, •NO2, and N2O3 coexist in a reaction mixture of NO and O2. Several studies have demonstrated that both •NO and N O can react with a variety of thiols (i.e., glutathione (GSH) and 2 2 3 cysteine) to produce S-nitroso compounds (RSNO). Yet, the mechanisms associated with RSNO- and N2O3-mediated thiol modification are proposed to be distinctively different. As depicted in Figure 11.2, thiol S-nitrosylation by •NO2 is radical-based, whereas S-nitrosylation by N2O3 follows a non-radical-based mechanism. A redox-sensitive S-nitrosylation site in H-Ras has been well characterized (46). Truncated H-Ras (residues 1–166) contains 3 cysteines. Among them, Cys118 (Ras-S118H) is solvent exposed and can be S-nitrosylated. We have recently reported that it is the process of Ras NO modification in the presence of O2, instead of the end product, RAS-SNO, that mediates Ras GNE (18). Moreover, we have proposed that a Ras thiyl-radical intermediate (Ras-S118•), formed by reaction of NO in the presence of O2, plays a key role facilitating Ras GNE (18). Ras Cys118 NO modification by one or both of the reaction products of NO and O2 (i.e., •NO2 or N2O3) could follow either the radical or non-radical acid–base mechanism depicted in Figure 11.2. Radical spin-trap reagents can be utilized to distinguish between these mechanisms, as radical spin-trap agents such as phenyl N-tert-butylnitrone (PBN) and 2-(4-Carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (carboxy-PTIO) can effectively scavenge NO and •NO2 (18). As illustrated in Figure 11.4, PBN and carboxy-PTIO effectively impede NO-mediated Ras S-nitrosylation in the presence of O2. These results suggest that NO/O2-mediated NO modification of H-Ras most likely

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 269 NO2− + H+

NO2

NO A. R

S H

R

R

S

S

NO

2 O + N N O

1

O B. R

S

NO2− + H+

3

H

R

S

NO

FIGURE 11.2 Possible mechanism of NO-mediated S-nitrosylation of thiol species in the presence of O2. R

O P-P

R

28

O Phe R R H O R OH OH H N H 415 O O Ser N H N HAla146 H H (i) P-P N H 119 O N7 OAsp H − + N1 −([NO2 ] + [H ]) O O O OH R H N N R 1/2 O2 O 118 NO NO2 S Cys H H

O R H OR R OH OH H N N H HN H H O H O N H O N7 N1 H O O OH N R HN O H

S +

(ii)

+ (H )

R

R

O

O H O P-P

H

OH OH H N H O N H N H H H N H O N7 O N1 H O O OH N R H N O + − (H ) ( i ii ) SH

R OR N H H O N H H H N H N H + O N7 N1 H O O OH N R H N O H

R R O

R

O

O R

OH OH

P-P

O

O R

H

R

O R

SH

R O H O

OH OH

P-P

O

H

R OR

R

N H H N H N H N7

N1 O R

O H H N H O O OH N N H O

O +( NO2 )

OH OH

P-P

N O

O

N1 NO2

R

P-P

OH OH N O

N7 O2N

O

H H N

N H NH

N H

O

(iv)

SH

O

H

N

N7

OH OH N

P-P O

N7 O2N

N N1

H N H

O

FIGURE 11.3 Proposed mechanism of NO-mediated Ras guanine nucleotide dissociation in the presence of O2. GDP is represented as in blue. The dotted line represents a putative hydrogen bond interaction between Ras residues and GDP. (Modified from Heo, J., Prutzman, K.C., Mocanu, V., and Campbell, S.L., J. Mol. Biol., 346, 1423–1440, 2005.)

270

Nitric Oxide, Cell Signaling, and Gene Expression 0.8 0.7

NO/O2 + wt Ras

Ras S-nitrosylation (mole)

0.6 0.5 0.4 0.3 NO only + wt Ras 0.2

NO/O2 + ascorbate + wt Ras

0.1 NO/O2 + PBN + wt Ras

0.0 −0.1

NO/O2 + wt Ras + Carboxy PTIO 0

1

2 NO (µM)

3

4

FIGURE 11.4 Quantification of Ras S-nitrosylation by NO in the presence and absence of O2. Various amounts of NO gas were introduced into anaerobically sealed assay cuvettes containing a transition metal-free buffer mixture (20 µM GDP, 5 mM MgCl2, and 50 mM NaCl in 20 mM mixed buffer pH 7.5). The quantity of NO adduct in the assay mixture was determined using the hemoglobin (Hb)-coupled NO assay. (From Heo, J., and Campbell, S. L. (2004), Biochemistry 43, 2314–22.) For reaction of NO in the presence of O2, the NO content in the assay mixture was determined by using the Hb-coupled NO assay, before addition of stoichiometric amounts of O2 (3NO:1O2, v:v) to produce a NO/O2 reaction mixture. The NO/O2 reaction mixture was then incubated for 10 min. When indicated, the spin-trapping reagent, Carboxy-PTIO (0.1 mM) or ascorbate (1 mM), and PBN (1 mM) was added to either the NO- or NO/O2-containing assay solution, and incubated for 1 min before transfer of wt Ras (0.5 µM). The content of PSNO was also measured using the Saville assay. (From Saville, B. (1958), Analyst 83, 670–2.) Saville reagent stocks were added to the PSNO sample mixtures in the following order: 1. N-(1-naphthyl)ethylenediamine dihydrochloride; 2. HgCl2; and 3. sulfanilamide (0.5 mL each). The mixture was incubated at room temperature for 10 min, and the absorbance was read at 540 nm. PSNO content was calculated according to a standard curve constructed with 0-5 µM NaNO2, and the plot fit to a simple exponential association. Values given in this figure represent mean values with standard errors obtained from measurements conducted in triplicate.

follows a radical-based mechanism but is insufficient to eliminate non-radicalbased N2O3-mediated Ras S-nitrosylation because, as noted elsewhere, N2O3 can be degraded into and equilibrate with •NO2 and NO. Thus, scavenging these radical species by spin-trap reagents may result in depletion of N2O3. Notably, the •NO2-mediated Ras S-nitrosylation mechanism suggests that a thiyl radical species, Ras-S118•, may be formed during the reaction process, which can further


react with NO to complete Ras S-nitrosylation. As depicted in Figure 11.2, if Ras NO modification follows a radical mechanism, a Ras-S118• intermediate is likely to be formed. The Ras-S118• formed in this reaction can react further with NO to produce S-nitrosylated Ras (Ras-SNO). Because ascorbate traps R-S• relative to other radical species (i.e., NO and •NO2), ascorbate should trap Ras-S118• and prevent Ras nitrosylation and NO-mediated Ras GNE in the presence of O2. Consistent with this premise, when ascorbate was present in a reaction mixture containing Ras, NO and O2, NO modification of Ras was blocked (Figure 11.4). This result supports a mechanism whereby reaction of Ras with a mixture of NO and O2 promotes Ras S-nitrosylation via a Ras-S118• intermediate, hence it is likely to follow a •NO2-mediated radical-based mechanism (Figure 11.2) (18). Moreover, previous studies utilizing small molecule thiols (e.g., GSH and CysSH) have demonstrated that S-nitrosylation of the GSH and Cys-SH occurs predominantly via thiyl-radical intermediates GS• and Cys-S• (19–21). Therefore, we proposed that in the presence of O2: 1. NO reacts with O2 to produce •NO2, which reacts with the redox-active Ras thiol to produce Ras-S•. 2. Ras-S• further reacts with NO to produce S-nitrosylated Ras (RasSNO). GSH is a cellularly abundant molecule that can react with NO produced from ROS to produce S-nitrosoglutathione (GSNO), and conceivably deliver NO to various target sites in cells (58). The mechanism of GSNO-mediated Ras S-nitrosylation is likely to be quite similar to that of NO-mediated Ras Snitrosylation in the presence of O2 because a GS• is formed by homolytic cleavage of GSNO (59). GS• then abstracts an electron from the solvent exposed redoxsensitive Ras thiol Ras-S118H to produce Ras-S118• and glutathione anion (GS−). Depending on the cellular pH, GS− may be then protonated to produce glutathione (GSH). It is likely that Ras-S118H is deprotonated prior to losing an electron to GS•, given the redox potentials of Ras-S118•/Ras-S118− and Ras-S118•/Ras-S118H with respect to GS•/GS−, (26). The formed Ras-S118•, in turn, could react with NO, a homolytic cleavage product of GSNO, to complete Ras S-nitrosylation. Consistent with this notion, ascorbate effectively inactivates GSNO-mediated and •NO - mediated GS• and Ras-S118• formation (vide supra) (18), and thus can block 2 radical exchange between GS• and the Ras Cys118 thiol (Figure 11.5). When the Ras Cys118 residue in the NKCD motif was replaced with serine (C118S), neither •NO2- nor GSNO-mediated Ras S-nitrosylation was observed (Figure 11.6), suggesting that Cys118 is indeed the target Ras S-nitrosylation site of •NO2 and GS• (derived from GSNO) (18). Thus, both •NO2- and GSNOmediated H-Ras (1–166) S-nitrosylation appears to occur through a Ras-S118• intermediate. Our proposed radical-based Ras S-nitrosylation mechanism (18, 22) should be applicable to other redox-active NKCD motif-containing GTPases, such as

272

Nitric Oxide, Cell Signaling, and Gene Expression 0.7

GSNO


0.5

GSNO + O2 0.3

GSNO + PBN 0.1

GSNO + ascorbate −0.1 0.0

0.5

1.0 GSNO (mM)

1.5

2.0

FIGURE 11.5 Quantification of Ras S-nitrosylation by NO and GSNO. Experimental conditions and data processing for GSNO-mediated Ras S-nitrosylation studies were identical to Figure 11.4, except that GSNO was used instead of NO, and an arbitrary amount of ambient O2 (100 µL) was added when indicated. GSNO-mediated S-nitrosylated wt Ras and unreacted GSNO have similar absorption intensity at 542 nm, so when GSNO was used, samples were applied to a size exclusion column (Sephadex G-25) to remove unreacted GSNO. (From Heo, J., and Campbell, S. L. (2004), Biochemistry 43, 2314–22.) Gel filtration was performed within 2 min under anaerobic conditions. The absorption intensity at 542 nm was scaled by a dilution factor because gel filtration causes dilution of the sample. The dilution factors (1.5–1.9) for each individual protein sample was determined using the Bradford protein assay (88) subsequent to gel filtration. The fraction of S-nitrosylated Ras was determined from the absorbance at 542 nm (18]) with the peak intensity plotted against various NO concentrations. The plot was fit to a simple exponential association. Values given in this figure represent mean values with standard errors obtained from measurements conducted in triplicate.

Rap1 and Rab3A. Consistent with this premise, ascorbate prevents •NO2- and GS•-mediated S-nitrosylation of Rap1 and Rab3A (data not shown). Although we favor a thiyl radical-based S-nitrosylation mechanism for Ras, other possible mechanisms cannot be discounted at this time. For example, it has been proposed that reaction of NO with a thiol (R-SH) in the absence of O2 can produce a R-S-•N-O-H radical intermediate, which in turn is converted to R-SNO in the presence of an electron acceptor (e.g., NAD+) (60). However, we had previously observed that a minimal amount of Ras-SNO was formed under anaerobic conditions in the presence of NAD+ and NO gas (18), and postulated this mechanism (60) is unlikely to contribute significantly to NO-mediated Ras

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 273 0.08


0.06 ΝΟ/Ο2 + wt Ras

0.04

0.02 ΝΟ/Ο2 + C118S Ras

0.00

0

1

2

3

4

NO (µM)

FIGURE 11.6 Quantification of wt and C118S Ras S-nitrosylation by NO in the presence of O2. Experimental conditions were identical to that of Figure 11.4, except that the Ras variant C118S (0.5 µM) was used in addition to wt Ras. (Modified from Heo, J., and Campbell, S. L. (2004), Biochemistry 43, 2314–22.)

nitrosylation. The small amount of S-nitrosylated Ras observed in our studies, is most likely due to trace amounts of O2 contamination.

11.6 CHARACTERIZATION OF NO-MEDIATED GUANINE NUCLEOTIDE EXCHANGE FOR REDOX-ACTIVE GTPASES NO modification of Ras has been proposed to activate Ras by promoting guanine nucleotide exchange thus populating Ras in its biologically active GTP-bound state in vivo (vide supra) (46). Our recent biochemical and biophysical studies provided substantive evidence that reaction of the Ras Cys118 thiol side chain with either •NO or GS• generates a Ras radical intermediate, Ras-S118•. It is the Ras thiyl 2 radical intermediate, rather than the S-nitrosylation end product (Ras-SNO), that promotes Ras GNE (vide supra) (18, 22). Although our studies indicate that RasSNO does not directly promote Ras GNE, the role of Ras-SNO is unclear. It is possible that formation of Ras SNO may prevent further radical reaction processes (vide infra) because reaction of the Ras radical intermediate Ras-S118• with NO to produce Ras-SNO will eliminate the Ras radical species (Figure 11.2). It is also possible that formation of Ras-SNO protects Ras from additional radical-mediated

274


Ras GNE because neither •NO2 nor GS• are likely to react with Ras-SNO to produce a Ras-radical species. Alternatively, Ras-SNO may be removed from Ras by reaction with cellular redox agents, such as abundant thiols (i.e., GSH), allowing regeneration of Ras for further redox-mediated regulation of Ras GNE. It is also conceivable that Ras-SNO may play a role in Ras-mediated cell signaling by serving as a target recognition site for protein modulators or downstream effectors. Further follow-up studies are necessary to understand the role of the SNO moiety in Ras function. As depicted in Figure 11.2, a H+ is released as a byproduct of •NO2- and GSNOmediated NO modification. It is possible that H+s produced from reaction of Ras with NO may also represent a possible Ras perturbation agent, in that production of H+s during the reaction may lower the local pH and interfere with Ras guanine nucleotide binding interactions ; however, NMR studies (61) indicated that pHdependent guanine nucleotide binding interactions were primarily associated with the Mg2+-binding residues instead of residues in the Ras NKCD and SAK motifs (Figure 11.7). These results suggest that it is the Ras-S118• intermediate, instead of the H+ byproduct or the Ras-Cys118-SNO end product of the reaction between NO/•NO2 and Ras, that plays a key role in NO-mediated Ras GNE. •NO can also facilitate GNE on other redox-active GTPases that contain the 2 NKCD motif. As illustrated in Figure 11.8, treatment of Rap1A and Rab3A with a reaction mixture containing NO and O2 facilitates dissociation of the GTPasebound guanine nucleotide. Similar to Ras, ascorbate impedes •NO2-mediated guanine nucleotide dissociation for both of these redox-active GTPases. Hence, our data indicates that the formation of a GTPase-thiyl radical (GTPase-S•) intermediate promotes NO-mediated GNE for Rap1A, Rab3A, as well as H-Ras NKCD-containing redox-active GTPases. In summary, we have proposed that the reaction of •NO2 with the redox-active cysteine in NKCD-containing GTPases leads to production of a radical GTPaseS• intermediate, which in turn promotes GNE. Thus, NO-mediated GNE of NKCD-containing Ras superfamily GTPases may represent a radical-based mechanism of GTPase activation in vivo, which is distinct from GEF-mediated GNE. The proposed mechanism consists of two steps: 1. The generation of a GTPase-S• radical intermediate that perturbs GTPase-guanine nucleotide binding interactions to enhance GTPase guanine nucleotide dissociation 2. Quenching of the GTPase-S• radical, allowing rebinding of guanine nucleotide ligands to the GTPase, hence completing Ras GNE GSH is present at high cellular concentrations and may serve as a GTPaseS• radical quenching agent, allowing reversible binding of guanine nucleotides. Yet, further studies are required to delineate the actual cellular quenching agents that allow rebinding of GTP after release of GDP to complete GNE and GTPase activation.


R

N R

R

57

O H Asp H O 58 Thr

N H O

O

H Ala

59 N

O

O

L

H

O H

Mg L

2+

O O OH P

P

35 H

H R

15

Gly N R O H N H R

O

R

O

OH

H Lys16 N

H

H

H

O

+ H

H

O

H O R

O

N R

145

H

− O Asp119 H N

N

H

H

O Ser H

N

O H

O N

N H

N

116 N Asn

R

H

R

N1

H H

28

O Ala146 R O R

H

N7

H 17 Lys N + H

Phe H

N

H

N H O

H

O

Thr

O

O

O

O

N

P O

OH O

Val29 N

H

N H

R

N R H Ser17 − O

O

N

O

N O

H

R

Asp30

−O

H

O

H

R

O

R

O

O

O H

O

N

S

118

Cys H

FIGURE 11.7 Schematic view of Ras-GDP (GTP) interactions. pH-sensitive amides within Ras are shown (0.20 ppm). Mg2+ ion and its coordination with water molecules are shown, whereas GDP is shown, and the γ-phosphate of GTP is shown. Hydrogen-bond interactions are shown as dotted lines. Aromatic–aromatic interaction between the Phe28 side chain and guanine nucleotide base is shown ≈, and putative electrostatic repulsive interactions between Val29-Asp30 are symbolized as. Mg2+ coordinates six ligands in the Ras NMR solution structure (PDB 1AA9); some of the Mg2+-coordinated ligands were expressed as L. Nucleotide interactions with Ras residues Lys117 and Lys147 were omitted for convenience. The schematic presentation of Ras residues involved in interactions with Mg2+•guanine nucleotide ligands (Mg2+•GDP and Mg2+•GTP) is based on the NMR solution structure PDB 1AA9 (GDP•Mg2+•Ras complex) and X-ray crystal structure 1QRA (GTP•Mg2+•Ras complex), although atomic distances and their relative positions were changed arbitrarily for presentation. (From Heo, J., Gao, G., and Campbell, S. (2004), Biochemistry 43, 10102–11.)

11.7 STRUCTURAL AND MECHANISTIC BASIS OF NO-MEDIATED GUANINE NUCLEOTIDE EXCHANGE ON REDOX-ACTIVE GTPASES IN THE PRESENCE OF O2 Ras binds guanine nucleotide ligands with high affinity due to multiple electrostatic (including hydrogen-bond interactions) and hydrophobic interactions. One of the interactions involves Phe28 (Ras numbering), which is strictly conserved in Ras superfamily GTPases. The Phe28 side chain ring packs perpendicularly with the guanine base via n–π stacking interactions. Reportedly, mutation of the Ras Phe28 residue to leucine drastically reduces the Ras guanine nucleotide-binding affinity

276


Fluorescence intensity (fraction)

1.0

0.9

0.8 Rap1A

Ras

Rab3A

0.7

0

500

1000 Time (sec)

1500

2000

FIGURE 11.8 Guanine nucleotide dissociation of Ras, Rap1A, and Rab3A in the presence of NO and O2. Fluorescence assay for NO-mediated Ras guanine nucleotide dissociation in the presence of O2, employing 2′-(or-3′)-O-(N-methylanthraniloyl)guanosine 5′-diphosphate (mant-GDP) as described previously (18).

and can promote Ras transformation in NIH 3T3 cells (62, 63). Hence, the n–π interaction between Phe28 and the GTPase bound guanine nucleotide base appears critical for high affinity Ras binding. Inspection of the available structures for NKCD motif-containing GTPases reveals that the center of the Phe28 side chain faces the sulfur atom of NKCD motif-containing Cys118 thiol (Ras numbering). Moreover, the distance, ~12 Å, between the center of Phe28 side chain and the sulfur atom associated with NKCD motif, is conserved in these NKCD motif-containing GTPases (Figure 11.9A). In many biological systems, redox coupling within and beyond this key distance is effective in promoting various cellular events (64). Intriguingly, when other non redox-active residues (e.g., valine or alanine) are present in the NKXD motif instead of the redox-active cysteine, the spatial orientation of the redox-inactive side chains does not face the Phe28 side chain (Figure 11.9B). We have recently investigated the role of Phe28 in NO-mediated Ras GNE. When Phe28 was replaced with leucine (F28L), •NO2- or GS•-mediated Ras guanine nucleotide dissociation was not observed (Figure 11.10). This kinetic result indicates that generation of the Ras-S118• intermediate promotes Ras guanine nucleotide dissociation through a mechanism involving Ras Phe28. Moreover, we have employed various biochemical, mass spectrometry, and UV visible spectroscopic analyses, and demonstrated that wt Ras-bound GDP is converted to a free 463.2 Da nitration product upon treatment of wt Ras with •NO2 (22). Based on these

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 277 A

B

F28 Nucleotide

F28 ~12 Å

Nucleotide C118

N116K117C118D119

N116K117V118D119

FIGURE 11.9 Spatial architecture of the Phe28 side chain and the NKCD motif of GTPases with respect to bound nucleotide. (A). The distance and spatial orientation of the Phe28 side chain, nucleotide ligand, and Cys118 thiol group in the NKCD motif of Rap1A is presented. (B). The spatial orientation of the Phe28 side chain, nucleotide ligand, and Val118 side chain in the NKVD motif of Rap2A is depicted. The scheme was generated by using RASMOL (86) with PDB 1C1Y for the NKCD motif-containing Rap1A GTPase and PDB 1KAO for the NKVD motif-containing Rap2A GTPase.

analyses, the 463.2 Da nitration product has been assigned as 5-guanidino-4nitroimidazole diphosphate (NIm-DP) (22). It has been previously reported that 5guanidino-4-nitroimidazole (NIm), which lacks the ribose and diphosphate compared with NIm-DP, can be produced by reaction of a radical form of the guanine base with •NO2 (65–70). In particular, a carbonate radical anion (CO3• −) can be formed by pulse radiolysis or laser photolysis, which in turn abstracts an electron from guanine base to produce a guanine nucleotide cation radical (G• +). G• + can be also produced by the treatment of peroxinitrite (ONOO−) with guanine (68). G• + is converted to a neural guanine nucleotide radical (G•) by elimination of a H+ from the N1 of G• +, and •NO2 can react with G• to produce a guanine base adduct, which is subsequently degraded to NIm (69). Given that the formation of NIm-DP is dependent on generation of a radicalized guanine nucleotide base moiety and that treatment of Ras-bound GDP with •NO2 generates NIm-DP, it is reasonable to propose that •NO2 facilitates Ras guanine nucleotide dissociation via radicalbased conversion of Ras-bound GDP into free NIm-DP (22). Consistent with this radical-based mechanism, formation of NIm-DP from wt Ras is blocked by addition of the radical quencher, ascorbate, before the treatment with •NO2. Moreover, NImDP was not detected when the Ras variants C118S and F28L were treated with •NO , suggesting that the Phe28 side chain as well as the Ras Cys118 thiol is involved 2 in radical-based conversion of Ras-bound GDP into free NIm-DP. Yet, the relative spatial orientation of the Ras residues Cys118 and Phe28 appears to be important because a treatment of •NO2 with a reaction mixture of free GDP, free phenylalanine, and free GSH (a source of cysteine) does not produce NIm-DP (22).


Flourescence intensity (fraction)

278 1.0

wt Ras C118S C118S + NO/O2

0.9

F28L F28L + NO/O2

0.8

0.7

wt Ras + NO/O2

0.6

0.5

0

100

200

300

Time (sec)

FIGURE 11.10 Dissociation of guanine nucleotide from wt Ras, F28L, and C118S by NO in the presence of O2. The assay procedure for NO-mediated GTPase guanine nucleotide dissociation in the presence of O2, employing 2′-(or-3′)-O-(N-methylanthraniloyl)guanosine 5′-diphosphate (mant-GDP) has been previously described (18). NO gas (~ 2 µM) was transferred into O2-free sealed assay cuvettes containing assay buffer at pH 7.5. The NO content was determined before its addition to protein samples followed by previous method (18). Where indicated, •NO2 is produced by addition of a stoichiometric amount of ambient O2 (2NO:1O2, mole:mole) into the NO-containing assay solution. Fluorescence mant-GDPloaded Ras (0.5 µM) was added and the decrease in fluorescence emission at 460 nm was recorded as a function of time. Rates of apparent NO/O2-mediated mant-GDP dissociation of 0.09 × 10−3, 1.82 × 10−3, and 3.34 × 10−3 s−1 for Ras C118S, Ras F28L, and wild-type Ras, respectively, were determined by fitting the data to a simple exponential decay. For control, intrinsic GDP dissociation rates from wt Ras and its variants C118S and F28L were measured. Rates of intrinsic GDP dissociation corresponding to 0.04 × 10−3, 1.38 × 10−3, and 0.02 × 10−3 s−1 for Ras C118S, Ras F28L, and wild-type Ras, respectively, were also determined by fitting the data to a simple exponential decay. (From Heo, J., Prutzman, K. C., Mocanu, V., and Campbell, S. L. (2005), J. Mol. Biol., 346, 1423–1440.)

Based on this work and our analyses, we have proposed the following mechanism of NO-mediated Ras guanine nucleotide dissociation in the presence of O2 (Figure 11.3): 1. NO reacts with O2 to produce •NO2, which in turn reacts with the Ras Cys118-SH to produce a Ras-S118• radical intermediate, and H+ may be released from Cys118-SH prior to reaction with •NO2.


2. Given the relative distance, orientation, and redox potential between the Ras-S118• and the Phe28 side chain, the Ras-S118• then withdraws an electron from the side chain of Phe28 to produce a Phe28 side chain cation radical and a Cys118 thiolate (Ras-S118−). Ras-S118− can be protonated to produce Ras-S118H at physiological pH. The thermodynamically unstable Phe28 side chain cation radical immediately abstracts an electron from the guanine nucleotide base to produce a guanine nucleotide cation radical intermediate (G• +-DP). 3. G• +-DP is then converted to a neural guanine nucleotide radical (G•DP) by an elimination of H+ from the N1 of G• +-DP, resulting in disruption of key hydrogen-bond interactions. We postulate that formation of a guanine nucleotide radical perturbs hydrogen bond interactions between the guanine N1 and Asp119 side chain, the G•-DP C6 oxygen and the Ala146 amide as well as the n–π interaction between the Phe28 side chain and the guanine base. Perturbations of these interactions are likely to trigger dissociation of the G•-DP ligand from Ras. 5. G•-DP can then react with •NO2 to produce a free GDP-NO2 adduct. The release of GDP-O2 adduct from Ras produces GDP-deficient Ras (apo Ras). 6. Given the cellular abundance of GTP, by mass action, apo Ras is likely to become GTP-bound, leading to Ras activation in situ. We have conducted our studies in the presence of excess •NO2 to aid in the characterization of Ras-released guanine products in the presence of NO gas, similar to studies previously conducted to characterize guanine base adducts exposed to •NO (65–70); however, amounts of •NO in the cell may be limited and will depend 2 2 on cellular conditions. Consequently, other guanine cation radical quenching agents, such as O2, may produce alternative adducts such as oxygenated-GDP adducts (i.e., GDP-O2). Another possibility is that, if ascorbate is present, G•-DP may be quenched by ascorbate to produce a GDP isomer, 2-amino-5H-purine-6-one ribose diphosphate (22); however, characterization of cellular-based •NO2-mediated Ras GNE and its end product(s) will be required to determine the fate of free GDP-O2 adduct released from GTPases in the cell. Given the structural conservation of Phe28 and the NKCD motif in other Ras superfamily GTPases, such as Rab3A and Rap1A, the mechanism by which the Phe28 side chain mediates electron transfer from CysS118• to the guanine nucleotide base to perturb Ras guanine nucleotide interactions may be common to all NKCD-containing GTPases.

11.8 SUPEROXIDE ANION RADICAL MEDIATES GUANINE NUCLEOTIDE EXCHANGE ON NKCD-CONTAINING GTPASES In addition to •NO2, O2• − can promote guanine nucleotide dissociation from NKCD motif-containing GTPases. When a fluorescent-GDP derivative, mant-

280


GDP was loaded onto various redox-active NKCD motif-containing GTPases (i.e., Ras, Rap1A, and Rab3A) and treated with O2• −, GTPase-bound mantGDP was dissociated and the dissociation rates were followed by fluorescence spectroscopy. These rates were found to be similar to •NO2-mediated Ras guanine nucleotide dissociation (71). In the assay, xanthine oxidase was used in the presence of the enzymatic reaction substrates, xanthine, and O2, to produce O2• − (72). The basic mechanism of O2• −-mediated guanine nucleotide dissociation from the NKCD motif-containing GTPases is similar to that of NO-mediated Ras guanine nucleotide dissociation, in that a GTPase-S118• radical intermediate is generated resulting in GNE; however, the reaction end product differs. Briefly, (i) O2• − reacts with the GTPase Cys118-SH to produce a GTPase-S118• radical intermediate. Cys118-SH may be deprotonated before reaction with O2• − to produce GTPase-S118•. For Steps (ii and iii), the GTPaseS118• then withdraws an electron from the guanine nucleotide base to produce G• +-DP. Similar to the mechanism described for NO-mediated Ras GNE in the presence of O2 (Figure 11.3), the Phe28 side chain serves as an electron conduit for this process. G• +-DP is thus expected to be formed and converted to G•-DP by elimination of H+ from the N1 atom of G• +-DP. The process disrupts key hydrogen-bond interactions as well as the n–π interaction between the GTPase and its ligand nucleotide, (iv) G•-DP can then react with O2• − to produce a GDP-OOH adduct, which may be further degraded into oxygenatednucleotide products (71). Similar to NO-mediated Ras GNE in the presence of O2, O2• −-mediated formation of GDP-OOH produces an apo form of the GTPase. Given the GTP/GDP ratios in cells, the apo GTPase may be exchanged with cellularly abundant GTP, leading to GTPase activation in situ. As described previously for •NO2-mediated Ras guanine nucleotide dissociation, depending on the cellular conditions, other radical quenching agents, such as O2 and ascorbate, may react with G•-DP to produce alternative products, such as a GDP-O2 adduct and a GDP isomer 2-amino-5H-purine6-one ribose diphosphate, respectively (22,71). To address the fate of free G•-DP released from Ras in the cell, cellular-based O2• −-mediated Ras GNE and its end product(s) will need to be characterized.

11.9 HYDROXYL RADICAL MEDIATES GUANINE NUCLEOTIDE EXCHANGE ON NKSD-CONTAINING GTPASES In addition to the •NO2/O2• −-sensitive NKCD-containing GTPases, OH•-sensitive NKSD-containing GTPases, such as Sec4, possess an identical structural configuration (i.e., distance and the spatial orientation) to that of the •NO2/O2• −-sensitive NKCD-containing GTPases, for redox coupling between Ser135 (Sec4 numbering: equivalent to Cys118 of Ras) and its counterpart Phe45 (Sec4 numbering: equivalent to Phe28 of Ras). Therefore, GTPases, such as Sec4, which contain a NKSD motif, may be sensitive to OH•-mediated GNE by a mechanism similar to •NO2/O2• −-


mediated GTPase GNE. The primary difference, however, is that OH•, instead of •NO /O • −, reacts with the Ser135 hydroxyl group (Sec4 numbering: Sec4-O135-H) 2 2 to produce a Sec4 hydroxyl radical (Sec4-O135•), which then interacts with the Phe45 side chain to produce a Phe45 cation radical. Similar to the NKCD-containing GTPases (e.g., Ras), the Phe45 side chain cation radical may interact with the Sec4bound guanine nucleotide base to produce G• +-DP, which in turn may be converted to G•-DP. Generation of both a Phe cation radical and subsequent conversion of G• +-DP to G•-DP is likely to disrupt critical hydrogen-bond interactions and the n–π interaction between the GTPase and its bound guanine nucleotide ligand, resulting in enhanced guanine nucleotide dissociation (vide supra). The freed G•DP may then react with OH• to product the GDP-OH adduct. Intriguingly, GNE associated with the Ras variant, Ras C118S, which contains a serine in place of Cys118 in the NKCD motif, is largely enhanced by OH•, but not by •NO2, indicating that the free radical specificity associated with GTPase guanine nucleotide dissociation is dependent on both the placement and type of redox-active residue contained within the GTPase. In particular, a protein-thiyl radical (Ras-S118•, in case of Ras) and a protein-hydroxyl radical (Sec4-O135•, in case of Sec4) may be generated by •NO2/O2• − and OH• free radicals, respectively. OH•-mediated modulation of GTPase activity may be operative in the cell under oxidative stress or pathophysiological conditions, as opposed to modulation of cell signaling activity under physiological conditions.

11.10 STRUCTURAL AND MECHANISTIC BASIS FOR •NO /O • −- AND OH•-MEDIATED GUANINE 2 2 NUCLEOTIDE EXCHANGE ON RAB AND RHO GTPASES As discussed previously, a distinct redox-sensitive motif, GxxxxGK(S/T)C motif, is found in Rho family GTPases. The redox-active thiol residue Cys18 in the GxxxxGK(S/T)C motif is ~3.6 Å away from Phe28 (Rac1 numbering, same as the Ras numbering for Phe28) in Rac1, Cdc42, and RhoA (Figure 11.11). In structures of Rho sub-family GTPases, the side chain of Phe28 interacts with the guanine nucleotide ligand and packs in a perpendicular arrangement with the guanine nucleotide base (73–78). We have recently generated evidence that GS• and •NO2/O2• − enhance guanine nucleotide dissociation for the GxxxxGK(S/T)C motif containing Rac1, RhoA and Cdc42 Rho family GTPases (data not shown), using assay methods similar to those described for NKCD-containing GTPases (18, 22, 71). Although, we can demonstrate reactivity of redox-active agents with redox sensitive GTPases in vitro, it is not clear whether these agents are co-localized with the GTPases at concentrations required for physiological regulation. We speculate that because the Rho family GTPase, Rac, co-localizes and modulates NADPH oxidase activity (51–54), O2• − may stimulate guanine nucleotide dissociation from Rac in vivo. The fundamental mechanistic process for free radical O2• −mediated GNE associated with these GxxxxGK(S/T)C motif-containing GTPases

282


C18 F28 ~3.6 Å

Nucleotide

T115K116L117D118

FIGURE 11.11 Spatial architecture of the Phe28 side chain, Cys18 side chain and the nucleotide substrate within the GxxxxGK(S/T)C-containing motif of Rac1. The scheme was generated by using RASMOL (86) with PDB 1MH1.

is anticipated to be similar to that of O2• −-mediated GNE of NKCD-containing GTPases (vide supra). The only difference lies in the initial thiyl-radical formation site of the GTPases. In the GxxxxGK(S/T)C motif-containing GTPases, the reaction of Cys18 thiol with O2• − produces a thiyl radical on the side chain of Cys18 (GTPase-S118•), whereas in NKCD motif containing GTPases, the reaction of the Cys118 thiol with O2• − produces a thiyl radical on the side chain of Cys118 (GTPase-S118•). The radical electron associated with the Cys18 side chain is likely to propagate to the GTPase-bound guanine nucleotide ligand via the Phe28 side chain to produce G•-DP. Formation of G• +-DP from G•-DP, in turn, disrupts hydrogen-bond interactions between the GTPase and its ligand G•-DP to produce the apo form of the GTPase and free G•-DP. The apo GTPase can be activated upon binding GTP, as GTP is present in excess of GDP in situ, whereas G•-DP reacts with O2• − to produce oxygenated-nucleotide products. Because the distance between the Phe28 side chain and the redox-active thiol of Cys18 in GxxxxGK(S/T)C motif containing GTPases is approximately fourfold shorter than the Phe28 side chain and redox-active thiol of Cys118 in the NKCD motif of Ras, redox coupling between the GxxxxGK(S/T)C motif in RhoA and Phe28 side chain is expected to be more efficient than observed for Ras. Therefore, an enhancement of •NO2/O2• −-mediated GNE for GxxxxGK(S/T)C-containing GTPases is expected relative to NKCD- or NKSD-containing GTPases. In support of this premise, the rate of GS • -enhanced GNE of Rac1, one of the GxxxxGK(S/T)C motif-containing GTPases, is at least twofold faster compared with that observed for Ras or Rap1A NKCD motif-containing GTPases (Figure 11.12).

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 283 1.0

Fluorescence intensity (fraction)

Rac1 and Ras

0.9

Rac1 + 1 mM GSNO

0.8 Ras + ~1 mM GSNO 0.7

0.6

0

100

200

300

Time (sec)

FIGURE 11.12 Guanine nucleotide dissociation from Rac1 in the presence of GSNO. The fluorescence assay for GSNO-mediated GTPase guanine nucleotide dissociation, employing 2′-(or-3′)-O-(N-methylanthraniloyl)guanosine 5′-diphosphate (mant-GDP) has been previously described (18).

In some Rab GTPases, a redox active Ser18 is found in the GxxxxGK(S/T)S motif. Because reaction of OH• should promote formation of GTPase-O• in NKSD-containing GTPases (vide supra), GNE of the redox-active Ser18-containing GTPases may also be enhanced by OH•.

11.11 ROLE AND FATE OF FREE RADICAL-MEDIATED GNE OF REDOX-ACTIVE GTPASES We have recently determined that treatment of the NKCD motif-containing GTPase, H-Ras with NO/O2 produces an end product, which appears to be NIm-DP, whereas addition of O2• − (which is produced by xanthine oxidase) to NKCD and GxxxxGK(S/T)S motif-containing GTPases produces oxygenated nucleotide products. NIm analogs have been useful for tumor diagnostics and suppression, as these compounds target and inhibit development of tumor hypoxic cells (79–83). It is intriguing to speculate that treatment of NO-sensitive GTPases with •NO2 may lead to production of NIm-DP, which may, in turn, promote or inhibit additional cellular processes. In contrast to the NIm analogs, the role(s) of oxygenatednucleotide products in cells have not been investigated. Further studies are required to assess whether production of these modified GDP products affects cellular function.

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11.12 REDOX SIGNALING AND GEFS Most Ras superfamily GTPases are regulated by specific GEFs, yet among them, only a subset of these GTPases are redox active. Although free radical-mediated regulation of GTPase activity may be an adjunctory regulatory mechanism for redox-active GTPases, cellular conditions required for redox- and GEF-mediated regulation of redox-sensitive GTPases are likely to differ, and may represent important yet distinct mechanisms for GTPase regulation. Thus, redox and GEFmediated regulation of redox-sensitive GTPases could act separately or synergistically, as discussed in more detail next. Mechanisms of GTPase activation by free radicals and GEFs are distinctively different. Analysis of Ras and other GTPase crystal structures indicate that binding of the GEF, SOS, to the Ras GTPase switch I and II regions, induces conformational changes in Ras leading to disruption of multiple interactions between Ras and its nucleotide ligands (84, 85). The Ras GTPase switch regions are involved in interactions with Mg2+-phosphate groups and the ribose of bound GDP and GTP. Thus, interactions between Ras and SOS are likely to primarily effect binding interactions between the Ras switch regions and the guanine nucleotide ligand to facilitate GNE. In contrast to GEFs, reaction of a free radical, such as ROS or RNS, with a redoxactive GTPase appears to generate a guanine nucleotide radical (G•-DP) that disrupts key binding interactions with the guanine nucleotide base. If both the GEF and free radical act together to promote GTPase activation, it is possible that either one of these activating agents may be dominant or they synergistically activate GTPase activity. The combined effects of both GEFs and free radicals on GTPase activity have not been investigated. Both GEFs and production of free radicals are known to be highly regulated, and free radical-mediated GEF activity may represent yet another level of regulation.

11.13 CONCLUDING REMARKS Various RNS and ROS can interact with target specific Ras superfamily GTPases to promote redox-regulation of GTPase activity in situ. Although reactive free radical species are well known for their role in oxidative damage and pathophysiological modulation of cellular processes, RNS and ROS may act as second messengers to regulate a multitude of GTPases and GTPase-mediated cellular processes. Unlike, GEFs, RNS and ROS are not enzymes and do not regulate GTPase targets through a “lock and key” recognition system but instead through a chemical recognition process. This chapter has described the structural and mechanistic basis for redox regulation of Ras superfamily GTPases. Future studies are needed to elucidate how the GTPases are modulated by redox agents in cells, and whether redox regulation of specific Ras superfamily GTPases act in conjunction with other regulatory proteins, such as GEFs, as well as GAPs and GDIs to modulate GTPase activity and GTPase-mediated processes in situ.


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Oxide and the 12 Nitric Hypoxia Inducible Factor-1 Transducing System Jie Zhou and Bernhard Brüne University of Frankfurt Medical School, Frankfurt, Germany

CONTENTS 12.1 Introduction ............................................................................................291 12.1.1 Abbreviations .............................................................................292 12.2 Formation and Signaling Properties of NO with Importance for Gene Activation ............................................................292 12.3 Stability Regulation of HIF-1α and Activation of HIF-1 .....................294 12.3.1 Lessons from Hypoxia ...............................................................294 12.3.2 NO: Stabilization of HIF-1α and Activation of HIF-1 under Normoxia .........................................................................298 12.3.3 Destabilization of HIF-1α: The Role of NO .............................301 12.4 Medical Implications: RNI and HIF-1 in Tumor Biology ...................303 12.5 Concluding Remarks .............................................................................304 12.6 Acknowledgment ...................................................................................305 References ...............................................................................................305

12.1 INTRODUCTION It is widely accepted that nitric oxide (NO) is an important signaling molecule that shapes several physiological and patho-physiological processes. Among multiple activities attributed to NO, we find regulation of gene expression. Although no evidence exists regarding whether direct NO-responsive DNA elements within promotor regions of eukaryotic genes, numerous transcription factors are affected either resulting in increased or decreased expression of target genes. In part, this is compatible with the interference of NO with signaling circuits upstream of transcription factors that, in turn, will modulate their activity. A characteristic feature of several transcription factors is their redox sensitivity as well as their 291

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low protein abundance in unstressed cells due to efficient 26S proteasomal degradation. One example is the hypoxia inducible factor-1α (HIF-1α) known as the master regulator allowing adaptation toward decreased oxygen availability. We now appreciate that NO under normoxic conditions mimics a hypoxic response by stabilizing HIF-1α. Under hypoxic conditions, however, NO destabilizes HIF-1α and thus reverses hypoxic adaptation. This chapter summarizes recent molecular understanding of how NO affects stability regulation of HIF1α under normoxia vs. hypoxia and discusses patho-physiological consequences. Targeting HIF-1α by NO expands the sphere of NO actions with medical-related consequences for conditions such as ischemia/reperfusion, angiogenesis, or tumor biology.

12.1.1 ABBREVIATIONS AhR Aryl hydrocarbon receptor ARNT AhR nuclear translocator cGMP cyclic guanosine monophosphate CTAD C-terminal transactivation domain DFX desferioxamie EDRF endothelium derived relaxing factor FIH factor inhibiting HIF GSNO S-nitrosoglutathione HIF-1 hypoxia inducible factor-1 HRE hypoxia-response element MAPK mitogen activated protein kinase Mdm2 murine double minute NOC-18 Z-1-1[2-aminoethyl-amino]diazen-1ium-1,2-diolate (i, n, e)NOS (inducible, neuronal, endothelial) nitric oxide synthase ODD oxygen-dependent degradation domain PHD prolyl hydroxylase domain-containing protein PI3K phosphatidyl inositol 3 kinase pVHL von Hippel-Lindau protein RNI reactive nitrogen intermediates ROI reactive oxygen intermediates SNP sodium nitroprusside VEGF vascular endothelial growth factor

12.2 FORMATION AND SIGNALING PROPERTIES OF NO WITH IMPORTANCE FOR GENE ACTIVATION Shortly after endothelium derived relaxing factor (EDRF) was discovered and identified as NO, it turned out that the small molecule, composed of nitrogen and oxygen, is a versatile messenger with signaling properties beyond the vascular system. We now appreciate that NO affects signal transmission in nearly all areas

Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System

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of life (1, 2). NO taught us that a radical stirs efficient patho-physiological signaling throughout biology/medicine, thereby eliminating traditional thinking that a radical is a destructive molecule only. Signaling qualities attributed to NO in biological systems are often determined by using compounds that mimic an endogenous response by administering a chemically diverse group of compounds that release NO, known as NO donors, by blocking NO formation with NOSinhibitors or by using knockout mice that lack isotype-specific NOS (3). Proteins converting L-arginine to citrulline and NO are known as NO-synthases (NOS) (4). Three isoenzymes named after the cell type from which they were first isolated and cloned are distinguished and known as neuronal NOS (nNOS), inducible NOS (iNOS), and endothelial NOS (eNOS). A major distinction between enzymes is a high (i.e., iNOS) vs. low (i.e., eNOS, nNOS) NO-output capacity. This is based on the regulation of nNOS as well as eNOS by a transient cytosolic calcium increase resulting in a pulsative enzyme activation vs. the cytokine-inducible and thus transcriptionally regulated iNOS, producing NO for hours or days at basal calcium until the enzyme is degraded (5). Following its production, NO is preserved in its molecular structure by a chemically heterogeneous group of compounds. This deposit stabilization contributes to tuning the biological activity of NO after its liberation. The chemistry of NO involves interrelated redox forms (NO-radical •NO, NO− and NO+) with different chemical reactivity toward distinct target groups, thus explaining some of the pleiotropic effects of NO in biology. In particular, •NO reacts with molecular oxygen, superoxide, and transition metals, leading to the formation of reactive nitrogen intermediates (RNI) that directly or indirectly support additional nitrosative chemistry. •NO coordinates with hemoproteins or iron–sulfur centers, the NO+ character is found in nitroso compounds and is involved in nitrosation/nitrosylation reactions with nucleophils, among others thiols, whereas NO− rapidly undergoes dimerization and dehydration. Thus, the term RNI comprises oxidation states and adducts of the products of NOS, including •NO, NO−, and NO+, as well as for the subsequent adducts of these species such as NO2, NO2−, NO3−, N2O3, N2O4, S-nitrosothiols, peroxynitrite, and nitrosyl-metal complexes (6, 7). Biological signaling attributed to RNI can simply be distinguished as either being cGMP-dependent or cGMP-independent (2). Binding of RNI to the heme moiety of soluble guanylyl cyclase, concomitant cGMP formation and signaling must be considered the classical RNI response in close association with EDRF-action and regulation of vascular homeostasis (8). On the contrary, alternative signaling pathways of RNI that are not mimicked by lipophilic cGMP analogs may refer to covalent modification of proteins or oxidation events that do not require attachment of the NO group (6, 9). Among those modifications, S-nitrosylation/S-nitrosation (10), tyrosine nitration, oxidation, or cGMP-independent phosphorylation gained considerable attention as signal transmission mechanisms that culminate in gene activation or suppression (11–13). Most, if not all, gene regulatory activities evoked by RNI are indirect. Until today, no evidence existed regarding the existence of DNA elements within promotor regions of eukaryotic genes that directly respond to RNI. Thus,

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to understand signaling qualities of RNI in regulating gene expression one needs to consider modification of transcription factors, their compartmentalization, their actions as transcriptional activators or inhibitors, stability of target mRNAs as well as the protein amount of individual transcription factors. Several more recent review articles addressed various aspects of gene regulation by RNI in general, with excellent coverage of primary literature (11–14). Although outside the focus of this article information summarized in Table 12.1 provides examples of eukaryotic transcription factors know to be affected by RNI. Interestingly, RNI often exert contradictory effects either with activating or inhibiting the same transcription factor. This may reflect the use or formation of RNI with different signaling properties, different concentrations of RNI, cell-free vs. intact cell systems or cell types that differ in their intracellular redox milieu. For example, in resting cells RNI increase NF-κB- or AP-1-dependent gene transcription whereas inhibition is noticed in activated cells (13). Signaling qualities of RNI depend on the biological milieu (i.e., the presence or absence of modulatory co-signals), which are often considered to be oxygen-derived radicals (i.e., superoxide) (15). Alternatively, appreciating the importance of protein thiol modification via S-nitros(yl)ation or oxidation for RNI-signaling points to the significance of redox sensitive thiol residues as potential targets during gene (in)activation. At the same time, redox-modulation of thiols may affect their accessibility as RNI targets. Unfortunately, a simple prediction on activation vs. inhibition of gene activation because of RNI formation still is missing. Cellular stress, such as reduced oxygen availability, is potentially harmful to every cell and requires initiation of appropriate defense responses. A first line defense system toward hypoxia is stabilization of the hypoxia inducible factor-1α (HIF-1α), which is a prerequisite to activate the transcription factor HIF-1, composed of HIF-1α and HIF-1β subunits (16–19). Among recent advances to understand molecular details of hypoxic signaling is the observation that HIF-1α can be stabilized by RNI under normoxia, whereas RNI destabilize HIF-1α under hypoxia.

12.3 STABILITY REGULATION OF HIF-1α AND ACTIVATION OF HIF-1 12.3.1 LESSONS

FROM

HYPOXIA

Cellular recognition of hypoxia (i.e., decreased oxygen tension) and an appropriate response to meet this stress condition is predominantly facilitated by the transcription factor known as hypoxia inducible factor-1 [HIF-1]). Pioneering work on the expression of erythropoietin, a classical hypoxia-responsive target gene, led to the discovery of HIF-1α more than 10 years ago (20). More recently integrating pictures of oxygen sensing emerged that advanced our understanding on molecular details of HIF-1 stability regulation and activation as well as pathophysiological consequences associated with HIF-1α overexpression (16, 17, 19, 21–23). HIF-1 is a heterodimer composed of one of the three alpha subunits (HIF-


295

TABLE 12.1 Transcription Factors under the Control of RNI Eukaryotic Notably Mammalian Factors Transcription Factor (TF) NF-κB

Modulation by RNI - Activation in resting cells, low-level RNI - Inhibition in stimulated cells, high-level RNI

AP-1

- Activation in unstimulated cells, low-level RNI - Inhibition in activated cells

Sp1, Egr-1 (zinc finger TFs)

- Rather uniform inhibition - Activation if Sp1 de-represses the TNFα promoter

VDR, RXR (nuclear hormone R)

- Inhibition of DNA-binding and reporter activity

PPARγ

- Activation at low level RNI - Inhibition at high level RNI

NFAT

- Inhibition in activated NK cells

HSFs

- Activation (HSP70 expression)

p53

- Activation

HIF-1

- Activation under normoxia (see text for details) - Inhibition under hypoxia (see text for details)

Note: Selected examples for the regulatory impact of RNI on eukaryotic transcription factors. In a very simplistic way, activation vs. inhibition by RNI are indicated. TF: transcription factor, R: receptor, NF-κB: nuclear factor-κB, AP-1: activator protein-1, Egr-1: early growth response-1, VDR: 1α,25-dihydroxy-vitamin D3 receptor, RXR: retinoid X receptor, PPAR: peroxisome proliferator-activated receptor γ, NFAT: nuclear factor of activated T-cells, HSF: heat shock factor, p53: tumor suppressor p53, HIF-1: hypoxia inducible factor-1.

1α, HIF-2α, or HIF-3α) and one HIF-1β subunit (24). HIF-1β is constitutively expressed and identical to the aryl hydrocarbon receptor (AhR), known as AhR nuclear translocator (ARNT). As implicated by its name, HIF should be active under hypoxic conditions, which is attributed to stabilization or expression of the alpha subunit (i.e., HIF-1α). Under normoxic conditions the alpha subunit is usually unstable and mostly undetectable due to polyubiquitination by an E3-ubiquitin ligase complex that is built among other proteins by the von Hippel Lindau protein (pVHL), followed by 26S proteasomal degradation (25–27). An oxygen-dependent prolyl-4-hydroxylase, similar to proline hydroxylation of collagens, covalently modifies a domain of HIF-1α known as the oxygen-dependent degradation domain (ODD) by hydroxylating proline residues 402 and 564 (28,

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29). Hydroxylases are known as orthologs of C. elegans Egl-9, designated as PH domain-containing enzymes (PHD) (i.e., prolyl hydroxylases [PHD1, PHD2, PHD3, and PHD4]) (30–32). Hydroxylated HIF-1α form hydrogen bonds with pVHL side chains, which promotes polyubiquitination of HIF-1α, followed by proteasomal degradation (25, 26, 33). Enzymes contain iron and require 2-oxoglutarate as well as oxygen as co-factors. This explains why “hypoxic-mimetics,” such as the iron chelator desferrioxamine, attenuates PHD activity and, in turn, stabilizes HIF-1α, or why replacing the loosely bound Fe(II) by cobaltous ions blocks PHD activity. Collagen prolyl hydroxylases share with PHDs the requirement for 2-oxogultarate and molecular oxygen and the dependence on ferrous iron and ascorbate, which implies some enzyme homology. X-ray crystallographic studies of other members of 2-oxoglutarate dependent oxygenases demonstrated common structural features based on a β-barrel jelly-roll confirmation in which the catalytic site is formed by a non-heme iron coordinated by a 2-histidine-1aspartate motif (34, 35). The kinetics of PHD reaction have been studied recently in vitro, which should be taken with some caution for cellular conditions, but demonstrated an unusual high Km for oxygen, close to the atmospheric concentration of oxygen (36). This suggests that small changes in oxygen supply affect enzyme activity, making PHDs well suited for a role as physiological oxygen sensors. Besides oxygen, it appears that enzyme activity may be limited by iron or ascorbate, at least under cell culture conditions (37, 38). A second transactivation domain, besides the one found in the ODD, is located in the extreme C-terminus of HIF-1α, known as C-terminal transactivation domain (CTAD). Hydroxylation of asparagine 803 by factor inhibiting HIF-1 (FIH) within the CTAD (28, 39) renders HIF-1α unable to bind to the p300/CBP co-activator thus preventing transactivation capabilities of HIF-1. FIH belongs to a distinct class of 2-oxoglutarate dependent dioxygenasaes that are, however, PHD related (40–42). Hypoxia attenuates Pro564/402 and Asn803 hydroxylation, which in turn provokes HIF-1α protein stabilization, HIF-1β association, co-activator recruitment, and subsequent activation of HIF-1, which results in expression of those targets that contain HRE (hypoxia responsive element) sites with the core DNA sequence 5′-ACGTG-3′ (23). A model description of HIF-1α stability regulation under hypoxia is shown in Figure 12.1. To date more than 60 putative direct HIF-1 target genes have been identified. Among these major groups can be categorized according to their signaling qualities with involvements in cell proliferation, cell survival, apoptosis, cytoskelatal structure, angiogenesis, vascular tone, transcriptional regulation, iron metabolism, glucose metabolism, or extracellular matrix metabolism (22, 43, 44). Although hypoxia activates HIF-1 in almost all cell types, the majority of target genes are regulated in a cell-type-specific manner. This suggests the functional interaction of HIF-1 with other transcription factors that determines the subgroup of activated genes in any particular cell. The most prominent target genes are erythropoietin (EPO), the first gene recognized to respond to hypoxia and vascular endothelial growth factor (VEGF), which is a key mediator of angiogenesis (17). Interestingly, PHDs (PHD2 and PHD3) are hypoxia-inducible target genes themselves, and this


297

O2 availability Pro HIF-1α Asn

2 oxoglutarate and O2 Succinate and CO2

PHD & FIH

Hypoxia Pro HIF-1α Asn

OH Pro HIF-1α HO Asn pVHL

HIF-1β

Pro HIF-1α HIF-1β Asn

OH Pro HIF-1α pVHL HO Asn Ub

HO Pro HIF-1α HIF-1β Asn HRE

OH Pro HIF-1α pVHL Asn Ub Ub Ub

Ub

26S proteasomal degradation

FIGURE 12.1 Stability regulation of HIF-1α and activation of HIF-1 by hypoxia. HIF1α is subjected to hydroxylation by PHDs and FIH, both requiring 2-oxoglutarate and oxygen as co-factors. This allows recruitment of pVHL, subsequent polyubiquitination, and concomitant 26S proteasomal degradation. Hypoxia attenuates PHD as well as FIH activity, thus abrogating HIF-1α hydroxylation. Binding of HIF-1β constitutes the active HIF-1 dimer, followed by binding to the HRE in target genes. For details, see text.

response is, at least in part, HIF-1 dependent (30, 45–48). One may speculate whether HIF-1-dependent PHD induction functions as a feedback mechanism to limit the activity of HIF-1 in hypoxia. This offers an explanation for the observation that exposure of cells to prolonged hypoxia accelerates HIF-1α destruction upon reoxygenation (49). Considering the variety of target genes, it is without surprise that we foresee the involvement of HIF-1 in developmental, physiological, and patho-physiological processes. In particular, this includes several major disease states such as ischemic cardiovascular disorders, pulmonary hypertension, stroke, pregnancy disorders, or cancer.

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12.3.2 NO: STABILIZATION OF HIF-1α HIF-1 UNDER NORMOXIA

AND

ACTIVATION

OF

Unquestionably, RNI stabilizes HIF-1α and causes transactivation of HIF-1 under normoxia (for references, see Reference 18). Independent lines of research in a variety of human, pig, or bovine cells ruled species-specific as well as cell-type-restricted effects out. With the use of chemically distinct NO donors, such as S-nitrosoglutathione (GSNO, considered the most physiological NO donor), NOC-18 (Z-1-1[2-aminoethyl-amino]diazen-1ium-1,2-diolate), NOC-5 (3-(hydroxy-1-(1-methylethyl)-2-nitrosohydazino)-1-propanamine, SNAP (S-nitroso-N-acetyl-D,L-penicillamine), or others, a direct involvement of RNI was assured (for references, see Reference 18). Supporting information was delivered by using NO donors with different half-lives to describe time- and concentration-dependent effects of RNI on HIF-1α accumulation (50). In addition, activation of the human VEGF promoter by RNI under normoxia and deletion as well as mutation analysis of the VEGF promoter indicated that the RNI-responsive cis-elements were the HIF-1 binding site and an adjacent ancillary sequence located immediately downstream within the HRE (51, 52). A hint on the NO species being involved came from experiments with GSNO, a nitrosonium donor, and observations that GSNO effects are reversed by dithiothreitol. This led to the proposal that S-nitrosylation stabilizes HIF-1α (53). Although S-nitrosation of HIF-1α was confirmed in vitro, the biological significance and causation of S-nitros(yl)ation in stabilizing HIF-1α remains unclear (54). Working with NO donors may raise questions on the biological significance of RNI concentrations being used. To overcome this potential drawback human iNOS was overexpressed, which succeeded in accumulating HIF-1α (50). Supporting evidence came from a transwell co-culture setup of lipopolysaccharide/interferon-γ-activated and thus NO/RNI-producing macrophages and tubular LLC-PK1 detector cells indicating that only NO/RNI-generating, but not resting macrophages, provoked a HIF-1α response in LLC-PK1 cells (55). These experiments suggest that autocrine or paracrine produced RNI stabilize HIF-1α under normoxia. HIF-1α accumulation in association with RNI production is supported under various experimental conditions. Cell-density-induced HRE activation in human prostate cells is facilitated via RNI formation, thus acting as paracrine and diffusible factors (56). In these cells, RNI use Ras, mitogen-activated protein kinase (MAPK), and HIF-1α signaling to activate HRE, suggesting a link between HIF1 activation and prostate tumor progression, thereby providing a survival or growth advantage of tumor cells. In some analogies, inhibition of iNOS blocked production of an angiogenic activity in thioglycolate-induced peritoneal and murine RAW264.7 macrophages, suggesting that VEGF contributes to macrophage-dependent angiogenic activity. In addition, modulation of the VEGF mRNA level is, at least in part, under the control of the iNOS pathway (57). Attenuating iNOS provokes formation of anti-angiogenic factors, which makes RNI likely players in the regulation of macrophage-dependent angiogenic activity in vivo,


299

in wound repair, and possibly in tumor development (57). It is interesting to note that iNOS and HIF-1 expression in macrophages are present in early wounds and thus contribute to VEGF production (58, 59). More important, an elegant conditional knockout of HIF-1α revealed that HIF-1α controls inflammatory responses through its regulation of the metabolic switch to glycolysis (60). Apparently, monocytes/macrophages rely on HIF-1α for regulating cell infiltration, edema formation, and tissue destruction and thus the orchestration of inflammation. The ability of NO donors, paracrine or autocrine, delivered RNI to stabilize HIF-1α under normoxia requires mechanistic explanations considering those concepts established for hypoxic stability regulation of HIF-1α. Analogous to hypoxia, RNI decreased ubiquitination of HIF-1α and dissociated binding of pVHL to HIF-1α (61). Knowing that prolyl hydroxylation of HIF-1α is a prerequisite for HIF-1α-pVHL interactions raises the possibility that RNI blocked HIF-1α prolyl hydroxylation. An in vitro HIF-1α-pVHL capture assay demonstrated a dose-dependent inhibition of PHD activity by the NO donor GSNO. Importantly, under the same experimental conditions the association of a synthetic peptide resembling a piece of the hydroxylated ODD-domain of HIF-1α with pVHL remained intact. The experiments suggest that hypoxia and RNI use overlapping signaling pathways to attenuate post-translational hydroxylation of HIF-1α by attenuating PHD activity. Figure 12.2 schematically proposes RNI actions that consequently provoke HIF-1α protein stabilization based on impaired proteasomal destruction. It is known that RNI interact with iron (II) in heme- or non-heme containing proteins (7), exemplified by spectroscopic studies when •NO directly coordinates the ferrous iron in protocatechuate 4,5-dioxygenase, catechol 2,3-dioxygenase (62) or in isopenicillin N synthase (35). These enzymes coordinate Fe2+ in their catalytic site in a 2-histidine-1-carboxylate facial triad which is the defining structural motif of mononuclear non-heme iron(II) enzymes (63). Taking into account that PHDs belong to a non-heme Fe2+-containing family of enzymes, it is rational to propose Fe2+-coordination by RNI in the catalytic site of PHD and thus enzyme inhibition. The concept on inhibition of PHD activity by RNI as the underlying mechanism to explain stabilization of HIF-1α was recently challenged by the observation that the NO donor NOC-18 did not inhibit HIF-1α hydroxylation, ubiquitination, and degradation (64). Instead, pulse-labeling studies in the presence of NOC-18 implied increased HIF-1α synthesis. Major conclusions in this study are derived from overexpression experiments of FLAG-HIF-1α and HApVHL and it remains open whether overexpressed proteins share regulatory features noticed for endogenous proteins. This becomes obvious, considering that expression of FLAG-tagged HIF-1α remained constant under the impact of NOC-18, and thus a difference in the HIF-1α/pVHL interaction would not be expected. Interestingly, when a GST-HIF-1α(429-608) fusion protein was incubated with in vitro translated pVHL in the presence of cell lysates from untreated or NO-donor-exposed cells, NOC-18 did not affect the interaction between HIF-1α and pVHL. On the contrary, lysate from GSNO-treated cells

300

Nitric Oxide, Cell Signaling, and Gene Expression NO/RNI

MAPK Pl3K

Akt

HIF-1α

Translation

mRNA

(S6K, elF-4E)

Pro HIF-1α Asn

PHD/FIH

Pro HIF-1α Asn HO

OH

26S proteasomal degradation

Pro HIF-1α HIF-1β Asn

HIF-1 transcriptional activity Regulation of HIF-1α synthesis

Regulation of HIF-1α stability

FIGURE 12.2 Synthesis vs. stability regulation of HIF-1α by NO/RNI. HIF-1α is subjected to hydroxylation by PHDs and FIH, which allows subsequent polyubiquitination and concomitant 26S proteasomal degradation. NO/RNI attenuate PHD as well as FIH activity under normoxia, thus abrogating HIF-1α hydroxylation. Binding of HIF-1β builds the active HIF-1 dimer, which allows HIF-1 transcriptional activity and thus accounts for increased HIF-1α stability regulation. Alternatively, NO may activate PI3K or MAPK to stimulate HIF-1α mRNA translation, which increases its protein content. For details, see text.

significantly inhibited the interaction of GST-HIF-1α(429-608) with pVHL, whereas lysate from (sodium nitroprusside) SNP-treated cells dramatically increased the interaction. This opens the possibility that different redox species derived from chemically distinct NO donors use divergent transmission systems to stabilize/express HIF-1α. By using LY294002, PD98059, or rapamycin, the authors went on to demonstrate that NOC-18 uses a PI3K, MAPK, and capdependent translation control system to express HIF-1α (64). Pathways provoking HIF-1α expression are used by various types of stimuli including growth factors, oncogenes, and inflammatory mediators (for references, see References 16 and 19). Considering the proposed role of PI3K in HIF-1α translational regulation, it may turn out that, in some cells, PI3K/Akt is stimulated by RNI, which in turn will provoke HIF-1α translation regulation. A similar scenario has also been reported for hypoxia (65). Under conditions where PI3K/Akt stimulation by hypoxia or RNI occurs, translational control mechanisms may overlap with pathways that stabilize HIF-1α to enhance HIF-1α protein appearance. It remains open whether activation of the PI3K/Akt pathway is restricted to distinct NO donors, such as NOC-18 but not GSNO.


12.3.3 DESTABILIZATION

OF

HIF-1α: THE ROLE

OF

301

NO

In keeping with the notion of the versatile reactivity of RNI with changing of the intracellular biological milieu, it appears logical that HIF-1α stabilization by RNI is subjected to multiple variables, among others, the redox environment (i.e., the rate of superoxide formation). Considering the near diffusion-controlled interaction between •NO and O2− one can predict that •NO signaling will be affected by the simultaneous formation of O2− which may shift signaling qualities of RNI or O2− toward other species (i.e., ONOO−). Experiments with the redox cycler DMNQ (2,3-dimethoxy-1,4-naphthoquinone) to generate O2− or H2O2 (derived from superoxide dismutase-triggered conversion of O2− to H2O2) attenuated RNIelicited HIF-1α accumulation (66). In human cerebral vascular smooth muscle cells, NOC-18 and GSNO stabilized HIF-1α alone and were synergistic with hypoxia, whereas SNP and SIN-1 inhibited basal HIF-1α levels and HIF-1α stabilization by GSNO and NOC-18 (67). The conclusion that inhibition was mediated by ROI, particularly ONOO−, was supported by experiments showing that superoxide dismutase (SOD) overcame the inhibitory effects of SNP/SIN-1, by demonstrating that SOD induced HIF-1α in the absence of hypoxia and by finding that the O2−-generating system of xanthine/xanthine oxidase inhibited HIF-1α stability in control cells, cells exposed to hypoxia, or NOC-18. These observations predict that the ability of RNI to stabilize HIF-1α depend to some extent on the generation of co-signals (i.e., superoxide); however, it remains to be seen whether O2− simply reduces effective concentrations of RNI and thereby eliminates the stimulus or whether O2−, regardless of its interaction with •NO, (in)directly destabilizes HIF-1α. Aside from the notion that the presence of O2− modulates RNI-evoked HIF-1α stabilization, seminal observations during 1998 and 1999 stated that carbon monoxide (CO) and RNI inhibit hypoxia-induced HIF-1α accumulation, although important differences with respect to either NO or CO action have been noticed (68–70). Sogawa et al. pointed out that different NO donors blocked an activation step of HIF-1α to a DNA-binding form under hypoxicand CoCl2-treatment (69). Liu and co-workers reported that NO and CO decreased HIF-1 DNA binding, although HIF-1 protein levels were unaffected by CO with the further notion that suppression of VEGF by these two molecules occurred via cGMP formation (68). Huang and colleagues observed that NO and CO abrogated hypoxia-induced accumulation of HIF-1α protein by targeting the ODD of HIF-1α and repressed the C-terminal transactivation domain of HIF-1α, whereas CO had no effect on CoCl2- or DFX-responses (70). In contrast, Srinivas et al. concluded that CO was not effective in suppressing hypoxia-inducible reporter gene activity (71). Although details among these studies vary, the fundamental observation that RNI attenuate hypoxia-evoked HIF-1α stabilization is consistent and contrasts RNI action under normoxia.

302


Considering the important role attributed to PHDs in stabilizing vs. degrading HIF-1α, this provided a basic understanding of oxygen sensing and allowed an acknowledgment of the role of co-factors such as iron, ascorbate, and 2oxoglutarate (28–30). Because hypoxia stabilizes HIF-1α by attenuating PHD activity, it appears attractive to speculate that RNI reverses inhibition (i.e., reactivate PHDs under hypoxia). This is in line with the notion by Huang et al., indicating that the ODD of HIF-1α, which accounts for protein stability, is involved in reversing HIF-1α stabilization by RNI (70). Additional support came from the more recent observations by Hagen and co-workers, indicating that expression of P402A/P564A-HIF-1α, a protein that lacks the hydroxylation and thus destabilization sites, is resistant to destabilization by DETA-NO under hypoxic conditions (72). Unpublished observations from our lab revealed that the hypoxic mimetic DFX abrogated pVHL-HIF-1α interactions, whereas NO donors restored this protein association, which in turn allowed proteasomal degradation (Callapina et al., unpublished). Participation of the proteasome pathway as well as PHD activity was further substantiated by employing the proteasome inhibitor MG 132 or the PHD blocking compound DMOG (dimethyloxalglycine), which antagonized the down-regulating behavior of RNI during DFX stimulation; however, the most important question of how PDH activity is restored by RNI under hypoxia still remains. Hagen and co-workers reported that inhibition of respiration promotes HIF-1α degradation in hypoxia and concluded that increased availability of non-respiratory oxygen consequently reactivates PHD activity (72). The increase in intracellular oxygen in association with blocked mitochondrial respiration was proven by targeting Renilla luciferase to the mitochondria of HeLa cells as a monitor of available oxygen. NO by blocking mitochondrial respiration via binding to cytochrome c oxidase may act as an endogenous regulator of oxygen availability in mammalian cells. It appears that HIF-1α suppressing actions under low RNI concentrations depend on inhibition of mitochondrial respiration because it is absent in p0-cells and is mimicked by inhibitors of mitochondrial respiration (73). The authors propose that destabilization of HIF-1α by RNI under hypoxia does not result from oxidative stress (i.e., ROI formation instead correlating with the inhibition of mitochondrial respiration by NO, which leaves more oxygen available for PHDs, thus allowing to regain PHD activity under hypoxia) (72, 73). Similar explanations may account for CO. Considering that the binding affinity of CO to cytochrome c oxidase is much lower compared with NO, however, may explain differences noticed in the literature regarding the efficacy of CO (70). CO may act more efficiently under an already limited oxygen supply, and is certainly dependent on the CO concentration being used; however, the concept of shifting oxygen from mitochondria to PHDs (i.e., the oxygen redistribution model) does not explain RNI actions during DFXor CoCl2-treatments because these experiments are performed under normoxia at 21% oxygen. A previous in vitro study reported that activation of the HIF-1αpVHL binding activity by NO donors (74). GST-HIF-1α(532-603) was incubated in the presence of 35S-pVHL with whole cell extracts generated from cells cultured in the presence of CoCl2 with or without NO donors. The authors


303

speculate on direct activation of PHD activity by NO without the addition of ferrous iron, which is surprising because many other authors do not detect basal activity in this assay without substitution of all co-factors, including ferrous iron. Searching for alternative explanations, aside from the oxygen redistribution model, indeed iron may offer some explanations. It has been known for some time that the addition of ferrous iron antagonized hypoxia as well as DFX-elicited HIF-1 responses (38). Moreover, it was reported that chronic exposure of cells for an extended period of 16 hours to NO donors (e.g., DETA-NO) decreased the activity and protein levels of complexes I, II, and IV. Inhibition of these respiratory complexes was accompanied by an increase in cellular S-nitrosothiol levels and an increase in the labile iron pool (75). As a working hypothesis, one may speculate whether the combination of hypoxia with RNI increases the pool of intracellular chelatable free iron (76), which may contribute to activity regulation of PHDs. Determination of free iron and PHD activity under conditions of O2− and RNI formation, as well as under conditions of co-treatment, will help to clarify these proposals.

12.4 MEDICAL IMPLICATIONS: RNI AND HIF-1 IN TUMOR BIOLOGY An important role of HIF-1 in tumor biology is supported by immunohistochemistry data indicating elevated levels of HIF-1α in a variety of primary malignant tumors or tumor metastases with low levels of HIF-1α in benign tumors (77). The interior of a growing tumor becomes progressively hypoxic as its size increases because oxygen only diffuses around 150–200 microns from capillaries. Thus, stabilization of HIF-1α is, in part, because PHD activity and thus proteasomal destruction are impaired. In addition, tumor-specific genetic alterations (i.e., mutations involving oncogenes and tumor suppressor genes) may enhance HIF-1 expression (19, 44). For example, loss of pVHL, PTEN, or p53 tumor suppressor genes correlate with HIF-1α expression as well as the transforming potential of the v-Src oncogene. The striking up-regulation of HIF-1α in many different tumors, by both physiologic and epigenetic mechanisms, raises the question how HIF-1 affects tumor biology. Apparently, HIF-1 allows metabolic adaptation to hypoxia, promotes angiogenesis, enhances survival, and stimulates proliferation. Therefore, HIF-1α overexpression in brain, breast, cervical, esophageal, or ovarian cancers is correlated with treatment failure and mortality. Aside from hypoxic responses, RNI are implicated in tumor growth and progression (78). Expression of NOS has been demonstrated in a variety of tumors including breast, head, neck, prostate, bladder, colon, and CNS tumors such as glioblastomas (for references, see Morbidelli [79]). RNI promote tumor growth by multiple actions such as regulating blood flow, maintaining the vasodilatory tone, promoting metastasis by increasing vascular permeability, as well as affecting matrix metalloproteinases and stimulating angiogenesis. The observation that

304


RNI and hypoxia share the ability to stabilize HIF-1α may be relevant for various aspects of tumor biology. It is also of interest that iNOS is a classical hypoxia-inducible gene product that opens the possibility of feed-back or feed-forward regulatory systems. In normoxic areas, infiltrating macrophages may produce NO to stabilize HIF-1α, which may elicit a proangiogenic response. In hypoxic regions, RNI may limit continuous HIF-1 signaling. This might be related to the observation that an increased production of RNI reduced tumor cell survival and induced tumor cell death (80). However, the role of RNI in affecting apoptosis is ambivalent and can be pro- as well as anti-apoptotic inasmuch as the role of RNI is linked to pro- as well as anti-tumor activities (for references, see Xu et al. [81]). As a rule of thumb, it can be postulated that high level of RNI formation acts as cytostatic or cytotoxic, whereas low-level RNI generation may promote tumor growth (82, 83). Regulation of tumor growth by RNI points to multiple facets of RNI signaling, such as HIF-1α stability regulation, which deserves consideration to determine the precise role of RNI in tumor biology and to understand contrasting observations of RNI in promoting or inhibiting the etiology of cancer.

12.5 CONCLUDING REMARKS Expression of gene regulation is one way to understand signaling by RNI. It appears relevant to fully acknowledge the sphere of RNI actions in coordinating inflammation, affecting proliferation, differentiation, and modulating cell survival decisions. Among the multiple transcriptional systems that are known to be under the influence of RNI, HIF-1 emerged as a relatively new target that increases our understanding of how RNI mimic or affect hypoxic conditions. This might refer to conditions of inflammation, wound healing, tumor biology, and vascular remodeling. At this point, we are beginning to understand molecular details of how RNI mimic a hypoxic response by attenuating prolyl hydroxylase activity under normoxia, whereas RNI destabilize HIF-1α under hypoxia. Figure 12.3 summarizes these aspects and implicates patho-physiological consequences by modulating the HIF-system. RNI have not been considered as classical activators of HIF-1α. This poses the question about the relevance of RNI in activating gene expression via HIF-1. Gene expression profiling may help to answer these questions in the future, as well as our search for medical symptoms associated with RNI formation and transcriptional regulation via HIF-1. Considering the numerous genes under the control of RNI, using multiple transcriptional regulators, we need to establish a hierarchy of gene activation processes that determines and allows the prediction of the signaling qualities of RNI.


305

RNI mimic a hypoxic response NO donors iNOS donor cells

Normoxia HIF-1α patho-physiological consequences: HIF-1β

NO donors iNOS

Hypoxia

-

Development Ischemic/hypoxic disease Neoplasia Inflammation

HIF-1α

RNI attenuate a hypoxic response

FIGURE 12.3 HIF-1α a target for NO/RNI under normoxia vs. hypoxia. Under normoxia, NO donors and an active iNOS or cells producing NO promote HIF-1α accumulation to elicit various patho-physiological responses. In contrast, under hypoxia, NO donors or an active iNOS attenuates HIF-1α accumulation, thereby attenuating a hypoxic response. For details, see text.

12.6 ACKNOWLEDGMENT The authors gratefully acknowledge researchers whose primary observations, which formed the basis for our current knowledge in this active field, may not have been cited in this chapter due to space limitations. Many researchers were acknowledged indirectly by citing review articles. Our work was supported by grants from Deutsche Forschungsgemeinschaft (Br 999), Deutsche Krebshilfe (10-2008-Br2), and Sander Foundation (2002.088.1).

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Cross Talk between 13 The Nitric Oxide and Ceramide: Coordinate Interactions among Signaling Pathways Regulating Cell Death, Survival, and Differentiation Cristiana Perrotta and Clara De Palma University of Calabria, Rende, Italy

Sestina Falcone and Emilio Clementi University of Milano, Milano, Italy E. Medea Scientific Institute, Bosisio Parini, Italy DIBIT H San Raffaele Scientific Institute, Milano, Italy

CONTENTS 13.1 13.2 13.3 13.4 13.5 13.6

Introduction ............................................................................................312 Activation of eNOS by Ceramide .........................................................312 NO Regulates Generation of Ceramide by SMases ..............................313 A Model for the Cross Talk among the NOS-SMases Pathways .........314 Conclusion .............................................................................................316 Acknowledgments ..................................................................................316 References ...............................................................................................316

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13.1 INTRODUCTION Sphingolipids, a family of phospho/glycolipids built upon sphingoid bases, comprise members of variable complexity, from simple ceramide and sphingosine to the elaborate sialic acid-containing gangliosides. They participate not only in the mechanical stabilization and chemical resistance of the outer leaflet of the plasma membrane but also in several intracellular signalling pathways presiding over complex cell functions, including growth, migration, differentiation, and death [1]. The action of sphingolipids in these intracellular pathways often takes place through coordinate interplays with other second messenger molecules and their generating enzymes [2–4]. One such molecule is the short-lived gaseous messenger nitric oxide (NO). NO is generated in cells by specific enzymes, the NO synthases (NOSs). Of these, the neuronal (nNOS) and the endothelial (eNOS) isoforms are expressed constitutively, operate under the control of second messengers and lead to generation of physiological concentrations of NO [5]. A third enzyme is inducible by a variety of stimuli, including cytokines and bacterial products, operates at constant rates, and often yields high NO concentrations that participate in the immune responses and may lead to cell damage [5]. Of importance, all NOS are endowed with specific signals that target them to cellular membranes. The Nterminus of nNOS contains a PDZ (post-synaptic density protein-95, discs-large, Z0-1) domain that allows interactions of the enzyme with other PDZ-containing proteins at the plasma membrane including α1-syntrophin, PSD-95, and PSD-93 [6]. eNOS can be myristoylated and palmitoylated, and these lipid modifications allow the enzyme to be localized at both the plasma membrane and the Golgi complex [7]. Finally, both eNOS and inducible NOS may interact with members of the caveolin family of proteins, which also contribute to localization of the enzymes at the plasma membrane [7, 8]. The localization at cellular membranes constitutes the structural basis of the functional interaction among sphingolipids, NO, and their generating enzymes. The long-term ability of sphingolipids to regulate the expression of NOS isoforms has been comprehensively described in a recent review [9]. This chapter concentrates on the short-term, two-way interactions involving ceramide, when synthesized ex novo by the acid and neutral sphingomyelinases (SMases), and NO, mainly when generated by eNOS.

13.2 ACTIVATION OF eNOS BY CERAMIDE Two modes of activation of eNOS by ceramide have been reported thus far. They involve ceramide generation by either the acid or the neutral SMases, the first stimulated by basic fibroblast growth factor (bFGF) in CHO-K1 cells [10], the second by tumor necrosis factor-alpha (TNF-α) in HeLa cell clones transfected with eNOS under a tetracycline-responsive element [11, 12]. These two activation modes have common properties. They originate at plasma membrane rafts, where receptors co-localize with eNOS, and result in rapid (within min) activation of

The Cross Talk between Nitric Oxide and Ceramide

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the enzyme. They, however, differ in terms of the molecular events leading to eNOS activation. Activation of the enzyme by TNF-α requires stimulation of the phosphatidylinositol 3’ kinase (PI3K)/Akt pathway that leads to phosphorylation of eNOS in its activating Ser 1179 residue. Preliminary evidence from our laboratory indicates that upstream to the PI3K/Akt step TNF-α activates sphingosine kinase, with conversion of ceramide into sphingosine 1 phosphate (S1P). The latter sphingolipid appears to be the one responsible for the stimulation PI3K/Akt, consistently with the role of S1P in eNOS activation described in endothelial cells [13–17]. In contrast, the mechanism of activation of eNOS by bFGF through acid SMase-generated ceramide, although not fully characterized, is independent of PI3K/Akt activation [10]. These different pathways of eNOS activation by TNFα and bFGF may explain also the difference in the biological effect of NO generated. The neutral SMase-dependent activation of eNOS regulates in a negative fashion the ability of TNF-α to induce apoptosis [11], whereas the acid SMasedependent NO generation cooperates to the proliferative effect of bFGF [10].

13.3 NO REGULATES GENERATION OF CERAMIDE BY SMASES The interplay between NO and the SMase/ceramide pathway not only leads to regulation of NO generation but also works the other way around because NO modulates the generation of ceramide. Low, physiological concentrations of NO, such as those produced by eNOS, inhibit apoptosis triggered by a variety of apoptogens, including those activating death receptors of the TNF-α (TNF-RI)/CD95 superfamily [18]. By contrast, at high concentrations NO can induce apoptosis per se. Various mechanisms have been proposed to account for these two, apparently conflicting effects of NO [18, 19]. Recent evidence indicates that a relevant mechanism resides in the ability of NO to regulate ceramide levels. Studies performed in the U937 monocytederived cells and in clones of γδ T lymphocytes have demonstrated that NO inhibits the apoptotic responses induced by CD95 and TNF-RI by reducing the ability of these receptors to generate ceramide [11, 20, 21]. Preliminary studies in the laboratory indicate that inhibition of ceramide generation by NO takes place also in human and murine dendritic cells exposed to apoptogenic concentrations of lipopolysaccharide (LPS) in vitro, and in a model of LPS-induced sepsis in vivo. In both conditions NO protects dendritic cells from apoptosis. In all the cell types mentioned previously, the inhibition of ceramide generation by NO is mediated through activation of soluble guanylyl cyclase and cGMP-dependent protein kinase, followed by the inhibition of acid SMase [22]. Ceramide generation by the latter enzyme is known to sustain the initial phase of deathreceptor triggered apoptosis, contributing to the increased receptor clustering with ensuing initiation of apoptotic signalling [23]. With NO, the ceramide-induced assembly of signalling molecules at death receptors is inhibited [24]. The fact

314


that acid SMase is a target of NO explains the efficacy of low concentrations of NO in yielding an anti-apoptotic effect. The situation appears completely different when higher concentrations of NO are employed. In human leukemia and mesangial cells treated for several hours with millimolar concentrations of various NO donors the concentration of ceramide does not decrease, but instead increases, with ensuing death by apoptosis [25, 26]. Interestingly, the degree of apoptosis correlates in a linear way with the concentration of ceramide accumulated within the cells. In mesangioblasts, these pro-apoptotic effects of high NO appear to result from stimulation of neutral and acid SMases [25], whereas in leukemia cells the Mg2+-dependent neutral SMase appears to be the only target [26]. The mechanisms involved in SMase activation by NO are distinct from those involved in their inhibition because, in the former case, they are independent of cyclic GMP and require a caspase-3-dependent step [25, 26]. Interestingly, when the concentrations of NO in leukemia cells are lowered, the NO-induced ceramide increases are no longer observed and the gaseous messenger becomes protective against apoptosis [26]. The inhibition of ceramide generation induced by low NO via the cyclic GMP pathway appears therefore as a general tuning mechanism protecting cells from apoptosis, whereas the ceramide increases induced by high NO have opposite effects. Regulation by NO of ceramide generation may have biological actions beyond control of apoptosis because many death receptors may also trigger differentiating and pro-inflammatory responses, depending on the biological context. For example, the TNF-RI interacting protein TRADD, where recruitment to TNF-RI is inhibited by NO via inhibition of ceramide generation [24], is involved not only in apoptosis (because of its ability to recruit FADD and caspase-8 to the receptor), but also in the initiation of proliferation, differentiation, and inflammation. This is due to the ability of TRADD to recruit to TNF-RI also RIP and TRAF-2, two proteins that lead to activation of NF-κB, JNK, p38, and AP-1 [27]. Thus, regulation by NO of TRADD recruitment might imply regulation of the nonapoptogenic effects of TNF-α. In addition, NO regulates activation, by TNF-α, of neutral SMase [22], an enzyme that plays a role also in proliferation and differentiation [1]. Finally, preliminary evidence in our laboratory indicates that NO inhibits the activation of acid SMase by LPS in dendritic cells. LPS is known to trigger dendritic cell differentiation toward a mature state [28]. Inhibition of acidic SMase activation by LPS might explain the ability of NO to regulate one key feature of maturing dendritic cells, namely endocytosis, which is a ceramidedependent process in these cells [29].

13.4 A MODEL FOR THE CROSS TALK AMONG THE NOS-SMASES PATHWAYS Figure 13.1 is a hypothetical model depicting the two-way relationship between NOS/NO and SMases/ceramide, based on the data presented previously and using


315

NO NO

S1P

RIP1/ TRAF2

EDG N-SMase

TRADD

FADD

Ceramide

FAN

TNF-R1

A-SMase

S1P

out eNOS

PI3K/ Akt

Ceramide

in

NO

NO

Apoptosis

Growth diﬀerentiation

FIGURE 13.1 The NO-ceramide interactions in the signalling activated by TNF-α at its p55 kDa receptor (TNF-RI). Signalling molecules are indicated in green; low and high concentrations of NO are indicated by matching font sizes, and metabolic pathways are indicated by blue arrows. Continuous and dotted arrows indicate stimulation and inhibition, respectively, of either enzymatic activities or final biological effects. Activation of TNF-RI leads to the recruitment of various adapter proteins. Among these are FAN, which activates the neutral SMase (N-SMase) [36], and TRADD, which leads to either survival/differentiation or apoptosis, depending on whether it assembles in a complex with RIP1/TRAF2 or FADD [27]. Activation of FADD is an obligatory step in the activation by TNF-RI of the acid SMase (A-SMase), an enzyme localized at the outer leaflet of the plasma membrane when active [23]. The steps leading from FADD/TNF-RI interaction to A-SMase activation are still elusive. The role of NO in TNF-RI signalling may vary, depending on its concentrations. When generated at low concentrations, following N-SMase-PI3K/Akt-dependent activation of eNOS, it regulates ceramide generation in an inhibitory fashion. A role in this pathway may be played by S1P, generated following activation of sphingosine kinase by TNF-α [30] and acting at its EDG receptors [13–17]. NO might thus cooperate with S1P to protection from apoptosis, and to cell differentiation [14, 17, 18, 22, 24]. When generated at high concentrations NO sitmulates SMases with ensuing increases in ceramide levels. This leads to enhanced apoptosis [25, 26].

TNF-RI as the receptor model. As outlined previously, however, the kinds of interactions described for this receptor appear valid for other receptor systems as well. TNF-RI stimulation by TNF-α leads to activation of SMases with generation of ceramide. The latter is converted to S1P, which is secreted in part [30] to trigger PI3K/Akt-dependent activation of eNOS via stimulation of EDG receptors. The functional coupling of TNF-RI and EDG with eNOS might be facilitated by their common sub-cellular localization at plasma membrane rafts [13, 31, 32].

316


Activation of eNOS with subsequent generation of NO initiates a feed-back loop leading to the inhibition of both SMases. Depending on the cell type and environment, this loop might regulate apoptosis in an inhibitory fashion, through the blockade of both acid SMase activation and TRADD recruitment to TNF-R1. The latter effect, however, together with the stimulation of eNOS by neutral SMase and S1P, might also contribute to regulation of non-apoptogenic effects of TNF-RI, such as cell growth and differentiation. In case signaling does not decline but keeps going, the inhibitory loop described previously might turn into stimulatory of TNF-RI signaling. In particular, prolonged stimulation of eNOS might yield high concentrations of NO sustaining not the inhibition but the activation of SMases. The high cellular levels of NO and ceramide reached under these conditions could then act on a variety of well-characterized apoptogenic signalling events [3, 19] and ultimately synergizes to stimulate apoptosis.

13.5 CONCLUSION The intricate, two-player games of ceramide and NO share properties common to other modulatory systems of cellular signalling, such as the feed-back regulation of NO and calcium signalling [33]. In particular, it induces changes of activities of enzymes involved in different signaling cascades, but acting coordinated to yield useful cellular responses. These different signaling effects appear to be primarily under the control of NO. At low levels, NO reduces the generation of ceramide and collaborates in the regulation of survival/differentiation; at high levels, it increases ceramide and contributes to cell death. This Janus face of NO opens new perspectives from a therapeutic standpoint, also in view of the recent developments of ceramide- and NO-based compounds as therapeutic approaches to several disorders [34, 35].

13.6 ACKNOWLEDGMENTS The authors are supported by the Italian Association for Cancer Research and the Italian Ministry for Instruction, University and Research.

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Signaling 14 S-Nitrosothiol and Gene Regulation in Pulmonary Pathophysiology Khalequz Zaman, Lisa A. Palmer, and Benjamin Gaston University of Virginia Health System, Charlottesville, Virginia

CONTENTS 14.1 14.2 14.3 14.4 14.5 14.6

Introduction ............................................................................................321 Hypoxia Inducible Factor 1 ...................................................................322 Specificity Factors 1 and 3 ....................................................................325 NF κB ....................................................................................................326 Summary ................................................................................................327 Acknowledgments ..................................................................................327 References ...............................................................................................328

14.1 INTRODUCTION Activation of nitric oxide synthase (NOS) isoforms can result in a remarkable diversity of cellular effects (1, 2). These effects are broadly classified as being guanylate cyclase/cyclic GMP (cGMP)-dependent and cGMP-independent. cGMP-independent effects can be cytotoxic/antimicrobial, such as those involving tyrosine nitration through peroxynitrite or nitrite, or they can involve thiol S-nitrosylation (2). This chapter focuses on NOS-dependent gene regulation through thiol S-nitrosylation reactions. The biochemistry downstream of NOS activation has been elegantly designed to provide a spectrum of different effects, depending on the concentrations, the cellular environment and the metabolism of NOS products (1, 2). Remarkably, these various different signaling pathways can produce opposite effects, depending on the molecular environment. For example, different levels of NOS activity can either promote or inhibit cellular apoptosis (3, 4). Further, eNOS activation in the pulmonary vascular endothelium, which classically causes acute 321

322


vascular smooth muscle relaxation through guanylate cyclase activation, may also contribute to pulmonary vascular remodeling and chronic pulmonary hypertension, in part through its S-nitrosothiol (SNO)-mediated gene regulatory effects (5, 6). SNOs can be formed in pulmonary and other tissues in association with NOS activation (8). Several mechanisms have been proposed; virtually all require an electron acceptor to oxidize NO to a nitrosonium (NO+) equivalent. NO+ does not exist independently as a cation in solution, but instead as a complex with electronegative species (1, 2). SNO synthesis can be catalyzed by ceruloplasmin, hemoglobin, albumin, and NOS itself (2, 8–12). Additionally, inorganic reactions involving a variety of electron acceptors and intermediates have been identified (2, 3, 14). In general, SNOs mediate bioactivities primarily through transnitrosation reactions, or transfer of an NO equivalent from one cysteine thiolate to another (1, 2, 20, 21). Depending on the function of the cysteine, the protein may be activated, may be inhibited, or may serve to stabilize and transfer the NO group from one cell or organ to another (1, 15, 16). Indeed, specific proteins have been identified that regulate a transfer of NO into and between cells (15, 16, 17, 19). Additionally, a variety of different enzyme systems have been characterized that degrade or bioactivate SNOs, regulating their concentrations and activities (1, 18, 22). Criteria have recently been proposed by which the biological relevance of S-nitrosylation reactions can be formally evaluated (13). No single study is likely to establish all these criteria for a given protein; however, they serve as a roadmap for SNO research (Table 14.1). Recently, it has become evident that SNOs are involved in several gene regulatory pathways relevant to the lung. In particular, this chapter focuses on regulation of the transcription factors, hypoxia inducible factor 1 (HIF 1), specificity protein 1 (Sp 1), Sp 3, and nuclear factor κB (NFκB).

14.2 HYPOXIA INDUCIBLE FACTOR 1 Hypoxia inducible factor 1 (HIF 1) is a heterodimer of HIF 1α and HIF 1β (7). Classically, its activity is regulated by post-translational regulation of the expression of HIF 1α. HIF 1α can be targeted for degradation in the presence of partial pressures of oxygen >10 mmHg by prolyl hydroxylases (PHDs). These cause the protein to be recognized and ubiquitinated by a protein von Hippel— Lindau (pvHL), an E3 ubiquitin ligase. The poly-ubiquitinated HIF 1α is then recognized by the proteasome and degraded, preventing both HIF 1 dimerization and subsequent transcription of hypoxia-associated genes. At oxygen tensions

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