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. 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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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]:

Nitric Oxide Controls Cell Respiration

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

Nitric Oxide, Cell Signaling, and Gene Expression

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.)

Nitric Oxide Controls Cell Respiration

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].

Nitric Oxide Controls Cell Respiration

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide Controls Cell Respiration

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

Nitric Oxide Controls Cell Respiration

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide Controls Cell Respiration

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

Nitric Oxide Controls Cell Respiration

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

Nitric Oxide Controls Cell Respiration

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

Nitric Oxide, Cell Signaling, and Gene Expression

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.

REFERENCES 1. Nisoli E, Clementi E, Moncada S, Carruba MO. Mitochondrial biogenesis as a cellular signaling framework. Biochem. Pharmacol. (2004) 67:1–15. 2. Brown GC, Bal-Price A. Inflammatory neurodegeneration mediated by nitric oxide, glutamate, and mitochondria. Mol. Neurobiol. (2003) 27:325–355. 3. Sarti P, Giuffrè A, Barone MC, Forte E, Mastronicola D, Brunori M. Nitric oxide and cytochrome oxidase: reaction mechanisms from the enzyme to the cell. Free Radic. Biol. Med. (2003) 34:509–520. 4. Stewart VC, Heales SJ. Nitric oxide-induced mitochondrial dysfunction: implications for neurodegeneration. Free Radic. Biol. Med. (2003) 34:287–303. 5. Shiva S, Brookes PS, Patel RP, Anderson PG, Darley-Usmar VM. Nitric oxide partitioning into mitochondrial membranes and the control of respiration at cytochrome c oxidase. Proc. Natl. Acad. Sci. USA (2001) 98:7212–7217. 6. Brown GC, Borutaite V. Nitric oxide inhibition of mitochondrial respiration and its role in cell death. Free Radic. Biol. Med. (2002) 33:1440–1450. 7. Richter C, Schweizer M. Introduction to serial reviews on nitric oxide in mitochondria. Free Radic. Biol. Med. (2002) 33:1439. 8. Brookes PS, Levonen AL, Shiva S, Sarti P, Darley-Usmar VM. Mitochondria: regulators of signal transduction by reactive oxygen and nitrogen species. Free Radic. Biol. Med. (2002) 33:755–764. 9. Moncada S, Erusalimsky JD. Does nitric oxide modulate mitochondrial energy generation and apoptosis? Nat. Rev. Mol. Cell. Biol. (2002) 3:214–220. 10. Boveris A, Alvarez S, Navarro A. The role of mitochondrial nitric oxide synthase in inflammation and septic shock. Free Radic. Biol. Med. (2002), 33:1186–1193. 11. Brown GC. Regulation of mitochondrial respiration by nitric oxide inhibition of cytochrome c oxidase. Biochim. Biophys. Acta. (2001) 1504:46–57. 12. Cadenas E, Poderoso JJ, Antunes F, Boveris A. Analysis of the pathways of nitric oxide utilization in mitochondria. Free Radic. Res. (2001) 33 :747–756. 13. Nathan C. Nitric oxide as a secretory product of mammalian cells. FASEB J. (1992) 6:3051–3064. 14. Lane P, Gross SS. Cell signalling by nitric oxide. Semin. Nephrol. (1999) 19:215–229.

Nitric Oxide Controls Cell Respiration

23

15. Ignarro LJ. Nitric oxide: a unique endogenous signalling molecule in vascular biology. Biosci. Rep. (1999) 19: 51–71. 16. Droge W. Free radicals in the physiological control of cell function. Physiol. Rev. (2002) 82:47–95. 17. Foster MW, McMahon TJ, Stamler JS. S-nitrosylation in health and disease. Trends Mol. Med. (2003) 9:160–168. 18. Palacios M, Knowles RG, Palmer RM, Moncada S. Nitric oxide from L-arginine stimulates the soluble guanylate cyclase in adrenal glands. Biochem. Biophys. Res. Commun. (1989) 165:802–809. 19. Bredt, DS and Snyder, SH. Isolation of nitric oxide synthase, a calmodulin requiring enzyme. Proc. Natl. Acad. Sci. USA (1990), 87: 682–685. 20. Mayer, B, John, M, Bohme, E. Purification of Ca++ calmoduline-dependent nitric oxide synthase from porcine cerebellum. Cofactor role of terahydrobiopterin. FEBS Lett. (1990) 277: 215–219. 21. Giulivi C. Characterization and function of mitochondrial nitric oxide synthase. Free Rad. Biol. Med. (2003) 34: 397–408. 22. Ghosh DK, Salerno JC. Nitric oxide synthases: domain structure and alignment in enzyme function and control Front. Biosci. (2003) 8:d193–209. 23. Kanai AJ, Pearce LL, Clemens PR, Birder LA, VanBibber MM, Choi SY, de Groat WC, Peterson J. Identification of a neuronal nitric oxide synthase in isolated cardiac mitochondria using electrochemical detection. Proc. Natl. Acad. Sci. USA (2001) 98: 14126–14131. 24. Elfering SL, Sarkela TM, Giulivi C. Biochemistry of mitochondrial nitric-oxide synthase. J. Biol. Chem. (2002) 277:38079–38086. 25. Kobzik L, Stringer B, Balligand JL, Reid MB, Stamler JS. Endothelial type nitric oxide synthase in skeletal muscle fibers: mitochondrial relationships. Biochem. Biophys. Res. Commun. (1995) 211: 375–381. 26. Bates TE, Loesch A, Burnstock G, Clark JB. Mitochondrial nitric oxide synthase: a ubiquitous regulator of oxidative phosphorylation? Biochem. Biophys. Res. Commun. (1996) 218:40–44. 27. Bates TE, Loesch A, Burnstock G, Clark JB. Immunocytochemical evidence for a mitochondrially located nitric oxide synthase in brain and liver. Biochem. Biophys. Res. Commun. (1995) 213: 896–900. 28. Frandsen U, Lopez-Figueroa M, Hellsten Y. Localization of nitric oxide synthase in human skeletal muscle. Biochem. Biophys. Res. Commun. (1996) 227: 88–93. 29. Ghafourifar, P, Richter, C. Nitric oxide synthase activity in mitochondria. FEBS Lett. (1997) 418: 291–296. 30. Tatoyan, A, Giulivi, C. Purification and characterization of a nitric-oxide synthase from rat liver mitochondria. J. Biol. Chem. (1998) 273: 11044–11048. 31. French, S, Giulivi, C, Balaban, RS. Nitric oxide synthase in porcine heart mitochondria: evidence for low physiological activity. Am. J. Physiol. Heart Circ. Physiol. (2001) 280: H2863–H2867. 32. Lores-Arnaiz, S, Coronel, MF, Boveris, A. Nitric oxide, superoxide and hydrogen peroxide production in brain mitochondria after haloperidol treatment. Nitric Oxide (1999) 3: 235–243. 33. Lacza Z, Puskar M, Figueroa JP, Zhang J, Rajapakse N, Busija DW. Mitochondrial nitric oxide synthase is constitutively active and is functionally upregulated in hypoxia. Free Radic. Biol. Med., (2001) 31:1609–1615.

24

Nitric Oxide, Cell Signaling, and Gene Expression 34. Bustamante J, Bersier G, Romero M, Badin RA, Boveris A. Nitric oxide production and mitochondrial dysfunction during rat thymocyte apoptosis. Arch. Biochem. Biophys. (2000) 376: 239–247. 35. Southan GJ, Szabo C. Selective pharmacological inhibition of distinct nitric oxide synthase isoforms. Biochem. Pharmacol. (1996), 51:383–394. 36. Wood J, Garthwaite J. Models of the diffusional spread of nitric oxide: implications for neural nitric oxide signalling and its pharmacological properties. Neuropharmacology (1994) 33:1235–1244. 37. Liu X, Miller MJ, Joshi MS, Thomas DD, Lancaster JR, Jr. Accelerated reaction of nitric oxide with O2 within the hydrophobic interior of biological membranes. Proc. Natl. Acad. Sci. USA (1998) 95:2175–2179. 38. Brown GC. Nitric oxide and mitochondrial respiration. Biochim. Biophys. Acta. (1999) 1411: 351–369. 39. Heales JR, Bolanos JP, Stewart VC, Brookes PS, Land JM, Clark JB. NO, mitochondria and neurological disease. Biochim. Biophys. Acta. (1999) 1410: 215–228. 40. Wolin MS,, Xie YW, Hintze TH. Nitric oxide as a regulator of tissue oxygen consumption. Curr. Opin. Nephrol. Hyperthens. (1999) 8: 97–103. 41. Johnson LV, Walsh ML, Chen LB. Localization of mitochondria in living cells with rhodamine 123. Proc. Natl. Acad. Sci. USA (1980) 77: 990–994. 42. Sarti P, Lendaro E, Ippoliti R, Bellelli A, Benedetti PA, Brunori M. Modulation of mitochondrial respiration by nitric oxide: investigation by single cell fluorescence microscopy. FASEB J. (1999) 13: 191–197. 43. Cleeter MWJ, Cooper JM, Darley-Usmar VM, Moncada S, Schapira AHV. Reversible inhibition of cytochrome c oxidase, the terminal enzyme of the mitochondrial respiratory chain, by nitric oxide. FEBS Lett. (1994) 345: 50–54. 44. Brown GC, Cooper CE. Nanomolar concentration of nitric oxide reversibly inhibit synaptosomal cytochrome oxidase respiration, by competing with oxygen at cytochrome oxidase. FEBS Lett. (1994) 356: 295–298. 45. HenryY, Lepoivre M, Drapie, JC, Ducrocq C, Boucher JL, Guissani A. EPR characterization of molecular targets for NO in mammalian cells and organelles. FASEB J. (1993) 7: 1124–1134. 46. Pearce LL, Epperly MW, Greenberger JS, Pit, BR, Peterson J. Identification of respiratory complexes I and III as mitochondrial sites of damage following exposure to ionizing radiation and nitric oxide. Nitric Oxide (2001) 5:128–136. 47. Ferguson-Miller S, Babcock GT. Heme-Copper terminal oxidases. Chem. Rev. (1996) 96: 2889–2907. 48. Iwata S, Ostermeier C, Ludwig B, Michel H. Structure at 2.8 Å resolution of cytochrome c oxidase from Paracoccus denitrificans. Nature (1995) 376: 660–669. 49. Yoshikawa S, Shinzawa-Itoh K, Tsukihara T. X-ray structure and the reaction mechanism of bovine heart cytochrome c oxidase. J. Inorg. Biochem. (2000) 82: 1–7. 50. Tsukihara T, Aoyama H, Yamashita E, Tomizaki T, Yamaguchi H, Shinzawa-Itoh K, Nakashima R, Yaono R, Yoshikawa S. Structures of metal sites of oxidized bovine heart cytochrome c oxidase at 2.8 Å. Science (1995) 269: 1069–1074. 51. Schmidt B, McCracken J, Ferguson-Miller S. A discrete water exit pathway in the membrane protein cytochrome c oxidase. Proc. Natl. Acad. Sci. USA (2003) 100:15539–15542.

Nitric Oxide Controls Cell Respiration

25

52. Abramson J, Riistama S, Larsson G, Jasaitis A, Svensson-Ek M, Laakkonen L, Puustinen A, Iwata S, Wikström M. The structure of the ubiquinol oxidase from Escherichia coli and its ubiquinone binding site. Nat. Struct. Biol. (2000) 7: 910–917. 53. H. Witt, V. Zickermann, B. Ludwig. Site-directed mutagenesis of cytochrome c oxidase reveals two acidic residues involved in the binding of cytochrome c. Biochim. Biophys. Acta. (1995) 1230: 74–76. 54. M. Saraste, J. Castresana, Cytochrome oxidase evolved by tinkering with denitrification enzymes. FEBS Lett. 341 (1994. 1–4. 55. van der Oost J, de Boer AP, de Gier JW, Zumft WG, Stouthamer AH, van Spanning RJ. The heme-copper oxidase family consists of three distinct types of terminal oxidases and is related to nitric oxide reductase. FEMS Microbiol. Lett. 121 (1994) 1–9. 56. Proshlyakov DA, Pressler MA, DeMaso C, Leykam JF, DeWitt DL, Babcock GT. Oxygen activation and reduction in respiration: involvement of redox-active tyrosine 244. Science (2000) 290: 1588–1591. 57. Crum J, Gruys KJ, Frey TG. Electron microscopy of cytochrome c oxidase crystals: labeling of subunit III with a monomaleimide undecagold cluster compound. Biochemistry (1994) 33 :13719–13726. 58. Malatesta F, Georgevich G, and Capaldi RA Structural studies on beef heart cytochrome c oxidase from which subunit III has been removed by chimotrypsin treatment. In Structure and Function of Membrane Proteins, Quagliarello E, Palmieri F, Eds., Elsevier 6, 223 (1983). 59. Pearce LL, Epperly MW, Greenberger JS, Pitt BR, Peterson J. Identification of respiratory complexes I and III as mitochondrial sites of damage following exposure to ionizing radiation and nitric oxide. Nitric Oxide (2001) 5: 128–136. 60. Giuffrè A, Barone MC, Brunori M, D' Itri E, Ludwig B, Malatesta F, Muller HW, and Sarti P. Nitric oxide reacts with the single-electron reduced active site of cytochrome c oxidase. J. Biol. Chem. (2002) 277: 22402–22406. 61. Giuffrè A, Barone MC, Mastronicola D, D’Itri E, Sarti P, Brunori M. Reaction of nitric oxide with the turnover intermediates of cytochrome c oxidase: reaction pathway and functional effects. Biochemistry (2000) 39: 15446–15453. 62. Chance B, Graham N, Legallais V. Low temperature trapping method for cytochrome oxidase oxygen intermediates. Anal. Biochem. (1975) 67: 552–579. 63. Van Eps N, Szundi I, Einarsdóttir O. A new approach for studying fast biological reactions involving dioxygen: the reaction of fully reduced cytochrome c oxidase with O2. Biochemistry (2000) 39:14576–14582. 64. Babcock GT, Wikström M. Oxygen activation and the conservation of energy in cell respiration. Nature (1992) 356: 301–309. 65. Sucheta A, Szundi I, Einarsdottir O. Intermediates in the reaction of fully reduced cytochrome c oxidase with dioxygen. Biochemistry (1998) 37:17905–17914. 66. Zaslavsky D, Gennis RB. Proton pumping by cytochrome oxidase: progress, problems and postulates. Biochim. Biophys. Acta. (2000) 1458:164–179. 67. Verkhovsky MI, Morgan JE, Wikström M. Control of electron delivery to the oxygen reduction site of cytochrome c oxidase: a role for protons. Biochemistry (1995) 34: 7483–7491. 68. Fabian M, Palmer G. Redox state of peroxy and ferryl intermediates in cytochrome c oxidase catalysis. Biochemistry (1999) 38:6270–5.

26

Nitric Oxide, Cell Signaling, and Gene Expression 69. Wanio WW. Reactions of cytochrome oxidase. J. Biol. Chem. (1955) 212: 723–733. 70. Gibson QH, Greenwood C. Reactions of cytochrome oxidase with oxygen and carbon monoxide. Biochem. J. (1963) 86: 541–554. 71. Brunori M, Giuffrè A, D’Itri E, Sarti P. Internal electron transfer in Cu-heme oxidases. Thermodynamic or kinetic control? J. Biol. Chem. (1997) 272: 19870–19874. 72. Brudvig GW, Stevens TH, Chan SI. Reactions of nitric oxide with cytochrome c oxidase. Biochemistry (1980) 19: 5275–5285. 73. Takehara Y, Kanno T, Yoshioka T, Inoue M, Utsumi K. Oxygen-dependent regulation of mitochondrial energy metabolism by nitric oxide. Arch. Biochem. Biophys. (1995) 323:27–32. 74. Torres J, Darley-Usmar VM, Wilson MT. Inhibition of cytochrome c oxidase in turnover by nitric oxide: mechanism and implications for control of respiration. Biochem. J. (1995) 312: 169–173. 75. Giuffrè A, Sarti P, D’Itri E, Buse G. Soulimane T, Brunori, M. On the mechanism of inhibition of cytochrome c oxidase by nitric oxide. J. Biol. Chem. (1996) 271: 33404–33408. 76. Cooper CE, Torres J, Sharpe MA, Wilson MT. Nitric oxide ejects electrons from the binuclear centre of cytochrome c oxidase by reacting with oxidised copper: a general mechanism for the interaction of copper proteins with nitric oxide? FEBS Lett. (1997) 414: 281–284. 77. Pearce LL, Kanai AJ, Birder LA, Pitt BR, Peterson J. The catabolic fate of nitric oxide: the nitric oxide oxidase and peroxynitrite reductase activities of cytochrome oxidase. J. Biol. Chem. (2002) 277:13556–13562. 78. Torres J, Cooper CE, Wilson MT. A common mechanism for the interaction of nitric oxide with the oxidised binuclear centre and oxygen intermediates of cytochrome c oxidase. J. Biol. Chem. (1998) 273: 8756–8766. 79. Giuffrè A, Stubauer G, Brunori M, Sarti P, Torres J, Wilson MT. Chloride bound to oxidised cytochrome c oxidase controls the reaction with nitric oxide. J. Biol. Chem. (1998) 273: 32475–32478. 80. Cooper CE, Torres J, Sharpe MA, Wilson MT. Nitric oxide ejects electrons from the binuclear centre of cytochrome c oxidase by reacting with oxidised copper: a general mechanism for the interaction of copper proteins with nitric oxide? FEBS Lett. (1997) 414: 281–284. 81. Torres J, Sharpe MA, Rosquist A, Cooper CE. Wilson MT. Cytochrome c oxidase rapidly metabolises nitric oxide to nitrite. FEBS Lett. (2000) 475: 263–266. 82. Blackmore RS, Greenwood C, Gibson QH. Studies of the primary oxygen intermediate in the reaction of fully reduced cytochrome oxidase. J. Bio.l Chem. (1991) 266:19245–19249. 83. Sarti P, Giuffrè A, Forte E, Mastronicola D, Barone MC, Brunori M. Nitric oxide and cytochrome c oxidase: mechanisms of inhibition and NO degradation. Biochem. Biophys. Res. Comm. (2000) 274:183–187. 84. Cooper CE. Nitric Oxide and Iron Proteins. Biochim. Biophys. Acta. (1999) 1411: 290–309. 85. Boelens R, Wever R, Van Gelder BF, Rademaker H. An EPR study of the photodissociation reactions of oxidised cytochrome c oxidase-nitric oxide complexes. Biochim. Biophys. Acta. (1983) 724: 176–183.

Nitric Oxide Controls Cell Respiration

27

86. Stubauer G, Giuffrè A, Brunori M, Sarti P. Cytochrome c oxidase does not catalyze the anaerobic reduction of NO. Biochem. Biophys. Res. Comm. (1998) 245: 459–465. 87. Pearce LL, Bominaar EL, Hill BC, Peterson J. Reversal of cyanide inhibition of cytochrome c oxidase by the auxiliary substrate nitric oxide: an endogenous antidote to cyanide poisoning? J. Biol. Chem. (2003) 278:52139–52145. 88. Ford PC, Wink DA, Stanbury DM. Autoxidation kinetics of aqueous nitric oxide. FEBS Lett. (1993) 326:1–3. 89. Brown GC. Nitric oxide regulates mitochondrial respiration and cell functions by inhibiting cytochrome oxidase. FEBS Lett. (1995) 369: 136–139. 90. Brunori M, Giuffrè A, Sarti P, Stubauer G, Wilson MT. Nitric oxide and cellular respiration. Cell Mol. Life Sci. (1999) 56:549–557. 91. Bellamy TC, Wood J, Goodwin DA, Garthwaite J. Rapid desensitization of the nitric oxide receptor, soluble guanylyl cyclase, underlies diversity of cellular cGMP responses. Proc. Natl. Acad. Sci. USA (2000) 97:2928–33. 92. Mastronicola D, Genova ML, Arese M, Barone MC, Giuffrè A, Bianchi C, Brunori M, Lenaz G, Sarti P. Control of respiration by nitric oxide in Keilin-Hartree particles, mitochondria and SH-SY5Y neuroblastoma cells. Cell Mol. Life Sci. (2003) 60:1752–1759. 93. Carreras MC, Franco MC, Peralta JG, Poderoso JJ. Nitric oxide, complex I, and the modulation of mitochondrial reactive species in biology and disease. Mol. Aspects Med. (2004) 25:125–139. 94. Hollis VS, Palacios-Callender M, Springett RJ, Delpy DT, Moncada S. Monitoring cytochrome redox changes in the mitochondria of intact cells using multi-wavelength visible light spectroscopy. Biochim. Biophys. Acta. (2003) 1607:191–202. 95. Borutaite V, Brown GC. Rapid reduction of nitric oxide by mitochondria, and reversible inhibition of mitochondrial respiration by nitric oxide. Biochem. J. (1996) 315:295–299. 96. Giuffrè A, Stubauer G, Sarti P, Brunori M, Zumft WG, Buse G, Soulimane T. The heme-copper oxidases of Thermus thermophilus catalyze the reduction of nitric oxide: evolutionary implications. Proc. Natl. Acad. Sci. USA (1999) 96: 14718–14723. 97. Forte E, Urbani A, Saraste M, Sarti P, Brunori M, Giuffrè A. The cytochrome cbb3 from Pseudomonas stutzeri displays nitric oxide reductase activity. Eur. J. Biochem. (2001) 268: 6486–6491. 98. Butler CS, Forte E, Scandurra FM, Arese M, Giuffrè A, Greenwood C, Sarti P. Cytochrome bo3 from Escherichia coli: the binding and turnover of nitric oxide. Biochem. Biophys. Res. Comm. (2002) 296: 1272–1278. 99. Liu X, Miller MJ, Joshi MS, Thomas DD, Lancaster JR, Jr. Accelerated reaction of nitric oxide with O2 within the hydrophobic interior of biological membranes. Proc. Natl. Acad. Sci. USA (1998) 95:2175–2179. 100. Shiva S, Crawford JH, Ramachandran A, Ceaser EK, Hillson T, Brookes PS, Patel RP, Darley-Usmar VM. Mechanisms of the interaction of nitroxyl with mitochondria. Biochem. J. (2004) 379:359–366. 101. Poderoso JJ, Lisdero C, Schopfer F, Riobo N, Carreras MC, Cadenas E, Boveris A. The regulation of mitochondrial oxygen uptake by redox reactions involving nitric oxide and ubiquinol. J. Biol. Chem. (1999), 274:37709–37716. 102. Cadenas E. Mitochondrial free radical production and cell signalling. Mol Aspects Med. (2004) 25:17–26.

28

Nitric Oxide, Cell Signaling, and Gene Expression

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

30

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Mitochondrial Nitric Oxide Signaling in Synaptic Plasticity and Cell Death

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Mitochondrial Nitric Oxide Signaling in Synaptic Plasticity and Cell Death

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

Mitochondrial Nitric Oxide Signaling in Synaptic Plasticity and Cell Death

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.

REFERENCES 1. Ignarro LJ, Ed., Nitric Oxide: Biology and Pathobiology. Academic Press, New York (2000). 2. Knowles RG, Moncada S. Nitric oxide synthases in mammals. Biochem. J. 298: 249–258 (1994). 3. Thomas DD, Liu X, Kantrow SP, Lancaster JR, Jr. The biological lifetime of nitric oxide: implications for the perivascular dynamics of NO and O2. Proc. Natl. Acad. Sci. USA 98: 355–360 (2001). 4. Kobzik L, Stringer B, Ballingand JL, Reid MB, Stamler JS. Endothelial type nitric oxide synthase in skeletal fibers: mitochondrial relationships. Biochem. Biophys. Res. Commun. 211: 375–381 (1995). 5. Bates TE, Loesch A, Burnstock G, Clark JB. Immunocytochemical evidence for a mitochondrially located nitric oxide synthase in brain and liver. Biochem. Biophys. Res. Commun. 213: 896–898 (1995). 6. Bates TE, Loesch A, Burnstock G, Clark JB. Mitochondrial nitric oxide synthase: a ubiquitous regulator of oxidative phosphorylation. Biochem. Biophys. Res. Commun. 218: 40–44 (1996). 7. Ghafourifar P, Richter C. Nitric oxide synthase activity in mitochondria. FEBS Lett. 418: 291–295 (1997). 8. Giulivi C, Poderoso JJ, Boveris A. Production of nitric oxide by mitochondria. J. Biol. Chem. 273: 11038–11043 (1998). 9. Lores-Arnaiz S, Coronel MF, Boveris A. Nitric oxide, superoxide, and hydrogen peroxide production in brain mitochondria after haloperidol treatment. Nitric Oxide. 3: 235–243 (1999). 10. Lacza Z, Puskar M, Figueroa JP, Zhang J, Rajapakse N, Busua DW. Mitochondrial nitric oxide synthase is constitutively active and is functionally upregulated in hypoxia. Free Radic. Biol. Med. 31: 1609–1615 (2001).

40

Nitric Oxide, Cell Signaling, and Gene Expression 11. Kanai AJ, Pearce LL, Clemens PR, Birder L, Van Bibber MM, Choi S-Y, de Groat WC, Peterson J. Identification of a neuronal nitric oxide synthase in isolated cardiac mitochondria using electrochemical detection. Proc. Natl. Acad. Sci. USA 98: 14126–14131 (2001). 12. Costa LE, La-Padula P, Lores-Arnaiz S, D’Amico G, Boveris A, Kurnjek ML, Basso N. Long-term angiotensin II inhibition increases mitochondrial nitric oxide synthase and not antioxidant enzyme activities in rat heart. J. Hypertens. 20: 2487–2494 (2002). 13. Bustamante J, Bersier G, Aron-Badin R, Cymeryng C, Parodi A, Boveris A. Sequential NO production by mitochondria and endoplasmic reticulum during induced apoptosis. Nitric Oxide 6: 333–341 (2002). 14. Boveris A, Valdez LB, Alvarez S, Zaobornyj T, Boveris AD, Navarro A. Kidney mitochondrial nitric oxide synthase. Antiox. Redox. Signal. 5: 265–271 (2003). 15. Boveris A, Alvarez S, Navarro A. The role of mitochondrial nitric oxide synthase in inflammation and septic shock. Free Radic. Biol. Med. 33: 1186–1193 (2002). 16. Tatoyan A, Giulivi C. Purification and characterization of a nitric oxide synthase from rat liver mitochondria. J. Biol. Chem. 273: 11044–11048 (1998). 17. Boveris A, Lores-Arnaiz S, Bustamante J, Alvarez S, Valdez LB, Boveris AD, Navarro A. Pharmacological regulation of mitochondrial nitric oxide synthase. Meth. Enzymol. 359: 328–339 (2002). 18. Elfering SL, Sarkela TM, Giulivi C. Biochemistry of nitric oxide synthase. J. Biol. Chem. 277: 38079–38086 (2002). 19. Riobo NA, Melani M, Sanjuan N, Fiszman NL, Gravielle MC, Carreras MC, Cadenas E, Poderoso JJ. The modulation of mitochondrial nitric oxide synthase activity in rat brain development. J. Biol. Chem. 277: 42447–42455 (2002). 20. Peralta JG, Finocchietto PV, Converso D, Shopfer F, Carreras MC, Poderoso JJ. The modulation of mitochondrial nitric oxide synthase and energy expenditure in rat cold acclimation. Am. J. Physiol. Heart Circ. Physiol. 284: 2375–2383 (2003). 21. Zaobornyj T, Valdez LB, La-Padula P, Costa LE, Boveris A. Effect of sustained hypobaric hypoxia during maturation and aging on rat myocardium. II. mtNOS acticity. J. Appl. Physiol., in press. 22. Carreras MC, Peralta JG, Converso DP, Finocchietto PV, Rebagliatti I, Zaninovich AA, Poderoso JJ. Modulation of liver mitochondrial NOS is implicated in thyroiddependent regulation of O2 uptake. Am. J. Physiol. Heart Circ. Physiol. 281: H2282–H2288 (2001). 23. Boveris A, D’Amico G, Lores-Arnaiz S, Costa LE. Enalapril increases mitochondrial nitric oxide synthase activity in heart and liver. Antiox. Redox. Signal. 5: 691–697 (2003). 24. Lores-Arnaiz S, D’Amico G, Czerniczyniec A, Bustamante A, Boveris A. Brain mitochondrial nitric oxide synthase: in vitro and in vivo inhibition by chlorpromazine. Arch. Biochem. Biophys. 430: 170–177 (2004). 25. Poderoso JJ, Lisdero CL, Schöpfer F, Riobó NA, Carreras MC, Cadenas E, Boveris A. The regulation of mitochondrial oxygen uptake by redox reactions involving nitric oxide and ubiquinol. J. Biol. Chem. 274: 37709–37716 (1999). 26. Valdez LB, Zaobornyj T, Boveris A. Mitochondrial NOS: a voltage-dependent enzyme activity. FEBS Lett., in press. 27. Alvarez S, Valdez LB, Zaobornyj T, Boveris A. Oxygen dependence of mitochondrial nitric oxide activity. Biochem. Biophys. Res. Commun. 305: 771–775 (2003).

Mitochondrial Nitric Oxide Signaling in Synaptic Plasticity and Cell Death

41

28. Cleeter MWJ, Cooper JM, Darley-Usmar VM, Moncada S, Shapira AHV. Reversible inhibition of cytochrome c oxidase, the terminal enzyme of the mitochondrial repiratory chain. Implications for neurodegenerative diseases. FEBS Lett. 345: 50–54 (1994). 29. Brown GC, Cooper CE. Nanomolar concentrations of nitric oxide reversibly inhibit synaptosomal respiration by competing with oxygen at cytochrome oxidase. FEBS Lett. 356: 295–298 (1994). 30. Takehara Y, Kanno T, Yoshioka T, Inoue M, Utsumi K. Oxygen-dependent regulation of mitochondrial energy metabolism. Arch. Biochem. Biophys. 323: 27–32 (1995). 31. Cassina A, Radi R. Differential inhibitory action of nitric oxide and peroxynitrite on mitochondrial electron transfer. Arch. Biochem. Biophys. 328: 309–316 (1996). 32. Poderoso JJ, Carreras MC, Lisdero CL, Riobó NA, Schöpfer F, Boveris A. Nitric oxide inhibits electron transfer and increases superoxide radical production in rat heart mitochondria and submitochondrial particles. Arch. Biochem. Biophys. 328: 85–92 (1996). 33. Koivisto A, Matthias A, Bronnikov G, Nedergard J. Kinetics of the inhibition of mitochondrial respiration by NO. FEBS Lett. 417: 75–80 (1997). 34. Antunes F, Boveris A, Cadenas E. On the mechanism and biology of cytochrome oxidase inhibition by nitric oxide. Proc Natl Acad Sci USA 101: 16774–16779 (2004). 35. Clementi E, Brown GC, Feelisch M, Moncada S. Persistent inhibition of cell respiration by nitric oxide: crucial role of S-nitrosylation of mitochondrial complex I and protective effect of glutathione. Proc. Natl. Acad. Sci. USA 95: 7631–7636 (1998). 36. Riobo NA, Clementi E, Melani M, Boveris A, Cadenas E, Moncada S, Poderoso JJ. Nitric oxide inhibits mitochondrial-ubquinone reductase activity through peroxynitrite formation. Biochem. J. 359: 139–145 (2001). 37. Giulivi C. Functional implications of nitric oxide produced by mitochondria in mitochondrial metabolism. Biochem. J. 332: 673–679 (1998). 38. Sarkela TM, Berthiaume J, Elfering S, Gybina AA, Giulivi C. The modulation of oxygen radical production by nitric oxide in mitochondria. J. Biol. Chem. 276: 6945–6949 (2001). 39. Giulivi C. Characterization and function of mitochondrial nitric oxide synthase. Free Radic. Biol. Med. 34: 397–408 (2003). 40. Brown GC. NO says yes to mitochondria. Science 299: 838–839 (2003). 41. Boveris A, Costa LE, Poderoso JJ. Regulation of mitochondrial respiration by oxygen and nitric oxide. Ann. NY Acad. Sci. 899: 121–135 (2000). 42. Brunori M, Giuffre A, Sarti P, Stubauer G, Wilson MT. Nitric oxide and cellular respiration. Cell Mol. Life Sc.i 56: 75–80 (1997). 43. Wang JH, Ko GY, Kelly PT. Cellular and molecular bases of memory: synaptic and neuronal plasticity. J Clin Neurophysiol 14: 264–293 (1997). 44. Kolb B. Brain plasticity and behavioral change. In: Advances in Psychological Sciences. XXVI International Congress of Psychology. Sabourin M, Craik F, Robert M, Eds., Psychology Press, Philadelphia, PA. 115–143 (1998). 45. von Bohlen-Halbach O, Albrecht D, Heinemann U, Schuchman S. Spatial nitric oxide imaging using 1,2-diaminoanthraquinone to investigate the involvement of nitric oxide in long-term potentiation in rat brain slices. Neuroimage 15: 633–639 (2002).

42

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).

Mitochondrial Nitric Oxide Signaling in Synaptic Plasticity and Cell Death

43

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

47

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.

48

Nitric Oxide, Cell Signaling, and Gene Expression

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).

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

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

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

51

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

52

Nitric Oxide, Cell Signaling, and Gene Expression

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).

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

53

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+

54

Nitric Oxide, Cell Signaling, and Gene Expression

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

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

55

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

56

Nitric Oxide, Cell Signaling, and Gene Expression

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).

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

57

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.

58

Nitric Oxide, Cell Signaling, and Gene Expression

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

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

59

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.

60

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

61

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

62

Nitric Oxide, Cell Signaling, and Gene Expression

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-

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

63

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

64

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

65

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.

REFERENCES 1. Boveris A, Costa L, Cadenas E, Poderoso JJ. 1999. Regulation of mitochondrial respiration by adenosine phosphate, oxygen, and nitric oxide. Methods Enzymol. 301:188–98. 2. Boveris A, Oshino N, Chance B. 1972. The cellular production of hydrogen peroxide. Biochem. J. 128:617–630. 3. Han D, Antunes F, Canali R, Rettori D, Cadenas E. 2003. Voltage-dependent anion channels control the release of the superoxide anion from mitochondria to cytosol. J. Biol Chem. 278:5557–5563. 4. Poderoso JJ, Carreras MC, Lisdero C, Riobó N, Schöpfer F, Boveris A. 1996. Nitric oxide inhibits electron transfer and increases superoxide radical production in rat heart mitochondria and submitochondrial particles. Arch. Biochem. Biophys. 328:85–92. 5. Poderoso JJ, Carreras MC, Schöpfer F, Lisdero C, Riobó NA, Giulivi C, Boveris AD, Boveris A, Cadenas E. 1999. The reaction of nitric oxide with ubiquinol: kinetic properties and biological significance. Free Rad. Biol. Med. 26:925–935. 6. Michel T, Feron O. Nitric oxide synthases: which, where, why? 1997. J. Clin. Invest. 100:2146–2152. 7. Giulivi C, Poderoso JJ, Boveris A. 1998. Production of nitric oxide by mitochondria. J. Biol. Chem. 273:11038–11043. 8. Ghafourifar P, Richter C. 1997. Nitric oxide synthase activity in mitochondria. FEBS Lett. 418:291–296. 9. Bustamante J, Bersier G, Romero M, Badin RA, Boveris A. 2000. Nitric oxide production and mitochondrial dysfunction during rat thymocyte apoptosis. Arch. Biochem. Biophys. 376:239–247. 10. Riobo NA, Melani M, Sanjuan N, Fiszman ML, Gravielle MC, Carreras MC, Cadenas E, Poderoso JJ. 2002. The modulation of mitochondrial nitric-oxide synthase activity in rat brain development. J. Biol. Chem. 277:42447–42455. 11. Christopherson KS, Bredt D. 1997. Nitric oxide in excitable tissues: physiological roles and disease. J. Clin.. Invest. 100:2424–2429. 12. Lamas S, Marsden PA, Lee GK, Tempst P, Michel T. 1992. Endothelial nitric oxide synthase: molecular cloning and characterization of a distinct enzyme isoform. Proc. Natl. Acad. Sci. 89:6348–6352. 13. Ganster R, Geller D. Molecular regulation of inducible nitric oxide synthase. 2000. In: Nitric oxide. biology and pathobiology. Lou Ignarro, Ed. pp. 129–156, Academic Press, San Diego. 14. Elfering SL, Sarkela TM, Giulivi C. 2002. Biochemistry of mitochondrial nitricoxide synthase. J. Biol. Chem. 277:38079–38086. 15. Boveris A, Arnaiz SL, Bustamante J, Alvarez S, Valdez L, Boveris AD, Navarro A. 2002. Pharmacological regulation of mitochondrial nitric oxide synthase. Methods Enzymol. 359:328–339.

66

Nitric Oxide, Cell Signaling, and Gene Expression 16. Carreras MC, Peralta JG, Converso DP, Finocchietto PV, Rebagliati I, Zaninovich AA, Poderoso JJ. 2001 Mitochondrial nitric oxide synthase is a final effector in the thyroid-dependent modulation of oxygen uptake. Am. J. Physiol. (Heart Circ. Physiol.) 281:H2282–2288. 17. Carreras MC, Converso DP, Lorenti AS, Barbich M, Levisman DM, Jaitovich A, Antico Arciuch VG, Galli S, Poderoso JJ. 2004. Mitochondrial nitric oxide synthase drives redox signals for proliferation and quiescence in rat liver development. Hepatology 40:157–166. 18. Wise DL, Houghton G. 1968. Diffusion of nitric oxide. Chem. Eng. Sci. 26:453–460. 19. Poderoso JJ, Peralta JG, Lisdero C, Carreras MC, Radisic M, Schopfer F, Cadenas E, Boveris A. 1998. Nitric oxide regulates oxygen uptake and hydrogen peroxide release by the isolated beating rat heart. Am. J. Physiol. (Cell Physiol.) 274:C112–119. 20. Carreras MC, Poderoso JJ, Cadenas E, Boveris A. 1996. Measurements of nitric oxide and hydrogen peroxide production from human neutrophils. Methods Enzymol. 269:65–75. 21. Boveris A and Poderoso JJ. Regulation of oxygen uptake by nitric oxide. 2000. In: Nitric Oxide, biology and pathobiology. Lou Ignarro, Ed. pp. 355–368, Academic Press, San Diego. 22. Boczkowski J, Lisdero C, Lanone S, Carreras MC, Boveris A, Aubier M, Poderoso JJ. 1999. Endogenous peroxynitrite mediates mitochondrial dysfunction in rat diaphram during endotoxemia. FASEB J. 13:1637–1646. 23. Boczkowski J, Lisdero C, Lanone S, Carreras MC, Aubier M, Poderoso JJ. 2001. Peroxynitrite-mediated mitochondrial dysfunction. Biol. Signals Recept. 10:66–80. 24. Giulivi C. 1998. Functional implications of nitric oxide produced by mitochondria in mitochondrial metabolism. Biochem. J. 332:673–679. 25. Cleeter MWJ, Cooper JM, Darley-Usmar VM, Moncada S, Schapira AHV. 1994. Reversible inhibition of cytochrome c oxidase, the terminal enzyme of the mitochondrial respiratory chain, by nitric oxide. FEBS Lett. 345:50–54. 26. Brown GC. 1995. Nitric oxide regulates mitochondrial respiration and cell functions by inhibiting cytochrome oxidase. FEBS Lett. 369:136–139. 27. Poderoso JJ, Lisdero C, Schöpfer F, Riobó N, Carreras MC, Cadenas E, Boveris A. 1999. The regulation of mitochondrial oxygen uptake by redox reactions involving nitric oxide and ubiquinol. J. Biol. Chem. 274:37709–37716. 28. Koivisto A, Matthias A, Bronnkov G, Nedergaard J. 1997. Kinetics of the inhibition of mitochondrial respiration by NO. FEBS Lett. 417:75–80. 29. Shen W, Xu X, Ochoa M, Zhao G. Wolin M, Hintze TH. 1994. Role of nitric oxide in the regulation of oxygen consumption in conscious dogs. Circ. Res. 75:1086–1095. 30. Clementi E, Brown GC, Feelish M, Moncada S. 1998. Persistent inhibition of cell respiration by nitric oxide: cruxial role of s-nitrosylation of mitochondrial complex I and protective action of glutathione. Proc. Natl. Acad. Sci. 95:7631–7636. 31. Riobó NA, Clementi E, Melani M, Boveris A, Cadenas E, Moncada S, Poderoso JJ. 2001. Nitric oxide inhibits mitochondrial NADH-ubiquinone reductase activity through peroxynitrite formation. Biochem.. J. 359:139–145.

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

67

32. Carreras MC, Franco MC, Peralta JG, Poderoso JJ. 2004. Nitric oxide, complex I, and the modulation of mitochondrial reactive species in biology and disease. Mol. Aspects Med. 25:125–139. 33. Stewart VC, Sharpe MA, Clark JB, Heales JR. 2000. Astrocyte-derived nitric oxide causes both reversible and irreversible damage to the neuronal mitochondrial respiratory chain. J. Neurochem. 75:694–700. 34. Cassina A, Radi R. 1996. Differential inhibitory action of nitric oxide and peroxynitrite on mitochondrial electron transport. Arch. Biochem. Biophys. 328:309–316. 35. Forman HJ, Fridovich I. 1973. Superoxide dismutase: a comparison of rate constants. Arch. Biochem. Biophys. 150:396–400. 36. Koppenol WH. 1998. The basic chemistry of nitrogen monoxide and peroxynitrite. Free Rad. Biol. Med. 25:381–391. 37. Cadenas E, Poderoso JJ, Antunes F, Boveris A. 2000. Analysis of the pathways of nitric oxide utilization in mitochondria. Free Rad. Res. 33:747–756. 38. Huang P, Feng L, Oldham EA, Keating MJ, Plunkett W. 2000. Superoxide dismutase as a target for the selective killing of cancer cells. Nature 407:390–395. 39. Allen RG, Tresini M. 2000. Oxidative stress and gene regulation. Free Rad. Biol. Med. 28:463–499. 40. Boveris A, Cadenas E. 1997. Cellular sources and steady-state of reactive oxygen species. In: Oxygen, gene expression and cellular function. Clerch L and Massaro D, Eds. pp. 3–27. Marcel Dekker, New York 41. Thayer WS. 1986. Role of catalase in metabolism of hydrogen peroxide by the perfused heart. FEBS Lett. 202:137–140. 42. Burk RF, Nishiki K, Lawrence RA, Chance B. 1978. Peroxide removal by selenium-dependent and selenium-independent gluthatione peroxidases in hemoglobin-free perfused rat liver. J. Biol. Chem. 253:43–46. 43. Kim MR, Chang HS, Kim BH, Kim S, Baek SH, Kim JH, Lee SR, Kim JR. 2003. Involvements of mitochondrial thioredoxin reductase (TrxR2) in cell proliferation. Biochem. Biophys. Res. Commun. 304:119–124. 44. Chance B, Sies H, Boveris A. 1979. Hydroperoxide metabolism in mammalian organs. Physiol. Rev. 59:527–605. 45. Turrens JF, Giulivi C, Boveris A. 1986. Increased spontaneous chemiluminiscence from liver homogenates and isolated hepatocytes upon inhibition of superoxide and hydrogen peroxide utilization. Free Rad. Biol. Med. 2:153–140. 46. Becker LB, Vanden Hoek TL, Shao ZH, Li CQ, Schumacker PT. 1999. Generation of superoxide in cardiomyocytes during ischemia before reperfusion. Am. J. Physiol. (Heart Circ Physiol). 277:H2240–H2246. 47. Desaint S, Luriau S, Aude JC, Rousselet G, Toledano MB. 2004. Mammalian antioxidant defenses are not inducible by H2O2. J. Biol. Chem. 279:31157–31163. 48. Davies KJA. 2000. The broad spectrum of responses to oxidants in proliferating cells: a new paradigm for oxidative stress. IUBMB Life 48:41–47. 49. Antunes F, Cadenas E. 2001. Cellular titration of apoptosis with steady-state concentrations of H2O2 induce apoptosis through Fenton chemistry independent of the cellular thiol state. Free Rad. Biol. Med. 30:1008–1018. 50. Li. J, Holbrook NJ. 2003. Common mechanisms for declines in oxidative stress tolerance and proliferation with aging. Free Rad. Biol. Med. 35:292–299.

68

Nitric Oxide, Cell Signaling, and Gene Expression 51. Lee FY, Li Y, Zhu H, Yang S, Lin HZ, et al. 1999. Tumor necrosis factor increases mitochondrial oxidant production and induces expression of uncoupling protein2 in the regenerating mice liver. Hepatology 29:677–687. 52. Diehl AM, Rai RM. 1996. Liver regeneration 3: Regulation of signal transduction during liver regeneration. FASEB J. 10:215–227. 53. Chung H-T, Pae H-O, Choi B-M, Billiar TR, Kim Y-M. 2001. Nitric oxide as a bioregulator of apoptosis. Biochem. Biophys. Res. Commun. 272:31138–31148. 54. Chae HJ, Kang JS, Han JI, Bang BG, Chae SW, Kim KW, Kim HM, Kim HR. 2000. Production of hydrogen peroxide by serum and its involvement in cell proliferation in ROS 172.8 osteoblasts. Immunopharmacol. Immunotoxicol. 22:317–337. 55. Brown MR, Miller FJ, Jr, Li W-H, Ellingson AN, Mozena JD, Chatterjee P, Engelhardt JF, Zwacka RM, Oberley LW, Fang X, Spector AA, Weintraub NL. 1999. Overexpression of human catalase inhibits proliferation and promotes apoptosis in vascular smooth muscle cells Circ. Res. 85:524–533. 56. Tsai JC, Jain M, Hsieh CM, Lee WS, Yoshizumi M, Patterson C, Perrella MA, Cooke C, Wang H, Haber E, Schlegel R, Lee ME. 1996. Induction of apoptosis by pyrrolidinedithiocarbamate and N-acetylcysteine in vascular smooth muscle cells. J. Biol. Chem. 271:3667–3670. 57. Preston TJ, Muller WJ, Singh G. 2001. Scavenging of extracellular H2O2 by catalase inhibits the proliferation of HER-2 Neu-transformed Rat-1 fibroblasts through the induction of a stress response. J. Biol. Chem. 276:9558–9564. 58. Alonso M, Melani M, Converso D, Jaitovich A, Paz C, Carreras MC, Medina JH, Poderoso JJ. 2004. Mitochondrial extracellular signal-regulated kinases 1/2 (ERK1/2) are modulated during brain development. J. Neurochem. 89:248–256. 59. Galli S, Labato MI, Bal de Kier Joffe E, Carreras MC, Poderoso JJ. 2003. Decreased mitochondrial nitric oxide synthase activity and hydrogen peroxide relate persistent tumoral proliferation to embryonic behavior. Cancer Res. 63:6370–6377. 60. Sundaresan M, Yu ZX, Ferrans VJ, Irani K, Finkel T. 1995. Requirement for generation of H2O2 for platelet-derived growth factor signal transduction. Science 270 (5234):296–299. 61. Irani K, Xia Y, Zweier JL, Sollot SJ, Der CJ, Fearon ER, Sundaresan M, Finkel T, Goldschmidt-Clermont PJ. 1997. Mitogenic signaling mediated by oxidants in Ras-transformed fibroblasts. Science 275 (5306):1567–1568. 62. Suh YA, Arnold RS, Lassegue B, Shi J, Xu X, Sorescu D, Chung AB, Griendling KK, Lambeth JD. 1999. Cell transformation by the superoxide-generating oxidase Nox1. Nature 401 (6748):79–82. 63. Arnold RS, Shi J, Murad E, Whalen AM, Sun CQ, Polavarapu R, Parthasarathy S, Petros JA, Lambeth JD. 2001. Hydrogen peroxide mediates the cell growth and transformation caused by the mitogenic oxidase Nox1. Proc. Natl. Acad. Sci. 98:5550–5555. 64. Huang RP, Peng A, Hossain MZ, Fan Y, Jagdale A, Boyton AL. 1999. Tumor promotion by hydrogen peroxide in rat liver epithelial cells. Carcinogenesis 20:485–492. 65. Wang X, McCullough KD, Franke TF, Holbrook NJ. 2000. Epidermal growth factor receptor-dependent Akt activation by oxidative stress enhances cell survival. J. Biol. Chem. 275:14624–14631.

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

69

66. Bange J, Zwick E, Ullrich A. 2001. Molecular targets for breast cancer therapy and prevention. Nat. Med. 7:548–552. 67. Bae YS, Kang SW, Seo MS, Baines IC, Tekle E, Chock PB, Rhee SG. 1997. Epidermal growth factor (EGF)-induced generation of hydrogen peroxide. Role in EGF receptor-mediated tyrosine phosphorylation. J. Biol. Chem. 272:217–221. 68. Hamada J, Nakata D, Nakae D, Kobayashi Y, Akai H, Konishi Y, Okada F, Shibata T, Hosokawa M, Moriuchi T. 2001. Increased oxidative DNA damage in mammary tumor cells by continuous epidermal growth factor. J. Natl. Cancer Inst. 93:214–219. 69. Hu X, Moscinski LC, Valkov NI, Fisher AB, Hill BJ, Zuckerman KS. 2000. Prolonged activation of the mitogen-activated protein kinase pathway is required for macrophage-like differentiation of a human myeloid leukemic cell line. Cell Growth Differ. 11:191–200. 70. Zhang P, Wang YZ, Kagan E, Bonner JC. 2000. Peroxynitrite targets the epidermal growth factor receptor, Raf-1, and MEK independently to activate MAPK. J. Biol. Chem. 275:22479–22486. 71. Kwon SH, Pimentel DR, Remondino A, Sawyer DB, Colucci WS. 2003. H2O2 regulates cardiac myocyte phenotype via concentration-dependent activation of distinct kinase pathways. J. Mol. Cell Cardiol. 35:615–621. 72. Wartemberg M, Diedershagen H, Hescheler J, Sauer H. 1999. Growth stimulation versus induction of cell quiescence by hydrogen peroxide in prostate tumor spheroids is encoded by the duration of the Ca2+ response. J. Biol. Chem. 274:27759–27767. 73. Herrera B, Murillo MM, Alvarez-Barrientos A, Beltran J, Fernandez M, Fabregat I. 2004. Source of early reactive oxygen species in the apoptosis induced by transforming growth factor-beta in fetal rat hepatocytes. Free Radic. Biol. Med. 36:16–26. 74. Evan GI, Vousden KH. 2001. Proliferation, cell cycle and apoptosis in cancer. Nature 411:342–347. 75. Xu Y, Nguyen Q, Lo DC, Czaja MJ. 1997. c-myc-dependent hepatoma cell apoptosis results from oxidative stress and not a deficiency of growth factors. J. Cell Physiol. 170:192–199. 76. Chang L, Karin M. 2001. Mammalian MAP kinase signaling cascades. Nature 410:37–40. 77. Kurata S. 2000. Selective activation of p38 MAPK cascade and mitotic arrest caused by low-level oxidative stress. J. Biol. Chem. 275:23413–23416. 78. Inanami O, Ohta T, Ito S, Kuwabara M. 1999. Elevation of intracellular calcium ions is essential for the H2O2-induced activation of SAPK/JNK but not for that of p38 and ERK in Chinese hamster V79 cells. Antioxid. Redox. Signal. 1:501–508. 79. Gaitanaki C, Konstantina S, Chrysa S, Beis I. 2003. Oxidative stress stimulates multiple MAPK signalling pathways and phosphorylation of the small HSP27 in the perfused amphibian heart. J. Exp. Biol. 206:2759–69. 80. Awad MM, Grappuso PA. 2000. Cell cycle control during liver development in the rat: evidence indicating a role for cyclin D1 post-transcriptional regulation. Cell Growth Differ. 11:325–334. 81. Tamagno G, Robino A, Obbili P, Bardini M, Aragno M, Parola Danni O. 2003. H2O2 and 4-hydroxynonenal mediate amyloid β-induced neuronal apoptosis by activating JNKs and p38MAPK. Exp. Neurol. 180:144–155.

70

Nitric Oxide, Cell Signaling, and Gene Expression 82. Ryoo SW, Kim DU, Won M, Chung KS, Jang YJ, Oh GT, Park SK, Maeng PJ, Yoo HS, Hoe KL. 2004. Native LDL induces interleukin-8 expression via H2O2, p38 Kinase, and activator protein-1 in human aortic smooth muscle cells. Cardiovasc. Res. 62:185–193. 83. Moriya R, Uehara T, Nomura Y. 2000. Mechanism of nitric oxide-induced apoptosis in human neuroblastoma SH-SY5Y cells. FEBS Lett. 484:253–260. 84. Ghatan S, Larner S, Kinoshita Y, Hetman S, Patel L, Xia Z, Youle RJ, Morrison RS. 2000. p38 MAP kinase mediates bax traslocation in nitric oxide-induced apoptosis. J. Cell Biol. 150:335–347. 85. Forman HJ, Fukuto JM, Torres M. 2004. Redox signaling: thiol chemistry defines which reactive oxygen and nitrogen species can act as second messengers. Am. J. Physiol. (Cell Physiol.) 287:C246–C256. 86. Torres M and Forman HJ. 2000. Nitric oxide, oxidative stress, and signal transduction. In: Nitric oxide. biology and pathobiology. Lou Ignarro, Ed. pp. 329–342, Academic Press, San Diego. 87. Foley TD, Armstrong JJ, Kupchak BR. 2004. Identification and H2O2 sensitivity of the major constitutive MAPK phosphatase from rat brain. Biochem. Biophys. Res. Commun. 315:568–574. 88. Bae G-U, Seo D-W, Kwon H-K, Lee HY, Hong S, Lee ZW, Ha KS, Lee HW, Han JW. 1999. Hydrogen peroxide activates p70S6k signaling pathway. J. Biol. Chem. 46:32596–32602. 89. Aguirre-Ghiso JA, Estrada Y, Liu D, Ossowski L. 2003. ERK (MAPK) activity as a determinant of tumor growth and dormancy; regulation by p38 (SAPK). Cancer Res. 63:1684–1695. 90. Lee KH, Hyun MS, Kim JR. 2003. Growth factor-dependent activation of the MAPK pathway in human pancreatic cancer: MEK/ERK and p38 MAP kinase interaction in uPA synthesis. Clin. Exp. Metastasis. 20:499–505. 91. Masuda K, Shima H, Katagiri C, Kikuchi K. 2003. Activation of ERK induces phosphorylation of MAPK phosphatase-7, a JNK specific phosphatase, at Ser446. J. Biol. Chem. 278:32448–32456. 82. Liu Q, Hofmann PA. 2004. Protein phosphatase 2A-mediated cross talk between p38 MAPK and ERK in apoptosis of cardiac myocytes. Am. J. Physiol. (Heart Circ. Physiol.) 286:H2204– H2212. 93. Deng Y, Ren X, Yang L, Lin Y, Wu X. 2003. A JNK-dependent pathway is required for TNF alpha-induced apoptosis. Cell 115:61–70. 94. Iles KE, Dickinson DA, Watanabe N, Iwamoto T, Forman HJ. 2002. AP-1 activation through endogenous H2O2 generation by alveolar macrophages. Free Radic. Biol. Med. 32:1304–1313. 95. Salh BS, Martens J, Hundal RS, Yoganathan N, Charest D, Mui A, Gomez-Muñoz A. 2000. PD98059 attenuates hydrogen peroxide-induced cell death through inhibition of Jun N-terminal kinase in HT29 cells. Mol. Cell Biol. Res. Commun. 4:158–165. 96. Nemoto S, Takeda K, Yu Z-X, Ferrans VJ, Finkel T. 2000. Role of mitochondrial oxidants as regulators of cellular metabolism. Mol. Cell Biol. 20:7311–7318. 97. Inanami O, Ohta T, Ito S, Kuwabara M. 1999. Elevation of intracellular calcium ions is essential for the H2O2-induced activation of SAPK/JNK but not for that of p38 and ERK in Chinese hamster V79 cells. Antioxid. Redox. Signal. 1:501–508.

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

71

98. Pantano C, Shrivastava P, McElhinney B, Janssen-Heininger Y. 2003. Hydrogen peroxide signaling through tumor necrosis factor receptor 1 leads to selective activation of c-Jun N-terminal Kinase. J. Biol. Chem. 278:44091–44096. 99. Buschmann T, Potapova O, Bar-Shira A, Ivanov VN, Fuchs SY, Henderson S, Fried VA, Minamoto T, Alarcon-Vargas D, Pincus MR, Gaarde WA, Holbrook NJ, Shiloh Y, Ronai Z. 2001. Jun NH2-terminal kinase phosphorylation of p53 on Thr-81 is important for p53 stabilization and transcriptional activities in response to stress. Mol. Cell Biol. 21:2743–2754. 100. Schroeter H, Boyd CS, Ahmed R, Spencer JP, Duncan RF, Rice-Evans C, Cadenas E. 2003. c-Jun N-terminal kinase (JNK)-mediated modulation of brain mitochondria function: new target proteins for JNK signalling in mitochondrion-dependent apoptosis. Biochem. J. 372:359–369. 101. Giardina C, Boulares H, Inan MS. 1999. NSAIDs and butyrate sensitize a human colorectal cancer cell line to TNF-alpha and Fas ligation: the role of reactive oxygen species. Biochim. Biophys. Acta 1448:425–438. 102. Cassarino DS, Halvorsen EM, Swerdlow RH, Abramova NN, Parker WD, Jr., Sturgill TW, Bennett JP, Jr. 2000. Interaction among mitochondria, mitogen. activated protein kinases, and nuclear factor-kappa B in cellular models of Parkinson’s disease. J. Neurochem. 74:1384–1392. 103. Wang HG, Rapp UR, Reed JC. 1996. Bcl-2 targets the protein kinase Raf-1 to mitochondria. Cell 87:589–92. 104. Casanovas O, Miró F, Estanyol JM, Itarte E, Agell N, Bachs O. 2000. Osmotic stress regulates the stability of cyclin D1 in a p38SAPK2-dependent manner. J. Biol. Chem. 275:35091–35097. 105. Frippiat C, Remacle J, Toussaint O. Down-regulation and decreased activity of cyclin-dependent kinase 2 in H2O2-induced premature senescence. 2003. Int. J. Biochem. Cell Biol. 35:246–254. 106. Shackelford RE, Kaufmann WK, Paules RS. 2000. Oxidative stress and cell cycle checkpoint function. Free Rad. Biol. Med. 28:1387–1404. 107. Rai RM, Lee FY, Rosen A, Yang SQ, Lin HZ, Koteish A, Liew FY, Zaragoza C, Lowenstein C, Diehl AM. 1998. Impaired liver regeneration in inducible nitric oxide synthase deficient mice. Proc. Natl. Acad. Sci. 95:13829–13834. 108. Hatano F, Bennett BL, Manning AM, Qian T, Lemasters JJ, Brenner DA. 2001. NF-B stimulates inducible nitric oxide synthase to protect mouse hepatocytes from TNF-α and Fas-mediated apoptosis. Gastroenterology 120:1251–1262. 109. Janssen-Heininger YMW, Poynter ME, Baeuerle PA. 2000. Recent advances towards understanding redox mechanisms in the activation of nuclear factor NFB. Free Rad. Biol. Med. 28:1317–1327. 110. Payne CM, Bernstein C, Bernstein H, Gerner EW, Garewal H. 1999. Reactive nitrogen species in colon carcinogenesis. Antioxid. Redox. Signal. 1:449–467. 111. Zachary I, Gliki G. 2001. Signaling transduction mechanisms mediating biological actions of the vascular endothelial growth factor family. Cardiovasc. Res. 49:568–581. 112. Ziche M, Morbidelly L. 2000. Nitric oxide and angiogenesis. J. Neurooncol. 50:139–148. 113. Pervin S, Singh R, Chaudhuri G. 2001. Nitric oxide-induced cytostasis and cell cycle arrest of a human breast cancer cell line (MDA-MB-231): Potential role of cyclin D1. Proc. Natl. Acad. Sci. 98:3583–3588.

72

Nitric Oxide, Cell Signaling, and Gene Expression

114. Swana HS, Smith SD, Perrotta PL, Saito N, Wheeler MA, Weiss RM. 1999. Inducible nitric oxide synthase with transitional cell carcinoma of the bladder. J. Urol. 161:630–634. 115. Morcos E, Jansson OT, Adolfsson J, Kratz G, Wiklund, NP. 1999. Endogenously formed nitric oxide modulates cell growth in bladder cancer cell lines. Urology 53:1252–1257. 116. Klotz T, Bloch W, Jacobs G, Niggemann S, Engelmann U, Addicks K. 1999. Immunolocalization of inducible and constitutive nitric oxide synthases in human bladder cancer. Urology 54:416–419. 117. Romanenko A, Morimura K, Wanibuchi H, Salim EI, Kinoshita A, Kaneko M, Vozianov A, Fukushima S. 2000. Increased oxidative stress with gene alteration in urinary bladder urothelium after the Chernobyl accident. Int. J. Cancer 86:790–798. 118. Chazotte-Aubert L, Hainaut P, Ohshima H. 2000. Nitric oxide nitrates tyrosine residues of tumor-supressor p53 protein in MCF-7 cells. Biochem. Biophys. Res. Commun. 267:609–613. 119. Edwards PD, Topping D, Kontaridis MI, Moldawer LL, Copeland E, 3rd, Lind DS. 1997. Arginine-enhanced enteral nutrition augments the growth of nitric oxide-producing tumor. J. Parent. Enteral Nutr. 21:215–219. 120. Orucevic A, Bechberger J, Green AM, Shapiro RA, Billiar TR, Lala PK. 1999. Nitric oxide production by murine mammary adenocarcinoma cells promotes tumor-cell invasiveness. Int. J. Cancer 81:889–896. 121. Reveneau S, Arnould L, Jolimoy G, Hilpert S, Lejeune P, Saint-Giorgio V, Belichard C, Jeannin JF. 1999. Nitric oxide synthase in human breast cancer is associated with tumor grade, proliferation rate, and expression of progesterone receptors. Lab. Invest. 79:1215–1225. 122. Tschugguel W, Schneeberger C, Unfried G, Czerwenka K, Weninger W, Mildner M, Gruber DM, Sator MO, Waldhor T, Huber JC. 1999. Expression of inducible nitric oxide synthase in human breast cancer depends on tumor grade. Breast Cancer Res. Treat. 56:145–151. 123. Ropponen KM, Kellokoski JK, Lipponen PK, Eskelinen MJ, Alanne L, Alhava EM, Kosma VM. 2000. Expression of inducible nitric oxide synthase in colorectal cancer and its association with prognosis. Scand. J. Gastroenterol. 35:1204–1211. 124. Duenas-Gonzalez A, Isales CM, del Mar Abad-Hernandez N, Gonza Sarmiento R, Sanguenza O, Rodriguez-Commes J. 1997. Expression of inducible nitric oxide synthase in breast cancer correlates with metastatic disease. Mod. Pathol. 10:645–649. 125. Vakkala M, Kahlos K, Lakari E, Pääkkö P, Kinnula V, Soini Y. 2000. Inducible nitric oxide synthase expression, apoptosis, and angiogenesis in in situ and invasive breast carcinomas. Clin. Cancer Res. 6:2408–2416. 126. Shi Q, Xiong Q, Wang B, Le X, Khan NA, Xie K. 2000. Influence of nitric oxide synthase ii gene disruption on tumor growth and metastasis. Cancer Res. 60:2579–2583. 127. Xie K, Huang S. 2003.Contribution of nitric oxide-mediated apoptosis to cancer metastasis inefficiency. Free Rad. Biol. Med. 34:969–986. 128. Renaudin K, Denis MG, Karam G, Vallette G, Buzelin F, Laboisse CL, Jarry A. 2004. Loss of NOS1 expression in high-grade renal cell carcinoma associated with a shift of NO signalling. Br. J. Cancer 90:2364–2369.

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

73

129. Aravind L, Dixit VM, Koonin EV. 1999. The domains of death: evolution of the apoptosis machinery. TIBS 24:47–53. 130. Orrenius S. 2004. Mitochondrial regulation of apoptotic cell death. Toxicol. Lett. 149:19–23. 131. Cardoso SM, Rego AC, Penacho N, Oliveira CR. 2004. Apoptotic cell death induced by hydrogen peroxide in NT2 parental and mitochondrial DNA depleted cells. Neurochem. Int. 45:693–698. 132. Hussain SP, Amstad P, He P, Robles A, Lupold S, Kaneko I, Ichimiya M, Sengupta S, Mechanic L, Okamura S, Hofseth LJ, Moake M, Nagashima M, Forrester KS, Harris CC. 2004. p53-induced up-regulation of MnSOD and GPx but not catalase increases oxidative stress and apoptosis. Cancer Res. 64:2350–2356. 133. Chung H-T, Pae H-O, Choi B-M, Billiar TR, Kim Y-M. 2001. Nitric oxide as a bioregulator of apoptosis. Biochem. Biophys. Res. Commun. 272:31138–31148. 134. Adams V, Gielen S, Hambrecht, Schuler G. 2001. Apoptosis in skeletal muscle. Frontiers Biosci. 6:D1–D11. 135. Nguyen T, Brunson D, Crespi CL, Penman BW, Wishnok JS, Tannenbaum SR. 1992. DNA damage and mutation in human cells exposed to nitric oxide in vitro. Proc. Natl. Acad. Sci. 89:3030–3034. 136. Raines KW, Cao GL, Porsuphatana S, Tsai P, Rosen GM, Shapiro P. 2004. Nitric oxide inhibition of ERK1/2 activity in cells expressing neuronal nitric-oxide synthase. J. Biol. Chem. 279:3933–4390. 137. Hortelano S, Alvarez AM, Bosca L. 1999. Nitric oxide induces tyrosine nitration and release of cytochrome c preceding an increase of mitochondrial transmembrane potential in macrophages. FASEB J. 13:2311–2317. 138. Patel RP, Moellering D, Murphy-Ullrich J, Jo H, Beckman JS, Darley-Usmar V. 2000. Cell signaling by reactive nitrogen and oxygen species in atherosclerosis. Free Rad. Biol. Med. 28:1780–1794. 139. Bonfoco E, Krainc D, Ankarcrona M, Nicotera P, Lipton SA. 1995. Apoptosis and necrosis: two distinct events induced, respectively, by mild and intense insults with N-methyl-D-aspartate or nitric oxide/superoxide in cortical cell cultures. Proc. Natl. Acad. Sci. 92:7162–7166. 140. Chandra J, Samali A, Orrenius S. 2000. Triggering and modulation of apoptosis by oxidative stress. Free Radic. Biol. Med. 29:323–333. 141. Peng HB, Spiecker M, Liao JK. 1998. Inducible nitric oxide: an autoregulatory feedback inhibitor of vascular inflammation. J. Immunol. 161:1970–1976. 142. Yamamoto Y, Gaynor RB. 2001. Therapeutic potential of inhibition of the NF-B pathway in the treatment of inflammation and cancer. J. Clin. Invest.107:135–142. 143. Giulivi C. 1998. Functional implications of nitric oxide produced by mitochondria in mitochondrial metabolism. Biochem. J. 332:673–679. 144. Sarkela TM, Berthiaume J, Elfering S, Gybina AA, Giulivi C. 2001.The modulation of oxygen radical production by nitric oxide in mitochondria. J. Biol. Chem. 276:6945–69459. 145. Barrera-Hernandez G, Park KS, Dace A, Zhan Q, Chen SI. 1999. Thyroid hormone-induced cell proliferation in GC cells is mediated by changes in G1 cyclin/cyclin-dependent kinase levels and activity. Endocrinology 140:5267–5274. 146. Peralta JG, Finocchietto PV, Converso D, Schopfer F, Carreras MC, Poderoso JJ. 2003. Modulation of mitochondrial nitric oxide synthase and energy expenditure in rats during cold acclimation. Am. J. Physiol (Heart Circ. Physiol.) 284:H2375– H2383.

74

Nitric Oxide, Cell Signaling, and Gene Expression

147. Ghafourifar P, Schenk U, Klein SD, Richter C. 1999. Mitochondrial nitric-oxide synthase stimulation causes cytochrome c release from isolated mitochondria. Evidence for intramitochondrial peroxynitrite formation. J. Biol. Chem. 274:31185–31188. 148. Dennis J, Bennett JP, Jr. 2003. Interactions among nitric oxide and Bcl-family proteins after MPP+ exposure of SH-SY5Y neural cells I: MPP+ increases mitochondrial NO and Bax protein. J. Neurosci. Res. 72:76–88. 149. Komarova SV, Ataullakhanov FI, Globus RK. 2000. Bioenergetics and mitochondrial transmembrane potential during differentiation of cultured osteoblasts. Am. J. Physiol. (Cell Physiol.) 279:C1220–C1229. 150. Rochard P, Rodier A, Casas F, Cassar-Malek I, Marchall-Victorion S, Daury L, Wrutniak C, Cabello G. 2000. Mitochondrial activity is involved in the regulation of myoblast differentiation through myogenin expression and activity of myogenic factors. J. Biol. Chem. 275:2733–2744. 151. Rochard P, Cassar-Malek I, Marchal S, Wrutniak C, Cabello G. 1996. Changes in mitochondrial activity during avian myoblast differentiation: influence of triiodothyronine or v-erb A expression. J. Cell Physiol. 168:239–247. 152. Simonnet H, Alazard N, Pfeiffer K, Gallou C, Beroud C, Demont J, Bouvier R, Schagger H, Godinot C. 2002. Low mitochondrial respiratory chain content correlates with tumor aggressiveness in renal cell carcinoma. Carcinogenesis 23:759–768. 153. Amuthan G, Biswas G, Ananadatheerthavarada HK, Vijayasarathy C, Shephard HM, Avadhani NG. 2002. Mitochondrial stress-induced calcium signaling, phenotypic changes and invasive behavior in human lung carcinoma A549 cells. Oncogene 21:7839–7849. 154. Torroni A, Stepien G, Hodge JA, Wallace DC. 1990. Neoplastic transformation is associated with coordinate induction of nuclear and cytosplasmic oxidative phosphorylation genes. J. Biol. Chem. 265:20589–20593. 155. Faure-Vigny H, Heddi A, Giraud S, Chautard D, Stepien G. 1996. Expression of oxidative phosphorylation gens in renal tumors and tumoral cell lines. Mol. Carcinog. 16:165–172. 156. Heddi A, Faure-Vigni H, Wallace DC, Stepien G. 1996. Coordinate expression of nuclear and mitochondrial genes involved in energy production in carcinoma and oncocytoma. Biochim. Biophys. Acta 1316:203–209. 157. Wang C, Youssef J, Saran B, Rothberg PG, Cunnigham ML, Molteni A, Badr M. 1999. Diminished energy metabolism and enhanced apoptosis in livers of B6C3F1 mice treated with the antihepatocarcinogen rotenone. Mol. Cell Biochem. 201:25–32. 158. Kowaltowski AJ, Vercesi AE. 1999. Mitochodrial damage induced by conditions of oxidative stress. Free Rad. Biol. Med. 26:463–471. 159. Aito H, Aalto TK, Raivio KO. 1999. Correlation of oxidant-induced acute ATP depletion with delayed cell death in human neuroblastoma cells. Am. J. Physiol. 277(Cell Physiol.) 46:C878–C883. 160. Midorikawa K, Kawanishi S. 2001. Superoxide dismutases enhance H2O2-induced DNA damage and alter its site specificity. FEBS Lett. 495:187–190. 161. Huang P, Feng L, Oldham EA, Keating MJ, Plunkett W. 2000. Superoxide dismutase as a target for the selective killing of cancer cells. Nature 407:390–395. 162. Nicholls DG, Budd SL. 2000. Mitochondria and neuronal survival. Physiol. Rev. 80:315–360.

Mitochondrial Nitric Oxide and Redox Signaling Modulation of Cell Behavior

75

163. von Wangenheim K-H, Peterson H-P. 1998. Control of cell proliferation by progress in differentiation: clues to mechanisms of aging, cancer causation and therapy. J. Theor. Biol. 193:663–678. 164. Cuezva JM, Ostronoff LK, Ricart J, López de Heredia M, Di Ligero CM, Izquierdo JM. 1997. Mitochondrial biogenesis in the liver during development and oncogenesis. J. Bioenerg. Biomembr. 29:365—377. 165. Nisoli E, Clementi E, Paolucci C, Cozzi V, Tonello C, Sciorati C, Bracale R, Valerio A, Francolini M, Moncada S, Carruba MO. 2003. Mitochondrial biogenesis in mammals: the role of endogenous nitric oxide. Science 299 (5608):896–899. 166. Betarbet R, Sherer TB, MacKenzie G, Garcia-Osuna M, Panov AV, Greenamyre JT. 2000. Chronic systemic pesticide exposure reproduces features of Parkinson’s disease. Nat. Neurosci. 3:1301–1306. 167. Estévez AG, Spear N, Manuel SM, Radi R, Henderson CE, Barbeito L, Beckman JS. 1998. Nitric oxide and superoxide contribute to motor neuron apoptosis induced by trophic factor deprivation. J. Neurosci. 18:923–931. 168. Schöpfer F, Riobó N, Carreras MC, Alvarez B, Radi R, Boveris A, Cadenas E, Poderoso JJ. 2000. Oxidation of ubiquinol by peroxynitrite: implications for protection of mitochondria against nitrosative damage. Biochem. J. 349:35–42.

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

78

Nitric Oxide, Cell Signaling, and Gene Expression

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

Functions of Mitochondrial Nitric Oxide Synthase

79

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

80

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Functions of Mitochondrial Nitric Oxide Synthase

81

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

82

Nitric Oxide, Cell Signaling, and Gene Expression

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

Functions of Mitochondrial Nitric Oxide Synthase

83

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).

84

Nitric Oxide, Cell Signaling, and Gene Expression

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

Α

B

80 ∗

∗

60

∗

40

∗

RCR (%)

RCR (%)

80

20 0

85

0.0

1.0

3.5

40

0

7.0

0.0

1.0 3.5 (NO) (µM)

100

C

80

7.0

D

80 ∗

∗

∗

40

∗

20

RCR (%)

RCR (%)

∗

20

100

0

∗

60

(NO) (µM)

60

∗

60 40

∗

∗

∗

20 0.0

1.0

3.5

(NO) (µM)

7.0

0

0.0

1.0

3.5

7.0

(NO) (µM)

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

86

Nitric Oxide, Cell Signaling, and Gene Expression

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

Functions of Mitochondrial Nitric Oxide Synthase

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).

88

Nitric Oxide, Cell Signaling, and Gene Expression

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

Functions of Mitochondrial Nitric Oxide Synthase

89

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

90

Nitric Oxide, Cell Signaling, and Gene Expression

Hypoxia Reoxygenation Normoxia

A

(NO)m (µmoles/mg prot)

0.40 0.35 0.30 0.25 0.20 0.15 0.10 0

5

10 15 20 25 30 Time (min)

9.0

10 15

100

B

80 (O2) (%)

8.0 7.0

60 40 20

6.0

0

5.0

Hypoxia

(NO) (µM)

5

4.0

0

4

8

12

16

20

24

Time (min)

3.0 2.0 1.0 0

0 2

6

10 36 40 44 Time (min)

48

52

56

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.

Functions of Mitochondrial Nitric Oxide Synthase

91

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.).

REFERENCES Aguilera-Aguirre L, González-Hernández JC, Pérez-Vázquez V, Ramírez J, ClementeGuerrero M, Villalobos-Molina R, Saavedra-Molina A. 2002. Role of intramitochondrial nitric oxide in rat heart and kidney during hypertension. Mitochondrion 1:413–413. Almeida A, Heales SJR, Bolanos JP, Medina JM. 1998. Glutamate neurotoxicity is associated with nitric oxide-mediated mitochondrial dysfunction and glutathione depletion. Brain Res. 790:209–216. Ames BN, Shigenaga MK, Hagen TM. 1993. Oxidants, antioxidants, and the degenerative diseases of aging. Proc. Natl. Acad. Sci. USA 90:7915–7922. Arnaiz SL, Coronel MF, Boveris A. 1999. Nitric oxide, superoxide, and hydrogen peroxide production in brain mitochondria after haloperidol treatment. Nitric Oxide 3:235–243. Ashraf M, Bloor CM. 1976. X-ray microanalysis of mitochondrial deposits in ischemic myocardium. Virchows Arch. B. Cell Pathol. 22:287–297. Baek JH, Lee YS, Kang CM, Kim JA, Kwon KS, Son HC, Kim KW. 1997. Intracellular Ca2+ release mediates ursolic acid-induced apoptosis in human leukemic HL-60 cells. Int. J. Cancer 73:725–728. Bates TE, Loesch A, Burnstock G, Clark JB. 1995. Immunocytochemical evidence for a mitochondrially located nitric oxide synthase in brain and liver. Biochem. Biophys. Res. Commun. 213:896–900.

92

Nitric Oxide, Cell Signaling, and Gene Expression

Bates TE, Loesch A, Burnstock G, Clark JB. 1996. Mitochondrial nitric oxide synthase: a ubiquitous regulator of oxidative phosphorylation? Biochem. Biophys. Res. Commun. 218:40–44. Borutaite V, Brown GC. 1996. Rapid reduction of nitric oxide by mitochondria, and reversible inhibition of mitochondrial respiration by nitric oxide. Biochem. J. 315:295–299. Borutaite V, Morkuniene R, Brown GC. 1999. Release of cytochrome c from heart mitochondria is induced by high Ca2+ and peroxynitrite and is responsible for Ca2+induced inhibition of substrate oxidation. Biochim. Biophys. Acta 1453:41–48. Bringold U, Ghafourifar P, Richter C. 2000. Peroxynitrite formed by mitochondrial NO synthase promotes mitochondrial Ca2+ release. Free Radic. Biol. Med. 29:343–348. Brookes P, Darley-Usmar VM. 2002. Hypothesis: the mitochondrial NO signaling pathway, and the transduction of nitrosative to oxidative cell signals: an alternative function for cytochrome C oxidase. Free Radic. Biol. Med. 32:370–374. Brown GC, Cooper CE. 1994. Nanomolar concentrations of nitric oxide reversibly inhibit synaptosomal respiration by competing with oxygen at cytochrome oxidase. FEBS Lett. 356:295–298. Brown GC. 1995. Nitric oxide regulates mitochondrial respiration and cell functions by inhibiting cytochrome oxidase. FEBS Lett. 369: 136–139. Brown GC, Bolaños JP, Heales SJ, Clark JB. 1995. Nitric oxide produced by activated astrocytes rapidly and reversibly inhibits cellular respiration. Neurosci. Lett. 193:201–204. Brown GC, Foxwell N, Moncada S. 1998. Transcellular regulation of cell respiration by nitric oxide generated by activated macrophages. FEBS Lett. 439:321–324. Brudvig GW, Stevens TH, Chan SI. 1980. Reactions of nitric oxide with cytochrome c oxidase. Biochemistry 19:5275–5285. Brune B, Gotz C, Messmer UK, Sandau K, Hirvonen MR, Lapetina EG. 1997. Superoxide formation and macrophage resistance to nitric oxide-mediated apoptosis. J. Biol. Chem. 272:7253–7258. Cadenas E, Davies KJ. 2000. Mitochondrial free radical generation, oxidative stress, and aging. Free Radic. Biol. Med. 29:222–230. Campo GM, Squadrito F, Campo S, Altavilla D, Quartarone C, Ceccarelli S, Ferlito M, Avenoso A, Squadrito G, Saitta A, Caputi AP. 1998. Beneficial effect of raxofelast, an hydrophilic vitamin E analogue, in the rat heart after ischemia and reperfusion injury. J. Mol. Cell. Cardiol. 30:1493–1503. Carafoli E. 1987. Intracellular calcium homeostasis. Annu. Rev. Biochem. 56:395–433. Carr GJ, Ferguson SJ. 1990. Nitric oxide formed by nitrite reductase of Paracoccus denitrificans is sufficiently stable to inhibit cytochrome oxidase activity and is reduced by its reductase under aerobic conditions. Biochim. Biophys. Acta 1017:57–62. Carreras MC, Peralta JG, Converso DP, Finocchietto PV, Rebagliati I, Zaninovich AA, Poderoso JJ. 2001. Modulation of liver mitochondrial NOS is implicated in thyroid-dependent regulation of O2 uptake. Am. J. Physiol. Heart Circ. Physiol. 281:H2282– H2288. Castrejón V, Parra C, Moreno R, Peña A, Uribe S. 1997. Potassium collapses the P in yeast mitochondria while the rate of ATP synthesis is inhibited only partially: modulation by phosphate. Arch. Biochem. Biophys. 346:37–34.

Functions of Mitochondrial Nitric Oxide Synthase

93

Chen Z, Siu B, Ho YS, Vincent R, Chua CC, Hamdy RC, Chua BH. 1998. Overexpression of MnSOD protects against myocardial ischemia/reperfusion injury in transgenic mice. J. Mol. Cell Cardiol. 30:2281–2289. Cleeter MW, Cooper JM, Darley-Usmar VM, Moncada S, Schapira AH. 1994. Reversible inhibition of cytochrome c oxidase, the terminal enzyme of the mitochondrial respiratory chain, by nitric oxide. Implications for neurodegenerative diseases. FEBS Lett. 345:50–54. Coll KE, Joseph SK, Corkey BE, Williamson JR. 1982. Determination of the matrix free Ca2+ concentration and kinetics of Ca2+ efflux in liver and heart mitochondria. J. Biol. Chem. 257:8696–8704. Das DK, Maulik N. 1994. Antioxidant effectiveness in ischemia-reperfusion tissue injury. Methods Enzymol. 233:601–610. Estevez AG, Spear N, Manuel SM, Barbeito L, Radi R, Beckman JS. 1998. Role of endogenous nitric oxide and peroxynitrite formation in the survival and death of motor neurons in culture. Prog. Brain Res. 118:269–280. Ferrante RJ, Hantraye P, Brouillet E, Beal MF. 1999. Increased nitrotyrosine immunoreactivity in substantia nigra neurons in MPTP treated baboons is blocked by inhibition of neuronal nitric oxide synthase. Brain Res. 823:177–182. Flatman PW. 1991. Mechanisms of magnesium transport. Annu Rev Physiol 53:259–271. Frandsen U, Lopez-Figueroa M, Hellsten Y. 1996. Localization of nitric oxide synthase in human skeletal muscle. Biochem. Biophys. Res. Commun. 227:88–93. French S, Giulivi C, Balaban RS. 2001. Nitric oxide synthase in porcine heart mitochondria: evidence for low physiological activity. Am. J.Physiol. Heart Circ. Physiol. 280:H2863– H2867. Furchgott RF, Zawadski JV. 1980. The obligatory role of endothelial cells in the relaxation of arterial smooth muscle by acetylcholine. Nature 288:373–376. Ghafourifar P, Richter C. 1997. Nitric oxide synthase activity in mitochondria. FEBS Lett. 418:291–296. Ghafourifar P, Richter C. 1999. Mitochondrial nitric oxide synthase regulates mitochondrial matrix pH. Biol. Chem. 380:1025–1028. Ghafourifar P, Schenk U, Klein SD, Richter C. 1999. Mitochondrial nitric-oxide synthase stimulation causes cytochrome c release from isolated mitochondria. Evidence for intramitochondrial peroxynitrite formation. J. Biol. Chem. 274:31185–31188. Ghafourifar P, Bringold U, Klein SD, Richter C. 2000. Mitochondrial nitric oxide synthase, oxidative stress and apoptosis. Biol. Sign. Recep. 10:57–65. Ghafourifar P, Richter C. 2001. Mitochondrial ubiquinone (Coenzyme Q10): Biochemical, Functional, Medical and Therapeutical Aspects in Human Health and Disease (Ebadi M, Marwah J, and Chopra R, Eds.), Prominent Press, AZ. Vol. 1, pp. 437–445. Ghafourifar P. 2002. Characterization of mitochondrial nitric oxide synthase. Methods Enzymol. 359:339–350. Giulivi C, Poderoso JJ, Boveris A. 1998. Production of nitric oxide by mitochondria. J. Biol. Chem. 273:11038–11043. González-Zulueta M, Ensz LM, Mukhina G, Lebovitz RM, Zwacka RM, Engelhardt JF, Oberley LW, Dawson VL, Dawson TM. 1998. Manganese superoxide dismutase protects nNOS neurons from NMDA and nitric oxide-mediated neurotoxicity. J. Neurosci. 18:2040–2055. Gottlieb RA, Engler RL. 1999. Apoptosis in myocardial ischemia-reperfusion. Ann. NY Acad. Sci. 874:412–426.

94

Nitric Oxide, Cell Signaling, and Gene Expression

Gottlieb RA. Programmed cell death. 2000. Drug News Perspect. 13:471–476. Granger DL, Lehninger AL. 1982. Sites of inhibition of mitochondrial electron transport in macrophage-injured neoplastic cells. J. Cell. Biol. 95:527–535. Green DR, Reed JC. 1998. Mitochondria and apoptosis. Science 281:1309–1312. Guerra G, Uribe S, Pardo JP. 1995. Reactivity of the H+-ATPase from Kluyveromyces lactis to sulfhydryl reagents. Arch. Biochem. Biophys. 321:101–107. Hajnoczky G, Csordas G, Madesh M, Pacher P. 2000. The machinery of local Ca2+ signaling between sarco-endoplasmic reticulum and mitochondria. J. Physiol. 529 (Pt 1):69–81. Haramaki N, Stewart DB, Aggarwal S, Ikeda H, Reznick AZ, Packer L. 1998. Networking antioxidants in the isolated rat heart are selectively depleted by ischemia-reperfusion. Free Radic. Biol. Med. 25:329–39. Hogg N, Kalyanaraman B. 1998. The use of NO gas in biological systems. Methods Mol. Biol. 100:231–236. Howard AB, Alexander RW, Taylor WR. 1995. Effects of magnesium on nitric oxide synthase activity in endothelial cells. Am. J. Physiol. 269:C612– C618. Hurst RD, Chowdhury R, Clark JB. 1996. Investigations into the mechanism of action of a novel nitric oxide generator on cellular respiration. J. Neurochem. 67:1200–1207. Ignarro LJ, Cirino G, Casini A, Napoli C. 1999. Nitric oxide as a signaling molecule in the vascular system: an overview. J. Cardiovasc. Pharmacol. 34:879–876. Ihnken K, Morita K, Buckberg GD, Winkelmann B, Schmitt M, Ignarro LJ, Sherman MP. 1997. Nitric-oxide-induced reoxygenation injury in the cyanotic immature heart is prevented by controlling oxygen content during initial reoxygenation. Angiology 48:189–202. Kanai AJ, Pearce LL, Clemens PR, Birder LA, VanBibber MM, Choi SY, de Groat WC, Peterson J. 2001. Identification of a neuronal nitric oxide synthase in isolated cardiac mitochondria using electrochemical detection. Proc. Natl. Acad. Sci. USA 98:14126–14131. Kang PM, Haunstetter A, Aoki H, Usheva A, Izumo S. 2000. Morphological and molecular characterization of adult cardiomyocyte apoptosis during hypoxia and reoxygenation. Circ. Res. 87:118–125. Karcsu S, Laszlo FA, Toth L, Jancso G, Bacsy E. 1983. Calcium-containing mitochondrial granules in neurohypophysial axon terminals disappear following vasopressin treatment of Brattleboro rats. Neurosci. Lett. 39:181–185. Keller JN, Kindy MS, Holtsberg FW, St Clair DK, Yen HC, Germeyer A, Steiner SM, Bruce-Keller AJ, Hutchins JB, Mattson MP. 1998. Mitochondrial manganese superoxide dismutase prevents neural apoptosis and reduces ischemic brain injury: suppression of peroxynitrite production, lipid peroxidation, and mitochondrial dysfunction. J. Neurosci. 18:687–697. Kerr JF, Wyllie AH, Currie AR. 1972. Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br. J. Cancer 26:239–257. Kharitonov VG, Sundquist AR, Sharma VS. 1994. Kinetics of nitric oxide autoxidation in aqueous solution. J. Biol. Chem. 269:5881–5883. Kissner R, Nauser T, Bugnon P, Lye PG, Koppenol WH. 1997. Formation and properties of peroxynitrite as studied by laser flash photolysis, high-pressure stopped-flow technique, and pulse radiolysis. Chem. Res. Toxicol. 10:1285–1292. Kloner RA, Przyklenk K, Whittaker P. 1989. Deleterious effects of oxygen radicals in ischemia/reperfusion. Resolved and unresolved issues. Circulation 80:1115–1127.

Functions of Mitochondrial Nitric Oxide Synthase

95

Kloner RA, Jennings RB. 2001. Consequences of brief ischemia: stunning, preconditioning, and their clinical implications: part 2. Circulation 104:3158–3167. Kobzik L, Stringer B, Balligand JL, Reid MB, Stamler JS. 1995. Endothelial type nitric oxide synthase in skeletal muscle fibers: mitochondrial relationships. Biochem. Biophys. Res. Commun. 211:375–381. Koshland DE, Jr. 1992. The molecule of the year. Science 258:1861. Kroemer G. 1999. Mitochondrial control of apoptosis: an overview. Biochem. Soc. Symp. 66:1–15. Kruman II, Nath A, Mattson MP. 1998. HIV-1 protein Tat induces apoptosis of hippocampal neurons by a mechanism involving caspase activation, calcium overload, and oxidative stress. Exp. Neurol. 154:276–288. Kruman II, Mattson MP. 1999. Pivotal role of mitochondrial calcium uptake in neural cell apoptosis and necrosis. J. Neurochem. 72:529–540. Lacza Z, Puskar M, Figueroa JP, Zhang J, Rajapakse N, Busija DW. 2001. Mitochondrial nitric oxide synthase is constitutively active and is functionally upregulated in hypoxia. Free Radic. Biol. Med. 31:1609–1615. Leist M, Fava E, Montecucco C, Nicotera P. 1997. Peroxynitrite and nitric oxide donors induce neuronal apoptosis by eliciting autocrine excitotoxicity. Eur. J. Neurosci. 9:1488–1498. Liang WY, Tang LX, Yang ZC, Huang YS. 2002. Calcium induced the damage of myocardial mitochondrial respiratory function in the early stage after severe burns. Burns 28:143–146. Lizasoain I, Moro MA, Knowles RG, Darley-Usmar V, Moncada S. 1996. Nitric oxide and peroxynitrite exert distinct effects on mitochondrial respiration which are differentially blocked by glutathione or glucose. Biochem. J. 314:877–880. López-Figueroa MO, Caamano C, Morano MI, Ronn LC, Akil H, Watson SJ. 2000. Direct evidence of nitric oxide presence within mitochondria. Biochem. Biophys. Res. Commun. 272:129–133. Loud AV. 1968. A quantitative stereological description of the ultrastructure of normal rat liver parenchymal cells. J. Cell Biol. 37:27–46. Ma XL, Gao F, López BL, Christopher TA, Vinten-Johansen J. 2000. Peroxynitrite, a twoedged sword in post-ischemic myocardial injury-dichotomy of action in crystalloid- versus blood-perfused hearts. J. Pharmacol. Exp. Ther. 292:912–920. Manzo-Ávalos S, Pérez-Vázquez V, Ramírez J, Aguilera-Aguirre L, González-Hernández JC, Clemente-Guerrero M, Villalobos-Molina R, Saavedra-Molina A. 2002. Regulation of the rate of synthesis of nitric oxide by Mg2+ and hypoxia. Studies in rat heart mitochondria. Amino Acids 22:381–389. McConkey DJ, Orrenius S. 1997. The role of calcium in the regulation of apoptosis. Biochem. Biophys. Res. Commun. 239:357–366. McKean TA. 1991. Calcium uptake by mitochondria isolated from muskrat and guinea pig hearts. J. Exp. Biol. 157:133–142. Messmer UK, Reimer DM, Reed JC, Brune B. 1996. Nitric oxide induced poly(ADPribose) polymerase cleavage in RAW 264.7 macrophage apoptosis is blocked by Bcl-2. FEBS Lett. 384:162–166. Mitchell, P, Vectorial, 1977. Prediction of subsequent ischemic heart disease using serial resting electrocardiograms. Annu. Rev. Biochem. 46:996–1005.

96

Nitric Oxide, Cell Signaling, and Gene Expression

Miyamoto Y, Saito Y, Kajiyama N, Yoshimura M, Shimasaki Y, Nakayama M, Kamitani S, Harada M, Ishikawa M, Kuwahara K, Ogawa E, Hamanaka I, Takahashi N, Kaneshige T, Teroaka H, Akamizu T, Azuma N, Yoshimasa Y, Takaaki Y, Itoh H, Masuda I, Yasue HY, Nakao K. 1998. Endothelial nitric oxide synthase gene is positively associated with essential hypertension. Hypertension 32:3–8. Miyata H, Silverman HS, Sollott SJ, Lakatta EG, Stern MD, Hansford RG. 1991. Measurement of mitochondrial free Ca2+ concentration in living single rat cardiac myocytes. Am. J. Physiol. 261:H1123–34. Molyneux CA, Glyn MC, Ward BJ. 2002. Oxidative stress and cardiac microvascular structure in ischemia and reperfusion: the protective effect of antioxidant vitamins. Microvasc. Res. 64:265–277. Moncada S, Palmer RM, Higgs EA. 1991. Nitric oxide: physiology, pathophysiology, and pharmacology. Pharmacol. Rev. 43:109–142. Nakamura M, Thourani VH, Ronson RS, Velez DA, Ma XL, Katzmark S, Robinson J, Schmarkey LS, Zhao ZQ, Wang NP, Guyton RA, Vinten-Johansen J. 2000. Glutathione reverses endothelial damage from peroxynitrite, the byproduct of nitric oxide degradation, in crystalloid cardioplegia. Circulation 102:III332– III338. Pacher P, Csordas G, Hajnoczky G. 2001. Mitochondrial Ca2+ signaling and cardiac apoptosis. Biol. Signals Recept. 10:200–223. Palmer RM, Ferrige AG, Moncada S. 1987. Nitric oxide release accounts for the biological activity of endothelium-derived relaxing factor. Nature (London) 327:524–526. Pepe S. 2000. Mitochondrial function in ischaemia and reperfusion of the ageing heart. Clin. Exp. Pharmacol. Physiol. 27:745–750. Pozzan T, Rizzuto R, Volpe P, Meldolesi J. 1994. Molecular and cellular physiology of intracellular calcium stores. Physiol. Rev. 74:595–636. Pozzan T, Magalhaes P, Rizzuto R. 2000. The comeback of mitochondria to calcium signalling. Cell Calcium 28:279–283. Qiu C, Muchant D, Beierwaltes WH, Racusen L, Baylis C. 1998. Evolution of chronic nitric oxide inhibition hypertension. Relationship to renal function. Hypertension 31:1–26. Ramachandran A, Ceaser E, Darley-Usmar VM. 2004. Chronic exposure to nitric oxide alters the free iron pool in endothelial cells: role of mitochondrial respiratory complexes and heat shock proteins. Proc. Natl. Acad. Sci. USA. 101:384–389. Richter C, Gogvadze V, Schlapbach R, Schweizer M, Schlegel J. 1994. Nitric oxide kills hepatocytes by mobilizing mitochondrial calcium. Biochem. Biophys. Res. Commun. 205:1143–1150. Richter C, Schweizer M, Ghafourifar P. 1999. Mitochondria, nitric oxide, and peroxynitrite. Methods Enzymol. 301:381–393. Rizzuto R, Pinton P, Carrington W, Fay FS, Fogarty KE, Lifshitz LM, Tuft RA, Pozzan T. 1998. Close contacts with the endoplasmic reticulum as determinants of mitochondrial Ca2+ responses. Science 280:1763–1766. Rizzuto R, Bernardi P, Pozzan T. 2000. Mitochondria as all-round players of the calcium game. J. Physiol. 529:37–47. Robb-Gaspers LD, Burnett P, Rutter GA, Denton RM, Rizzuto R, Thomas A. 1998. Integrating cytosolic calcium signals into mitochondrial metabolic responses. EMBO J. 17:4987–5000. Rodríguez-Zavala JS, Saavedra-Molina A, Moreno-Sánchez R. 1997. Effect of intramitochondrial Mg2+ on citrulline synthesis in rat liver mitochondria. Biochem. Mol. Biol. Int. 41:179–177.

Functions of Mitochondrial Nitric Oxide Synthase

97

Rodríguez-Zavala JS, Moreno-Sánchez R. 1998. Modulation of oxidative phosphorylation by Mg2+ in rat heart mitochondria. J. Biol. Chem. 273:7850–7855. Romani A, Dowell E, Scarpa A. 1991. Cyclic AMP-induced Mg2+ release from rat hepatocytes,permeabilized hepatocytes, and isolated mitochondria. J. Biol. Chem. 266:24376–24384. Ronson RS, Nakamura M, Vinten-Johansen J. 1999. The cardiovascular effects and implications of peroxynitrite. Cardiovasc. Res. 44:47–59. Saavedra-Molina A, Devlin TM. 1997. Effect of extra- and intra-mitochondrial calcium on citrulline synthesis in rat liver. Amino Acids 12:293–298. Saavedra-Molina A, Ramírez-Emiliano J, Clemente-Guerrero M, Pérez-Vázquez V, Aguilera-Aguirre L, González-Hernández JC. 2003. Mitochondrial nitric oxide inhibits ATP synthesis. Effect of free calcium in rat heart. Amino Acids 24:95–102. Santos DJ, Moreno AJ. 2001. Inhibition of heart mitochondrial lipid peroxidation by nontoxic concentrations of carvedilol and its analog BM-910228. Biochem. Pharmacol. 61:155–164. Schweizer M, Richter C. 1994. Nitric oxide potently and reversibly deenergizes mitochondria at low oxygen tension. Biochem. Biophys. Res. Commun. 204:169–175. Shesely EG, Maeda N, Kim HS, Deasai KM, Krege JH, Laubach VE, Sherman PA, Sessa WC, Smithies O. 1996. Elevated blood pressures in mice lacking endothelial nitric oxide synthase. Proc. Natl. Acad. Sci. USA 93:13176–13171. Sheu SS, Sharma VK. 1999. Rapid report: a novel technique for quantitative measurement of free Ca2+ concentration in rat heart mitochondria. J. Physiol. 518:577–584. Smaili SS, Hsu YT, Youle RJ, Russell JT. 2000. Mitochondria in Ca2+ signaling and apoptosis. J. Bioenerg. Biomembr. 32:35–46. Steinmann CM, Storm M. 1997. The conservation of mitochondrial function by ischaemia-the ischaemia paradox. Acta Physiol. Scand. 159:171–173. Stout A, Raphael HM, Kanterewicz BI, Klann E, Reynolds IJ. 1998. Glutamate-induced neuron death requires mitochondrial calcium uptake. Nat. Neurosci. 1:366–373. Szabo C, Day BJ, Salzman AL. 1996. Evaluation of the relative contribution of nitric oxide and peroxynitrite to the suppression of mitochondrial respiration in immunostimulated macrophages using a manganese mesoporphyrin superoxide dismutase mimetic and peroxynitrite scavenger. FEBS Lett. 381:82–86. Szibor M, Richter C, Ghafourifar P. 2001. Redox control of mitochondrial functions. Antioxid. Redox. Signal. 3:515–524. Takabayashi A, Kawai Y, Iwata S, Kanai M, Denno R, Kawada K, Obama K, Taki Y. 2000. Nitric oxide induces a decrease in the mitochondrial membrane potential of peripheral blood lymphocytes, especially in natural killer cells. Antioxid. Redox. Signal. 2:673–680. Takehara Y, Kanno T, Yoshioka T, Inoue M, Utsumi K. 1995. Oxygen-dependent regulation of mitochondrial energy metabolism by nitric oxide. Arch. Biochem. Biophys. 323:27–32. Takehara Y, Nakahara H, Inai Y, Yabuki M, Hamazaki K, Yoshioka T, Inoue M, Horton AA, Utsumi K. 1996. Oxygen-dependent reversible inhibition of mitochondrial respiration by nitric oxide. Cell Struct. Funct. 21:251–258. Tatoyan A, Giulivi C. 1998. Purification and characterization of a nitric-oxide synthase from rat liver mitochondria. J. Biol. Chem. 273:11044–11048. Torres J, Darley-Usmar V, Wilson MT. 1995. Inhibition of cytochrome c oxidase in turnover by nitric oxide: mechanism and implications for control of respiration. Biochem. J. 312:169–173.

98

Nitric Oxide, Cell Signaling, and Gene Expression

Toyo-oka T, Arisaka H, Sanma H, Shin WS, Dan Y, Sugimoto T. 1989. Synergistic deleterious effect of micromolar Ca ions and free radicals on respiratory function of heart mitochondria at cytochrome C and its salvage trial. Biochem. Biophys. Res. Commun. 163:1397–1403. Tsuda T, Kogure K, Nishioka K, Watanabe T. 1991. Synergistic deleterious effect of micromolar Ca ions and free radicals on respiratory function of heart mitochondria at cytochrome c and its salvage trial. Neuroscience 44:335–341. Tyler DD. 1992. Metabolite Transporting systems of mitochondria. In: The mitochondrion in health and disease, Wiley–VCH, Weinheim, Germany. pp 403–441. Umansky V, Schirrmacher V. 2001. Nitric oxide-induced apoptosis in tumor cells. Adv. Cancer Res. 82:107–131. Vanden Hoek TL, Qin Y, Wojcik K, Li CQ, Shao ZH, Anderson T, Becker LB, Hamann KJ. 2003. Reperfusion, not simulated ischemia, initiates intrinsic apoptosis injury in chick cardiomyocytes. Am. J. Physiol. Heart Circ. Physiol. 284:H141– H150. Vander Heide RS, Schwartz LM, Reimer KA. 1994. The novel calcium antagonist Ro 405967 limits myocardial infarct size in the dog. Cardiovasc. Res. 28:1526–1532. Vergun O, Sobolevsky AI, Yelshansky MV, Keelan J, Khodorov BI, Duchen MR. 2001. Exploration of the role of reactive oxygen species in glutamate neurotoxicity in rat hippocampal neurones in culture. J. Physiol. 531:147–163. Votyakova TV, Bazhenova EN, Zvjagilskaya RA. 1993. Yeast mitochondrial calcium uptake: regulation by polyamines and magnesium ions. J. Bioenerg. Biomembr. 25:569–574. Weiss JN, Korge P, Honda HM, Ping P. 2003. Role of the mitochondrial permeability transition in myocardial disease. Circ. Res. 93:292–301. Xie YW, Wolin MS. 1996. Role of nitric oxide and its interaction with superoxide in the suppression of cardiac muscle mitochondrial respiration. Involvement in response to hypoxia/reoxygenation. Circulation 94:2580–2586. Xie YW, Kaminski PM, Wolin MS. 1998. Inhibition of rat cardiac muscle contraction and mitochondrial respiration by endogenous peroxynitrite formation during posthypoxic reoxygenation. Circ. Res. 82:891–897. Yang CS, Chen WY, Tsai PJ, Kuo JS. 1999. Alpha-tocopherol acetate significantly suppressed the increase in heart interstitial 8-hydroxydeoxyguanosine following myocardial ischemia and reperfusion in anesthetized rats. Clin. Chim. Acta 285:163–168. Ylitalo KV, Ala-Rami A, Liimatta EV, Peuhkurinen KJ, Hassinen IE. 2000. Intracellular free calcium and mitochondrial membrane potential in ischemia/reperfusion and preconditioning. J. Mol. Cell. Cardiol. 32:1223–1238.

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

99

100

5.4

5.5 5.6 5.7

Nitric Oxide, Cell Signaling, and Gene Expression

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

102

Nitric Oxide, Cell Signaling, and Gene Expression

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

104

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

114

Nitric Oxide, Cell Signaling, and Gene Expression

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].

Peroxynitrite

115

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

116

Nitric Oxide, Cell Signaling, and Gene Expression

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,

Peroxynitrite

117

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],

118

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Peroxynitrite

119

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,

120

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Peroxynitrite

121

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

122

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Peroxynitrite

123

REFERENCES 1. Radi, R.D., A. Alvarez, B. Ferrer-Sueta, G. Rubbo, H., The biological chemistry of peroxynitrite, in Nitric oxide biology and pathobiology, J.L. Ignarro, Ed. 2000. Academic Press: San Diego. p. 57–82. 2. Huie, R.E. and S. Padmaja, The reaction of NO with superoxide. Free Radic. Res. Commun., 1993. 18(4): 195–199. 3. Goldstein, S. and G. Czapski, The reaction of NO with O2.− and HO2: a pulse radiolysis study. Free Radic. Biol. Med., 1995. 19(4): 505–510. 4. Kissner, R., T. Nauser, P. Bugnon, P.G. Lye, and W.H. Koppenol, Formation and properties of peroxynitrite as studied by laser flash photolysis, high-pressure stopped-flow technique, and pulse radiolysis. Chem. Res. Toxicol., 1997. 10(11): 1285–1292. 5. Turrens, J.F., Mitochondrial formation of reactive oxygen species. J. Physiol., 2003. 522.2: 335–344. 6. Barja, G., Mitochondrial free radical production and aging in mammals and birds. Ann. NY Acad. Sci., 1998. 854: 224–238. 7. Turrens, J.F., A. Alexandre, and A.L. Lehninger, Ubisemiquinone is the electron donor for superoxide formation by complex III of heart mitochondria. Arch. Biochem. Biophys., 1985. 237(2): 408–414. 8. Turrens, J.F. and A. Boveris, Generation of superoxide anion by the NADH dehydrogenase of bovine heart mitochondria. Biochem. J., 1980. 191(2): 421–427. 9. Fiskum, G., A. Starkov, B.M. Polster, and C. Chinopoulos, Mitochondrial mechanisms of neural cell death and neuroprotective interventions in Parkinson’s disease. Ann. NY Acad. Sci., 2003. 991: 111–119. 10. Cadenas, E. and K.J. Davies, Mitochondrial free radical generation, oxidative stress, and aging. Free Radic. Biol. Med., 2000. 29(3-4): 222–230. 11. Hsu, J.L., Y. Hsieh, C. Tu, D. O’Connor, H.S. Nick, and D.N. Silverman, Catalytic properties of human manganese superoxide dismutase. J. Biol. Chem., 1996. 271(30): 17687–17691. 12. Gardner, P.R. and I. Fridovich, Inactivation-reactivation of aconitase in Escherichia coli. A sensitive measure of superoxide radical. J. Biol. Chem., 1992. 267(13): 8757–8763. 13. Han, D., E. Williams, and E. Cadenas, Mitochondrial respiratory chain-dependent generation of superoxide anion and its release into the intermembrane space. Biochem. J., 2001. 353(Pt 2): 411–416. 14. Okado-Matsumoto, A. and I. Fridovich, Subcellular distribution of superoxide dismutases (SOD) in rat liver: Cu,Zn-SOD in mitochondria. J. Biol. Chem., 2001. 276(42): 38388–38393. 15. Sturtz, L.A., K. Diekert, L.T. Jensen, R. Lill, and V.C. Culotta, A fraction of yeast Cu,Zn-superoxide dismutase and its metallochaperone, CCS, localize to the intermembrane space of mitochondria. A physiological role for SOD1 in guarding against mitochondrial oxidative damage. J. Biol. Chem., 2001. 276(41): 38084–38089. 16. Inarrea, P., Purification and determination of activity of mitochondrial cyanidesensitive superoxide dismutase in rat tissue extract. Methods Enzymol., 2002. 349: 106–114.

124

Nitric Oxide, Cell Signaling, and Gene Expression

17. Gardner, P.R., D.D. Nguyen, and C.W. White, Aconitase is a sensitive and critical target of oxygen poisoning in cultured mammalian cells and in rat lungs. Proc. Natl. Acad. Sci. USA, 1994. 91(25): 12248–12252. 18. Du, Y., C.M. Miller, and T.S. Kern, Hyperglycemia increases mitochondrial superoxide in retina and retinal cells. Free Radic. Biol. Med., 2003. 35(11): 1491–1499. 19. Bindokas, V.P., A. Kuznetsov, S. Sreenan, K.S. Polonsky, M.W. Roe, and L.H. Philipson, Visualizing superoxide production in normal and diabetic rat islets of Langerhans. J. Biol. Chem., 2003. 278(11): 9796–9801. 20. Dugan, L.L., S.L. Sensi, L.M. Canzoniero, S.D. Handran, S.M. Rothman, T.S. Lin, M.P. Goldberg, and D.W. Choi, Mitochondrial production of reactive oxygen species in cortical neurons following exposure to N-methyl-D-aspartate. J. Neurosci., 1995. 15(10): 6377–6388. 21. Li, Q.Y., C. Pedersen, B.J. Day, and M. Patel, Dependence of excitotoxic neurodegeneration on mitochondrial aconitase inactivation. J. Neurochem., 2001. 78(4): 746–755. 22. Luetjens, C.M., N.T. Bui, B. Sengpiel, G. Munstermann, M. Poppe, A.J. Krohn, E. Bauerbach, J. Krieglstein, and J.H. Prehn, Delayed mitochondrial dysfunction in excitotoxic neuron death: cytochrome c release and a secondary increase in superoxide production. J. Neurosci., 2000. 20(15): 5715–5723. 23. Henry, Y., M. Lepoivre, J.C. Drapier, C. Ducrocq, J.L. Boucher, and A. Guissani, EPR characterization of molecular targets for NO in mammalian cells and organelles. FASEB J., 1993. 7(12): 1124–1134. 24. Boveris, A., S. Alvarez, and A. Navarro, The role of mitochondrial nitric oxide synthase in inflammation and septic shock. Free Radic. Biol. Med., 2002. 33(9): 1186–1193. 25. Khadour, F.H., D. Panas, P. Ferdinandy, C. Schulze, T. Csont, M.M. Lalu, S.M. Wildhirt, and R. Schulz, Enhanced NO and superoxide generation in dysfunctional hearts from endotoxemic rats. Am. J. Physiol. Heart Circ. Physiol., 2002. 283(3): H1108– H1115. 26. Boczkowski, J., S. Lanone, D. Ungureanu-Longrois, G. Danialou, T. Fournier, and M. Aubier, Induction of diaphragmatic nitric oxide synthase after endotoxin administration in rats: role on diaphragmatic contractile dysfunction. J. Clin. Invest., 1996. 98(7): 1550–1559. 27. Boczkowski, J., C.L. Lisdero, S. Lanone, A. Samb, M.C. Carreras, A. Boveris, M. Aubier, and J.J. Poderoso, Endogenous peroxynitrite mediates mitochondrial dysfunction in rat diaphragm during endotoxemia. FASEB J., 1999. 13(12): 1637–1646. 28. Eliasson, M.J., Z. Huang, R.J. Ferrante, M. Sasamata, M.E. Molliver, S.H. Snyder, and M.A. Moskowitz, Neuronal nitric oxide synthase activation and peroxynitrite formation in ischemic stroke linked to neural damage. J. Neurosci., 1999. 19(14): 5910–5918. 29. Heales, S.J., J.P. Bolanos, V.C. Stewart, P.S. Brookes, J.M. Land, and J.B. Clark, Nitric oxide, mitochondria and neurological disease. Biochim. Biophys. Acta, 1999. 1410(2): 215–228. 30. Johnson, A.W., J.M. Land, E.J. Thompson, J.P. Bolanos, J.B. Clark, and S.J. Heales, Evidence for increased nitric oxide production in multiple sclerosis. J Neurol. Neurosurg. Psychiatry, 1995. 58(1): 107.

Peroxynitrite

125

31. Ghafourifar, P., U. Schenk, S.D. Klein, and C. Richter, Mitochondrial nitric-oxide synthase stimulation causes cytochrome c release from isolated mitochondria. Evidence for intramitochondrial peroxynitrite formation. J. Biol. Chem., 1999. 274(44): 31185–31188. 32. Ghafourifar, P. and C. Richter, Nitric oxide synthase activity in mitochondria. FEBS Lett., 1997. 418(3): 291–296. 33. Kanai, A.J., L.L. Pearce, P.R. Clemens, L.A. Birder, M.M. VanBibber, S.Y. Choi, W.C. de Groat, and J. Peterson, Identification of a neuronal nitric oxide synthase in isolated cardiac mitochondria using electrochemical detection. Proc. Natl. Acad. Sci. USA, 2001. 98(24): 14126–14131. 34. French, S., C. Giulivi, and R.S. Balaban, Nitric oxide synthase in porcine heart mitochondria: evidence for low physiological activity. Am. J. Physiol. Heart Circ. Physiol., 2001. 280(6): H2863– H2867. 35. Tatoyan, A. and C. Giulivi, Purification and characterization of a nitric-oxide synthase from rat liver mitochondria. J. Biol. Chem., 1998. 273(18): 11044–11048. 36. Giulivi, C., Characterization and function of mitochondrial nitric-oxide synthase. Free Radic. Biol. Med., 2003. 34(4): 397–408. 37. Kanai, A., M. Epperly, L. Pearce, L. Birder, M. Zeidel, S. Meyers, J. Greenberger, W. de Groat, G. Apodaca, and J. Peterson, Differing roles of mitochondrial nitric oxide synthase in cardiomyocytes and urothelial cells. Am. J. Physiol. Heart Circ. Physiol., 2004. 286(1): H13–21. 38. Giulivi, C., Functional implications of nitric oxide produced by mitochondria in mitochondrial metabolism. Biochem. J., 1998. 332 (Pt 3): 673–679. 39. Sarkela, T.M., J. Berthiaume, S. Elfering, A.A. Gybina, and C. Giulivi, The modulation of oxygen radical production by nitric oxide in mitochondria. J. Biol. Chem., 2001. 276(10): 6945–6949. 40. Kozlov, A.V., K. Staniek, and H. Nohl, Nitrite reductase activity is a novel function of mammalian mitochondria. FEBS Lett., 1999. 454(1-2): 127–130. 40a. Quijano, C., N. Romero, and R. Radi, Tyrosine nitration by superoxide and nitric oxide fluxes in biological systems: Modeling the impact of superoxide dismutase and nitric oxide diffusion. Free Radic. Biol. Med., 2005. In press. 41. Takahashi, M.A. and K. Asada, Superoxide anion permeability of phospholipid membranes and chloroplast thylakoids. Arch. Biochem. Biophys., 1983. 226(2): 558–566. 42. Mao, G.D. and M.J. Poznansky, Electron spin resonance study on the permeability of superoxide radicals in lipid bilayers and biological membranes. FEBS Lett., 1992. 305(3): 233–236. 43. Lynch, R.E. and I. Fridovich, Permeation of the erythrocyte stroma by superoxide radical. J. Biol. Chem., 1978. 253(13): 4697–4699. 44. Han, D., F. Antunes, R. Canali, D. Rettori, and E. Cadenas, Voltage-dependent anion channels control the release of the superoxide anion from mitochondria to cytosol. J. Biol. Chem., 2003. 278(8): 5557–5563. 45. Malinski, T., Z. Taha, S. Grunfeld, S. Patton, M. Kapturczak, and P. Tomboulian, Diffusion of nitric oxide in the aorta wall monitored in situ by porphyrinic microsensors. Biochem. Biophys. Res. Commun., 1993. 193(3): 1076–1082. 46. Denicola, A., J.M. Souza, R. Radi, and E. Lissi, Nitric oxide diffusion in membranes determined by fluorescence quenching. Arch. Biochem. Biophys., 1996. 328(1): 208–212.

126

Nitric Oxide, Cell Signaling, and Gene Expression

47. Vanderkooi, J.M., W.W. Wright, and M. Erecinska, Nitric oxide diffusion coefficients in solutions, proteins and membranes determined by phosphorescence. Biochim. Biophys. Acta, 1994. 1207(2): 249–254. 48. Clementi, E., G.C. Brown, N. Foxwell, and S. Moncada, On the mechanism by which vascular endothelial cells regulate their oxygen consumption. Proc. Natl. Acad. Sci. USA, 1999. 96(4): 1559–1562. 49. Clementi, E., G.C. Brown, M. Feelisch, and S. Moncada, Persistent inhibition of cell respiration by nitric oxide: crucial role of S-nitrosylation of mitochondrial complex I and protective action of glutathione. Proc. Natl. Acad. Sci. USA, 1998. 95(13): 7631–7636. 50. Miranda, K.M., M.G. Espey, D. Jourd’heuil, M.B. Grisham, J.M. Fukuto, M. Feelisch, and D.A. Wink, The Chemical Biology of Nitric Oxide, in Nitric oxide biology and pathobiology, J.L. Ignarro, Ed. 2000, Academic Press: San Diego. p. 41–55. 51. Brown, G.C. and C.E. Cooper, Nanomolar concentrations of nitric oxide reversibly inhibit synaptosomal respiration by competing with oxygen at cytochrome oxidase. FEBS Lett., 1994. 356(2-3): 295–298. 52. Cleeter, M.W., J.M. Cooper, V.M. Darley-Usmar, S. Moncada, and A.H. Schapira, Reversible inhibition of cytochrome c oxidase, the terminal enzyme of the mitochondrial respiratory chain, by nitric oxide. Implications for neurodegenerative diseases. FEBS Lett., 1994. 345(1): 50–54. 53. Cassina, A. and R. Radi, Differential inhibitory action of nitric oxide and peroxynitrite on mitochondrial electron transport. Arch. Biochem. Biophys., 1996. 328(2): 309–316. 54. Poderoso, J.J., M.C. Carreras, C. Lisdero, N. Riobo, F. Schopfer, and A. Boveris, Nitric oxide inhibits electron transfer and increases superoxide radical production in rat heart mitochondria and submitochondrial particles. Arch. Biochem. Biophys., 1996. 328(1): 85–92. 55. Pearce, L.L., A.J. Kanai, L.A. Birder, B.R. Pitt, and J. Peterson, The catabolic fate of nitric oxide: the nitric oxide oxidase and peroxynitrite reductase activities of cytochrome oxidase. J. Biol. Chem., 2002. 277(16): 13556–13562. 56. Elfering, S.L., V.L. Haynes, N.J. Traaseth, A. Ettl, and C. Giulivi, Aspects, mechanism, and biological relevance of mitochondrial protein nitration sustained by mitochondrial nitric oxide synthase. Am. J. Physiol. Heart Circ. Physiol., 2004. 286(1): H22–H29. 57. Radi, R., A. Cassina, R. Hodara, C. Quijano, and L. Castro, Peroxynitrite reactions and formation in mitochondria. Free Radic. Biol. Med., 2002. 33(11): 1451–1464. 58. Hogg, N., R.J. Singh, and B. Kalyanaraman, The role of glutathione in the transport and catabolism of nitric oxide. FEBS Lett., 1996. 382(3): 223–238. 59. Shafirovich, V. and S.V. Lymar, Nitroxyl and its anion in aqueous solutions: spin states, protic equilibria, and reactivities toward oxygen and nitric oxide. Proc. Natl. Acad. Sci. USA, 2002. 99(11): 7340–7345. 60. Poderoso, J.J., M.C. Carreras, F. Schopfer, C.L. Lisdero, N.A. Riobo, C. Giulivi, A.D. Boveris, A. Boveris, and E. Cadenas, The reaction of nitric oxide with ubiquinol: kinetic properties and biological significance. Free Radic. Biol. Med., 1999. 26(7-8): 925–935. 61. Sharpe, M.A. and C.E. Cooper, Reactions of nitric oxide with mitochondrial cytochrome c: a novel mechanism for the formation of nitroxyl anion and peroxynitrite. Biochem. J., 1998. 332 (Pt 1): 9–19.

Peroxynitrite

127

62. Lymar, S.V. and J.K. Hurst, Rapid reaction between peroxynitrite ion and carbon dioxide: implications for biological activity. J. Am. Chem. Soc., 1995. 117: 8867–8868. 63. Denicola, A., B.A. Freeman, M. Trujillo, and R. Radi, Peroxynitrite reaction with carbon dioxide/bicarbonate: kinetics and influence on peroxynitrite-mediated oxidations. Arch. Biochem. Biophys., 1996. 333(1): 49–58. 64. Bonini, M.G., R. Radi, G. Ferrer-Sueta, A.M. Ferreira, and O. Augusto, Direct EPR detection of the carbonate radical anion produced from peroxynitrite and carbon dioxide. J. Biol. Chem., 1999. 274(16): 10802–10806. 65. Yamakura, F., H. Taka, T. Fujimura, and K. Murayama, Inactivation of human manganese-superoxide dismutase by peroxynitrite is caused by exclusive nitration of tyrosine 34 to 3-nitrotyrosine. J. Biol. Chem., 1998. 273(23): 14085–14089. 66. MacMillan-Crow, L.A., J.P. Crow, and J.A. Thompson, Peroxynitrite-mediated inactivation of manganese superoxide dismutase involves nitration and oxidation of critical tyrosine residues. Biochemistry, 1998. 37(6): 1613–1622. 67. Quijano, C., D. Hernandez-Saavedra, L. Castro, J.M. McCord, B.A. Freeman, and R. Radi, Reaction of peroxynitrite with Mn-superoxide dismutase. Role of the metal center in decomposition kinetics and nitration. J. Biol. Chem., 2001. 276(15): 11631–11638. 68. Lancaster, J.J. Nitroxidative stress: predicting the metal-independent oxidative chemistry of reactive nitrogen oxide species under physiological conditions, in SFRR’s 12th Biennal Meeting. 2004. Buenos Aires, Argentina: Free Radic. Biol. Med. 36(1): S4. 69. Radi, R., J.S. Beckman, K.M. Bush, and B.A. Freeman, Peroxynitrite oxidation of sulfhydryls. The cytotoxic potential of superoxide and nitric oxide. J. Biol. Chem., 1991. 266(7): 4244–4250. 70. Beckman, J.S., H. Ischiropoulos, L. Zhu, M. van der Woerd, C. Smith, J. Chen, J. Harrison, J.C. Martin, and M. Tsai, Kinetics of superoxide dismutase- and ironcatalyzed nitration of phenolics by peroxynitrite. Arch. Biochem. Biophys., 1992. 298(2): 438–445. 71. Alvarez, B., H. Rubbo, M. Kirk, S. Barnes, B.A. Freeman, and R. Radi, Peroxynitrite-dependent tryptophan nitration. Chem. Res. Toxicol., 1996. 9(2): 390–396. 72. Brown, A.S., M.A. Moro, J.M. Masse, E.M. Cramer, M. Radomski, and V. DarleyUsmar, Nitric oxide-dependent and independent effects on human platelets treated with peroxynitrite. Cardiovasc. Res., 1998. 40(2): 380–388. 73. Thomson, L., A. Denicola, and R. Radi, The trypanothione-thiol system in Trypanosoma cruzi as a key antioxidant mechanism against peroxynitrite-mediated cytotoxicity. Arch. Biochem. Biophys., 2003. 412(1): 55–64. 74. Lin, T.K., G. Hughes, A. Muratovska, F.H. Blaikie, P.S. Brookes, V. Darley-Usmar, R.A. Smith, and M.P. Murphy, Specific modification of mitochondrial protein thiols in response to oxidative stress: a proteomics approach. J. Biol. Chem., 2002. 277(19): 17048–17056. 75. Scarlett, J.L., M.A. Packer, C.M. Porteous, and M.P. Murphy, Alterations to glutathione and nicotinamide nucleotides during the mitochondrial permeability transition induced by peroxynitrite. Biochem. Pharmacol., 1996. 52(7): 1047–1055. 76. Ramezanian, M.S., S. Padmaja, and W.H. Koppenol, Nitration and hydroxylation of phenolic compounds by peroxynitrite. Chem. Res. Toxicol., 1996. 9(1): 232–240.

128

Nitric Oxide, Cell Signaling, and Gene Expression

77. Alvarez, B. and R. Radi, Peroxynitrite reactivity with amino acids and proteins. Amino Acids, 2003. 25(3-4): 295–311. 78. Lymar, S.V., Q. Jiang, and J.K. Hurst, Mechanism of carbon dioxide-catalyzed oxidation of tyrosine by peroxynitrite. Biochemistry, 1996. 35(24): 7855–7861. 79. Lymar, S.V. and J.K. Hurst, CO2-Catalyzed one-electron oxidations by peroxynitrite: properties of the reactive intermediate. Inorg. Chem., 1998. 37: 294–301. 80. Ferrer-Sueta, G., L. Ruiz-Ramirez, and R. Radi, Ternary copper complexes and manganese (III) tetrakis(4-benzoic acid) porphyrin catalyze peroxynitrite-dependent nitration of aromatics. Chem. Res. Toxicol., 1997. 10(12): 1338–1344. 81. Ferrer-Sueta, G., I. Batinic-Haberle, I. Spasojevic, I. Fridovich, and R. Radi, Catalytic scavenging of peroxynitrite by isomeric Mn(III) N-methylpyridylporphyrins in the presence of reductants. Chem. Res. Toxicol., 1999. 12(5): 442–429. 82. Crow, J.P., Manganese and iron porphyrins catalyze peroxynitrite decomposition and simultaneously increase nitration and oxidant yield: implications for their use as peroxynitrite scavengers in vivo. Arch. Biochem. Biophys., 1999. 371(1): 41–52. 83. Alvarez, B., V. Demicheli, R. Durán, M. Trujillo, C. Cerveñansky, B.A. Freeman, and R. Radi, Inactivation of human Cu,Zn Superoxide dismutase by peroxynitrite and formation of hystidinyl radical. Free Radic. Biol. Med., 2004. 37(6): 813–822. 84. Ischiropoulos, H., L. Zhu, J. Chen, M. Tsai, J.C. Martin, C.D. Smith, and J.S. Beckman, Peroxynitrite-mediated tyrosine nitration catalyzed by superoxide dismutase. Arch. Biochem. Biophys., 1992. 298(2): 431–437. 85. Kono, Y., T. Yamasaki, A. Ueda, and H. Shibata, Catalase catalyzes of peroxynitrite-mediated phenolic nitration. Biosci. Biotechnol. Biochem., 1998. 62(3): 448–452. 86. Radi, R., Nitric oxide, oxidants, and protein tyrosine nitration. Proc. Natl. Acad. Sci. USA, 2004. 101(12): 4003–4008. 87. Baldus, S., J.P. Eiserich, M.L. Brennan, R.M. Jackson, C.B. Alexander, and B.A. Freeman, Spatial mapping of pulmonary and vascular nitrotyrosine reveals the pivotal role of myeloperoxidase as a catalyst for tyrosine nitration in inflammatory diseases. Free Radic. Biol. Med., 2002. 33(7): 1010–1019. 88. Eiserich, J.P., M. Hristova, C.E. Cross, A.D. Jones, B.A. Freeman, B. Halliwell, and A. van der Vliet, Formation of nitric oxide-derived inflammatory oxidants by myeloperoxidase in neutrophils. Nature, 1998. 391(6665): 393–397. 89. Baldus, S., J.P. Eiserich, A. Mani, L. Castro, M. Figueroa, P. Chumley, W. Ma, A. Tousson, C.R. White, D.C. Bullard, M.L. Brennan, A.J. Lusis, K.P. Moore, and B.A. Freeman, Endothelial transcytosis of myeloperoxidase confers specificity to vascular ECM proteins as targets of tyrosine nitration. J. Clin. Invest., 2001. 108(12): 1759–1770. 90. Brennan, M.L., W. Wu, X. Fu, Z. Shen, W. Song, H. Frost, C. Vadseth, L. Narine, E. Lenkiewicz, M.T. Borchers, A.J. Lusis, J.J. Lee, N.A. Lee, H.M. Abu-Soud, H. Ischiropoulos, and S.L. Hazen, A tale of two controversies: defining both the role of peroxidases in nitrotyrosine formation in vivo using eosinophil peroxidase and myeloperoxidase-deficient mice, and the nature of peroxidase-generated reactive nitrogen species. J. Biol. Chem., 2002. 277(20): 17415–17427. 91. Gaut, J.P., J. Byun, H.D. Tran, W.M. Lauber, J.A. Carroll, R.S. Hotchkiss, A. Belaaouaj, and J.W. Heinecke, Myeloperoxidase produces nitrating oxidants in vivo. J. Clin. Invest., 2002. 109(10): 1311–139.

Peroxynitrite

129

92. van der Vliet, A., J.P. Eiserich, B. Halliwell, and C.E. Cross, Formation of reactive nitrogen species during peroxidase-catalyzed oxidation of nitrite. A potential additional mechanism of nitric oxide-dependent toxicity. J. Biol. Chem., 1997. 272(12): 7617–7625. 93. Prutz, W.A., H. Monig, J. Butler, and E.J. Land, Reactions of nitrogen dioxide in aqueous model systems: oxidation of tyrosine units in peptides and proteins. Arch. Biochem. Biophys., 1985. 243(1): 125–134. 94. Ford, E., M.N. Hughes, and P. Wardman, Kinetics of the reactions of nitrogen dioxide with glutathione, cysteine, and uric acid at physiological pH. Free Radic. Biol. Med., 2002. 32(12): 1314–1323. 95. Radi, R.A., H. Rubbo, and E. Prodanov, Comparison of the effects of superoxide dismutase and cytochrome c on luminol chemiluminescence produced by xanthine oxidase-catalyzed reactions. Biochim. Biophys. Acta, 1989. 994(1): 89–93. 96. Radi, R., J.F. Turrens, and B.A. Freeman, Cytochrome c-catalyzed membrane lipid peroxidation by hydrogen peroxide. Arch. Biochem. Biophys., 1991. 288(1): 118–125. 97. Radi, R., L. Thomson, H. Rubbo, and E. Prodanov, Cytochrome c-catalyzed oxidation of organic molecules by hydrogen peroxide. Arch. Biochem. Biophys., 1991. 288(1): 112–117. 98. Radi, R., S. Sims, A. Cassina, and J.F. Turrens, Roles of catalase and cytochrome c in hydroperoxide-dependent lipid peroxidation and chemiluminescence in rat heart and kidney mitochondria. Free Radic. Biol. Med., 1993. 15(6): 653–659. 99. Radi, R., K.M. Bush, and B.A. Freeman, The role of cytochrome c and mitochondrial catalase in hydroperoxide-induced heart mitochondrial lipid peroxidation. Arch. Biochem. Biophys., 1993. 300(1): 409–415. 100. Cassina, A.M., R. Hodara, J.M. Souza, L. Thomson, L. Castro, H. Ischiropoulos, B.A. Freeman, and R. Radi, Cytochrome c nitration by peroxynitrite. J. Biol. Chem., 2000. 275(28): 21409–21415. 100a. Batthyány, C., J.M. Souta, R. Durán, A. Cassina, C. Cerveñansky, and R. Radi, Time course and site(s) of cytochrome c tyrosine nitration by peroxynitrate. Biochemistry, 2005. 44:8038–8046. 101. Castro, L., J.P. Eiserich, S. Sweeney, R. Radi, and B.A. Freeman, Cytochrome c: a catalyst and target of nitrite-hydrogen peroxide-dependent protein nitration. Arch. Biochem. Biophys., 2004. 421(1): 99–107. 102. Koppenol, W.H., The centennial of the Fenton reaction. Free Radic. Biol. Med., 1993. 15(6): 645–651. 103. Welch, K.D., T.Z. Davis, M.E. Van Eden, and S.D. Aust, Deleterious iron-mediated oxidation of biomolecules. Free Radic. Biol. Med., 2002. 32(7): 577–583. 104. Bian, K., Z. Gao, N. Weisbrodt, and F. Murad, The nature of heme/iron-induced protein tyrosine nitration. Proc. Natl. Acad. Sci. USA, 2003. 100(10): 5712–5717. 105. Thomas, D.D., M.G. Espey, M.P. Vitek, K.M. Miranda, and D.A. Wink, Protein nitration is mediated by heme and free metals through Fenton-type chemistry: an alternative to the NO/O2- reaction. Proc. Natl. Acad. Sci. USA, 2002. 99(20): 12691–12696. 106. Hausladen, A. and I. Fridovich, Superoxide and peroxynitrite inactivate aconitases, but nitric oxide does not. J. Biol. Chem., 1994. 269(47): 29405–29408.

130

Nitric Oxide, Cell Signaling, and Gene Expression

107. Castro, L.A., R.L. Robalinho, A. Cayota, R. Meneghini, and R. Radi, Nitric oxide and peroxynitrite-dependent aconitase inactivation and iron-regulatory protein-1 activation in mammalian fibroblasts. Arch. Biochem. Biophys., 1998. 359(2): 215–224. 108. Ramachandran, A., E. Ceaser, and V.M. Darley-Usmar, Chronic exposure to nitric oxide alters the free iron pool in endothelial cells: role of mitochondrial respiratory complexes and heat shock proteins. Proc. Natl. Acad. Sci. USA, 2004. 101(1): 384–389. 109. Gunther, M.R., L.C. Hsi, J.F. Curtis, J.K. Gierse, L.J. Marnett, T.E. Eling, and R.P. Mason, Nitric oxide trapping of the tyrosyl radical of prostaglandin H synthase-2 leads to tyrosine iminoxyl radical and nitrotyrosine formation. J. Biol. Chem., 1997. 272(27): 17086–17090. 110. Chen, Y.R., C.L. Chen, W. Chen, J.L. Zweier, O. Augusto, R. Radi, and R.P. Mason, Formation of protein tyrosine ortho-semiquinone radical and nitrotyrosine from cytochrome c-derived tyrosyl radical. J. Biol. Chem., 2004. 279(17): 18054–18062. 111. Goldstein, S., G. Czapski, J. Lind, and G. Merenyi, Tyrosine nitration by simultaneous generation of (.)NO and O-(2) under physiological conditions. How the radicals do the job. J. Biol. Chem., 2000. 275(5): 3031–3036. 112. Turko, I.V. and F. Murad, Protein nitration in cardiovascular diseases. Pharmacol. Rev., 2002. 54(4): 619–634. 113. Aulak, K.S., M. Miyagi, L. Yan, K.A. West, D. Massillon, J.W. Crabb, and D.J. Stuehr, Proteomic method identifies proteins nitrated in vivo during inflammatory challenge. Proc. Natl. Acad. Sci. USA, 2001. 98(21): 12056–12061. 114. Turko, I.V., L. Li, K.S. Aulak, D.J. Stuehr, J.Y. Chang, and F. Murad, Protein tyrosine nitration in the mitochondria from diabetic mouse heart. Implications to dysfunctional mitochondria in diabetes. J. Biol. Chem., 2003. 278(36): 33972–33977. 115. Radi, R., M. Rodriguez, L. Castro, and R. Telleri, Inhibition of mitochondrial electron transport by peroxynitrite. Arch. Biochem. Biophys., 1994. 308(1): 89–95. 116. Riobo, N.A., E. Clementi, M. Melani, A. Boveris, E. Cadenas, S. Moncada, and J.J. Poderoso, Nitric oxide inhibits mitochondrial NADH: ubiquinone reductase activity through peroxynitrite formation. Biochem. J., 2001. 359(Pt 1): 139–145. 117. Bolanos, J.P., S.J. Heales, J.M. Land, and J.B. Clark, Effect of peroxynitrite on the mitochondrial respiratory chain: differential susceptibility of neurones and astrocytes in primary culture. J. Neurochem., 1995. 64(5): 1965–1972. 118. Xie, Y.W. and M.S. Wolin, Role of nitric oxide and its interaction with superoxide in the suppression of cardiac muscle mitochondrial respiration. Involvement in response to hypoxia/reoxygenation. Circulation, 1996. 94(10): 2580–2586. 119. Murray, J., S.W. Taylor, B. Zhang, S.S. Ghosh, and R.A. Capaldi, Oxidative damage to mitochondrial complex I due to peroxynitrite: identification of reactive tyrosines by mass spectrometry. J. Biol. Chem., 2003. 278(39): 37223–37230. 120. Yamamoto, T., W. Maruyama, Y. Kato, H. Yi, M. Shamoto-Nagai, M. Tanaka, Y. Sato, and M. Naoi, Selective nitration of mitochondrial complex I by peroxynitrite: involvement in mitochondria dysfunction and cell death of dopaminergic SHSY5Y cells. J. Neural. Transm., 2002. 109(1): 1–13. 121. Brealey, D., M. Brand, I. Hargreaves, S. Heales, J. Land, R. Smolenski, N.A. Davies, C.E. Cooper, and M. Singer, Association between mitochondrial dysfunction and severity and outcome of septic shock. Lancet, 2002. 360(9328): 219–223.

Peroxynitrite

131

122. Tomita, M., S. Mukae, E. Geshi, K. Umetsu, M. Nakatani, and T. Katagiri, Mitochondrial respiratory impairment in streptozotocin-induced diabetic rat heart. Jpn. Circ. J., 1996. 60(9): 673–682. 123. Hederstedt, L. and T. Oshnishi, Progressin succinate: quinone oxidoreductase research, in Bioenergetics, L. Ernster, ed. 1992, Elsevier Science B.V.: Amsterdam. pp. 199–216. 124. Rubbo, H., A. Denicola, and R. Radi, Peroxynitrite inactivates thiol-containing enzymes of Trypanosoma cruzi energetic metabolism and inhibits cell respiration. Arch. Biochem. Biophys., 1994. 308(1): 96–102. 125. Aulak, K.S., T. Koeck, J.W. Crabb, and D.J. Stuehr, Dynamics of protein nitration in cells and mitochondria. Am. J. Physiol. Heart Circ. Physiol., 2004. 286(1): H30–38. 126. Kagawa, Y., Proton motive ATP synthesis, in Bioenergetics, L. Ernster, ed. 1984, Elsevier Science B.V.: Amsterdam. pp. 149–186. 127. Thomson, L., M. Trujillo, R. Telleri, and R. Radi, Kinetics of cytochrome c2+ oxidation by peroxynitrite: implications for superoxide measurements in nitric oxide-producing biological systems. Arch. Biochem. Biophys., 1995. 319(2): 491–497. 128. MacMillan-Crow, L.A., D.L. Cruthirds, K.M. Ahki, P.W. Sanders, and J.A. Thompson, Mitochondrial tyrosine nitration precedes chronic allograft nephropathy. Free Radic. Biol. Med., 2001. 31(12): 1603–1608. 129. Cruthirds, D.L., L. Novak, K.M. Akhi, P.W. Sanders, J.A. Thompson, and L.A. MacMillan-Crow, Mitochondrial targets of oxidative stress during renal ischemia/reperfusion. Arch. Biochem. Biophys., 2003. 412(1): 27–33. 130. Alonso, D., J.M. Encinas, L.O. Uttenthal, L. Bosca, J. Serrano, A.P. Fernandez, S. Castro-Blanco, M. Santacana, M.L. Bentura, A. Richart, P. Fernandez-Vizarra, and J. Rodrigo, Coexistence of translocated cytochrome c and nitrated protein in neurons of the rat cerebral cortex after oxygen and glucose deprivation. Neuroscience, 2002. 111(1): 47–56. 131. Cooper, C.E., N.A. Davies, M. Psychoulis, L. Canevari, T.E. Bates, M.S. Dobbie, C.S. Casley, and M.A. Sharpe, 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(1): 27–34. 132. Sharpe, M.A. and C.E. Cooper, Interaction of peroxynitrite with mitochondrial cytochrome oxidase. Catalytic production of nitric oxide and irreversible inhibition of enzyme activity. J. Biol. Chem., 1998. 273(47): 30961–30972. 133. Pearce, L.L., B.R. Pitt, and J. Peterson, The peroxynitrite reductase activity of cytochrome c oxidase involves a two-electron redox reaction at the heme a(3)Cu(B) site. J. Biol. Chem., 1999. 274(50): 35763–35767. 134. Spindler, M., K.W. Saupe, R. Tian, S. Ahmed, M.A. Matlib, and J.S. Ingwall, Altered creatine kinase enzyme kinetics in diabetic cardiomyopathy. A(31)P NMR magnetization transfer study of the intact beating rat heart. J. Mol. Cell Cardiol., 1999. 31(12): 2175–2189. 135. Soboll, S., D. Brdiczka, D. Jahnke, A. Schmidt, U. Schlattner, S. Wendt, M. Wyss, and T. Wallimann, Octamer-dimer transitions of mitochondrial creatine kinase in heart disease. J. Mol. Cell Cardiol., 1999. 31(4): 857–866. 136. Wendt, S., A. Dedeoglu, O. Speer, T. Wallimann, M.F. Beal, and O.A. Andreassen, Reduced creatine kinase activity in transgenic amyotrophic lateral sclerosis mice. Free Radic. Biol. Med., 2002. 32(9): 920–926.

132

Nitric Oxide, Cell Signaling, and Gene Expression

137. Stachowiak, O., M. Dolder, T. Wallimann, and C. Richter, Mitochondrial creatine kinase is a prime target of peroxynitrite-induced modification and inactivation. J. Biol. Chem., 1998. 273(27): 16694–16699. 138. Wendt, S., U. Schlattner, and T. Wallimann, Differential effects of peroxynitrite on human mitochondrial creatine kinase isoenzymes. Inactivation, octamer destabilization, and identification of involved residues. J. Biol. Chem., 2003. 278(2): 1125–1130. 139. Konorev, E.A., N. Hogg, and B. Kalyanaraman, Rapid and irreversible inhibition of creatine kinase by peroxynitrite. FEBS Lett., 1998. 427(2): 171–174. 140. Dolder, M., S. Wendt, and T. Wallimann, Mitochondrial creatine kinase in contact sites: interaction with porin and adenine nucleotide translocase, role in permeability transition and sensitivity to oxidative damage. Biol. Signals Recept., 2001. 10(1-2): 93–111. 141. Fukumoto, K., A. Pierro, L. Spitz, and S. Eaton, Differential effects of neonatal endotoxemia on heart and kidney carnitine palmitoyl transferase I. J. Pediatr. Surg., 2002. 37(5): 723–726. 142. Eaton, S., K. Fukumoto, G. Stefanutti, L. Spitz, V.A. Zammit, and A. Pierro, Myocardial carnitine palmitoyltransferase I as a target for oxidative modification in inflammation and sepsis. Biochem. Soc. Trans., 2003. 31(Pt 6): 1133–1136. 143. Castro, L., M. Rodriguez, and R. Radi, Aconitase is readily inactivated by peroxynitrite, but not by its precursor, nitric oxide. J. Biol. Chem., 1994. 269(47): 29409–29415. 144. Gardner, P.R., G. Costantino, C. Szabo, and A.L. Salzman, Nitric oxide sensitivity of the aconitases. J. Biol. Chem., 1997. 272(40): 25071–25076. 145. Lee, J.H., E.S. Yang, and J.W. Park, Inactivation of NADP+-dependent isocitrate dehydrogenase by peroxynitrite. Implications for cytotoxicity and alcohol-induced liver injury. J. Biol. Chem., 2003. 278(51): 51360–51371. 146. Park, L.C., H. Zhang, K.F. Sheu, N.Y. Calingasan, B.S. Kristal, J.G. Lindsay, and G.E. Gibson, Metabolic impairment induces oxidative stress, compromises inflammatory responses, and inactivates a key mitochondrial enzyme in microglia. J. Neurochem., 1999. 72(5): 1948–1958. 147. Mastrogiacomo, F., C. Bergeron, and S.J. Kish, Brain alpha-ketoglutarate dehydrogenase complex activity in Alzheimer’s disease. J. Neurochem., 1993. 61(6): 2007–2014. 148. Mizuno, Y., S. Matuda, H. Yoshino, H. Mori, N. Hattori, and S. Ikebe, An immunohistochemical study on alpha-ketoglutarate dehydrogenase complex in Parkinson’s disease. Ann. Neurol., 1994. 35(2): 204–210. 149. Marcondes, S., I.V. Turko, and F. Murad, Nitration of succinyl-CoA:3-oxoacid CoA-transferase in rats after endotoxin administration. Proc. Natl. Acad. Sci. USA, 2001. 98(13): 7146–7151. 150. Turko, I.V., S. Marcondes, and F. Murad, Diabetes-associated nitration of tyrosine and inactivation of succinyl-CoA:3-oxoacid CoA-transferase. Am. J. Physiol. Heart Circ. Physiol., 2001. 281(6): H2289– H2294. 151. Crompton, M., The mitochondrial permeability transition pore and its role in cell death. Biochem. J., 1999. 341 (Pt 2): 233–249. 152. Rego, A.C. and C.R. Oliveira, Mitochondrial dysfunction and reactive oxygen species in excitotoxicity and apoptosis: implications for the pathogenesis of neurodegenerative diseases. Neurochem. Res., 2003. 28(10): 1563–1574.

Peroxynitrite

133

153. Iverson, S.L. and S. Orrenius, The cardiolipin-cytochrome c interaction and the mitochondrial regulation of apoptosis. Arch. Biochem. Biophys., 2004. 423(1): 37–46. 154. Estevez, A.G., R. Radi, L. Barbeito, J.T. Shin, J.A. Thompson, and J.S. Beckman, Peroxynitrite-induced cytotoxicity in PC12 cells: evidence for an apoptotic mechanism differentially modulated by neurotrophic factors. J. Neurochem., 1995. 65(4): 1543–1550. 155. Salgo, M.G., E. Bermudez, G.L. Squadrito, and W.A. Pryor, Peroxynitrite causes DNA damage and oxidation of thiols in rat thymocytes [corrected]. Arch. Biochem. Biophys., 1995. 322(2): 500–505. 156. Salgo, M.G., G.L. Squadrito, and W.A. Pryor, Peroxynitrite causes apoptosis in rat thymocytes. Biochem. Biophys. Res. Commun., 1995. 215(3): 1111–1118. 157. Keller, J.N., M.S. Kindy, F.W. Holtsberg, D.K. St. Clair, H.C. Yen, A. Germeyer, S.M. Steiner, A.J. Bruce-Keller, J.B. Hutchins, and M.P. Mattson, Mitochondrial manganese superoxide dismutase prevents neural apoptosis and reduces ischemic brain injury: suppression of peroxynitrite production, lipid peroxidation, and mitochondrial dysfunction. J. Neurosci., 1998. 18(2): 687–697. 158. Estevez, A.G., N. Spear, S.M. Manuel, R. Radi, C.E. Henderson, L. Barbeito, and J.S. Beckman, Nitric oxide and superoxide contribute to motor neuron apoptosis induced by trophic factor deprivation. J. Neurosci., 1998. 18(3): 923–931. 159. Hortelano, S., A.M. Alvarez, and L. Bosca, Nitric oxide induces tyrosine nitration and release of cytochrome c preceding an increase of mitochondrial transmembrane potential in macrophages. FASEB J., 1999. 13(15): 2311–2317. 160. Hortelano, S., A. Castrillo, A.M. Alvarez, and L. Bosca, Contribution of cyclopentenone prostaglandins to the resolution of inflammation through the potentiation of apoptosis in activated macrophages. J. Immunol., 2000. 165(11): 6525–6531. 161. Zou, M.H., C. Shi, and R.A. Cohen, High glucose via peroxynitrite causes tyrosine nitration and inactivation of prostacyclin synthase that is associated with thromboxane/prostaglandin H(2) receptor-mediated apoptosis and adhesion molecule expression in cultured human aortic endothelial cells. Diabetes, 2002. 51(1): 198–203. 162. Gadelha, F.R., L. Thomson, M.M. Fagian, A.D. Costa, R. Radi, and A.E. Vercesi, Ca2+-independent permeabilization of the inner mitochondrial membrane by peroxynitrite is mediated by membrane protein thiol cross-linking and lipid peroxidation. Arch. Biochem. Biophys., 1997. 345(2): 243–250. 163. Borutaite, V., R. Morkuniene, and G.C. Brown, Release of cytochrome c from heart mitochondria is induced by high Ca2+ and peroxynitrite and is responsible for Ca(2+)-induced inhibition of substrate oxidation. Biochim. Biophys. Acta, 1999. 1453(1): 41–48. 164. Vieira, H.L., A.S. Belzacq, D. Haouzi, F. Bernassola, I. Cohen, E. Jacotot, K.F. Ferri, C. El Hamel, L.M. Bartle, G. Melino, C. Brenner, V. Goldmacher, and G. Kroemer, The adenine nucleotide translocator: a target of nitric oxide, peroxynitrite, and 4-hydroxynonenal. Oncogene, 2001. 20(32): 4305–4316. 165. Brookes, P.S. and V.M. Darley-Usmar, Role of calcium and superoxide dismutase in sensitizing mitochondria to peroxynitrite-induced permeability transition. Am. J. Physiol. Heart Circ. Physiol., 2004. 286(1): H39–46.

134

Nitric Oxide, Cell Signaling, and Gene Expression

166. Schweizer, M. and C. Richter, Peroxynitrite stimulates the pyridine nucleotidelinked Ca2+ release from intact rat liver mitochondria. Biochemistry, 1996. 35(14): 4524–4528. 167 Guo, W., T. Adachi, R. Matsui, S. Xu, B. Jiang, M.H. Zou, M. Kirber, W. Lieberthal, and R.A. Cohen, Quantitative assessment of tyrosine nitration of manganese superoxide dismutase in angiotensin II-infused rat kidney. Am. J. Physiol. Heart Circ. Physiol., 2003. 285(4): H1396–1403. 168. van der Loo, B., R. Labugger, J.N. Skepper, M. Bachschmid, J. Kilo, J.M. Powell, M. Palacios-Callender, J.D. Erusalimsky, T. Quaschning, T. Malinski, D. Gygi, V. Ullrich, and T.F. Luscher, Enhanced peroxynitrite formation is associated with vascular aging. J. Exp. Med., 2000. 192(12): 1731–1744. 169. Pittman, K.M., L.A. MacMillan-Crow, B.P. Peters, and J.B. Allen, Nitration of manganese superoxide dismutase during ocular inflammation. Exp. Eye Res., 2002. 74(4): 463–471. 170. Knight-Lozano, C.A., C.G. Young, D.L. Burow, Z.Y. Hu, D. Uyeminami, K.E. Pinkerton, H. Ischiropoulos, and S.W. Ballinger, Cigarette smoke exposure and hypercholesterolemia increase mitochondrial damage in cardiovascular tissues. Circulation, 2002. 105(7): 849–854. 171. MacMillan-Crow, L.A., J.P. Crow, J.D. Kerby, J.S. Beckman, and J.A. Thompson, Nitration and inactivation of manganese superoxide dismutase in chronic rejection of human renal allografts. Proc. Natl. Acad. Sci. USA, 1996. 93(21): 11853–11858. 172. Aoyama, K., K. Matsubara, Y. Fujikawa, Y. Nagahiro, K. Shimizu, N. Umegae, N. Hayase, H. Shiono, and S. Kobayashi, Nitration of manganese superoxide dismutase in cerebrospinal fluids is a marker for peroxynitrite-mediated oxidative stress in neurodegenerative diseases. Ann. Neurol., 2000. 47(4): 524–527. 172a. Casoni, F., M. Basso, T. Massighan, E. Gianazza, C. Cheroni, M. Salmona, C. Bendotti, and V. Bonetto, Protein nitration in a mouse model of familial amyotrophic lateral sclerosis. J. Biol. Chem., 2005. 280(16): 16295–16304. 173. Meister, A., Glutathione metabolism and its selective modification. J. Biol. Chem., 1988. 263(33): 17205–17208. 174. Pryor, W.A. and G.L. Squadrito, The chemistry of peroxynitrite: a product from the reaction of nitric oxide with superoxide. Am. J. Physiol., 1995. 268(5 Pt 1): L699–722. 175. Koppenol, W.H., J. Moreno, W.A. Pryor, H. Ischiropoulos, and J.A. Beckman, Peroxynitrite, a cloacked oxidant formed by nitric oxide and superoxide. Chem. Res. Toxicol., 1992. 5: 834–842. 176. Quijano, C., B. Alvarez, R.M. Gatti, O. Augusto, and R. Radi, Pathways of peroxynitrite oxidation of thiol groups. Biochem. J., 1997. 322 (Pt 1): 167–173. 177. Bonini, M.G. and O. Augusto, Carbon dioxide stimulates the production of thiyl, sulfinyl, and disulfide radical anion from thiol oxidation by peroxynitrite. J. Biol. Chem., 2001. 276(13): 9749–9754. 178. Sies, H., V.S. Sharov, L.O. Klotz, and K. Briviba, Glutathione peroxidase protects against peroxynitrite-mediated oxidations. A new function for selenoproteins as peroxynitrite reductase. J. Biol. Chem., 1997. 272(44): 27812–27817. 179. Fu, Y., H. Sies, and X.G. Lei, Opposite roles of selenium-dependent glutathione peroxidase-1 in superoxide generator diquat- and peroxynitrite-induced apoptosis and signaling. J. Biol. Chem., 2001. 276(46): 43004–43009.

Peroxynitrite

135

180. Christen, S., A.A. Woodall, M.K. Shigenaga, P.T. Southwell-Keely, M.W. Duncan, and B.N. Ames, Gamma-tocopherol traps mutagenic electrophiles such as NO(X) and complements alpha-tocopherol: physiological implications. Proc. Natl. Acad. Sci. USA, 1997. 94(7): 3217–3222. 181. Hoglen, N.C., S.C. Waller, I.G. Sipes, and D.C. Liebler, Reactions of peroxynitrite with gamma-tocopherol. Chem. Res. Toxicol., 1997. 10(4): 401–407. 182. Botti, H., C. Batthyany, A. Trostchansky, R. Radi, B.A. Freeman, and H. Rubbo, Peroxynitrite-mediated alpha-tocopherol oxidation in low-density lipoprotein: a mechanistic approach. Free Radic. Biol. Med., 2004. 36(2): 152–162. 183. Botti, H., M. Trujillo, C. Batthyány, H. Rubbo, G. Ferrer-Sueta, and R. Radi, Homolytic pathways drive peroxynitrite-dependent Trolox C oxidation. Chem. Res. Toxicol., 2004. 17, 1377–1384. 184. Halliwell, B. and J.M.C. Gutteridge, Free Radicals in Biology and Medicine. 3rd ed. 1999, New York: Oxford University Press. 185. Rubbo, H., R. Radi, M. Trujillo, R. Telleri, B. Kalyanaraman, S. Barnes, M. Kirk, and B.A. Freeman, Nitric oxide regulation of superoxide and peroxynitrite-dependent lipid peroxidation. Formation of novel nitrogen-containing oxidized lipid derivatives. J. Biol. Chem., 1994. 269(42): 26066–26075. 186. Niki, E., Action of ascorbic acid as a scavenger of active and stable oxygen radicals. Am. J. Clin. Nutr., 1991. 54(6 Suppl): 1119S–1124S. 187. Squadrito, G.L., X. Jin, and W.A. Pryor, Stopped-flow kinetic study of the reaction of ascorbic acid with peroxynitrite. Arch. Biochem. Biophys., 1995. 322(1): 53–59. 188. Kirsch, M. and H. de Groot, Ascorbate is a potent antioxidant against peroxynitrite-induced oxidation reactions. Evidence that ascorbate acts by re-reducing substrate radicals produced by peroxynitrite. J. Biol. Chem., 2000. 275(22): 16702–16708. 189. Sturgeon, B.E., H.J. Sipe, Jr., D.P. Barr, J.T. Corbett, J.G. Martinez, and R.P. Mason, The fate of the oxidizing tyrosyl radical in the presence of glutathione and ascorbate. Implications for the radical sink hypothesis. J. Biol. Chem., 1998. 273(46): 30116–30121. 190. Vatassery, G.T., J.C. Lai, E.G. DeMaster, W.E. Smith, and H.T. Quach, Oxidation of vitamin E and vitamin C and inhibition of brain mitochondrial oxidative phosphorylation by peroxynitrite. J. Neurosci. Res, 2004. 75(6): 845–853. 191. Vatassery, G.T., W.E. Smith, and H.T. Quach, Alpha-tocopherol in rat brain subcellular fractions is oxidized rapidly during incubations with low concentrations of peroxynitrite. J. Nutr., 1998. 128(2): 152–157. 192. Hogg, N., J. Joseph, and B. Kalyanaraman, The oxidation of alpha-tocopherol and trolox by peroxynitrite. Arch. Biochem. Biophys., 1994. 314(1): 153–158. 193. Schopfer, F., N. Riobo, M.C. Carreras, B. Alvarez, R. Radi, A. Boveris, E. Cadenas, and J.J. Poderoso, Oxidation of ubiquinol by peroxynitrite: implications for protection of mitochondria against nitrosative damage. Biochem. J., 2000. 349(Pt 1): 35–42. 194. Souza, J.M., I. Choi, Q. Chen, M. Weisse, E. Daikhin, M. Yudkoff, M. Obin, J. Ara, J. Horwitz, and H. Ischiropoulos, Proteolytic degradation of tyrosine nitrated proteins. Arch. Biochem. Biophys., 2000. 380(2): 360–366. 195. Koeck, T., X. Fu, S.L. Hazen, J.W. Crabb, D.J. Stuehr, and K.S. Aulak, Rapid and selective oxygen regulated protein tyrosine “denitration” and nitration in mitochondria. J. Biol. Chem., 2004. 279(26): 27257–27262.

136

Nitric Oxide, Cell Signaling, and Gene Expression

196. Reusch, J.E., Diabetes, microvascular complications, and cardiovascular complications: what is it about glucose? J. Clin. Invest., 2003. 112(7): 986–988. 197. Sheetz, M.J. and G.L. King, Molecular understanding of hyperglycemia’s adverse effects for diabetic complications. JAMA, 2002. 288(20): 2579–2588. 198. Brownlee, M., Biochemistry and molecular cell biology of diabetic complications. Nature, 2001. 414(6865): 813–820. 199. Nishikawa, T., D. Edelstein, X.L. Du, S. Yamagishi, T. Matsumura, Y. Kaneda, M.A. Yorek, D. Beebe, P.J. Oates, H.P. Hammes, I. Giardino, and M. Brownlee, Normalizing mitochondrial superoxide production blocks three pathways of hyperglycaemic damage. Nature, 2000. 404(6779): 787–790. 200. Du, X.L., D. Edelstein, L. Rossetti, I.G. Fantus, H. Goldberg, F. Ziyadeh, J. Wu, and M. Brownlee, Hyperglycemia-induced mitochondrial superoxide overproduction activates the hexosamine pathway and induces plasminogen activator inhibitor-1 expression by increasing Sp1 glycosylation. Proc. Natl. Acad. Sci. USA, 2000. 97(22): 12222–12226. 201. Zou, M.H., C. Shi, and R.A. Cohen, Oxidation of the zinc-thiolate complex and uncoupling of endothelial nitric oxide synthase by peroxynitrite. J. Clin. Invest., 2002. 109(6): 817–826. 202. El-Remessy, A.B., G. Abou-Mohamed, R.W. Caldwell, and R.B. Caldwell, High glucose-induced tyrosine nitration in endothelial cells: role of eNOS uncoupling and aldose reductase activation. Invest. Ophthalmol. Vis. Sci., 2003. 44(7): 3135–3143. 203. Christ, M., J. Bauersachs, C. Liebetrau, M. Heck, A. Gunther, and M. Wehling, Glucose increases endothelial-dependent superoxide formation in coronary arteries by NAD(P)H oxidase activation: attenuation by the 3-hydroxy-3-methylglutaryl coenzyme A reductase inhibitor atorvastatin. Diabetes, 2002. 51(8): 2648–2652. 204. Desco, M.C., M. Asensi, R. Marquez, J. Martinez-Valls, M. Vento, F.V. Pallardo, J. Sastre, and J. Vina, Xanthine oxidase is involved in free radical production in type 1 diabetes: protection by allopurinol. Diabetes, 2002. 51(4): 1118–1124. 205. Du, X.L., D. Edelstein, S. Dimmeler, Q. Ju, C. Sui, and M. Brownlee, Hyperglycemia inhibits endothelial nitric oxide synthase activity by posttranslational modification at the Akt site. J. Clin. Invest., 2001. 108(9): 1341–1348. 206. Pieper, G.M. and G.J. Gross, Oxygen free radicals abolish endothelium-dependent relaxation in diabetic rat aorta. Am. J. Physiol., 1988. 255(4 Pt 2): H825– H833. 207. Hoeldtke, R.D., K.D. Bryner, D.R. McNeill, G.R. Hobbs, and C. Baylis, Peroxynitrite versus nitric oxide in early diabetes. Am. J. Hypertens., 2003. 16(9 Pt 1): 761–766. 208. Timimi, F.K., H.H. Ting, E.A. Haley, M.A. Roddy, P. Ganz, and M.A. Creager, Vitamin C improves endothelium-dependent vasodilation in patients with insulindependent diabetes mellitus. J. Am. Coll. Cardiol., 1998. 31(3): 552–557. 209. Ting, H.H., F.K. Timimi, K.S. Boles, S.J. Creager, P. Ganz, and M.A. Creager, Vitamin C improves endothelium-dependent vasodilation in patients with noninsulin-dependent diabetes mellitus. J. Clin. Invest., 1996. 97(1): 22–28. 210. Johnstone, M.T., S.J. Creager, K.M. Scales, J.A. Cusco, B.K. Lee, and M.A. Creager, Impaired endothelium-dependent vasodilation in patients with insulindependent diabetes mellitus. Circulation, 1993. 88(6): 2510–2516.

Peroxynitrite

137

211. McVeigh, G.E., G.M. Brennan, G.D. Johnston, B.J. McDermott, L.T. McGrath, W.R. Henry, J.W. Andrews, and J.R. Hayes, Impaired endothelium-dependent and independent vasodilation in patients with type 2 (non-insulin-dependent) diabetes mellitus. Diabetologia, 1992. 35(8): 771–776. 212. Title, L.M., P.M. Cummings, K. Giddens, and B.A. Nassar, Oral glucose loading acutely attenuates endothelium-dependent vasodilation in healthy adults without diabetes: an effect prevented by vitamins C and E. J. Am. Coll. Cardiol., 2000. 36(7): 2185–2191. 213. Beckman, J.A., A.B. Goldfine, M.B. Gordon, and M.A. Creager, Ascorbate restores endothelium-dependent vasodilation impaired by acute hyperglycemia in humans. Circulation, 2001. 103(12): 1618–1623. 214. Ceriello, A., F. Mercuri, L. Quagliaro, R. Assaloni, E. Motz, L. Tonutti, and C. Taboga, Detection of nitrotyrosine in the diabetic plasma: evidence of oxidative stress. Diabetologia, 2001. 44(7): 834–838. 215. Marfella, R., L. Quagliaro, F. Nappo, A. Ceriello, and D. Giugliano, Acute hyperglycemia induces an oxidative stress in healthy subjects. J. Clin. Invest., 2001. 108(4): 635–636. 216. Ceriello, A., L. Quagliaro, B. Catone, R. Pascon, M. Piazzola, B. Bais, G. Marra, L. Tonutti, C. Taboga, and E. Motz, Role of hyperglycemia in nitrotyrosine postprandial generation. Diabetes Care, 2002. 25(8): 1439–1443. 217. Lin, K.Y., A. Ito, T. Asagami, P.S. Tsao, S. Adimoolam, M. Kimoto, H. Tsuji, G.M. Reaven, and J.P. Cooke, Impaired nitric oxide synthase pathway in diabetes mellitus: role of asymmetric dimethylarginine and dimethylarginine dimethylaminohydrolase. Circulation, 2002. 106(8): 987–992. 218. Hayden, M.R. and S.C. Tyagi, Is type 2 diabetes mellitus a vascular disease (atheroscleropathy) with hyperglycemia a late manifestation? The role of NOS, NO, and redox stress. Cardiovasc. Diabetol., 2003. 2(1): 2. 219. Cosentino, F., M. Eto, P. De Paolis, B. van der Loo, M. Bachschmid, V. Ullrich, A. Kouroedov, C. Delli Gatti, H. Joch, M. Volpe, and T.F. Luscher, High glucose causes upregulation of cyclooxygenase-2 and alters prostanoid profile in human endothelial cells: role of protein kinase C and reactive oxygen species. Circulation, 2003. 107(7): 1017–1023. 220. Brodsky, S.V., S. Gao, H. Li, and M.S. Goligorsky, Hyperglycemic switch from mitochondrial nitric oxide to superoxide production in endothelial cells. Am. J. Physiol. Heart Circ. Physiol., 2002. 283(5): H2130– H2139. 221. Frustaci, A., J. Kajstura, C. Chimenti, I. Jakoniuk, A. Leri, A. Maseri, B. NadalGinard, and P. Anversa, Myocardial cell death in human diabetes. Circ. Res., 2000. 87(12): 1123–1132. 222. Kajstura, J., F. Fiordaliso, A.M. Andreoli, B. Li, S. Chimenti, M.S. Medow, F. Limana, B. Nadal-Ginard, A. Leri, and P. Anversa, IGF-1 overexpression inhibits the development of diabetic cardiomyopathy and angiotensin II-mediated oxidative stress. Diabetes, 2001. 50(6): 1414–1424. 223. Ceriello, A., L. Quagliaro, M. D’Amico, C. Di Filippo, R. Marfella, F. Nappo, L. Berrino, F. Rossi, and D. Giugliano, Acute hyperglycemia induces nitrotyrosine formation and apoptosis in perfused heart from rat. Diabetes, 2002. 51(4): 1076–1082.

138

Nitric Oxide, Cell Signaling, and Gene Expression

224. Oliveira, P.J., R. Seica, P.M. Coxito, A.P. Rolo, C.M. Palmeira, M.S. Santos, and A.J. Moreno, Enhanced permeability transition explains the reduced calcium uptake in cardiac mitochondria from streptozotocin-induced diabetic rats. FEBS Lett., 2003. 554(3): 511–514. 225. Pierce, G.N. and N.S. Dhalla, Heart mitochondrial function in chronic experimental diabetes in rats. Can. J. Cardiol., 1985. 1(1): 48–54. 226. Kucharska, J., Z. Braunova, O. Ulicna, L. Zlatos, and A. Gvozdjakova, Deficit of coenzyme Q in heart and liver mitochondria of rats with streptozotocin-induced diabetes. Physiol. Res., 2000. 49(4): 411–418. 227. Mihm, M.J. and J.A. Bauer, Peroxynitrite-induced inhibition and nitration of cardiac myofibrillar creatine kinase. Biochimie, 2002. 84(10): 1013–1019. 228. Kern, T.S. and R.L. Engerman, Pharmacological inhibition of diabetic retinopathy: aminoguanidine and aspirin. Diabetes, 2001. 50(7): 1636–1642. 229. El-Remessy, A.B., M.A. Behzadian, G. Abou-Mohamed, T. Franklin, R.W. Caldwell, and R.B. Caldwell, Experimental diabetes causes breakdown of the bloodretina barrier by a mechanism involving tyrosine nitration and increases in expression of vascular endothelial growth factor and urokinase plasminogen activator receptor. Am. J. Pathol., 2003. 162(6): 1995–2004. 230. Podesta, F., G. Romeo, W.H. Liu, S. Krajewski, J.C. Reed, C. Gerhardinger, and M. Lorenzi, Bax is increased in the retina of diabetic subjects and is associated with pericyte apoptosis in vivo and in vitro. Am. J. Pathol., 2000. 156(3): 1025–1032. 231. Kowluru, R.A. and S.N. Abbas, Diabetes-induced mitochondrial dysfunction in the retina. Invest. Ophthalmol. Vis. Sci., 2003. 44(12): 5327–5334. 232. Ruetten, H. and C. Thiemermann, Nitric oxide and septic shock, in Nitric Oxide Biology and Pathobiology, J.L. Ignarro, Ed. 2000, Academic Press: San Diego, CA. 233. Lanone, S., A. Mebazaa, C. Heymes, D. Henin, J.J. Poderoso, Y. Panis, C. Zedda, T. Billiar, D. Payen, M. Aubier, and J. Boczkowski, Muscular contractile failure in septic patients: role of the inducible nitric oxide synthase pathway. Am. J. Respir. Crit. Care Med., 2000. 162(6): 2308–2315. 234. Preiser, J.C., H. Zhang, B. Vray, A. Hrabak, and J.L. Vincent, Time course of inducible nitric oxide synthase activity following endotoxin administration in dogs. Nitric Oxide, 2001. 5(2): 208–211. 235. el-Dwairi, Q., A. Comtois, Y. Guo, and S.N. Hussain, Endotoxin-induced skeletal muscle contractile dysfunction: contribution of nitric oxide synthases. Am. J. Physiol., 1998. 274(3 Pt 1): C770– C779. 236. Escames, G., J. Leon, M. Macias, H. Khaldy, and D. Acuna-Castroviejo, Melatonin counteracts lipopolysaccharide-induced expression and activity of mitochondrial nitric oxide synthase in rats. FASEB J., 2003. 17(8): 932–934. 237. Thiemermann, C. and J. Vane, Inhibition of nitric oxide synthesis reduces the hypotension induced by bacterial lipopolysaccharides in the rat in vivo. Eur. J. Pharmacol., 1990. 182(3): 591–595. 238. Szabo, C., J.A. Mitchell, C. Thiemermann, and J.R. Vane, Nitric oxide-mediated hyporeactivity to noradrenaline precedes the induction of nitric oxide synthase in endotoxin shock. Br. J. Pharmacol., 1993. 108(3): 786–792. 239. Nethery, D., A. DiMarco, D. Stofan, and G. Supinski, Sepsis increases contractionrelated generation of reactive oxygen species in the diaphragm. J. Appl. Physiol., 1999. 87(4): 1279–1286.

Peroxynitrite

139

240. Callahan, L.A., D. Nethery, D. Stofan, A. DiMarco, and G. Supinski, Free radicalinduced contractile protein dysfunction in endotoxin-induced sepsis. Am. J. Respir. Cell Mol. Biol., 2001. 24(2): 210–217. 241. Callahan, L.A., D.A. Stofan, L.I. Szweda, D.E. Nethery, and G.S. Supinski, Free radicals alter maximal diaphragmatic mitochondrial oxygen consumption in endotoxin-induced sepsis. Free Radic. Biol. Med., 2001. 30(1): 129–138. 242. Llesuy, S., P. Evelson, B. Gonzalez-Flecha, J. Peralta, M.C. Carreras, J.J. Poderoso, and A. Boveris, Oxidative stress in muscle and liver of rats with septic syndrome. Free Radic. Biol. Med., 1994. 16(4): 445–451. 243. Javesghani, D., S.A. Magder, E. Barreiro, M.T. Quinn, and S.N. Hussain, Molecular characterization of a superoxide-generating NAD(P)H oxidase in the ventilatory muscles. Am. J. Respir. Crit. Care Med., 2002. 165(3): 412–418. 244. Nin, N., A. Cassina, J. Boggia, A. Alfonso, H. Botti, G. Peluffo, A. Trostchansky, C. Batthyány, R. Radi, R. Rubbo, and F. Hurtado, Septic diaphragmatic dysfunction is prevented by Mn(III)porphyrin therapy and iNOS inhibition. Crit. Care Med., 2004. 30: 2271–2278. 245. Ruff, R.L. and D. Secrist, Inhibitors of prostaglandin synthesis or cathepsin B prevent muscle wasting due to sepsis in the rat. J. Clin. Invest., 1984. 73(5): 1483–1486. 246. Crouser, E.D., M.W. Julian, and P.M. Dorinsky, Ileal VO(2)-O(2) alterations induced by endotoxin correlate with severity of mitochondrial injury. Am. J. Respir. Crit. Care Med., 1999. 160(4): 1347–1353. 247. Barreiro, E., A.S. Comtois, J. Gea, V.E. Laubach, and S.N. Hussain, Protein tyrosine nitration in the ventilatory muscles: role of nitric oxide synthases. Am. J. Respir. Cell Mol. Biol., 2002. 26(4): 438–446. 248. Kooy, N.W., S.J. Lewis, J.A. Royall, Y.Z. Ye, D.R. Kelly, and J.S. Beckman, Extensive tyrosine nitration in human myocardial inflammation: evidence for the presence of peroxynitrite. Crit. Care Med., 1997. 25(5): 812–819. 249. Watts, J.A., J.A. Kline, L.R. Thornton, R.M. Grattan, and S.S. Brar, Metabolic dysfunction and depletion of mitochondria in hearts of septic rats. J. Mol. Cell Cardiol., 2004. 36(1): 141–150. 250. Brealey, D., S. Karyampudi, T.S. Jacques, M. Novelli, R. Stidwill, V. Taylor, R.T. Smolenski, and M. Singer, Mitochondrial dysfunction in a long-term rodent model of sepsis and organ failure. Am. J. Physiol. Regul. Integr. Comp. Physiol., 2004. 286(3): R491– R497. 251. Kantrow, S.P., D.E. Taylor, M.S. Carraway, and C.A. Piantadosi, Oxidative metabolism in rat hepatocytes and mitochondria during sepsis. Arch. Biochem. Biophys., 1997. 345(2): 278–288. 252. Hantraye, P., E. Brouillet, R. Ferrante, S. Palfi, R. Dolan, R.T. Matthews, and M.F. Beal, Inhibition of neuronal nitric oxide synthase prevents MPTP-induced parkinsonism in baboons. Nat. Med., 1996. 2(9): 1017–1021. 253. Przedborski, S., V. Jackson-Lewis, R. Yokoyama, T. Shibata, V.L. Dawson, and T.M. Dawson, Role of neuronal nitric oxide in 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP)-induced dopaminergic neurotoxicity. Proc. Natl. Acad. Sci. USA, 1996. 93(10): 4565–4571. 254. Liberatore, G.T., V. Jackson-Lewis, S. Vukosavic, A.S. Mandir, M. Vila, W.G. McAuliffe, V.L. Dawson, T.M. Dawson, and S. Przedborski, Inducible nitric oxide synthase stimulates dopaminergic neurodegeneration in the MPTP model of Parkinson’s disease. Nat. Med., 1999. 5(12): 1403–1409.

140

Nitric Oxide, Cell Signaling, and Gene Expression

255. Dehmer, T., J. Lindenau, S. Haid, J. Dichgans, and J.B. Schulz, Deficiency of inducible nitric oxide synthase protects against MPTP toxicity in vivo. J. Neurochem., 2000. 74(5): 2213–2216. 256. Heath, P.R. and P.J. Shaw, Update on the glutamatergic neurotransmitter system and the role of excitotoxicity in amyotrophic lateral sclerosis. Muscle Nerve, 2002. 26(4): 438–458. 257. Ebadi, M. and S.K. Sharma, Peroxynitrite and mitochondrial dysfunction in the pathogenesis of Parkinson’s disease. Antioxid. Redox. Signal, 2003. 5(3): 319–335. 258. Dawson, V.L. and T.M. Dawson, Nitric oxide neurotoxicity. J. Chem. Neuroanat., 1996. 10(3-4): 179–190. 259. Haas, J., B. Storch-Hagenlocher, A. Biessmann, and B. Wildemann, Inducible nitric oxide synthase and argininosuccinate synthetase: co-induction in brain tissue of patients with Alzheimer’s dementia and following stimulation with beta-amyloid 1-42 in vitro. Neurosci Lett., 2002. 322(2): 121–125. 260. Stewart, V.C. and S.J. Heales, Nitric oxide-induced mitochondrial dysfunction: implications for neurodegeneration. Free Radic. Biol. Med., 2003. 34(3): 287–303. 261. Iadecola, C., F. Zhang, S. Xu, R. Casey, and M.E. Ross, Inducible nitric oxide synthase gene expression in brain following cerebral ischemia. J. Cereb. Blood Flow Metab., 1995. 15(3): 378–384. 262. Bagasra, O., F.H. Michaels, Y.M. Zheng, L.E. Bobroski, S.V. Spitsin, Z.F. Fu, R. Tawadros, and H. Koprowski, Activation of the inducible form of nitric oxide synthase in the brains of patients with multiple sclerosis. Proc. Natl. Acad. Sci. USA, 1995. 92(26): 12041–12045. 263. Lebovitz, R.M., H. Zhang, H. Vogel, J. Cartwright, Jr., L. Dionne, N. Lu, S. Huang, and M.M. Matzuk, Neurodegeneration, myocardial injury, and perinatal death in mitochondrial superoxide dismutase-deficient mice. Proc. Natl. Acad. Sci. USA, 1996. 93(18): 9782–9787. 264. Melov, S., J.A. Schneider, B.J. Day, D. Hinerfeld, P. Coskun, S.S. Mirra, J.D. Crapo, and D.C. Wallace, A novel neurological phenotype in mice lacking mitochondrial manganese superoxide dismutase. Nat. Genet., 1998. 18(2): 159–163. 265. Kim, G.W., T. Kondo, N. Noshita, and P.H. Chan, Manganese superoxide dismutase deficiency exacerbates cerebral infarction after focal cerebral ischemia/reperfusion in mice: implications for the production and role of superoxide radicals. Stroke, 2002. 33(3): 809–815. 266. Andreassen, O.A., R.J. Ferrante, P. Klivenyi, A.M. Klein, L.A. Shinobu, C.J. Epstein, and M.F. Beal, Partial deficiency of manganese superoxide dismutase exacerbates a transgenic mouse model of amyotrophic lateral sclerosis. Ann. Neurol., 2000. 47(4): 447–455. 267. Klivenyi, P., D. St. Clair, M. Wermer, H.C. Yen, T. Oberley, L. Yang, and M. Flint Beal, Manganese superoxide dismutase overexpression attenuates MPTP toxicity. Neurobiol. Dis., 1998. 5(4): 253–258. 268. Patel, M., B.J. Day, J.D. Crapo, I. Fridovich, and J.O. McNamara, Requirement for superoxide in excitotoxic cell death. Neuron, 1996. 16(2): 345–355. 269. Abe, K., L.H. Pan, M. Watanabe, T. Kato, and Y. Itoyama, Induction of nitrotyrosine-like immunoreactivity in the lower motor neuron of amyotrophic lateral sclerosis. Neurosci. Lett., 1995. 199(2): 152–154. 270. Abe, K., L.H. Pan, M. Watanabe, H. Konno, T. Kato, and Y. Itoyama, Upregulation of protein-tyrosine nitration in the anterior horn cells of amyotrophic lateral sclerosis. Neurol Res, 1997. 19(2): 124–128.

Peroxynitrite

141

271. Smith, M.A., P.L. Richey-Harris, L.M. Sayre, J.S. Beckman, and G. Perry, Widespread peroxynitrite-mediated damage in Alzheimer’s disease. J. Neurosci., 1997. 17(8): 2653–2657. 272. Good, P.F., A. Hsu, P. Werner, D.P. Perl, and C.W. Olanow, Protein nitration in Parkinson’s disease. J. Neuropathol. Exp. Neurol., 1998. 57(4): 338–342. 273. Hattori, F., N. Murayama, T. Noshita, and S. Oikawa, Mitochondrial peroxiredoxin-3 protects hippocampal neurons from excitotoxic injury in vivo. J. Neurochem., 2003. 86(4): 860–868. 274. Sasaki, S., H. Warita, K. Abe, and M. Iwata, Inducible nitric oxide synthase (iNOS) and nitrotyrosine immunoreactivity in the spinal cords of transgenic mice with a G93A mutant SOD1 gene. J. Neuropathol. Exp. Neurol., 2001. 60(9): 839–846. 275. Gonzalez-Zulueta, M., L.M. Ensz, G. Mukhina, R.M. Lebovitz, R.M. Zwacka, J.F. Engelhardt, L.W. Oberley, V.L. Dawson, and T.M. Dawson, Manganese superoxide dismutase protects nNOS neurons from NMDA and nitric oxide-mediated neurotoxicity. J. Neurosci., 1998. 18(6): 2040–2055. 276. Kim, G.W. and P.H. Chan, Involvement of superoxide in excitotoxicity and DNA fragmentation in striatal vulnerability in mice after treatment with the mitochondrial toxin, 3-nitropropionic acid. J. Cereb. Blood Flow Metab., 2002. 22(7): 798–809. 277. Lipton, P., Ischemic cell death in brain neurons. Physiol. Rev., 1999. 79(4): 1431–1568. 278. Forman, L.J., P. Liu, R.G. Nagele, K. Yin, and P.Y. Wong, Augmentation of nitric oxide, superoxide, and peroxynitrite production during cerebral ischemia and reperfusion in the rat. Neurochem. Res., 1998. 23(2): 141–148. 279. Yu, Z.F., A.J. Bruce-Keller, Y. Goodman, and M.P. Mattson, Uric acid protects neurons against excitotoxic and metabolic insults in cell culture, and against focal ischemic brain injury in vivo. J. Neurosci. Res., 1998. 53(5): 613–625. 280. Piantadosi, C.A. and J. Zhang, Mitochondrial generation of reactive oxygen species after brain ischemia in the rat. Stroke, 1996. 27(2): 327–332. 281. Fujimura, M., Y. Morita-Fujimura, M. Kawase, J.C. Copin, B. Calagui, C.J. Epstein, and P.H. Chan, Manganese superoxide dismutase mediates the early release of mitochondrial cytochrome c and subsequent DNA fragmentation after permanent focal cerebral ischemia in mice. J. Neurosci., 1999. 19(9): 3414–3422. 282. Orth, M. and A.H. Schapira, Mitochondrial involvement in Parkinson’s disease. Neurochem. Int., 2002. 40(6): 533–541. 283. Andreassen, O.A., R.J. Ferrante, A. Dedeoglu, D.W. Albers, P. Klivenyi, E.J. Carlson, C.J. Epstein, and M.F. Beal, Mice with a partial deficiency of manganese superoxide dismutase show increased vulnerability to the mitochondrial toxins malonate, 3-nitropropionic acid, and MPTP. Exp. Neurol., 2001. 167(1): 189–195. 284. Beal, M.F., Mitochondria, oxidative damage, and inflammation in Parkinson’s disease. Ann. NY Acad. Sci., 2003. 991: 120–131. 285. Giasson, B.I., J.E. Duda, I.V. Murray, Q. Chen, J.M. Souza, H.I. Hurtig, H. Ischiropoulos, J.Q. Trojanowski, and V.M. Lee, Oxidative damage linked to neurodegeneration by selective alpha-synuclein nitration in synucleinopathy lesions. Science, 2000. 290(5493): 985–989. 286. Ara, J., S. Przedborski, A.B. Naini, V. Jackson-Lewis, R.R. Trifiletti, J. Horwitz, and H. Ischiropoulos, Inactivation of tyrosine hydroxylase by nitration following exposure to peroxynitrite and 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP). Proc. Natl. Acad. Sci. USA, 1998. 95(13): 7659–7663.

142

Nitric Oxide, Cell Signaling, and Gene Expression

287. Ischiropoulos, H. and J.S. Beckman, Oxidative stress and nitration in neurodegeneration: cause, effect, or association? J. Clin. Invest., 2003. 111(2): 163–169. 288. Cassarino, D.S., C.P. Fall, T.S. Smith, and J.P. Bennett, Jr., Pramipexole reduces reactive oxygen species production in vivo and in vitro and inhibits the mitochondrial permeability transition produced by the Parkinsonian neurotoxin methylpyridinium ion. J. Neurochem., 1998. 71(1): 295–301. 289. Mochizuki, H., K. Goto, H. Mori, and Y. Mizuno, Histochemical detection of apoptosis in Parkinson’s disease. J. Neurol. Sci., 1996. 137(2): 120–123. 290. Tatton, N.A., A. Maclean-Fraser, W.G. Tatton, D.P. Perl, and C.W. Olanow, A fluorescent double-labeling method to detect and confirm apoptotic nuclei in Parkinson’s disease. Ann. Neurol., 1998. 44(3 Suppl 1): S142– S148. 291. Tatton, N.A. and S.J. Kish, In situ detection of apoptotic nuclei in the substantia nigra compacta of 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine-treated mice using terminal deoxynucleotidyl transferase labelling and acridine orange staining. Neuroscience, 1997. 77(4): 1037–1048. 292. Cassarino, D.S., J.K. Parks, W.D. Parker, Jr., and J.P. Bennett, Jr., The Parkinsonian neurotoxin MPP+ opens the mitochondrial permeability transition pore and releases cytochrome c in isolated mitochondria via an oxidative mechanism. Biochim. Biophys. Acta, 1999. 1453(1): 49–62. 293. Vila, M., V. Jackson-Lewis, S. Vukosavic, R. Djaldetti, G. Liberatore, D. Offen, S.J. Korsmeyer, and S. Przedborski, Bax ablation prevents dopaminergic neurodegeneration in the 1-methyl- 4-phenyl-1,2,3,6-tetrahydropyridine mouse model of Parkinson’s disease. Proc. Natl. Acad. Sci. USA, 2001. 98(5): 2837–2842. 294. Wispe, J.R., B.B. Warner, J.C. Clark, C.R. Dey, J. Neuman, S.W. Glasser, J.D. Crapo, L.Y. Chang, and J.A. Whitsett, Human Mn-superoxide dismutase in pulmonary epithelial cells of transgenic mice confers protection from oxygen injury. J. Biol. Chem., 1992. 267(33): 23937–23941. 295. Zwacka, R.M., W. Zhou, Y. Zhang, C.J. Darby, L. Dudus, J. Halldorson, L. Oberley, and J.F. Engelhardt, Redox gene therapy for ischemia/reperfusion injury of the liver reduces AP1 and NF-kappaB activation. Nat. Med., 1998. 4(6): 698–704. 296. Bertera, S., M.L. Crawford, A.M. Alexander, G.D. Papworth, S.C. Watkins, P.D. Robbins, and M. Trucco, Gene transfer of manganese superoxide dismutase extends islet graft function in a mouse model of autoimmune diabetes. Diabetes, 2003. 52(2): 387–393. 297. Riley, D., Functional mimics of superoxide dismutase enzymes as therapeutic agents. Chem. Rev., 1999. 99: 2573–2587. 298. Kelso, G.F., C.M. Porteous, C.V. Coulter, G. Hughes, W.K. Porteous, E.C. Ledgerwood, R.A. Smith, and M.P. Murphy, Selective targeting of a redox-active ubiquinone to mitochondria within cells: antioxidant and antiapoptotic properties. J. Biol. Chem., 2001. 276(7): 4588–4596. 299. Smith, R.A., C.M. Porteous, C.V. Coulter, and M.P. Murphy, Selective targeting of an antioxidant to mitochondria. Eur. J. Biochem., 1999. 263(3): 709–716. 300. Dessolin, J., M. Schuler, A. Quinart, F. De Giorgi, L. Ghosez, and F. Ichas, Selective targeting of synthetic antioxidants to mitochondria: towards a mitochondrial medicine for neurodegenerative diseases? Eur. J. Pharmacol., 2002. 447(2-3): 155–161. 301. Coulter, C.V., G.F. Kelso, T.K. Lin, R.A. Smith, and M.P. Murphy, Mitochondrially targeted antioxidants and thiol reagents. Free Radic. Biol. Med., 2000. 28(10): 1547–1554.

Peroxynitrite

143

302. Batinic-Haberle, I., Manganese porphyrins and related compounds as mimics of superoxide dismutase. Methods Enzymol., 2002. 349: 223–233. 303. Ferrer-Sueta, G., C. Quijano, B. Alvarez, and R. Radi, Reactions of manganese porphyrins and manganese-superoxide dismutase with peroxynitrite. Methods Enzymol., 2002. 349: 23–37. 304. Ferrer-Sueta, G., D. Vitturi, I. Batinic-Haberle, I. Fridovich, S. Goldstein, G. Czapski, and R. Radi, Reactions of manganese porphyrins with peroxynitrite and carbonate radical anion. J. Biol. Chem., 2003. 278(30): 27432–27438. 305. Mackensen, G.B., M. Patel, H. Sheng, C.L. Calvi, I. Batinic-Haberle, B.J. Day, L.P. Liang, I. Fridovich, J.D. Crapo, R.D. Pearlstein, and D.S. Warner, Neuroprotection from delayed postischemic administration of a metalloporphyrin catalytic antioxidant. J. Neurosci., 2001. 21(13): 4582–4592. 306. Zingarelli, B., B.J. Day, J.D. Crapo, A.L. Salzman, and C. Szabo, The potential role of peroxynitrite in the vascular contractile and cellular energetic failure in endotoxic shock. Br. J. Pharmacol., 1997. 120(2): 259–267. 307. Piganelli, J.D., S.C. Flores, C. Cruz, J. Koepp, I. Batinic-Haberle, J. Crapo, B. Day, R. Kachadourian, R. Young, B. Bradley, and K. Haskins, A metalloporphyrinbased superoxide dismutase mimic inhibits adoptive transfer of autoimmune diabetes by a diabetogenic T-cell clone. Diabetes, 2002. 51(2): 347–355. 308. Chaloin, L., P. Bigey, C. Loup, M. Marin, N. Galeotti, M. Piechaczyk, F. Heitz, and B. Meunier, Improvement of porphyrin cellular delivery and activity by conjugation to a carrier peptide. Bioconjug. Chem., 2001. 12(5): 691–700. 309. Trujillo, M. and R. Radi, Peroxynitrite reaction with the reduced and the oxidized forms of lipoic acid: new insights into the reaction of peroxynitrite with thiols. Arch. Biochem. Biophys., 2002. 397(1): 91–98. 310. Bryk, R., P. Griffin, and C. Nathan, Peroxynitrite reductase activity of bacterial peroxiredoxins. Nature, 2000. 407(6801): 211–215. 311. Jaeger, T., H. Budde, L. Flohe, U. Menge, M. Singh, M. Trujillo, and R. Radi, Multiple thioredoxin-mediated routes to detoxify hydroperoxides in Mycobacterium tuberculosis. Arch. Biochem. Biophys., 2004. 423(1): 182–191.

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

145

146

Nitric Oxide, Cell Signaling, and Gene Expression

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].

148

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 149

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

150

Nitric Oxide, Cell Signaling, and Gene Expression

(a) Astrocytes

(b) Skeletal muscle

Inflammation, ischemia, sepsis

Muscle contraction AMP:ATP

.NO

PO43−

AMPK

AMPK

Cytochrome c oxidase

AMP:ATP

PO43−

eNOS

eNOS

PO43−

AMPK

AMPK

GLUT4-containing vesicles

.NO

? Guanylyl cGMP cyclase

GTP

3

UT GL

Glucose

GLUT4 Glucose

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

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 151

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.

152

Nitric Oxide, Cell Signaling, and Gene Expression

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

Inflammation, ischemia, sepsis

iNOS

Glutamate

iNOS

nNOS

G6PD . NO Cytoprotection

−

O2• GSSG

GSH

ONOO− NADP+

NADPH

G6PD

+

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

154

Nitric Oxide, Cell Signaling, and Gene Expression

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:

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 155

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

156

Nitric Oxide, Cell Signaling, and Gene Expression

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

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 157

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

158

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

Astrocytes

iNOS arginine

O2 Cytochrome c oxidase

.NO

ATP

ADP

AMP

PO43−

AMPK

PO43− 3−

PO4

AMPK

F2,6P2 PO 3− 4

PFK2 + PFK2

PFK1

PO43−

F6P PO43−

PO43−

F1,6P2 NAD+ Survival

ATP

pyruvate NADH(H+) + lactate NAD

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).

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 159

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.

REFERENCES 1. Clarke, D.D. and Sokoloff, L. (1994) in Basic Neurochemistry (Siegel, G.J., Agranoff, B.W., Albers, R.W., and Molinoff, P.B., Eds.), pp. 645–680, Raven Press, New York.

160

Nitric Oxide, Cell Signaling, and Gene Expression

2. Hawkins, R. (1985) in Cerebral Energy Metabolism and Metabolic Encephalopathy (McCandless, D.W., Ed.), pp. 3–17, Plenum Press, New York. 3. Golgi, C. (1886) Sulla fine anatomia degli organi centrali del sisterna nervoso, Milan, Italy. 4. Morgello, S., Uson, R.R., Schwartz, E.J., and Haber, R.S. (1995) Glia 14, 43–54. 5. Kacem, K., Lacombe, P., Seylaz, J., and Bonvento, G. (1998) Glia 23, 1–10. 6. Blumcke, I., Eggli, P., and Celio, M.R. (1995) Glia 15, 131–140. 7. Grosche, J., Matyash, V., Moller, T., Verkhratsky, A., Reichenbach, A., and Kettenmann, H. (1999) Nat. Neurosci. 2, 139–143. 8. Mennerick, S. and Zorumski, C.F. (1994) Nature 368, 59–62. 9. Magistretti, P.J., Hof, P.R., and Martin, J.L. (1986) J. Neuroscience 6, 2558–2562. 10. Tsacopoulos, M. and Magistretti, P.J. (1996) J. Neurosci. 16, 877–885. 11. Pardridge, W.M., Boado, R.J., and Farrell, C.R. (1990) J. Biol. Chem. 265, 18035–18040. 12. Maher, F., Vannucci, S.J., and Simpson, I.A. (1993) J. Cereb. Blood Flow Metab. 13, 342–345. 13. Garthwaite, J., Charles, S.L., and Chess-Williams, R. (1988) Nature 336, 385–387. 14. Palmer, R.M., Ashton, D.S., and Moncada, S. (1988) Nature 333, 664–666. 15. Bredt, D.S. and Snyder, S.H. (1990) Proc. Natl. Acad. Sci. USA 87, 682–685. 16. Knowles, R.G. and Moncada, S. (1994) Biochem. J. 298, 249–258. 17. Knowles, R.G., Palacios, M., Palmer, R.M.J., and Moncada, S. (1989) Proc. Natl. Acad. Sci. USA 86, 5159–5162. 18. Bredt, D.S. and Snyder, S.H. (1989) Proc. Natl. Acad. Sci. USA 86, 9030–9033. 19. Stuehr, D.J. and Nathan, C.F. (1989) J. Exp. Med. 169, 1543–1555. 20. Cleeter, M.W.J., Cooper, J.M., Darley-Usmar, V.M., Moncada, S., and Schapira, A.H. (1994) FEBS Lett. 345, 50–54. 21. Brown, G.C. and Cooper, C.E. (1994) FEBS Lett. 356, 295–298. 22. Schweizer, M. and Richter, C. (1994) Biochem. Biophys. Res. Commun. 204, 169–175. 23. Bolaños, J.P., Peuchen, S., Heales, S.J.R., Land, J.M., and Clark, J.B. (1994) J. Neurochem. 63, 910–916. 24. Cooper, C.E. (2002) Trends Biochem. Sci. 27, 489–492. 25. Moncada, S. and Erusalimsky, J.D. (2002) Nature Rev. 3, 214–220. 26. Nomura, Y. and Kitamura, Y. (1993) Neurosci. Res. 18, 103–107. 27. Murphy, S., Simmons, M.L., Agulló, L., García, A., Feinstein, D.L., Galea, E., Reis, D.J., Minc-Golomb, D., and Schwartz, J.P. (1993) Trends Neurosci. 16, 323–328. 28. Murphy, S. and Grzybicki, D. (1996) Neuroscientist 2, 90–99. 29. Merrill, J.E., Murphy, S.P., Mitrovic, B., Mackenzie-Graham, A., Dopp, J.C., Ding, M., Griscavage, J., Ignarro, L.J., and Lowenstein, C.J. (1997) J. Neurosci. Res. 48, 372–384. 30. Agulló, L. and García, A. (1992) Biochem. Biophys. Res. Commun. 182, 1362–1368. 31. Agulló, L. and García, A. (1992) Biochem. J. 288, 619–624. 32. Simmons, M.L. and Murphy, S. (1992) J. Neurochem. 59, 897–905. 33. Galea, E., Feinstein, D.L., and Reis, D.J. (1992) Proc. Natl. Acad. Sci. USA 89, 10945–10949. 34. Kilbourn, R.G. and Belloni, P. (1990) J. Natl. Cancer Inst. 82, 772–776.

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 161 35. Minc-Golomb, D., Yadid, G., Tsarfaty, I., Resau, J.H., and Schwartz, J.P. (1996) J. Neurochem. 66, 1504–1509. 36. Blough, N.V. and Zafiriou, O.C. (1985) Inorg. Chem. 24, 3502–3504. 37. Beckman, J.S., Beckman, T.W., Chen, J., Marshall, P.A., and Freeman, B.A. (1990) Proc. Natl. Acad. Sci. USA 87, 1620–1624. 38. Lipton, S.A., Choi, Y.B., Pan, Z.H., Lei, S.Z., Chen, H.S.V., Sucher, N.J., Loscalzo, J., Singel, D.J., and Stamler, J.S. (1993) Nature 364, 626–632. 39. Bonfoco, E., Krainc, C., Ankarcrona, M., Nicotera, P., and Lipton, S.A. (1995) Proc. Natl. Acad. Sci. USA 92, 7162–7166. 40. Bolaños, J.P., Almeida, A., Stewart, V., Peuchen, S., Land, J.M., Clark, J.B., and Heales, S.J.R. (1997) J. Neurochem. 68, 2227–2240. 41. Bolaños, J.P., García-Nogales, P., and Almeida, A. (2004) Curr. Pharm. Design 10, 867–877. 42. Dawson, V.L., Dawson, T.M., London, E.D., Bredt, D.S., and Snyder, S.H. (1991) Proc. Natl. Acad. Sci. USA 88, 6368–6371. 43. Dawson, V.L., Dawson, T.M., Bartley, D.A., Uhl, G.R., and Snyder, S.H. (1993) J. Neurosci. 13, 2651–2661. 44. Dawson, V.L. and Dawson, T.M. (1996) J. Chem. Neuroanatom. 10, 179–190. 45. Turski, L. and Turski, W.A. (1993) Experientia 49, 1064–1072. 46. Ikonomidou, C. and Turski, L. (1995) Curr. Opin. Neurol. 8, 487–497. 47. Schulz, J.B., Matthews, R.T., and Beal, M.F. (1995) Curr. Opin. Neurol. 8, 480–486. 48. Bolaños, J.P. and Almeida, A. (1999) Biochim. Biophys. Acta 1411, 415–436. 49. Bolaños, J.P., Heales, S.J.R., Peuchen, S., Barker, J.E., Land, J.M., and Clark, J.B. (1996) Free Rad. Biol. Med. 21, 995–1001. 50. Zhang, J., Dawson, V.L., Dawson, T.M., and Snyder, S.H. (1994) Science 263, 687–689. 51. Radons, J., Heller, B., Bürkle, A., Hartmann, B., Rodriguez, M.-L., Kröncke, K.D., Burkart, V., and Kolb, H. (1994) Biochem. Biophys. Res. Commun. 199, 1270–1277. 52. Zingarelli, B., O’Connor, M., Wong, H., Salzman, A.L., and Szabó, C. (1996) J. Immunol. 156, 350–358. 53. Szabó, C., Zingarelli, B., O’Connor, M., and Salzman, A.L. (1996) Proc. Natl. Acad. Sci. USA 93, 1753–1758. 54. Szabó, C., Zingarelli, B., and Salzman, A.L. (1996) Neuroscience 78, 1051–1063. 55. Hausladen, A. and Fridovich, I. (1994) J. Biol. Chem. 269, 29405–29408. 56. Castro, L., Rodríguez, M., and Radi, R. (1994) J. Biol. Chem. 269, 29409–29415. 57. Radi, R., Rodríguez, M., Castro, L., and Telleri, R. (1994) Arch. Biochem. Biophys. 308, 89–95. 58. Bolaños, J.P., Heales, S.J.R., Land, J.M., and Clark, J.B. (1995) J. Neurochem. 64, 1965–1972. 59. Lizasoain, I., Moro, M.A., Knowles, R.G., Darley-Usmar, V., and Moncada, S. (1996) Biochem. J. 314, 877–880. 60. Sakurai-Yamashita, Y., Yamashita, K., Kataoka, Y., Himeno, A., Niwa, M., and Taniyama, K. (1997) Brain Res. 756, 61–67. 61. Mollace, V., Muscoli, C., Rotiroti, D., and Nistico, G. (1997) Biochem. Biophys. Res. Commun. 238, 916–919. 62. Rothwell, K.J. and Relton, K.J. (1993) Cerebrovasc. Brain Met. 5, 178–198.

162

Nitric Oxide, Cell Signaling, and Gene Expression

63. Reis, D.J., Feinstein, D., Galea, E., and Golanov, E.V. (1997) Fundam. Clin. Pharm. 11, S39–S43. 64. Maeda, Y., Matsumoto, M., Hori, O., Kuwabara, K., Ogawa, S., Yan, S.D., Ohtsuki, T., Kamada, T., and Stern, S.D. (1994) J. Exp. Med. 180, 2297–2308. 65. Maiese, K., Boniece, I., Demeo, D., and Wagner, J.A. (1993) J. Neurosci. 13, 3034–3040. 66. Tagami, M., Ikeda, K., Nara, Y., Fujino, H., Kubota, A., Numano, F., and Yamori, Y. (1997) Lab. Invest. 76, 613–617. 67. Wang, Y., Lin, S.Z., Chiou, A.L., Williams, L.R., and Hoffer, B.J. (1997) J. Neurosci. 17, 4341–4348. 68. Henrichnoack, P., Prehn, J.H.M., and Krieglstein, J. (1996) Stroke 27, 1609–1614. 69. Faraci, F.M. and Brian, J.E. (1994) Stroke 25, 692–703. 70. Wood, P.L. (1995) Neurol. Res. 17, 242–248. 71. Gehrmann, J., Banati, R.B., Wiessnert, C., Hossmann, K.A., and Kreutzberg, G.W. (1995) Neuropathol. Appl. Neurobiol. 21, 277–289. 72. Bidmon, H.J., Wu, J., I, B., Mayer, B., Witte, O.W., and Zilles, K. (1998) Neuroscience 82, 377–395. 73. Iadecola, C. (1997) Trends Neurosci. 20, 132–139. 74. Cataldo, A.M. and Broadwell, R.D. (1986) J. Electron Microsc. Tech. 3, 413–437. 75. Medina, J.M., Cuezva, J.M., and Mayor, F. (1980) FEBS Lett. 114, 132–134. 76. Medina, J.M., Fernández, E., Bolaños, J.P., Vicario, C., and Arizmendi, C. (1990) in Endocrine and Biochemical Development of the Fetus and Neonate (Cuezva, J.M., Pascual-Leone, A.M. and Patel, M.S., Eds.), pp. 175–194, Plenum Press, New York. 77. Vicario, C., Arizmendi, C., Malloch, G., Clark, J.B., and Medina, J.M. (1991) J. Neurochem. 57, 1700–1707. 78. Almeida, A., Bolaños, J.P., and Medina, J.M. (1992) Pediatr. Res. 31, 415–418. 79. Tabernero, A., Bolaños, J.P., and Medina, J.M. (1993) Biochem. J. 294, 635–638. 80. Pellerin, L. (2003) Neurochem. Int. 43, 331–338. 81. Andriezen, W.L. (1893) Int. Monatsschr. Anat. Physiol. 10, 532–540. 82. Tabernero, A., Giaume, C., and Medina, J.M. (1996) Glia 16, 187–195. 83. Giaume, C., Tabernero, A., and Medina, J.M. (1997) Glia 21, 114–123. 84. Wiesinger, H., Hamprecht, B., and Dringen, R. (1997) Glia 21, 22–34. 85. Dringen, R. and Hamprecht, B. (1992) J. Neurochem. 58, 511–517. 86. Dringen, R., Gebhardt, R., and Hamprecht, B. (1993) Brain Res. 623, 208–214. 87. Pellerin, L. and Magistretti, P.J. (1994) Proc. Natl. Acad. Sci. USA 91, 10625–10629. 88. Schurr, A., Payne, R.S., Miller, J.J., and Rigor, B.M. (1997) J. Neurochem. 69, 423–426. 89. Schurr, A., West, C.A., and Rigor, B.M. (1988) Science 240, 1326–1328. 90. Wood, I.S. and Trayhurn, P. (2003) Br J. Nutr. 89, 3–9. 91. Gorovits, N. and Charron, M.J. (2003) Biochem. Mol. Biol. Education 13, 163–172. 92. Mueckler, M., Caruso, C., Baldwin, S.A., Panico, M., Blench, I., Morris, H.R., Allard, W.J., Lienhard, G.E., and Lodish, H.F. (1985) Science 229, 941–945. 93. Birnbaum, M.J., Haspel, H.C., and Rosen, O.M. (1986) Proc. Natl. Acad. Sci. USA 83, 5784–5788. 94. Joost, H.G. and Thorens, B. (2001) Mol. Membr. Biol. 18, 247–256. 95. Wu, X. and Freeze, H.H. (2002) Genomics 80, 553–557.

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 163 96. Maher, F. (1995) J. Neurosci. Res. 42, 459–469. 97. Maher, F., Vannucci, S.J., and Simpson, I.A. (1994) FASEB J. 8, 1003–1011. 98. Nagamatsu, S., Sawa, H., Kamada, K., Nakamichi, Y., Yoshimoto, K., and Hoshino, T. (1993) FEBS Lett. 334, 289–295. 99. Choeiri, C., Staines, W., and Messier, C. (2002) Neuroscience 111, 19–34. 100. Gould, G.W., Thomas, H.M., Jess, T.J., and Bell, G.I. (1991) Biochemistry 30, 5139–5145. 101. Pessin, J.E. and Bell, G.I. (1992) Annu. Rev. Physiol. 54, 911–930. 102. Apelt, J., Mehlhorn, G., and Schliebs, R. (1999) J. Neurosci. Res. 57, 693–705. 103. Dijkmans, R. and Billiau, A. (1991) Eur. J. Biochem. 202, 151–159. 104. Welsh, N. and Sandler, S. (1992) Biochem. Biophys. Res. Commun. 182, 333–340. 105. Geng, Y.-J., Hansson, G.K., and Holme, E. (1992) Circ. Res. 71, 1268–1276. 106. Albina, J.E. and Mastrofrancesco, B. (1993) Am. J. Physiol. 264, C1594–C1599. 107. Spolarics, Z. and Spitzer, J.J. (1993) Hepatology 17, 615–620. 108. Le Goffe, C., Vallette, G., Charrier, L., Candelon, T., Bou-Hanna, C., Bouhours, J.F., and Laboisse, C.L. (2002) Biochem. J. 364, 349–359. 109. García-Nogales, P., Almeida, A., Fernández, E., Medina, J.M., and Bolaños, J.P. (1999) J. Neurochem. 72, 1750–1758. 110. Cidad, P., García-Nogales, P., Almeida, A., and Bolaños, J.P. (2001) J. Neurochem. 79, 17–24. 111. Bell, G.I., Burant, C.F., Takeda, J., and Gould, G.W. (1993) J. Biol. Chem. 268, 19161–19164. 112. Xie, Q.-W., Kashiwabara, Y., and Nathan, C. (1994) J. Biol. Chem. 269, 4705–4708. 113. Etgen, G.J., Jr., Fryburg, D.A., and Gibbs, E.M. (1997) Diabetes 46, 1915–1919. 114. Roberts, C.K., Barnard, R.J., Scheck, S.H., and Balon, T.W. (1997) Am. J. Physiol. 273, E220-E225. 115. Fryer, L.G., Hajduch, E., Rencurel, F., Salt, I.P., Hundal, H.S., Hardie, D.G., and Carling, D. (2000) Diabetes 49, 1978–1985. 116. Hardie, D.G., Carling, D., and Sim, A.T.R. (1989) Trends Biochem. Sci. 14, 20–23. 117. Carling, D. (2004) Trends Biochem. Sci. 29, 18–24. 118. Chen, Z.P., Mitchelhill, K.I., Michell, B.J., Stapleton, D., Rodriguez-Crespo, I., Witters, L.A., Power, D.A., Ortiz de Montellano, P.R., and Kemp, B.E. (1999) FEBS Lett. 443, 285–289. 119. Shi, Y., Liu, H., Vanderburg, G., Samuel, S.J., Ismail-Beigi, F., and Jung, C.Y. (1995) J. Biol. Chem. 270, 21772–21778. 120. Abbud, W., Habinowski, S., Zhang, J.Z., Kendrew, J., Elkairi, F.S., Kemp, B.E., Witters, L.A., and Ismail-Beigi, F. (2000) Arch. Biochem. Biophy. 380, 347–352. 121. Barnes, K., Ingram, J.C., Porras, O.H., Barros, L.F., Hudson, E.R., Fryer, L.G., Foufelle, F., Carling, D., Hardie, D.G., and Baldwin, S.A. (2002) J. Cell Sci. 115, 2433–2442. 122. Radi, R., Beckman, J.S., Bush, K.M., and Freeman, B.A. (1991) J. Biol. Chem. 266, 4244–4250. 123. Clancy, R.M., Levartovsky, D., Leszczynska-Piziak, J., Yegudin, J., and Abramson, S. (1994) Proc. Natl. Acad. Sci. USA 91, 3680–3684. 124. Clementi, E., Brown, G.C., Feelisch, M., and Moncada, S. (1998) Proc. Natl. Acad. Sci. USA 95, 7631–7636. 125. Bal-Price, A. and Brown, G.C. (2000) J. Neurochem. 75, 1455–1464. 126. Spolarics, Z., Stein, D.S., and García, Z.C. (1996) Hepatology 24, 691–696.

164

Nitric Oxide, Cell Signaling, and Gene Expression

127. Ursini, M.V., Parrella, A., Rosa, G., Salzano, S., and Martini, G. (1997) Biochem. J. 323, 801–806. 128. Ben-Yoseph, O., Boxer, P.A., and Ross, B.D. (1996) J. Neurochem. 66, 2329–2337. 129. Kussmaul, L., Hamprecht, B., and Dringen, R. (1999) J. Neurochem. 73, 1246–1253. 130. Kletzien, R.F., Harris, P.K.W., and Foellmi, L.A. (1994) FASEB J. 8, 174–181. 131. Salvemini, F., Franzé, A., Iervolino, A., Filosa, S., Salzano, S., and Ursini, M.V. (1999) J. Biol. Chem. 274, 2750–2757. 132. Xia, Y., Roman, I.J., Masters, B.S., and Zweier, J.L. (1998) J. Biol. Chem. 273, 22635–22639. 133. García-Nogales, P., Almeida, A., and Bolaños, J.P. (2003) J. Biol. Chem. 278, 864–874. 134. Almeida, A. and Bolaños, J.P. (2001) J. Neurochem. 77, 676–690. 135. Almeida, A., Almeida, J., Bolaños, J.P., and Moncada, S. (2001) Proc. Natl. Acad. Sci. USA 98, 15294–15299. 136. Eu, J.P., Liu, L., Zeng, M., and Stamler, J.S. (2000) Biochemistry 39, 1040–1047. 137. Sastry, P.S. and Rao, K.S. (2000) J. Neurochem. 74, 1–20. 138. Stanton, R.C., Seifter, J.L., Boxer, D.C.E., and Cantley, L.C. (1991) J. Biol. Chem. 266, 12442–12448. 139. Oury, T.D., Ho, Y.S., Piantadosi, C.A., and Crapo, J.D. (1992) Proc. Natl. Acad. Sci. USA 89, 9715–9719. 140. Lafon-Cazal, M., Pietri, S., Culcasi, M., and Bockaert, J. (1993) Nature 364, 535–537. 141. Rauhala, P., Lin, A.M., and Chiueh, C.C. (1998) FASEB J. 12, 165–173. 142. Mohanakumar, K.P., Thomas, B., Sharma, S.M., Muralikrishnan, D., Chowdhury, R., and Chiueh, C.C. (2002) Ann. NY. Acad. Sci. 962, 389–401. 143. Chang, J., Rao, N.V., Markewitz, B.A., Hoidal, J.R., and Michael, J.R. (1996) Am. J. Physiol. 270, L931–L949. 144. Wink, D.A., Cook, J.A., Pacelli, R., DeGraff, W., Gamson, J., Liebmann, J., Krishna, M.C., and Mitchell, J.B. (1996) Arch. Biochem. Biophys. 331, 241–248. 145. Delgado-Esteban, M., Almeida, A., and Bolaños, J.P. (2000) J. Neurochem. 75, 1618–1624. 146. Molina y Vedia, L., McDonald, B., Reep, B., Brüne, B., Di Silvio, M., Billiar, T.R., and Lapentina, E.G. (1992) J. Biol. Chem. 267, 24929–24932. 147. Zhang, J. and Snyder, S.H. (1992) Proc. Natl. Acad. Sci. USA 89, 9382–9385. 148. Lehninger, A.L., Nelson, D.L., and Cox, M.M. (1995) Principles of Biochemistry, Worth Publishers, New York. 149. Mateo, R.B., Reichner, J.S., Mastrofrancesco, B., Kraft-Stolar, D., and Albina, J.E. (1995) Am. J. Physiol. 268, C669–C675. 150. Messmer, U.K. and Brune, B. (1996) Eur. J. Pharmacol. 302, 171–182. 151. Erecinska, M., Nelson, D., and Vanderkooi, J.M. (1995) J. Neurochem. 65, 2699–2705. 152. Galli, F., Rossi, R., Di Simplicio, P., Floridi, A., and Canestrari, F. (2002) Nitric Oxide 6, 186–199. 153. Mohr, S., Stamler, J.S., and Brune, B. (1996) J. Biol. Chem. 271, 4209–4214. 154. Wu, K., Aoki, C., Elste, A., Rogalski-Wilk, A.A., and Siekevitz, P. (1997) Proc. Natl. Acad. Sci. 94, 13273–13278. 155. Mallozzi, C., Di Stasi, A.M.M., and Minetti, M. (1997) FASEB J. 11, 1281–1290. 156. Pauwels, P.J., Opperdoes, F.R., and Trouet, A. (1985) J. Neurochem. 44, 143–148.

Modulation of Glucose Metabolism by Nitric Oxide in Astrocytes and Neurons 165 157. 158. 159. 160.

Walz, W. and Mukerji, S. (1988) Glia 1, 366–370. Almeida, A., Moncada, S., and Bolaños, J.P. (2004) Nature Cell Biol. 6, 45–51. Hers, H.G. and Van Schaftingen, E. (1982) Biochem. J. 206, 1–12. Manzano, A., Rosa, J.L., Ventura, F., Perez, J.X., Nadal, M., Estivill, X., Ambrosio, S., Gil, J., and Bartrons, R. (1998) Cytogenet. Cell Genet. 83, 214–217. 161. Marsin, A.S., Bertrand, L., Rider, M.H., Deprez, J., Beauloye, C., Vincent, M.F., Van den Berghe, G., Carling, D., and Hue, L. (2000) Curr. Biol. 10, 1247–1255.

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

168

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Nitric Oxide Cell Signaling Mediated by cGMP

169

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

170

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide Cell Signaling Mediated by cGMP

171

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

172

Nitric Oxide, Cell Signaling, and Gene Expression

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].

Nitric Oxide Cell Signaling Mediated by cGMP

173

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.

174

Nitric Oxide, Cell Signaling, and Gene Expression

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].

Nitric Oxide Cell Signaling Mediated by cGMP

175

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

176

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide Cell Signaling Mediated by cGMP

177

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

178

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide Cell Signaling Mediated by cGMP

179

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

180

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide Cell Signaling Mediated by cGMP

181

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

182

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide Cell Signaling Mediated by cGMP

183

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

184

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide Cell Signaling Mediated by cGMP

185

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

186

Nitric Oxide, Cell Signaling, and Gene Expression

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].

Nitric Oxide Cell Signaling Mediated by cGMP

187

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.

7.3.3 NO/CGMP SIGNALING

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

188

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Nitric Oxide Cell Signaling Mediated by cGMP

189

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.

7.3.4 NO/CGMP SIGNALING

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

190

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Nitric Oxide Cell Signaling Mediated by cGMP

7.3.7 NO/CGMP SIGNALING

AND

191

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.

7.3.8 NO/CGMP SIGNALING

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

192

Nitric Oxide, Cell Signaling, and Gene Expression

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).

Nitric Oxide Cell Signaling Mediated by cGMP

193

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.

REFERENCES 1. Greene, N.E. and J.H. Shaw, The identification of atmospheric nitric oxide by a spectroscopic scan. J. Air Pollut. Control Assoc., 1972. 22(6): 468–470. 2. Murad, F., et al., Properties and regulation of guanylate cyclase and some proposed functions for cyclic GMP. Adv. Cyclic Nucleotide Res., 1979. 11: 175–204. 3. Murad, F., Cyclic guanosine monophosphate as a mediator of vasodilation. J. Clin. Invest., 1986. 78(1): 1–5. 4. Ignarro, L.J., et al., Endothelium-derived relaxing factor produced and released from artery and vein is nitric oxide. Proc. Natl. Acad. Sci. USA, 1987. 84(24): 9265–9269. 5. Palmer, R.M., A.G. Ferrige, and S. Moncada, Nitric oxide release accounts for the biological activity of endothelium-derived relaxing factor. Nature, 1987. 327(6122): 524–526.

194

Nitric Oxide, Cell Signaling, and Gene Expression

6. Wink, D.A., et al., The multifaceted roles of nitric oxide in cancer. Carcinogenesis, 1998. 19(5): 711–721. 7. Wink, D.A. and J.B. Mitchell, Chemical biology of nitric oxide: Insights into regulatory, cytotoxic, and cytoprotective mechanisms of nitric oxide. Free Radic. Biol. Med., 1998. 25(4-5): 434–456. 8. Wink, D.A., K.M. Miranda, and M.G. Espey, Cytotoxicity related to oxidative and nitrosative stress by nitric oxide. Exp. Biol. Med. (Maywood), 2001. 226(7): 621–623. 9 Abe, K., et al., Induction of nitrotyrosine-like immunoreactivity in the lower motor neuron of amyotrophic lateral sclerosis. Neurosci. Lett., 1995. 199(2): 152–154. 10. Jang, D. and G.A. Murrell, Nitric oxide in arthritis. Free Radic. Biol. Med., 1998. 24(9): 1511–1519. 11. Kooy, N.W., et al., Extensive tyrosine nitration in human myocardial inflammation: evidence for the presence of peroxynitrite. Crit. Care Med., 1997. 25(5): 812–819. 12. Iwashita, E., et al., High nitric oxide synthase activity in endothelial cells in ulcerative colitis. J. Gastroenterol., 1995. 30(4): 551–554. 13. Kelly, C.J. and D.P. Gold, Nitric oxide in interstitial nephritis and other autoimmune diseases. Semin. Nephrol., 1999. 19(3): 288–295. 14. Chan, N.N., P. Vallance, and H.M. Colhoun, Nitric oxide and vascular responses in Type I diabetes. Diabetologia, 2000. 43(2): 137–147. 15. Schulz, J.B., et al., The role of mitochondrial dysfunction and neuronal nitric oxide in animal models of neurodegenerative diseases. Mol. Cell Biochem., 1997. 174(1-2): 193–197. 16. Davis, K.L., et al., Novel effects of nitric oxide. Annu. Rev. Pharmacol. Toxicol., 2001. 41: 203–236. 17. Brown, G.C. and C.E. Cooper, Nanomolar concentrations of nitric oxide reversibly inhibit synaptosomal respiration by competing with oxygen at cytochrome oxidase. FEBS Lett., 1994. 356(2-3): 295–298. 18. Schweizer, M. and C. Richter, Nitric oxide potently and reversibly deenergizes mitochondria at low oxygen tension. Biochem. Biophys. Res. Commun., 1994. 204(1): 169–175. 19. Cleeter, M.W., et al., Reversible inhibition of cytochrome c oxidase, the terminal enzyme of the mitochondrial respiratory chain, by nitric oxide. Implications for neurodegenerative diseases. FEBS Lett., 1994. 345(1): 50–54. 20. Arnold, W.P., et al., Nitric oxide activates guanylate cyclase and increases guanosine 3′:5′-cyclic monophosphate levels in various tissue preparations. Proc. Natl. Acad. Sci. USA, 1977. 74(8): 3203–3207. 21. MacMicking, J., Q.W. Xie, and C. Nathan, Nitric oxide and macrophage function. Annu Rev Immunol, 1997. 15: 323–350. 22. Chao, D.S., et al., Localization of neuronal nitric oxide synthase. Methods Enzymol., 1996. 268: 488–496. 23. Bredt, D.S. and S.H. Snyder, Nitric oxide mediates glutamate-linked enhancement of cGMP levels in the cerebellum. Proc. Natl. Acad. Sci. USA, 1989. 86(22): 9030–9033. 24. Brenman, J.E., et al., Cloning and characterization of postsynaptic density 93, a nitric oxide synthase interacting protein. J. Neurosci., 1996. 16(23): 7407–7415. 25. Brenman, J.E., et al., Interaction of nitric oxide synthase with the postsynaptic density protein PSD-95 and alpha1-syntrophin mediated by PDZ domains. Cell, 1996. 84(5): 757–767.

Nitric Oxide Cell Signaling Mediated by cGMP

195

26. Kornau, H.C., et al., Domain interaction between NMDA receptor subunits and the postsynaptic density protein PSD-95. Science, 1995. 269(5231): 1737–1740. 27. Jaffrey, S.R. and S.H. Snyder, PIN: an associated protein inhibitor of neuronal nitric oxide synthase. Science, 1996. 274(5288): 774–777. 28. Rodriguez-Crespo, I., et al., Binding of dynein light chain (PIN) to neuronal nitric oxide synthase in the absence of inhibition. Arch. Biochem. Biophys., 1998. 359(2): 297–304. 29. Venema, V.J., et al., Interaction of neuronal nitric-oxide synthase with caveolin3 in skeletal muscle. Identification of a novel caveolin scaffolding/inhibitory domain. J. Biol. Chem., 1997. 272(45): 28187–28190. 30. Garcia-Cardena, G., et al., Dissecting the interaction between nitric oxide synthase (NOS) and caveolin. Functional significance of the nos caveolin binding domain in vivo. J. Biol. Chem., 1997. 272(41): 25437–25440. 31. Pollock, J.S., et al., Particulate and soluble bovine endothelial nitric oxide synthases are structurally similar proteins yet different from soluble brain nitric oxide synthase. J. Cardiovasc. Pharmacol., 1992. 20 Suppl. 12: S50–53. 32. Shaul, P.W., et al., Acylation targets endothelial nitric-oxide synthase to plasmalemmal caveolae. J. Biol. Chem., 1996. 271(11): 6518–6522. 33. Feron, O., et al., Endothelial nitric oxide synthase targeting to caveolae. Specific interactions with caveolin isoforms in cardiac myocytes and endothelial cells. J. Biol. Chem., 1996. 271(37): 22810–22814. 34. Pollock, J.S., et al., Endothelial nitric oxide synthase is myristylated. FEBS Lett., 1992. 309(3): 402–404. 35. Robinson, L.J., L. Busconi, and T. Michel, Agonist-modulated palmitoylation of endothelial nitric oxide synthase. J. Biol. Chem., 1995. 270(3): 995–998. 36. Feron, O., et al., The endothelial nitric-oxide synthase-caveolin regulatory cycle. J. Biol. Chem., 1998. 273(6): 3125–3128. 37. Schmidt, H.H., et al., Regulation and subcellular location of nitrogen oxide synthases in RAW264.7 macrophages. Mol. Pharmacol., 1992. 41(4): 615–624. 38. Forstermann, U., et al., Induced RAW 264.7 macrophages express soluble and particulate nitric oxide synthase: inhibition by transforming growth factor-beta. Eur. J. Pharmacol., 1992. 225(2): 161–165. 39. Vodovotz, Y., et al., Vesicle membrane association of nitric oxide synthase in primary mouse macrophages. J. Immunol., 1995. 154(6): 2914–2925. 40. Lucas, K.A., et al., Guanylyl cyclases and signaling by cyclic GMP. Pharmacol. Rev., 2000. 52(3): 375–414. 41. Kimura, H. and F. Murad, Two forms of guanylate cyclase in mammalian tissues and possible mechanisms for their regulation. Metabolism, 1975. 24(3): 439–445. 42. Katsuki, S., et al., Stimulation of guanylate cyclase by sodium nitroprusside, nitroglycerin and nitric oxide in various tissue preparations and comparison to the effects of sodium azide and hydroxylamine. J. Cyclic Nucleotide Res., 1977. 3(1): 23–35. 43. Craven, P.A. and F.R. DeRubertis, Restoration of the responsiveness of purified guanylate cyclase to nitrosoguanidine, nitric oxide, and related activators by heme and hemeproteins. Evidence for involvement of the paramagnetic nitrosyl-heme complex in enzyme activation. J. Biol. Chem., 1978. 253(23): 8433–8443.

196

Nitric Oxide, Cell Signaling, and Gene Expression

44. Ignarro, L.J., et al., Mechanism of vascular smooth muscle relaxation by organic nitrates, nitrites, nitroprusside and nitric oxide: evidence for the involvement of S-nitrosothiols as active intermediates. J. Pharmacol. Exp. Ther., 1981. 218(3): 739–749. 45. Stone, J.R. and M.A. Marletta, Soluble guanylate cyclase from bovine lung: activation with nitric oxide and carbon monoxide and spectral characterization of the ferrous and ferric states. Biochemistry, 1994. 33(18): 5636–5640. 46. Martin, E., et al., A constitutively activated mutant of human soluble guanylyl cyclase (sGC): implication for the mechanism of sGC activation. Proc. Natl. Acad. Sci. USA, 2003. 100(16): 9208–9213. 47. Koesling, D., Studying the structure and regulation of soluble guanylyl cyclase. Methods, 1999. 19(4): 485–493. 48. Kamisaki, Y., et al., Soluble guanylate cyclase from rat lung exists as a heterodimer. J. Biol. Chem., 1986. 261(16): 7236–7241. 49. Zabel, U., et al., Human soluble guanylate cyclase: functional expression and revised isoenzyme family. Biochem. J., 1998. 335(Pt 1): 51–57. 50. Andreopoulos, S. and A. Papapetropoulos, Molecular aspects of soluble guanylyl cyclase regulation. Gen. Pharmacol., 2000. 34(3): 147–157. 51. Russwurm, M., et al., Functional properties of a naturally occurring isoform of soluble guanylyl cyclase. Biochem. J., 1998. 335(Pt 1): 125–130. 52. Yuen, P.S., L.R. Potter, and D.L. Garbers, A new form of guanylyl cyclase is preferentially expressed in rat kidney. Biochemistry, 1990. 29(49): 10872–10878. 53. Buechler, W.A., M. Nakane, and F. Murad, Expression of soluble guanylate cyclase activity requires both enzyme subunits. Biochem. Biophys. Res. Commun., 1991. 174(1): 351–357. 54. Zabel, U., et al., Homodimerization of soluble guanylyl cyclase subunits. Dimerization analysis using a glutathione s-transferase affinity tag. J. Biol. Chem., 1999. 274(26): 18149–52. 55. Koglin, M., et al., Nitric oxide activates the beta 2 subunit of soluble guanylyl cyclase in the absence of a second subunit. J. Biol. Chem., 2001. 276(33): 30737–30743. 56. Garbers, D.L., The guanylyl cyclase receptors. Methods, 1999. 19(4): 477–484. 57. Sunahara, R.K., et al., Crystal structure of the adenylyl cyclase activator Gsalpha [see comments]. Science, 1997. 278(5345): 1943–1947. 58. Sunahara, R.K., et al., Exchange of substrate and inhibitor specificities between adenylyl and guanylyl cyclases. J. Biol. Chem., 1998. 273(26): 16332–6338. 59. Wedel, B., et al., Functional domains of soluble guanylyl cyclase. J. Biol. Chem., 1995. 270(42): 24871–24875. 60. Foerster, J., et al., A functional heme-binding site of soluble guanylyl cyclase requires intact N-termini of alpha 1 and beta 1 subunits. Eur. J. Biochem., 1996. 240(2): 380–386. 61. Gerzer, R., et al., Soluble guanylate cyclase purified from bovine lung contains heme and copper. FEBS Lett., 1981. 132(1): 71–74. 62. Stone, J.R. and M.A. Marletta, Spectral and kinetic studies on the activation of soluble guanylate cyclase by nitric oxide. Biochemistry, 1996. 35(4): 1093–1099. 63. Stone, J.R., et al., Electron paramagnetic resonance spectral evidence for the formation of a pentacoordinate nitrosyl-heme complex on soluble guanylate cyclase. Biochem. Biophys. Res. Commun., 1995. 207(2): 572–577.

Nitric Oxide Cell Signaling Mediated by cGMP

197

64. Tomita, T., et al., Effects of GTP on bound nitric oxide of soluble guanylate cyclase probed by resonance Raman spectroscopy. Biochemistry, 1997. 36(33): 10155–10160. 65. Deinum, G., et al., Binding of nitric oxide and carbon monoxide to soluble guanylate cyclase as observed with Resonance raman spectroscopy. Biochemistry, 1996. 35(5): 1540–1547. 66. Zhao, Y., et al., A molecular basis for nitric oxide sensing by soluble guanylate cyclase. Proc. Natl. Acad. Sci. USA, 1999. 96(26): 14753–14758. 67. Kharitonov, V.G., et al., Kinetics of nitric oxide dissociation from five- and sixcoordinate nitrosyl hemes and heme proteins, including soluble guanylate cyclase. Biochemistry, 1997. 36(22): 6814–6818. 68. Ignarro, L.J., K.S. Wood, and M.S. Wolin, Activation of purified soluble guanylate cyclase by protoporphyrin IX. Proc. Natl. Acad. Sci. USA, 1982. 79(9): 2870–2873. 69. Zhao, Y., et al., Identification of histidine 105 in the beta1 subunit of soluble guanylate cyclase as the heme proximal ligand. Biochemistry, 1998. 37(13): 4502–4509. 70. Wedel, B., et al., Mutation of His-105 in the beta 1 subunit yields a nitric oxideinsensitive form of soluble guanylyl cyclase. Proc. Natl. Acad. Sci. USA, 1994. 91(7): 2592–2596. 71. Tesmer, J.J., et al., Crystal structure of the catalytic domains of adenylyl cyclase in a complex with Gsα.GTPγS [see comments]. Science, 1997. 278(5345): 1907–1916. 72. Russwurm, M., N. Wittau, and D. Koesling, Guanylyl cyclase/PSD-95 interaction: targeting of the nitric oxide-sensitive alpha2beta1 guanylyl cyclase to synaptic membranes. J. Biol. Chem., 2001. 276(48): 44647–44652. 73. Zabel, U., et al., Calcium-dependent membrane association sensitizes soluble guanylyl cyclase to nitric oxide. Nat. Cell Biol., 2002. 4(4): 307–311. 74. Fesenko, E.E., S.S. Kolesnikov, and A.L. Lyubarsky, Induction by cyclic GMP of cationic conductance in plasma membrane of retinal rod outer segment. Nature, 1985. 313(6000): 310–313. 75. Haynes, L. and K.W. Yau, Cyclic GMP-sensitive conductance in outer segment membrane of catfish cones. Nature, 1985. 317(6032): 61–64. 76. Nakamura, T. and G.H. Gold, A cyclic nucleotide-gated conductance in olfactory receptor cilia. Nature, 1987. 325(6103): 442–444. 77. Dryer, S.E. and D. Henderson, A cyclic GMP-activated channel in dissociated cells of the chick pineal gland. Nature, 1991. 353(6346): 756–758. 78. Hofmann, F., M. Biel, and U.B. Kaupp, International Union of Pharmacology. XLII. Compendium of voltage-gated ion channels: cyclic nucleotide-modulated channels. Pharmacol. Rev., 2003. 55(4): 587–589. 79. Bradley, J., et al., Nomenclature for ion channel subunits. Science, 2001. 294(5549): 2095–2096. 80. Chen, T.Y., et al., A new subunit of the cyclic nucleotide-gated cation channel in retinal rods. Nature, 1993. 362(6422): 764–767. 81. Korschen, H.G., et al., A 240 kDa protein represents the complete beta subunit of the cyclic nucleotide-gated channel from rod photoreceptor. Neuron, 1995. 15(3): 627–636. 82. Varnum, M.D. and W.N. Zagotta, Subunit interactions in the activation of cyclic nucleotide-gated ion channels. Biophys. J., 1996. 70(6): 2667–2679.

198

Nitric Oxide, Cell Signaling, and Gene Expression

83. Liu, D.T., G.R. Tibbs, and S.A. Siegelbaum, Subunit stoichiometry of cyclic nucleotide-gated channels and effects of subunit order on channel function. Neuron, 1996. 16(5): 983–990. 84. Gordon, S.E. and W.N. Zagotta, Subunit interactions in coordination of Ni2+ in cyclic nucleotide-gated channels. Proc. Natl. Acad. Sci. USA, 1995. 92(22): 10222–10226. 85. Zheng, J., M.C. Trudeau, and W.N. Zagotta, Rod cyclic nucleotide-gated channels have a stoichiometry of three CNGA1 subunits and one CNGB1 subunit. Neuron, 2002. 36(5): 891–896. 86. Zhong, H., et al., The heteromeric cyclic nucleotide-gated channel adopts a 3A:1B stoichiometry. Nature, 2002. 420(6912): 193–198. 87. Weitz, D., et al., Subunit stoichiometry of the CNG channel of rod photoreceptors. Neuron, 2002. 36(5): 881–889. 88. Gerstner, A., et al., Molecular cloning and functional characterization of a new modulatory cyclic nucleotide-gated channel subunit from mouse retina. J. Neurosci., 2000. 20(4): 1324–1332. 89. Bonigk, W., et al., Rod and cone photoreceptor cells express distinct genes for cGMP-gated channels. Neuron, 1993. 10(5): 865–877. 90. Sautter, A., et al., An isoform of the rod photoreceptor cyclic nucleotide-gated channel beta subunit expressed in olfactory neurons. Proc. Natl. Acad. Sci. USA, 1998. 95(8): 4696–4701. 91. Picco, C., P. Gavazzo, and A. Menini, Co-expression of wild-type and mutant olfactory cyclic nucleotide-gated channels: restoration of the native sensitivity to Ca(2+) and Mg(2+) blockage. Neuroreport, 2001. 12(11): 2363–2367. 92. Bonigk, W., et al., The native rat olfactory cyclic nucleotide-gated channel is composed of three distinct subunits. J. Neurosci., 1999. 19(13): 5332–5347. 93. Liman, E.R. and L.B. Buck, A second subunit of the olfactory cyclic nucleotidegated channel confers high sensitivity to cAMP. Neuron, 1994. 13(3): 611–621. 94. Dhallan, R.S., et al., Primary structure and functional expression of a cyclic nucleotide-activated channel from olfactory neurons. Nature, 1990. 347(6289): 184–187. 95. Bradley, J., et al., Heteromeric olfactory cyclic nucleotide-gated channels: a subunit that confers increased sensitivity to cAMP. Proc. Natl. Acad. Sci. USA, 1994. 91(19): 8890–8894. 96. Bradley, J., et al., Functional expression of the heteromeric “olfactory” cyclic nucleotide-gated channel in the hippocampus: a potential effector of synaptic plasticity in brain neurons. J. Neurosci., 1997. 17(6): 1993–2005. 97. Biel, M., et al., Primary structure and functional expression of a cyclic nucleotidegated channel from rabbit aorta. FEBS Lett., 1993. 329(1-2): 134–138. 98. Biel, M., et al., Another member of the cyclic nucleotide-gated channel family, expressed in testis, kidney, and heart. Proc. Natl. Acad. Sci. USA, 1994. 91(9): 3505–3509. 99. Biel, M., et al., Molecular cloning and expression of the modulatory subunit of the cyclic nucleotide-gated cation channel. J. Biol. Chem., 1996. 271(11): 6349–6355. 100. Distler, M., et al., Expression of cyclic nucleotide-gated cation channels in nonsensory tissues and cells. Neuropharmacology, 1994. 33(11): 1275–1282.

Nitric Oxide Cell Signaling Mediated by cGMP

199

101. Kingston, P.A., F. Zufall, and C.J. Barnstable, Rat hippocampal neurons express genes for both rod retinal and olfactory cyclic nucleotide-gated channels: novel targets for cAMP/cGMP function. Proc. Natl. Acad. Sci. USA, 1996. 93(19): 10440–10445. 102. Weyand, I., et al., Cloning and functional expression of a cyclic-nucleotide-gated channel from mammalian sperm. Nature, 1994. 368(6474): 859–863. 103. Kaupp, U.B., et al., Primary structure and functional expression from complementary DNA of the rod photoreceptor cyclic GMP-gated channel. Nature, 1989. 342(6251): 762–766. 104. Henn, D.K., A. Baumann, and U.B. Kaupp, Probing the transmembrane topology of cyclic nucleotide-gated ion channels with a gene fusion approach. Proc. Natl. Acad. Sci. USA, 1995. 92(16): 7425–7429. 105. Molday, R.S., et al., The cGMP-gated channel of the rod photoreceptor cell characterization and orientation of the amino terminus. J. Biol. Chem., 1991. 266(32): 21917–21922. 106. Wohlfart, P., et al., Antibodies against synthetic peptides used to determine the topology and site of glycosylation of the cGMP-gated channel from bovine rod photoreceptors. J. Biol. Chem., 1992. 267(1): 644–648. 107. Doyle, D.A., et al., The structure of the potassium channel: molecular basis of K+ conduction and selectivity. Science, 1998. 280(5360): 69–77. 108. Su, Y., et al., Regulatory subunit of protein kinase A: structure of deletion mutant with cAMP binding domains. Science, 1995. 269(5225): 807–813. 109. Varnum, M.D., K.D. Black, and W.N. Zagotta, Molecular mechanism for ligand discrimination of cyclic nucleotide-gated channels. Neuron, 1995. 15(3): 619–625. 110. Gordon, S.E. and W.N. Zagotta, A histidine residue associated with the gate of the cyclic nucleotide-activated channels in rod photoreceptors. Neuron, 1995. 14(1): 177–183. 111. Sunderman, E.R. and W.N. Zagotta, Mechanism of allosteric modulation of rod cyclic nucleotide-gated channels. J. Gen. Physiol., 1999. 113(5): 601–620. 112. Shapiro, M.S. and W.N. Zagotta, Structural basis for ligand selectivity of heteromeric olfactory cyclic nucleotide-gated channels. Biophys. J., 2000. 78(5): 2307–2320. 113. Anholt, R.R., Molecular neurobiology of olfaction. Crit. Rev. Neurobiol., 1993. 7(1): 1–22. 114. Ildefonse, M. and N. Bennett, Single-channel study of the cGMP-dependent conductance of retinal rods from incorporation of native vesicles into planar lipid bilayers. J. Membr. Biol., 1991. 123(2): 133–147. 115. Karpen, J.W., et al., Interactions between divalent cations and the gating machinery of cyclic GMP-activated channels in salamander retinal rods. J. Gen. Physiol., 1993. 101(1): 1–25. 116. Molokanova, E., et al., Activity-dependent modulation of rod photoreceptor cyclic nucleotide-gated channels mediated by phosphorylation of a specific tyrosine residue. J. Neurosci., 1999. 19(12): 4786–4795. 117. Molokanova, E., et al., Modulation of rod photoreceptor cyclic nucleotide-gated channels by tyrosine phosphorylation. J. Neurosci., 1997. 17(23): 9068–9076. 118. Muller, F., et al., Ligand sensitivity of the 2 subunit from the bovine cone cGMPgated channel is modulated by protein kinase C but not by calmodulin. J. Physiol., 2001. 532(Pt 2): 399–409.

200

Nitric Oxide, Cell Signaling, and Gene Expression

119. Gordon, S.E., D.L. Brautigan, and A.L. Zimmerman, Protein phosphatases modulate the apparent agonist affinity of the light-regulated ion channel in retinal rods. Neuron, 1992. 9(4): 739–748. 120. Muller, F., et al., Phosphorylation of mammalian olfactory cyclic nucleotide-gated channels increases ligand sensitivity. J. Neurosci., 1998. 18(1): 164–173. 121. Wohlfart, P., H. Muller, and N.J. Cook, Lectin binding and enzymatic deglycosylation of the cGMP-gated channel from bovine rod photoreceptors. J. Biol. Chem., 1989. 264(35): 20934–20939. 122. Womack, K.B., et al., Do phosphatidylinositides modulate vertebrate phototransduction? J. Neurosci., 2000. 20(8): 2792–2799. 123. Crary, J.I., et al., Mechanism of inhibition of cyclic nucleotide-gated ion channels by diacylglycerol. J. Gen. Physiol., 2000. 116(6): 755–768. 124. Rebrik, T.I. and J.I. Korenbrot, In intact cone photoreceptors, a Ca2+-dependent, diffusible factor modulates the cGMP-gated ion channels differently than in rods. J. Gen. Physiol., 1998. 112(5): 537–548. 125. Ko, G.Y., M.L. Ko, and S.E. Dryer, Circadian regulation of cGMP-gated cationic channels of chick retinal cones. Erk MAP Kinase and Ca2+/calmodulin-dependent protein kinase II. Neuron, 2001. 29(1): 255–266. 126. Yau, K.W. and D.A. Baylor, Cyclic GMP-activated conductance of retinal photoreceptor cells. Annu. Rev. Neurosci., 1989. 12: 289–327. 127. Cobbs, W.H. and E.N. Pugh, Jr., Kinetics and components of the flash photocurrent of isolated retinal rods of the larval salamander, Ambystoma tigrinum. J. Physiol., 1987. 394: 529–572. 128. Yau, K.W. and K. Nakatani, Light-suppressible, cyclic GMP-sensitive conductance in the plasma membrane of a truncated rod outer segment. Nature, 1985. 317(6034): 252–255. 129. Koch, K.W. and L. Stryer, Highly cooperative feedback control of retinal rod guanylate cyclase by calcium ions. Nature, 1988. 334(6177): 64–66. 130. Lolley, R.N. and E. Racz, Calcium modulation of cyclic GMP synthesis in rat visual cells. Vision Res., 1982. 22(12): 1481–1486. 131. Hsu, Y.T. and R.S. Molday, Modulation of the cGMP-gated channel of rod photoreceptor cells by calmodulin. Nature, 1993. 361(6407): 76–79. 132. Kawamura, S., et al., Recoverin has S-modulin activity in frog rods. J. Biol. Chem., 1993. 268(20): 14579–14582. 133. Lagnado, L. and D.A. Baylor, Calcium controls light-triggered formation of catalytically active rhodopsin. Nature, 1994. 367(6460): 273–277. 134. Frings, S., et al., Profoundly different calcium permeation and blockage determine the specific function of distinct cyclic nucleotide-gated channels. Neuron, 1995. 15(1): 169–179. 135. Dzeja, C., et al., Ca2+ permeation in cyclic nucleotide-gated channels. EMBO J., 1999. 18(1): 131–144. 136. Harbeck, B., et al., Phosphorylation of the vasodilator-stimulated phosphoprotein regulates its interaction with actin. J. Biol. Chem., 2000. 275(40): 30817–30825. 137. Varnum, M.D. and W.N. Zagotta, Interdomain interactions underlying activation of cyclic nucleotide-gated channels. Science, 1997. 278(5335): 110–113. 138. Kurahashi, T. and A. Menini, Mechanism of odorant adaptation in the olfactory receptor cell. Nature, 1997. 385(6618): 725–729. 139. Liu, M., et al., Calcium-calmodulin modulation of the olfactory cyclic nucleotidegated cation channel. Science, 1994. 266(5189): 1348–1354.

Nitric Oxide Cell Signaling Mediated by cGMP

201

140. McNaughton, P.A., Light response of vertebrate photoreceptors. Physiol. Rev., 1990. 70(3): 847–883. 141. Pugh, E.N., Jr. and T.D. Lamb, Cyclic GMP and calcium: the internal messengers of excitation and adaptation in vertebrate photoreceptors. Vision Res., 1990. 30(12): 1923–1948. 142. Dryja, T.P., et al., Mutations in the gene encoding the alpha subunit of the rod cGMP-gated channel in autosomal recessive retinitis pigmentosa. Proc. Natl. Acad. Sci. USA, 1995. 92(22): 10177–10181. 143. Kohl, S., et al., Total colourblindness is caused by mutations in the gene encoding the alpha-subunit of the cone photoreceptor cGMP-gated cation channel. Nat. Genet., 1998. 19(3): 257–259. 144. Kohl, S., et al., Mutations in the CNGB3 gene encoding the beta-subunit of the cone photoreceptor cGMP-gated channel are responsible for achromatopsia (ACHM3) linked to chromosome 8q21. Hum. Mol. Genet., 2000. 9(14): 2107–2116. 145. Sundin, O.H., et al., Genetic basis of total colourblindness among the Pingelapese islanders. Nat. Genet., 2000. 25(3): 289–293. 146. Brunet, L.J., G.H. Gold, and J. Ngai, General anosmia caused by a targeted disruption of the mouse olfactory cyclic nucleotide-gated cation channel. Neuron, 1996. 17(4): 681–693. 147. Baker, H., et al., Targeted deletion of a cyclic nucleotide-gated channel subunit (OCNC1): biochemical and morphological consequences in adult mice. J. Neurosci., 1999. 19(21): 9313–9321. 148. Parent, A., et al., Synaptic transmission and hippocampal long-term potentiation in olfactory cyclic nucleotide-gated channel type 1 null mouse. J. Neurophysiol., 1998. 79(6): 3295–3301. 149. Biel, M., et al., Selective loss of cone function in mice lacking the cyclic nucleotide-gated channel CNG3. Proc. Natl. Acad. Sci. USA, 1999. 96(13): 7553–7557. 150. Munger, S.D., et al., Central role of the CNGA4 channel subunit in Ca2+-calmodulin-dependent odor adaptation. Science, 2001. 294(5549): 2172–2175. 151. Kurenny, D.E., et al., Modulation of ion channels in rod photoreceptors by nitric oxide. Neuron, 1994. 13(2): 315–324. 152. Koch, K.W., et al., Functional coupling of a Ca2+/calmodulin-dependent nitric oxide synthase and a soluble guanylyl cyclase in vertebrate photoreceptor cells. EMBO J., 1994. 13(14): 3312–3320. 153. Liepe, B.A., et al., Nitric oxide synthase in Muller cells and neurons of salamander and fish retina. J. Neurosci., 1994. 14(12): 7641–7654. 154. Savchenko, A., S. Barnes, and R.H. Kramer, Cyclic-nucleotide-gated channels mediate synaptic feedback by nitric oxide. Nature, 1997. 390(6661): 694–698. 155. Broillet, M.C. and S. Firestein, Direct activation of the olfactory cyclic nucleotidegated channel through modification of sulfhydryl groups by NO compounds. Neuron, 1996. 16(2): 377–385. 156. Brown, R.L., S.D. Snow, and T.L. Haley, Movement of gating machinery during the activation of rod cyclic nucleotide-gated channels. Biophys. J., 1998. 75(2): 825–833. 157. Komatsu, H., et al., Functional reconstitution of a heteromeric cyclic nucleotidegated channel of Caenorhabditis elegans in cultured cells. Brain Res., 1999. 821(1): 160–168.

202

Nitric Oxide, Cell Signaling, and Gene Expression

158. Trivedi, B. and R.H. Kramer, Real-time patch-cram detection of intracellular cGMP reveals long-term suppression of responses to NO and muscarinic agonists. Neuron, 1998. 21(4): 895–906. 159. Breer, H., T. Klemm, and I. Boekhoff, Nitric oxide mediated formation of cyclic GMP in the olfactory system. Neuroreport, 1992. 3(11): 1030–1032. 160. Lischka, F.W. and D. Schild, Effects of nitric oxide upon olfactory receptor neurones in Xenopus laevis. Neuroreport, 1993. 4(5): 582–584. 161. Inamura, K., M. Kashiwayanagi, and K. Kurihara, Effects of cGMP and sodium nitroprusside on odor responses in turtle olfactory sensory neurons. Am. J. Physiol., 1998. 275(5 Pt 1): C1201– C1206. 162. Kawai, F. and P. Sterling, AMPA receptor activates a G-protein that suppresses a cGMP-gated current. J. Neurosci., 1999. 19(8): 2954–2959. 163. Kishimoto, J., et al., Localization of nitric oxide synthase in the mouse olfactory and vomeronasal system: a histochemical, immunological and in situ hybridization study. Eur. J. Neurosci., 1993. 5(12): 1684–1694. 164. Bredt, D.S. and S.H. Snyder, Nitric oxide: a physiologic messenger molecule. Annu. Rev. Biochem., 1994. 63: 175–195. 165. Kulkarni, A.P., T.V. Getchell, and M.L. Getchell, Neuronal nitric oxide synthase is localized in extrinsic nerves regulating perireceptor processes in the chemosensory nasal mucosae of rats and humans. J. Comp. Neurol., 1994. 345(1): 125–138. 166. Roskams, A.J., et al., Nitric oxide mediates the formation of synaptic connections in developing and regenerating olfactory receptor neurons. Neuron, 1994. 13(2): 289–299. 167. Marks, G.S., et al., Does carbon monoxide have a physiological function? Trends Pharmacol. Sci., 1991. 12(5): 185–18. 168. Goldberg, N.D., et al., 18O-Labeling of guanosine monophosphate upon hydrolysis of cyclic guanosine 3′:5′-monophosphate by phosphodiesterase. J. Biol. Chem., 1980. 255(21): 10344–10347. 169. Francis, S.H., et al., Zinc interactions and conserved motifs of the cGMP-binding cGMP-specific phosphodiesterase suggest that it is a zinc hydrolase. J. Biol. Chem., 1994. 269(36): 22477–22480. 170. Butcher, R.W. and E.W. Sutherland, Adenosine 3′,5′-phosphate in biological materials. I. Purification and properties of cyclic 3',5'-nucleotide phosphodiesterase and use of this enzyme to characterize adenosine 3′,5′-phosphate in human urine. J. Biol. Chem., 1962. 237: 1244–1250. 171. Rall, T.W. and E.W. Sutherland, Formation of a cyclic adenine ribonucleotide by tissue particles. J. Biol. Chem., 1958. 232(2): 1065–1076. 172. Maurice, D.H., et al., Cyclic nucleotide phosphodiesterase activity, expression, and targeting in cells of the cardiovascular system. Mol. Pharmacol., 2003. 64(3): 533–546. 173. Sheth, S.B., et al., Isolation and regulation of the cGMP-inhibited cAMP phosphodiesterase in human erythroleukemia cells. Thromb. Haemost., 1997. 77(1): 155–162. 174. Kakkar, R., R.V. Raju, and R.K. Sharma, Calmodulin-dependent cyclic nucleotide phosphodiesterase (PDE1). Cell Mol. Life Sci., 1999. 55(8-9): 1164–1186. 175. Zhao, A.Z., et al., Recent advances in the study of Ca2+/CaM-activated phosphodiesterases: expression and physiological functions. Adv. Second Messenger Phosphoprotein Res., 1997. 31: 237–251.

Nitric Oxide Cell Signaling Mediated by cGMP

203

176. Sonnenburg, W.K., D. Seger, and J.A. Beavo, Molecular cloning of a cDNA encoding the “61-kDa” calmodulin-stimulated cyclic nucleotide phosphodiesterase. Tissue-specific expression of structurally related isoforms. J. Biol. Chem., 1993. 268(1): 645–652. 177. Chiu, P.J., et al., Comparative effects of vinpocetine and 8-Br-cyclic GMP on the contraction and 45Ca-fluxes in the rabbit aorta. Am. J. Hypertens., 1988. 1(3 Pt 1): 262–268. 178. Rybalkin, S.D., et al., Calmodulin-stimulated cyclic nucleotide phosphodiesterase (PDE1C) is induced in human arterial smooth muscle cells of the synthetic, proliferative phenotype. J. Clin. Invest., 1997. 100(10): 2611–2621. 179. Rybalkin, S.D., et al., Cyclic nucleotide phosphodiesterase 1C promotes human arterial smooth muscle cell proliferation. Circ. Res., 2002. 90(2): 151–157. 180. Han, P., et al., The calcium/calmodulin-dependent phosphodiesterase PDE1C down-regulates glucose-induced insulin secretion. J. Biol. Chem., 1999. 274(32): 22337–22344. 181. Bode, D.C., J.R. Kanter, and L.L. Brunton, Cellular distribution of phosphodiesterase isoforms in rat cardiac tissue. Circ. Res., 1991. 68(4):1070–1079. 182. Ashikaga, T., S.J. Strada, and W.J. Thompson, Altered expression of cyclic nucleotide phosphodiesterase isozymes during culture of aortic endothelial cells. Biochem. Pharmacol., 1997. 54(10): 1071–1079. 183. Martinez, S.E., et al., The two GAF domains in phosphodiesterase 2A have distinct roles in dimerization and in cGMP binding. Proc. Natl. Acad. Sci. USA, 2002. 99(20): 13260–13265. 184. Juilfs, D.M., et al., A subset of olfactory neurons that selectively express cGMPstimulated phosphodiesterase (PDE2) and guanylyl cyclase-D define a unique olfactory signal transduction pathway. Proc. Natl. Acad. Sci. USA, 1997. 94(7): 3388–3395. 185. Sadhu, K., et al., Differential expression of the cyclic GMP-stimulated phosphodiesterase PDE2A in human venous and capillary endothelial cells. J. Histochem. Cytochem., 1999. 47(7): 895–906. 186. Dickinson, N.T., E.K. Jang, and R.J. Haslam, Activation of cGMP-stimulated phosphodiesterase by nitroprusside limits cAMP accumulation in human platelets: effects on platelet aggregation. Biochem. J., 1997. 323 (Pt 2): 371–377. 187. Wechsler, J., et al., Isoforms of cyclic nucleotide phosphodiesterase PDE3A in cardiac myocytes. J. Biol. Chem., 2002. 277(41): 38072–38078. 188. Reinhardt, R.R., et al., Distinctive anatomical patterns of gene expression for cGMP-inhibited cyclic nucleotide phosphodiesterases. J. Clin. Invest., 1995. 95(4): 1528–1538. 189. Shakur, Y., et al., Regulation and function of the cyclic nucleotide phosphodiesterase (PDE3) gene family. Prog. Nucleic Acid Res. Mol. Biol., 2001. 66: 241–277. 190. Vandecasteele, G., et al., Cyclic GMP regulation of the L-type Ca(2+) channel current in human atrial myocytes. J. Physiol., 2001. 533(Pt 2): 329–340. 191. Loughney, K., et al., Isolation and characterization of cDNAs encoding PDE5A, a human cGMP-binding, cGMP-specific 3′,5′-cyclic nucleotide phosphodiesterase. Gene, 1998. 216(1): 139–147. 192. Corbin, J.D. and S.H. Francis, Cyclic GMP phosphodiesterase-5: target of sildenafil. J. Biol. Chem., 1999. 274(20): 13729–13732.

204

Nitric Oxide, Cell Signaling, and Gene Expression

193. Michelakis, E., et al., Oral sildenafil is an effective and specific pulmonary vasodilator in patients with pulmonary arterial hypertension: comparison with inhaled nitric oxide. Circulation, 2002. 105(20): 2398–2403. 194. Mullershausen, F., et al., Rapid nitric oxide-induced desensitization of the cGMP response is caused by increased activity of phosphodiesterase type 5 paralleled by phosphorylation of the enzyme. J. Cell Biol., 2001. 155(2): 271–278. 195. Mullershausen, F., et al., Direct activation of PDE5 by cGMP: long-term effects within NO/cGMP signaling. J. Cell Biol., 2003. 160(5): 719–727. 196. Yarfitz, S. and J.B. Hurley, Transduction mechanisms of vertebrate and invertebrate photoreceptors. J. Biol. Chem., 1994. 269(20): 14329–14332. 197. Soderling, S.H. and J.A. Beavo, Regulation of cAMP and cGMP signaling: new phosphodiesterases and new functions. Curr. Opin. Cell Biol., 2000. 12(2): 174–179. 198. van Staveren, W.C., et al., Cloning and localization of the cGMP-specific phosphodiesterase type 9 in the rat brain. J. Neurocytol., 2002. 31(8-9): 729–741. 199. Soderling, S.H., S.J. Bayuga, and J.A. Beavo, Identification and characterization of a novel family of cyclic nucleotide phosphodiesterases. J. Biol. Chem., 1998. 273(25): 15553–15558. 200. Soderling, S.H., S.J. Bayuga, and J.A. Beavo, Isolation and characterization of a dual-substrate phosphodiesterase gene family: PDE10A. Proc. Natl. Acad. Sci. USA, 1999. 96(12): 7071–7076. 201. Fujishige, K., et al., Cloning and characterization of a novel human phosphodiesterase that hydrolyzes both cAMP and cGMP (PDE10A). J. Biol. Chem., 1999. 274(26): 18438–18445. 202. Hetman, J.M., et al., Cloning and characterization of two splice variants of human phosphodiesterase 11A. Proc. Natl. Acad. Sci. USA, 2000. 97(23): 12891–12895. 203. Yuasa, K., et al., Genomic organization of the human phosphodiesterase PDE11A gene. Evolutionary relatedness with other PDEs containing GAF domains. Eur. J. Biochem., 2001. 268(1): 168–178. 204. Fawcett, L., et al., Molecular cloning and characterization of a distinct human phosphodiesterase gene family: PDE11A. Proc. Natl. Acad. Sci. USA, 2000. 97(7): 3702–3707. 205. Francis, S.H. and J.D. Corbin, Cyclic nucleotide-dependent protein kinases: intracellular receptors for cAMP and cGMP action. Crit. Rev. Clin. Lab. Sci., 1999. 36(4): 275–328. 206. Francis, S.H., et al., Types I alpha and I beta isozymes of cGMP-dependent protein kinase: alternative mRNA splicing may produce different inhibitory domains. Second Messengers Phosphoproteins, 1988. 12(5-6): 301–310. 207. Wernet, W., V. Flockerzi, and F. Hofmann, The cDNA of the two isoforms of bovine cGMP-dependent protein kinase. FEBS Lett., 1989. 251(1-2): 191–196. 208. Uhler, M.D., Cloning and expression of a novel cyclic GMP-dependent protein kinase from mouse brain. J. Biol. Chem., 1993. 268(18): 13586–13591. 209. Lohmann, S.M., et al., Distinct and specific functions of cGMP-dependent protein kinases. Trends Biochem. Sci., 1997. 22(8): 307–312. 210. Pfeifer, A., et al., Defective smooth muscle regulation in cGMP kinase I-deficient mice. EMBO J., 1998. 17(11): 3045–3051. 211. Feil, R., et al., Cyclic GMP-dependent protein kinases and the cardiovascular system: insights from genetically modified mice. Circ. Res., 2003. 93(10): 907–916.

Nitric Oxide Cell Signaling Mediated by cGMP

205

212. Wall, M.E., et al., Mechanisms associated with cGMP binding and activation of cGMP-dependent protein kinase. Proc. Natl. Acad. Sci. USA, 2003. 100(5): 2380–2385. 213. Chu, D.M., et al., Activation by cyclic GMP binding causes an apparent conformational change in cGMP-dependent protein kinase. J. Biol. Chem., 1997. 272(50): 31922–31928. 214. Aitken, A., B.A. Hemmings, and F. Hofmann, Identification of the residues on cyclic GMP-dependent protein kinase that are autophosphorylated in the presence of cyclic AMP and cyclic GMP. Biochim. Biophys. Acta, 1984. 790(3): 219–225. 215. Smith, J.A., et al., Autophosphorylation of type Ibeta cGMP-dependent protein kinase increases basal catalytic activity and enhances allosteric activation by cGMP or cAMP. J. Biol. Chem., 1996. 271(34): 20756–20762. 216. Francis, S.H., et al., Arginine 75 in the pseudosubstrate sequence of type Ibeta cGMP-dependent protein kinase is critical for autoinhibition, although autophosphorylated serine 63 is outside this sequence. J. Biol. Chem., 1996. 271(34): 20748–20755. 217. Landgraf, W. and F. Hofmann, The amino terminus regulates binding to and activation of cGMP-dependent protein kinase. Eur. J. Biochem., 1989. 181(3): 643–650. 218. de Jonge, H.R. and O.M. Rosen, Self-phosphorylation of cyclic guanosine 3′:5′monophosphate-dependent protein kinase from bovine lung. Effect of cyclic adenosine 3′:5′-monophosphate, cyclic guanosine 3′:5′-monophosphate and histone. J. Biol. Chem., 1977. 252(8): 2780–2783. 219. Gamm, D.M., et al., The type II isoform of cGMP-dependent protein kinase is dimeric and possesses regulatory and catalytic properties distinct from the type I isoforms. J. Biol. Chem., 1995. 270(45): 27380–27388. 220. Monken, C.E. and G.N. Gill, Structural analysis of cGMP-dependent protein kinase using limited proteolysis. J. Biol. Chem., 1980. 255(15): 7067–7070. 221. Wolfe, L., S.H. Francis, and J.D. Corbin, Properties of a cGMP-dependent monomeric protein kinase from bovine aorta. J. Biol. Chem., 1989. 264(7): 4157–4162. 222. Sinnaeve, P., et al., Overexpression of a constitutively active protein kinase G mutant reduces neointima formation and in-stent restenosis. Circulation, 2002. 105(24): 2911–2916. 223. Pussard, G., et al., Endothelin-1 modulates cyclic GMP production and relaxation in human pulmonary vessels. J. Pharmacol. Exp. Ther., 1995. 274(2): 969–975. 224. Iranami, H., et al., A beta-adrenoceptor agonist evokes a nitric oxide-cGMP relaxation mechanism modulated by adenylyl cyclase in rat aorta. Halothane does not inhibit this mechanism. Anesthesiology, 1996. 85(5): 1129–1138. 225. Trovati, M., et al., Insulin increases cyclic nucleotide content in human vascular smooth muscle cells: a mechanism potentially involved in insulin-induced modulation of vascular tone. Diabetologia, 1995. 38(8): 936–941. 226. Trovati, M., et al., Studies on the influence of insulin on cyclic adenosine monophosphate in human vascular smooth muscle cells: dependence on cyclic guanosine monophosphate and modulation of catecholamine effects. Diabetologia, 1996. 39(10): 1156–1164. 227. Rosenfeld, C.R., et al., Nitric oxide contributes to estrogen-induced vasodilation of the ovine uterine circulation. J. Clin. Invest., 1996. 98(9): 2158–2166.

206

Nitric Oxide, Cell Signaling, and Gene Expression

228. White, R.E., D.J. Darkow, and J.L. Lang, Estrogen relaxes coronary arteries by opening BKCa channels through a cGMP-dependent mechanism. Circ. Res., 1995. 77(5): 936–942. 229. Clifton, V.L., et al., Corticotropin-releasing hormone-induced vasodilatation in the human fetal-placental circulation: involvement of the nitric oxide-cyclic guanosine 3′,5′-monophosphate-mediated pathway. J. Clin. Endocrinol. Metab., 1995. 80(10): 2888–2893. 230. Huang, P.L., et al., Hypertension in mice lacking the gene for endothelial nitric oxide synthase. Nature, 1995. 377(6546): 239–242. 231. Sausbier, M., et al., Mechanisms of NO/cGMP-dependent vasorelaxation. Circ. Res., 2000. 87(9): 825–830. 232. Massberg, S., et al., Increased adhesion and aggregation of platelets lacking cyclic guanosine 3′,5′-monophosphate kinase I. J. Exp. Med., 1999. 189(8): 1255–1264. 233. Furukawa, K., Y. Tawada, and M. Shigekawa, Regulation of the plasma membrane Ca2+ pump by cyclic nucleotides in cultured vascular smooth muscle cells. J. Biol. Chem., 1988. 263(17): 8058–8065. 234. Vrolix, M., et al., Cyclic GMP-dependent protein kinase stimulates the plasmalemmal Ca2+ pump of smooth muscle via phosphorylation of phosphatidylinositol. Biochem. J., 1988. 255(3): 855–863. 235. Yoshida, Y., et al., Cyclic GMP-dependent protein kinase stimulates the plasma membrane Ca2+ pump ATPase of vascular smooth muscle via phosphorylation of a 240-kDa protein. J. Biol. Chem., 1991. 266(29): 19819–19825. 236. Cornwell, T.L., et al., Regulation of sarcoplasmic reticulum protein phosphorylation by localized cyclic GMP-dependent protein kinase in vascular smooth muscle cells. Mol. Pharmacol., 1991. 40(6): 923–931. 237. Quignard, J.F., et al., Voltage-gated calcium channel currents in human coronary myocytes. Regulation by cyclic GMP and nitric oxide. J. Clin. Invest., 1997. 99(2): 185–193. 238. Tewari, K. and J.M. Simard, Sodium nitroprusside and cGMP decrease Ca2+ channel availability in basilar artery smooth muscle cells. Pflugers Arch., 1997. 433(3): 304–311. 239. Clapp, L.H. and A.M. Gurney, Modulation of calcium movements by nitroprusside in isolated vascular smooth muscle cells. Pflugers Arch., 1991. 418(5): 462–470. 240. Yamakage, M., C.A. Hirshman, and T.L. Croxton, Sodium nitroprusside stimulates Ca2+-activated K+ channels in porcine tracheal smooth muscle cells. Am. J. Physiol., 1996. 270(3 Pt 1): L338– L345. 241. Mikawa, K., H. Kume, and K. Takagi, Effects of atrial natriuretic peptide and 8brom cyclic guanosine monophosphate on human tracheal smooth muscle. Arzneimittelforschung, 1998. 48(9): 914–918. 242. Tanaka, Y., et al., Involvement of maxi-K(Ca) channel activation in atrial natriuretic peptide-induced vasorelaxation. Naunyn Schmiedebergs Arch. Pharmacol., 1998. 357(6): 705–708. 243. Zhou, X.B., et al., Protein phosphatase 2A is essential for the activation of Ca2+activated K+ currents by cGMP-dependent protein kinase in tracheal smooth muscle and Chinese hamster ovary cells. J. Biol. Chem., 1996. 271(33): 19760–19767. 244. Komalavilas, P. and T.M. Lincoln, Phosphorylation of the inositol 1,4,5-trisphosphate receptor by cyclic GMP-dependent protein kinase. J. Biol. Chem., 1994. 269(12): 8701–8707.

Nitric Oxide Cell Signaling Mediated by cGMP

207

245. Komalavilas, P. and T.M. Lincoln, Phosphorylation of the inositol 1,4,5-trisphosphate receptor. Cyclic GMP-dependent protein kinase mediates cAMP and cGMP dependent phosphorylation in the intact rat aorta. J. Biol. Chem., 1996. 271(36): 21933–21938. 246. Schlossmann, J., et al., Regulation of intracellular calcium by a signalling complex of IRAG, IP3 receptor and cGMP kinase Ibeta. Nature, 2000. 404(6774): 197–201. 247. Lincoln, T.M. and T.L. Cornwell, Intracellular cyclic GMP receptor proteins. FASEB J., 1993. 7(2): 328–338. 248. Feil, R., et al., Functional reconstitution of vascular smooth muscle cells with cGMP-dependent protein kinase I isoforms. Circ. Res., 2002. 90(10): 1080–1086. 249. Chen, X.L. and C.M. Rembold, Cyclic nucleotide-dependent regulation of Mn2+ influx, [Ca2+]i, and arterial smooth muscle relaxation. Am. J. Physiol., 1992. 263(2 Pt 1): C468– C473. 250. Lee, M.R., L. Li, and T. Kitazawa, Cyclic GMP causes Ca2+ desensitization in vascular smooth muscle by activating the myosin light chain phosphatase. J. Biol. Chem., 1997. 272(8): 5063–5068. 251. Wu, X., A.V. Somlyo, and A.P. Somlyo, Cyclic GMP-dependent stimulation reverses G-protein-coupled inhibition of smooth muscle myosin light chain phosphate. Biochem. Biophys. Res. Commun., 1996. 220(3): 658–663. 252. Surks, H.K., et al., Regulation of myosin phosphatase by a specific interaction with cGMP-dependent protein kinase Ialpha. Science, 1999. 286(5444): 1583–1587. 253. Walker, L.A., et al., Site-specific phosphorylation and point mutations of telokin modulate its Ca2+-desensitizing effect in smooth muscle. J. Biol. Chem., 2001. 276(27): 24519–24524. 254. Nishikawa, M., et al., Phosphorylation of mammalian myosin light chain kinases by the catalytic subunit of cyclic AMP-dependent protein kinase and by cyclic GMP-dependent protein kinase. J. Biol. Chem., 1984. 259(13): 8429–8436. 255. Sauzeau, V., et al., Cyclic GMP-dependent protein kinase signaling pathway inhibits RhoA-induced Ca2+ sensitization of contraction in vascular smooth muscle. J. Biol. Chem., 2000. 275(28): 21722–21729. 256. Sauzeau, V., et al., RhoA expression is controlled by nitric oxide through cGMPdependent protein kinase activation. J. Biol. Chem., 2003. 278(11): 9472–9480. 257. Reinhard, M., et al., The 46/50 kDa phosphoprotein VASP purified from human platelets is a novel protein associated with actin filaments and focal contacts. EMBO J., 1992. 11(6): 2063–2070. 258. Smolenski, A., et al., Regulation of human endothelial cell focal adhesion sites and migration by cGMP-dependent protein kinase I. J. Biol. Chem., 2000. 275(33): 25723–25732. 259. Murphy-Ullrich, J.E., et al., Cyclic GMP-dependent protein kinase is required for thrombospondin and tenascin mediated focal adhesion disassembly. J. Cell Sci., 1996. 109 (Pt 10): 2499–2508. 260. Draijer, R., et al., Expression of cGMP-dependent protein kinase I and phosphorylation of its substrate, vasodilator-stimulated phosphoprotein, in human endothelial cells of different origin. Circ. Res., 1995. 77(5): 897–905. 261. Aszodi, A., et al., The vasodilator-stimulated phosphoprotein (VASP) is involved in cGMP- and cAMP-mediated inhibition of agonist-induced platelet aggregation, but is dispensable for smooth muscle function. EMBO J., 1999. 18(1): 37–48.

208

Nitric Oxide, Cell Signaling, and Gene Expression

262. Jerius, H., et al., Endothelial-dependent vasodilation is associated with increases in the phosphorylation of a small heat shock protein (HSP20). J. Vasc. Surg., 1999. 29(4): 678–684. 263. Beall, A., et al., The small heat shock-related protein, HSP20, is phosphorylated on serine 16 during cyclic nucleotide-dependent relaxation. J. Biol. Chem., 1999. 274(16): 11344–11351. 264. Brophy, C.M., S. Lamb, and A. Graham, The small heat shock-related protein-20 is an actin-associated protein. J. Vasc. Surg., 1999. 29(2): 326–333. 265. Tessier, D.J., et al., The small heat shock protein (HSP) 20 is dynamically associated with the actin cross-linking protein actinin. J. Surg. Res., 2003. 111(1): 152–157. 266. Beall, A.C., et al., Cyclic nucleotide-dependent vasorelaxation is associated with the phosphorylation of a small heat shock-related protein. J. Biol. Chem., 1997. 272(17): 11283–11287. 267. Brophy, C.M., et al., Small heat shock proteins and vasospasm in human umbilical artery smooth muscle. Biol. Reprod., 1997. 57(6): 1354–1359. 268. Radomski, M.W., R.M. Palmer, and S. Moncada, Comparative pharmacology of endothelium-derived relaxing factor, nitric oxide and prostacyclin in platelets. Br. J. Pharmacol., 1987. 92(1): 181–187. 269. Radomski, M.W., R.M. Palmer, and S. Moncada, Endogenous nitric oxide inhibits human platelet adhesion to vascular endothelium. Lancet, 1987. 2(8567): 1057–1058. 270. Radomski, M.W., R.M. Palmer, and S. Moncada, The anti-aggregating properties of vascular endothelium: interactions between prostacyclin and nitric oxide. Br. J. Pharmacol., 1987. 92(3): 639–646. 271. Moro, M.A., et al., cGMP mediates the vascular and platelet actions of nitric oxide: confirmation using an inhibitor of the soluble guanylyl cyclase. Proc. Natl. Acad. Sci. USA, 1996. 93(4): 1480–1485. 272. Radomski, M.W., R.M. Palmer, and S. Moncada, An L-arginine/nitric oxide pathway present in human platelets regulates aggregation. Proc. Natl. Acad. Sci. USA, 1990. 87(13): 5193–5197. 273. Buechler, W.A., et al., Soluble guanylyl cyclase and platelet function. Ann. NY Acad. Sci., 1994. 714: 151–157. 274. Haslam, R.J., N.T. Dickinson, and E.K. Jang, Cyclic nucleotides and phosphodiesterases in platelets. Thromb. Haemost., 1999. 82(2): 412–423. 275. Nakashima, S., et al., Inhibitory action of cyclic GMP on secretion, polyphosphoinositide hydrolysis and calcium mobilization in thrombin-stimulated human platelets. Biochem. Biophys. Res. Commun., 1986. 135(3): 1099–1104. 276. Geiger, J., et al., Role of cGMP and cGMP-dependent protein kinase in nitrovasodilator inhibition of agonist-evoked calcium elevation in human platelets. Proc. Natl. Acad. Sci. USA, 1992. 89(3): 1031–1035. 277. Johansson, J.S. and D.H. Haynes, Cyclic GMP increases the rate of the calcium extrusion pump in intact human platelets but has no direct effect on the dense tubular calcium accumulation system. Biochim. Biophys. Acta, 1992. 1105(1): 40–50. 278. Morgan, R.O. and A.C. Newby, Nitroprusside differentially inhibits ADP-stimulated calcium influx and mobilization in human platelets. Biochem. J., 1989. 258(2): 447–454.

Nitric Oxide Cell Signaling Mediated by cGMP

209

279. Trepakova, E.S., R.A. Cohen, and V.M. Bolotina, Nitric oxide inhibits capacitative cation influx in human platelets by promoting sarcoplasmic/endoplasmic reticulum Ca2+-ATPase-dependent refilling of Ca2+ stores. Circ. Res., 1999. 84(2): 201–209. 280. Cavallini, L., et al., Prostacyclin and sodium nitroprusside inhibit the activity of the platelet inositol 1,4,5-trisphosphate receptor and promote its phosphorylation. J. Biol. Chem., 1996. 271(10): 5545–5551. 281. Ryningen, A., B. Olav Jensen, and H. Holmsen, Elevation of cyclic AMP decreases phosphoinositide turnover and inhibits thrombin-induced secretion in human platelets. Biochim. Biophys. Acta, 1998. 1394(2-3): 235–248. 282. Murohara, T., et al., Inhibition of nitric oxide biosynthesis promotes P-selectin expression in platelets. Role of protein kinase C. Arterioscler. Thromb. Vasc. Biol., 1995. 15(11): 2068–2075. 283. Michelson, A.D., et al., Effects of nitric oxide/EDRF on platelet surface glycoproteins. Am. J. Physiol., 1996. 270(5 Pt 2): H1640– H1648. 284. Mendelsohn, M.E., et al., Inhibition of fibrinogen binding to human platelets by S-nitroso-N-acetylcysteine. J. Biol. Chem., 1990. 265(31): 19028–19034. 285. Wang, G.R., et al., Mechanism of platelet inhibition by nitric oxide: in vivo phosphorylation of thromboxane receptor by cyclic GMP-dependent protein kinase. Proc. Natl. Acad. Sci. USA, 1998. 95(9): 4888–4893. 286. Aktas, B., et al., Inhibition of platelet P2Y12 and alpha2A receptor signaling by cGMP-dependent protein kinase. Biochem. Pharmacol., 2002. 64(3): 433–439. 287. Reinhard, M., T. Jarchau, and U. Walter, Actin-based motility: stop and go with Ena/VASP proteins. Trends Biochem. Sci., 2001. 26(4): 243–249. 288. Butt, E., et al., Heat shock protein 27 is a substrate of cGMP-dependent protein kinase in intact human platelets: phosphorylation-induced actin polymerization caused by HSP27 mutants. J. Biol. Chem., 2001. 276(10): 7108–7113. 289. Butt, E., et al., Actin binding of human LIM and SH3 protein is regulated by cGMP- and cAMP-dependent protein kinase phosphorylation on serine 146. J. Biol. Chem., 2003. 278(18): 15601–15607. 290. Saklatvala, J., et al., Role for p38 mitogen-activated protein kinase in platelet aggregation caused by collagen or a thromboxane analogue. J. Biol. Chem., 1996. 271(12): 6586–6589. 291. Schwarz, U.R., et al., Inhibition of agonist-induced p42 and p38 mitogen-activated protein kinase phosphorylation and CD40 ligand/P-selectin expression by cyclic nucleotide-regulated pathways in human platelets. Biochem. Pharmacol., 2000. 60(9): 1399–1407. 292. Reep, B.R. and E.G. Lapetina, Nitric oxide stimulates the phosphorylation of rap1b in human platelets and acts synergistically with iloprost. Biochem. Biophys. Res. Commun., 1996. 219(1): 1–5. 293. Miura, Y., et al., Phosphorylation of smg p21B/rap1B p21 by cyclic GMP-dependent protein kinase. FEBS Lett., 1992. 297(1-2): 171–174. 294. Garthwaite, J., S.L. Charles, and R. Chess-Williams, Endothelium-derived relaxing factor release on activation of NMDA receptors suggests role as intercellular messenger in the brain. Nature, 1988. 336(6197): 385–388. 295. Murphy, S., Production of nitric oxide by glial cells: regulation and potential roles in the CNS. Glia, 2000. 29(1): 1–13.

210

Nitric Oxide, Cell Signaling, and Gene Expression

296. Dinerman, J.L., et al., Endothelial nitric oxide synthase localized to hippocampal pyramidal cells: implications for synaptic plasticity. Proc. Natl. Acad. Sci. USA, 1994. 91(10): 4214–4218. 297. Wiencken, A.E. and V.A. Casagrande, Endothelial nitric oxide synthetase (eNOS) in astrocytes: another source of nitric oxide in neocortex. Glia, 1999. 26(4): 280–290. 298. Bohme, G.A., et al., Possible involvement of nitric oxide in long-term potentiation. Eur. J. Pharmacol., 1991. 199(3): 379–381. 299. O'Dell, T.J., et al., Tests of the roles of two diffusible substances in long-term potentiation: evidence for nitric oxide as a possible early retrograde messenger. Proc. Natl. Acad. Sci. USA, 1991. 88(24): 11285–11289. 300. Schuman, E.M. and D.V. Madison, A requirement for the intercellular messenger nitric oxide in long-term potentiation. Science, 1991. 254(5037): 1503–1506. 301. Haley, J.E., G.L. Wilcox, and P.F. Chapman, The role of nitric oxide in hippocampal long-term potentiation. Neuron, 1992. 8(2): 211–216. 302. O'Dell, T.J., et al., Endothelial NOS and the blockade of LTP by NOS inhibitors in mice lacking neuronal NOS. Science, 1994. 265(5171): 542–546. 303. Son, H., et al., Long-term potentiation is reduced in mice that are doubly mutant in endothelial and neuronal nitric oxide synthase. Cell, 1996. 87(6): 1015–1023. 304. Zhuo, M., et al., Role of guanylyl cyclase and cGMP-dependent protein kinase in long-term potentiation. Nature, 1994. 368(6472): 635–639. 305. El-Husseini, A.E., et al., Localization of the cGMP-dependent protein kinases in relation to nitric oxide synthase in the brain. J. Chem. Neuroanat., 1999. 17(1): 45–55. 306. Kleppisch, T., et al., Long-term potentiation in the hippocampal CA1 region of mice lacking cGMP-dependent kinases is normal and susceptible to inhibition of nitric oxide synthase. J. Neurosci., 1999. 19(1): 48–55. 307. Shibuki, K. and D. Okada, Endogenous nitric oxide release required for long-term synaptic depression in the cerebellum. Nature, 1991. 349(6307): 326–328. 308. Lev-Ram, V., et al., Absence of cerebellar long-term depression in mice lacking neuronal nitric oxide synthase. Learn. Mem., 1997. 4(1): 169–177. 309. Lev-Ram, V., et al., Synergies and coincidence requirements between NO, cGMP, and Ca2+ in the induction of cerebellar long-term depression. Neuron, 1997. 18(6): 1025–1038. 310. Linden, D.J. and J.A. Connor, Long-term depression of glutamate currents in cultured cerebellar purkinje neurons does not require nitric oxide signalling. Eur J. Neurosci., 1992. 4(1): 10–15. 311. Feil, R., et al., Impairment of LTD and cerebellar learning by Purkinje cell-specific ablation of cGMP-dependent protein kinase I. J. Cell Biol., 2003. 163(2): 295–302. 312. Boxall, A.R. and J. Garthwaite, Long-term depression in rat cerebellum requires both NO synthase and NO-sensitive guanylyl cyclase. Eur J. Neurosci., 1996. 8(10): 2209–2212. 313. Hartell, N.A., Inhibition of cGMP breakdown promotes the induction of cerebellar long-term depression. J. Neurosci., 1996. 16(9): 2881–2890. 314. Meller, S.T. and G.F. Gebhart, Nitric oxide (NO) and nociceptive processing in the spinal cord. Pain, 1993. 52(2): 127–136. 315. Sluka, K.A. and W.D. Willis, Increased spinal release of excitatory amino acids following intradermal injection of capsaicin is reduced by a protein kinase G inhibitor. Brain Res., 1998. 798(1-2): 281–286.

Nitric Oxide Cell Signaling Mediated by cGMP

211

316. Kitto, K.F., J.E. Haley, and G.L. Wilcox, Involvement of nitric oxide in spinally mediated hyperalgesia in the mouse. Neurosci. Lett., 1992. 148(1-2): 1–5. 317. Malmberg, A.B. and T.L. Yaksh, Spinal nitric oxide synthesis inhibition blocks NMDA-induced thermal hyperalgesia and produces antinociception in the formalin test in rats. Pain, 1993. 54(3): 291–300. 318. Zhuo, M., S.T. Meller, and G.F. Gebhart, Endogenous nitric oxide is required for tonic cholinergic inhibition of spinal mechanical transmission. Pain, 1993. 54(1): 71–78. 319. Gillespie, J.S., Searching for non-adrenergic, non-cholinergic autonomic transmitter, in Pharmacology, M.J. Rand, Raper, C., eds.. 1987, Excerta Medica: Amsterdam. 161–170. 320. Moncada, S., A. Higgs, and R. Furchgott, International Union of Pharmacology Nomenclature in Nitric Oxide Research. Pharmacol. Rev., 1997. 49(2): 137–142. 321. Fykse, E.M., C. Li, and T.C. Sudhof, Phosphorylation of rabphilin-3A by Ca2+/calmodulin- and cAMP-dependent protein kinases in vitro. J. Neurosci., 1995. 15(3 Pt 2): 2385–2395. 322. Smith, S.L. and T.S. Otis, Persistent changes in spontaneous firing of Purkinje neurons triggered by the nitric oxide signaling cascade. J. Neurosci., 2003. 23(2): 367–372. 323. Wang, T., Z. Xie, and B. Lu, Nitric oxide mediates activity-dependent synaptic suppression at developing neuromuscular synapses. Nature, 1995. 374(6519): 262–266. 324. Huganir, R.L. and K. Miles, Protein phosphorylation of nicotinic acetylcholine receptors. Crit. Rev. Biochem. Mol. Biol., 1989. 24(3): 183–215. 325. Luise, M., et al., Dystrophin is phosphorylated by endogenous protein kinases. Biochem. J., 1993. 293 (Pt. 1): 243–247. 326. Huang, C.C., S.H. Chan, and K.S. Hsu, cGMP/protein kinase G-dependent potentiation of glutamatergic transmission induced by nitric oxide in immature rat rostral ventrolateral medulla neurons in vitro. Mol. Pharmacol., 2003. 64(2): 521–532. 327. Stanton, P.K., et al., Long-term depression of presynaptic release from the readily releasable vesicle pool induced by NMDA receptor-dependent retrograde nitric oxide. J. Neurosci., 2003. 23(13): 5936–5944. 328. Kone, B.C. and C. Baylis, Biosynthesis and homeostatic roles of nitric oxide in the normal kidney. Am. J. Physiol., 1997. 272(5 Pt 2): F561– F578. 329. Ren, Y.L., J.L. Garvin, and O.A. Carretero, Role of macula densa nitric oxide and cGMP in the regulation of tubuloglomerular feedback. Kidney Int., 2000. 58(5): 2053–2060. 330. Wilcox, C.S., Role of macula densa NOS in tubuloglomerular feedback. Curr. Opin. Nephrol. Hypertens., 1998. 7(4): 443–449. 331. Kurtz, A. and C. Wagner, Role of nitric oxide in the control of renin secretion. Am. J. Physiol., 1998. 275(6 Pt 2): F849– F862. 332. Kurtz, A., et al., Mode of nitric oxide action on the renal vasculature. Acta Physiol. Scand., 2000. 168(1): 41–45. 333. Lahera, V., et al., Effects of NG-monomethyl-L-arginine and L-arginine on acetylcholine renal response. Hypertension, 1990. 15(6 Pt 1): 659–663. 334. Majid, D.S., et al., Renal responses to intra-arterial administration of nitric oxide donor in dogs. Hypertension, 1993. 22(4): 535–541.

212

Nitric Oxide, Cell Signaling, and Gene Expression

335. Lahera, V., et al., Exogenous cGMP prevents decrease in diuresis and natriuresis induced by inhibition of NO synthesis. Am. J. Physiol., 1993. 264(2 Pt 2): F344– F347. 336. Lahera, V., et al., Effects of NG-nitro-L-arginine methyl ester on renal function and blood pressure. Am. J. Physiol., 1991. 261(6 Pt 2): F1033– F1037. 337. Wagner, C., et al., Role of cGMP-kinase II in the control of renin secretion and renin expression. J. Clin. Invest., 1998. 102(8): 1576–1582. 338. Moreland, R.B., I. Goldstein, and A. Traish, Sildenafil, a novel inhibitor of phosphodiesterase type 5 in human corpus cavernosum smooth muscle cells. Life Sci., 1998. 62(20): PL 309–318. 339. Trigo-Rocha, F., et al., The role of cyclic adenosine monophosphate, cyclic guanosine monophosphate, endothelium and nonadrenergic, noncholinergic neurotransmission in canine penile erection. J. Urol., 1993. 149(4): 872–877. 340. Maytom, M.C., et al., A two-part pilot study of sildenafil (VIAGRA) in men with erectile dysfunction caused by spinal cord injury. Spinal Cord, 1999. 37(2): 110–116. 341. Hedlund, P., et al., Erectile dysfunction in cyclic GMP-dependent kinase I-deficient mice. Proc. Natl. Acad. Sci. USA, 2000. 97(5): 2349–2354. 342. Yallampalli, C., R.E. Garfield, and M. Byam-Smith, Nitric oxide inhibits uterine contractility during pregnancy but not during delivery. Endocrinology, 1993. 133(4): 1899–1902. 343. Izumi, H., C. Yallampalli, and R.E. Garfield, Gestational changes in L-arginineinduced relaxation of pregnant rat and human myometrial smooth muscle. Am. J. Obstet. Gynecol., 1993. 169(5): 1327–1337. 344. Weiner, C.P. and L.P. Thompson, Nitric oxide and pregnancy. Semin. Perinatol., 1997. 21(5): 367–380. 345. Amin, A.R. and S.B. Abramson, The role of nitric oxide in articular cartilage breakdown in osteoarthritis. Curr. Opin. Rheumatol., 1998. 10(3): 263–268. 346. Stern, P.H. and J. Diamond, Sodium nitroprusside increases cyclic GMP in fetal rat bone cells and inhibits resorption of fetal rat limb bones. Res. Commun. Chem. Pathol. Pharmacol., 1992. 75(1): 19–28. 347. Holliday, L.S., et al., Low NO concentrations inhibit osteoclast formation in mouse marrow cultures by cGMP-dependent mechanism. Am. J. Physiol., 1997. 272(3 Pt 2): F283–291. 348. Van Epps-Fung, C., et al., Regulation of osteoclastic acid secretion by cGMPdependent protein kinase. Biochem. Biophys. Res. Commun., 1994. 204(2): 565–571. 349. Otsuka, E., et al., Effects of nitric oxide from exogenous nitric oxide donors on osteoblastic metabolism. Eur. J. Pharmacol., 1998. 349(2-3): 345–350. 350. Kalra, D., et al., Nitric oxide provokes tumor necrosis factor-alpha expression in adult feline myocardium through a cGMP-dependent pathway. Circulation, 2000. 102(11): 1302–1307. 351. Perez-Sala, D. and S. Lamas, Regulation of cyclooxygenase-2 expression by nitric oxide in cells. Antioxid. Redox. Signal, 2001. 3(2): 231–248. 352. Tetsuka, T., et al., Nitric oxide amplifies interleukin 1-induced cyclooxygenase-2 expression in rat mesangial cells. J. Clin. Invest., 1996. 97(9): 2051–2056.

Nitric Oxide Cell Signaling Mediated by cGMP

213

353. Bouchie, J.L., H. Hansen, and E.P. Feener, Natriuretic factors and nitric oxide suppress plasminogen activator inhibitor-1 expression in vascular smooth muscle cells. Role of cGMP in the regulation of the plasminogen system. Arterioscler. Thromb. Vasc. Biol., 1998. 18(11): 1771–1779. 354. Gudi, T., S.M. Lohmann, and R.B. Pilz, Regulation of gene expression by cyclic GMP-dependent protein kinase requires nuclear translocation of the kinase: identification of a nuclear localization signal. Mol. Cell Biol., 1997. 17(9): 5244–5254. 355. Gudi, T., et al., NO activation of fos promoter elements requires nuclear translocation of G-kinase I and CREB phosphorylation but is independent of MAP kinase activation. Oncogene, 2000. 19(54): 6324–6333. 356. Gudi, T., et al., cGMP-dependent protein kinase inhibits serum-response elementdependent transcription by inhibiting rho activation and functions. J. Biol. Chem., 2002. 277(40): 37382–37393. 357. Pilz, R.B., et al., Nitric oxide and cGMP analogs activate transcription from AP1-responsive promoters in mammalian cells. FASEB J., 1995. 9(7): 552–558. 358. Haby, C., et al., Stimulation of the cyclic GMP pathway by NO induces expression of the immediate early genes c-fos and junB in PC12 cells. J. Neurochem., 1994. 62(2): 496–501. 359. Johnston, H.M. and B.J. Morris, N-methyl-D-aspartate and nitric oxide regulate the expression of calcium/calmodulin-dependent kinase II in the hippocampal dentate gyrus. Brain Res. Mol. Brain Res., 1995. 31(1-2): 141–150. 360. Gudi, T., et al., Regulation of gene expression by cGMP-dependent protein kinase. Transactivation of the c-fos promoter. J. Biol. Chem., 1996. 271(9): 4597–4600. 361. Casteel, D.E., et al., cGMP-dependent protein kinase I beta physically and functionally interacts with the transcriptional regulator TFII-I. J. Biol. Chem., 2002. 277(35): 32003–32014. 362. Cibelli, G., et al., Nitric oxide-induced programmed cell death in human neuroblastoma cells is accompanied by the synthesis of Egr-1, a zinc finger transcription factor. J. Neurosci. Res., 2002. 67(4): 450–460. 363. He, B. and G.F. Weber, Phosphorylation of NF-kappaB proteins by cyclic GMPdependent kinase. A noncanonical pathway to NF-kappaB activation. Eur. J. Biochem., 2003. 270(10): 2174–2185. 364. Fiedler, B., et al., Inhibition of calcineurin-NFAT hypertrophy signaling by cGMPdependent protein kinase type I in cardiac myocytes. Proc. Natl. Acad. Sci. USA, 2002. 99(17): 11363–11368. 365. Wu, C.F., N.H. Bishopric, and R.E. Pratt, Atrial natriuretic peptide induces apoptosis in neonatal rat cardiac myocytes. J. Biol. Chem., 1997. 272(23): 14860–14866. 366. Li, J., et al., Nitric oxide reversibly inhibits seven members of the caspase family via S-nitrosylation. Biochem. Biophys. Res. Commun., 1997. 240(2): 419–424. 367. Kim, Y.M., et al., Nitric oxide prevents tumor necrosis factor alpha-induced rat hepatocyte apoptosis by the interruption of mitochondrial apoptotic signaling through S-nitrosylation of caspase-8. Hepatology, 2000. 32(4 Pt 1): 770–778. 368. Ciani, E., et al., Nitric oxide protects neuroblastoma cells from apoptosis induced by serum deprivation through cAMP-response element-binding protein (CREB) activation. J. Biol. Chem., 2002. 277(51): 49896–49902.

214

Nitric Oxide, Cell Signaling, and Gene Expression

369. Andoh, T., C.C. Chiueh, and P.B. Chock, Cyclic GMP-dependent protein kinase regulates the expression of thioredoxin and thioredoxin peroxidase-1 during hormesis in response to oxidative stress-induced apoptosis. J. Biol. Chem., 2003. 278(2): 885–890. 370. Akahori, M., et al., Nitric oxide ameliorates actinomycin D/endotoxin-induced apoptotic liver failure in mice. J. Surg. Res., 1999. 85(2): 286–293. 371. Kim, Y.M., R.V. Talanian, and T.R. Billiar, Nitric oxide inhibits apoptosis by preventing increases in caspase-3-like activity via two distinct mechanisms. J. Biol. Chem., 1997. 272(49): 31138–31148. 372. Desai, K.M., W.C. Sessa, and J.R. Vane, Involvement of nitric oxide in the reflex relaxation of the stomach to accommodate food or fluid. Nature, 1991. 351(6326): 477–479. 373. Burns, A.J., et al., Interstitial cells of Cajal mediate inhibitory neurotransmission in the stomach. Proc. Natl. Acad. Sci. USA, 1996. 93(21): 12008–12013. 374. Huber, A., et al., Protein kinase G expression in the small intestine and functional importance for smooth muscle relaxation. Am. J. Physiol., 1998. 275(4 Pt 1): G629– G637. 375. Huang, P.L., et al., Targeted disruption of the neuronal nitric oxide synthase gene. Cell, 1993. 75(7): 1273–1286. 376. Rogers, N.E. and L.J. Ignarro, Constitutive nitric oxide synthase from cerebellum is reversibly inhibited by nitric oxide formed from L-arginine. Biochem. Biophys. Res. Commun., 1992. 189(1): 242–249. 377. Griscavage, J.M., et al., Nitric oxide inhibits neuronal nitric oxide synthase by interacting with the heme prosthetic group. Role of tetrahydrobiopterin in modulating the inhibitory action of nitric oxide. J. Biol. Chem., 1994. 269(34): 21644–21649. 378. Abu-Soud, H.M., et al., Neuronal nitric oxide synthase self-inactivates by forming a ferrous-nitrosyl complex during aerobic catalysis. J. Biol. Chem., 1995. 270(39): 22997–13006. 379. Lee, Y.C., E. Martin, and F. Murad, Human recombinant soluble guanylyl cyclase: expression, purification, and regulation. Proc. Natl. Acad. Sci. USA, 2000. 97(20): 10763–10768. 380. Papapetropoulos, A., et al., Downregulation of nitrovasodilator-induced cyclic GMP accumulation in cells exposed to endotoxin or interleukin-1 beta. Br. J. Pharmacol., 1996. 118(6): 1359–1366. 381. Filippov, G., D.B. Bloch, and K.D. Bloch, Nitric oxide decreases stability of mRNAs encoding soluble guanylate cyclase subunits in rat pulmonary artery smooth muscle cells. J. Clin. Invest., 1997. 100(4): 942–948. 382. Soff, G.A., et al., Smooth muscle cell expression of type I cyclic GMP-dependent protein kinase is suppressed by continuous exposure to nitrovasodilators, theophylline, cyclic GMP, and cyclic AMP. J. Clin. Invest., 1997. 100(10): 2580–2587. 383. Ferrero, R., et al., Nitric oxide-sensitive guanylyl cyclase activity inhibition through cyclic GMP-dependent dephosphorylation. J. Neurochem., 2000. 75(5): 2029–2039. 384. Wu, C.C., Ko, F.N., Kuo, S.E., Lee, F.Y., and Teng, C.M., VC-1 inhibited human platelet aggregation through NO-independent activation at soluble guanylate cyclase. Br. J. Pharmacol,, 1995. 166: 1973–1978.

Nitric Oxide Cell Signaling Mediated by cGMP

215

385. Stasch, J.P., Dembrowsky, K., Perzborn, E., Stahl, E., and Schramm, M., Cardiovascular action of a novel NO-independent guanylyl cyclase stimulator, BAY418543: in vivo studies. Br. J. Pharmacol., 2002. 135: 344–355.

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

218

Nitric Oxide, Cell Signaling, and Gene Expression

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

219

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

220

Nitric Oxide, Cell Signaling, and Gene Expression

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

Regulation of Cell Signaling by Protein Nitrosylation/ Denitrosylation

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

222

Nitric Oxide, Cell Signaling, and Gene Expression

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

Regulation of Cell Signaling by Protein Nitrosylation/ Denitrosylation

223

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).

224

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

Regulation of Cell Signaling by Protein Nitrosylation/ Denitrosylation

225

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.

226

Nitric Oxide, Cell Signaling, and Gene Expression

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.

REFERENCES 1. Stamler JS, Lamas S, Fang FC. Nitrosylation. The prototypic redox-based signaling mechanism. Cell 2001;106(6):675–683. 2. Eu JP, Sun J, Xu L, Stamler JS, Meissner G. The skeletal muscle calcium release channel: coupled O2 sensor and NO signaling functions. Cell 2000;102(4):499–509. 3. Sun J, Xin C, Eu JP, Stamler JS, Meissner G. Cysteine-3635 is responsible for skeletal muscle ryanodine receptor modulation by NO. Proc. Natl. Acad. Sci. USA 2001;98(20):11158–11162. 4. Brenman JE, Chao DS, Gee SH, McGee AW, Craven SE, Santillano DR, et al. Interaction of nitric oxide synthase with the postsynaptic density protein PSD-95 and alpha1-syntrophin mediated by PDZ domains. Cell 1996;84(5):757–767. 5. Jaffrey SR, Snowman AM, Eliasson MJ, Cohen NA, Snyder SH. CAPON: a protein associated with neuronal nitric oxide synthase that regulates its interactions with PSD95. Neuron 1998;20(1):115–124. 6. Russwurm M, Wittau N, Koesling D. Guanylyl cyclase/PSD-95 interaction: targeting of the nitric oxide-sensitive alpha2beta1 guanylyl cyclase to synaptic membranes. J. Biol. Chem. 2001;276(48):44647–44652. 7. Christopherson KS, Hillier BJ, Lim WA, Bredt DS. PSD-95 assembles a ternary complex with the N-methyl-D-aspartic acid receptor and a bivalent neuronal NO synthase PDZ domain. J. Biol. Chem. 1999;274(39):27467–27473. 8. Fang M, Jaffrey SR, Sawa A, Ye K, Luo X, Snyder SH. Dexras1: a G protein specifically coupled to neuronal nitric oxide synthase via CAPON. Neuron 2000;28(1):183–193. 9. Xu KY, Huso DL, Dawson TM, Bredt DS, Becker LC. Nitric oxide synthase in cardiac sarcoplasmic reticulum. Proc. Natl. Acad. Sci. USA 1999;96(2):657–662. 10. Matsumoto A, Comatas KE, Liu L, Stamler JS. Screening for nitric oxide-dependent protein-protein interactions. Science 2003;301(5633):657–661. 11. Stamler JS, Toone EJ, Lipton SA, Sucher NJ. (S)NO signals: translocation, regulation, and a consensus motif. Neuron 1997;18(5):691–696.

Regulation of Cell Signaling by Protein Nitrosylation/ Denitrosylation

227

12. Ascenzi P, Colasanti M, Persichini T, Muolo M, Polticelli F, Venturini G, et al. Re-evaluation of amino acid sequence and structural consensus rules for cysteinenitric oxide reactivity. Biol. Chem. 2000;381(7):623–627. 13. Hess DT, Matsumoto A, Nudelman R, Stamler JS. S-nitrosylation: spectrum and specificity. Nat. Cell Biol. 2001;3(2):E46–49. 14. Avila MA, Mingorance J, Martinez-Chantar ML, Casado M, Martin-Sanz P, Bosca L, et al. Regulation of rat liver S-adenosylmethionine synthetase during septic shock: role of nitric oxide. Hepatology 1997;25(2):391–396. 15. Perez-Mato I, Castro C, Ruiz FA, Corrales FJ, Mato JM. Methionine adenosyltransferase S-nitrosylation is regulated by the basic and acidic amino acids surrounding the target thiol. J. Biol. Chem. 1999;274(24):17075–17079. 16. Liu X, Miller MJ, Joshi MS, Thomas DD, Lancaster JR. Accelerated reaction of nitric oxide with O2 within the hydrophobic interior of biological membranes. Proc. Natl. Acad. Sci. USA 1998;95(5):2175–2179. 17. Stamler JS. Redox signaling: nitrosylation and related target interactions of nitric oxide. Cell 1994;78(6):931–936. 18. Liu L, Hausladen A, Zeng M, Que L, Heitman J, Stamler JS. A metabolic enzyme for S-nitrosothiol conserved from bacteria to humans. Nature 2001;410(6827):490–494. 19. Liu L, Yan Y, Zeng M, Zhang J, Hanes MA, Ahearn G, et al. Essential roles of S-nitrosothiols in vascular homeostasis and endotoxic shock. Cell 2004;116(4):617–628. 20. Muzio M, Chinnaiyan AM, Kischkel FC, O'Rourke K, Shevchenko A, Ni J, et al. FLICE, a novel FADD-homologous ICE/CED-3-like protease, is recruited to the CD95 (Fas/APO-1) death-inducing signaling complex. Cell 1996;85(6):817–827. 21. Kischkel FC, Hellbardt S, Behrmann I, Germer M, Pawlita M, Krammer PH, et al. Cytotoxicity-dependent APO-1 (Fas/CD95)-associated proteins form a deathinducing signaling complex (DISC) with the receptor. EMBO J. 1995;14(22):5579–5588. 22. Boldin MP, Goncharov TM, Goltsev YV, Wallach D. Involvement of MACH, a novel MORT1/FADD-interacting protease, in Fas/APO-1- and TNF receptorinduced cell death. Cell 1996;85(6):803–815. 23. Saleh A, Srinivasula SM, Acharya S, Fishel R, Alnemri ES. Cytochrome c and dATP-mediated oligomerization of Apaf-1 is a prerequisite for procaspase-9 activation. J. Biol. Chem. 1999;274(25):17941–17945. 24. Zou H, Henzel WJ, Liu X, Lutschg A, Wang X. Apaf-1, a human protein homologous to C. elegans CED-4, participates in cytochrome c-dependent activation of caspase-3. Cell 1997;90(3):405–413. 25. Zou H, Li Y, Liu X, Wang X. An APAF-1.cytochrome c multimeric complex is a functional apoptosome that activates procaspase-9. J. Biol. Chem. 1999;274(17):11549–11556. 26. Liu X, Kim CN, Yang J, Jemmerson R, Wang X. Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell 1996;86(1):147–157. 27. Li P, Nijhawan D, Budihardjo I, Srinivasula SM, Ahmad M, Alnemri ES, et al. Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade. Cell 1997;91(4):479489. 28. Li J, Billiar TR, Talanian RV, Kim YM. Nitric oxide reversibly inhibits seven members of the caspase family via S-nitrosylation. Biochem. Biophys. Res. Commun. 1997;240(2):419–424.

228

Nitric Oxide, Cell Signaling, and Gene Expression

29. Mannick JB, Hausladen A, Liu L, Hess DT, Zeng M, Miao QX, et al. Fas-induced caspase denitrosylation. Science 1999;284(5414):651–654. 30. Mannick JB, Schonhoff C, Papeta N, Ghafourifar P, Szibor M, Fang K, et al. SNitrosylation of mitochondrial caspases. J. Cell Biol. 2001;154(6):1111–1116. 31. Nunez G, Benedict MA, Hu Y, Inohara N. Caspases: the proteases of the apoptotic pathway. Oncogene 1998;17(25):3237–3245. 32. Schonhoff CM, Gaston B, Mannick JB. Nitrosylation of cytochrome c during apoptosis. J. Biol. Chem. 2003;278(20):18265–18270. 33. Hollmann M, Heinemann S. Cloned glutamate receptors. Annu. Rev. Neurosci. 1994;17:31–108. 34. McBain CJ, Mayer ML. N-methyl-D-aspartic acid receptor structure and function. Physiol. Rev. 1994;74(3):723–760. 35. Beal MF. Does impairment of energy metabolism result in excitotoxic neuronal death in neurodegenerative illnesses? Ann. Neurol. 1992;31(2):119–130. 36. Garthwaite J, Charles SL, Chess-Williams R. Endothelium-derived relaxing factor release on activation of NMDA receptors suggests role as intercellular messenger in the brain. Nature 1988;336(6197):385–388. 37. Lipton SA, Choi YB, Pan ZH, Lei SZ, Chen HS, Sucher NJ, et al. A redox-based mechanism for the neuroprotective and neurodestructive effects of nitric oxide and related nitroso-compounds. Nature 1993;364(6438):626–632. 38. Choi YB, Tenneti L, Le DA, Ortiz J, Bai G, Chen HS, et al. Molecular basis of NMDA receptor-coupled ion channel modulation by S-nitrosylation. Nat. Neurosci. 2000;3(1):15–21. 39. Lander HM, Ogiste JS, Pearce SF, Levi R, Novogrodsky A. Nitric oxide-stimulated guanine nucleotide exchange on p21ras. J. Biol. Chem. 1995;270(13):7017–7020. 40. Lander HM, Hajjar DP, Hempstead BL, Mirza UA, Chait BT, Campbell S, et al. A molecular redox switch on p21(ras). Structural basis for the nitric oxide-p21(ras) interaction. J. Biol. Chem. 1997;272(7):4323–4326. 41. Mott HR, Carpenter JW, Campbell SL. Structural and functional analysis of a mutant Ras protein that is insensitive to nitric oxide activation. Biochemistry 1997;36(12):3640–3644. 42. Williams JG, Pappu K, Campbell SL. Structural and biochemical studies of p21Ras S-nitrosylation and nitric oxide-mediated guanine nucleotide exchange. Proc. Natl. Acad. Sci. USA 2003;100(11):6376–6381. 43. Heo J, Campbell SL. Mechanism of p21Ras S-nitrosylation and kinetics of nitric oxide-mediated guanine nucleotide exchange. Biochemistry 2004;43(8):2314–2322. 44. Humbert P, Niroomand F, Fischer G, Mayer B, Koesling D, Hinsch KD, et al. Purification of soluble guanylyl cyclase from bovine lung by a new immunoaffinity chromatographic method. Eur. J. Biochem. 1990;190(2):273–278. 45. Stone JR, Marletta MA. Spectral and kinetic studies on the activation of soluble guanylate cyclase by nitric oxide. Biochemistry 1996;35(4):1093–1099. 46. Colledge M, Scott JD. AKAPs: from structure to function. Trends Cell Biol. 1999;9(6):216–221. 47. Xu L, Eu JP, Meissner G, Stamler JS. Activation of the cardiac calcium release channel (ryanodine receptor) by poly-S-nitrosylation. Science 1998;279(5348):234–237. 48. Eu JP, Hare JM, Hess DT, Skaf M, Sun J, Cardenas-Navina I, et al. Concerted regulation of skeletal muscle contractility by oxygen tension and endogenous nitric oxide. Proc. Natl. Acad. Sci. USA 2003;100(25):15229–15234.

Regulation of Cell Signaling by Protein Nitrosylation/ Denitrosylation

229

49. Nordberg J, Arner ES. Reactive oxygen species, antioxidants, and the mammalian thioredoxin system. Free Radic. Biol. Med. 2001;31(11):1287–1312. 50. Haendeler J, Hoffmann J, Tischler V, Berk BC, Zeiher AM, Dimmeler S. Redox regulatory and anti-apoptotic functions of thioredoxin depend on S-nitrosylation at cysteine 69. Nat. Cell Biol. 2002;4(10):743–749. 51. Hoffmann J, Haendeler J, Zeiher AM, Dimmeler S. TNFalpha and oxLDL reduce protein S-nitrosylation in endothelial cells. J. Biol. Chem. 2001;276(44):41383–41387. 52. Garg AK, Aggarwal BB. Reactive oxygen intermediates in TNF signaling. Mol. Immunol. 2002;39(9):509–517.

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

232

Nitric Oxide, Cell Signaling, and Gene Expression

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

233

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

234

Nitric Oxide, Cell Signaling, and Gene Expression

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-

Nitric Oxide and Caspase Activation

235

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

236

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide and Caspase Activation

237

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

238

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Nitric Oxide and Caspase Activation

239

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.

REFERENCES Adak, S., Crooks, C., Wang, Q., Crane, B. R., Tainer, J. A., Getzoff, E. D., and Stuehr, D. J. (1999). Tryptophan 409 controls the activity of neuronal nitric-oxide synthase by regulating nitric oxide feedback inhibition. J. Biol. Chem. 274, 26907–26911. Adler, V., Yin, Z., Fuchs, S. Y., Benezra, M., Rosario, L., Tew, K. D., Pincus, M. R., Sardana, M., Henderson, C. J., Wolf, C. R., et al. (1999). Regulation of JNK signaling by GSTp. EMBO J. 18, 1321–1334. Andreopoulos, S., and Papapetropoulos, A. (2000). Molecular aspects of soluble guanylyl cyclase regulation. Gen. Pharmacol. 34, 147–157. Andrew, P. J., and Mayer, B. (1999). Enzymatic function of nitric oxide synthases. Cardiovasc. Res. 43, 521–531. Archer, S. L., Huang, J. M., Hampl, V., Nelson, D. P., Shultz, P. J., and Weir, E. K. (1994). Nitric oxide and cGMP cause vasorelaxation by activation of a charybdotoxinsensitive K channel by cGMP-dependent protein kinase. Proc. Natl. Acad. Sci. USA, 91, 7583–7587. Asai, K., Kudej, R. K., Shen, Y. T., Yang, G. P., Takagi, G., Kudej, A. B., Geng, Y. J., Sato, N., Nazareno, J. B., Vatner, D. E., et al. (2000). Peripheral vascular endothelial dysfunction and apoptosis in old monkeys. Arterioscler. Thromb. Vasc. Biol. 20, 1493–1499. Boveris, A., and Cadenas, E. (2000). Mitochondrial production of hydrogen peroxide regulation by nitric oxide and the role of ubisemiquinone. IUBMB Life 50, 245–250. Brookes, P., and Darley-Usmar, V. M. (2002). Hypothesis: the mitochondrial NO(*) signaling pathway, and the transduction of nitrosative to oxidative cell signals: an alternative function for cytochrome c oxidase. Free Radic. Biol. Med. 32, 370–374. Brown, G. C., and Borutaite, V. (2002). Nitric oxide inhibition of mitochondrial respiration and its role in cell death. Free Radic. Biol. Med. 33, 1440–1450. Burney, S., Caulfield, J. L., Niles, J. C., Wishnok, J. S., and Tannenbaum, S. R. (1999). The chemistry of DNA damage from nitric oxide and peroxynitrite. Mutat. Res. 424, 37–49. Cadenas, E. (2004). Mitochondrial free radical production and cell signaling. Mol. Aspects Med. 25, 17–26. Castro, L., Rodriguez, M., and Radi, R. (1994). Aconitase is readily inactivated by peroxynitrite, but not by its precursor, nitric oxide. J. Biol. Chem. 269, 29409–29415. Ceneviva, G. D., Tzeng, E., Hoyt, D. G., Yee, E., Gallagher, A., Engelhardt, J. F., Kim, Y. M., Billiar, T. R., Watkins, S. A., and Pitt, B. R. (1998). Nitric oxide inhibits lipopolysaccharide-induced apoptosis in pulmonary artery endothelial cells. Am. J. Physiol. 275, L717–728. Dietrich, N., Thastrup, J., Holmberg, C., Gyrd-Hansen, M., Fehrenbacher, N., Lademann, U., Lerdrup, M., Herdegen, T., Jaattela, M., and Kallunki, T. (2004). JNK2 mediates TNF-induced cell death in mouse embryonic fibroblasts via regulation of both caspase and cathepsin protease pathways. Cell Death Differ. 11, 301–313.

240

Nitric Oxide, Cell Signaling, and Gene Expression

Dimmeler, S., Fisslthaler, B., Fleming, I., Hermann, C., Busse, R., and Zeiher, A. M. (1999). Activation of nitric oxide synthase in endothelial cells via Akt-dependent phosphorylation. Nature 399, 601–605. Dimmeler, S., Haendeler, J., Galle, J., and Zeiher, A. M. (1997a). Oxidized low density lipoprotein induces apoptosis of human endothelial cells by activation of CPP32like proteases: A mechanistic clue to the response to injury hypothesis. Circulation 95, 1760–1763. Dimmeler, S., Haendeler, J., Nehls, M., and Zeiher, A. M. (1997b). Suppression of apoptosis by nitric oxide via inhibition of ICE-like and CPP32-like proteases. J. Exp. Med. 185, 601–608. Dimmeler, S., Rippmann, V., Weiland, U., Haendeler, J., and Zeiher, A. M. (1997c). Angiotensin II induces apoptosis of human endothelial cells. Protective effect of nitric oxide. Circ. Res. 81, 970–976. Espey, M. G., Thomas, D. D., Miranda, K. M., and Wink, D. A. (2002). Focusing of nitric oxide mediated nitrosation and oxidative nitrosylation as a consequence of reaction with superoxide. Proc. Natl. Acad. Sci. USA 99, 11127–11132 Epub 12002 Aug 11112. Estevez, A. G., and Jordan, J. (2002). Nitric oxide and superoxide, a deadly cocktail. Ann. NY Acad. Sci. 962, 207–211. Eu, J. P., Sun, J., Xu, L., Stamler, J. S., and Meissner, G. (2000). The skeletal muscle calcium release channel: coupled O2 sensor and NO signaling functions. Cell 102, 499–509. Fiorucci, S., Mencarelli, A., Mannucci, R., Distrutti, E., Morelli, A., del Soldato, P., and Moncada, S. (2002). NCX-4016, a nitric oxide-releasing aspirin, protects endothelial cells against apoptosis by modulating mitochondrial function. FASEB J. 16, 1645–1647 Epub 2002 Aug 1647. Fujita, N., Manabe, H., Yoshida, N., Matsumoto, N., Ochiai, J., Masui, Y., Uemura, M., Naito, Y., and Yoshikawa, T. (2000). Inhibition of angiotensin-converting enzyme protects endothelial cell against hypoxia/reoxygenation injury. Biofactors 11, 257–266. Fulton, D., Gratton, J. P., McCabe, T. J., Fontana, J., Fujio, Y., Walsh, K., Franke, T. F., Papapetropoulos, A., and Sessa, W. C. (1999). Regulation of endothelium-derived nitric oxide production by the protein kinase Akt. Nature 399, 597–601. Haendeler, J., Hoffmann, J., Tischler, V., Berk, B. C., Zeiher, A. M., and Dimmeler, S. (2002). Redox regulatory and anti-apoptotic functions of thioredoxin depend on Snitrosylation at cysteine 69. Nat. Cell Biol. 4, 743–749. Haendeler, J., Weiland, U., Zeiher, A. M., and Dimmeler, S. (1997). Effects of redoxrelated congeners on apoptosis and caspase-3 activity. Nitric Oxide 1, 282–293. Harada-Shiba, M., Kinoshita, M., Kamido, H., and Shimokado, K. (1998). Oxidized low density lipoprotein induces apoptosis in cultured human umbilical vein endothelial cells by common and unique mechanisms. J. Biol. Chem. 273, 9681–9687. Hare, J. M., Keaney, J. F., Jr., Balligand, J. L., Loscalzo, J., Smith, T. W., and Colucci, W. S. (1995). Role of nitric oxide in parasympathetic modulation of beta-adrenergic myocardial contractility in normal dogs. J. Clin. Invest. 95, 360–366. Hengartner, M. O. (2000). The biochemistry of apoptosis. Nature 407, 770–776. Hoffmann, J., Dimmeler, S., and Haendeler, J. (2003). Shear stress increases the amount of S-nitrosylated molecules in endothelial cells: Important role for signal transduction. FEBS Lett. 551, 153–158.

Nitric Oxide and Caspase Activation

241

Hoffmann, J., Haendeler, J., Aicher, A., Rossig, L., Vasa, M., Zeiher, A. M., and Dimmeler, S. (2001). Aging enhances the sensitivity of endothelial cells toward apoptotic stimuli: important role of nitric oxide. Circ. Res. 89, 709–715. Huang, P. L., Huang, Z., Mashimo, H., Bloch, K. D., Moskowitz, M. A., Bevan, J. A., and Fishman, M. C. (1995). Hypertension in mice lacking the gene for endothelial nitric oxide synthase. Nature 377, 239–242. Ignarro, L. J. (1989). Endothelium-derived nitric oxide: actions and properties. FASEB J. 3, 31–36. Ignarro, L. J. (1990). Haem-dependent activation of guanylate cyclase and cyclic GMP formation by endogenous nitric oxide: a unique transduction mechanism for transcellular signaling. Pharmacol. Toxicol. 67, 1–7. Jourd’heuil, D., Jourd’heuil, F. L., and Feelisch, M. (2003). Oxidation and nitrosation of thiols at low micromolar exposure to nitric oxide. Evidence for a free radical mechanism. J. Biol. Chem. 278, 15720—15726, Epub 12003 Feb 15720. Keller, J. N., Kindy, M. S., Holtsberg, F. W., St. Clair, D. K., Yen, H. C., Germeyer, A., Steiner, S. M., Bruce-Keller, A. J., Hutchins, J. B., and Mattson, M. P. (1998). Mitochondrial manganese superoxide dismutase prevents neural apoptosis and reduces ischemic brain injury: suppression of peroxynitrite production, lipid peroxidation, and mitochondrial dysfunction. J. Neurosci. 18, 687–697. Kim, Y. M., Chung, H. T., Kim, S. S., Han, J. A., Yoo, Y. M., Kim, K. M., Lee, G. H., Yun, H. Y., Green, A., Li, J., et al. (1999). Nitric oxide protects PC12 cells from serum deprivation-induced apoptosis by cGMP-dependent inhibition of caspase signaling. J. Neurosci. 19, 6740–6747. Kim, Y.-M., de Vera, M. E., Watkins, S. C., and Billiar, T. R. (1997). Nitric oxide protects cultured rat hepatocytes from tumor necrosis factor-alpha-induced apoptosis by inducting heat shock protein 70 expression. J. Biol. Chem. 272, 1402–1411. Kim, Y. M., Kim, T. H., Chung, H. T., Talanian, R. V., Yin, X. M., and Billiar, T. R. (2000). Nitric oxide prevents tumor necrosis factor alpha-induced rat hepatocyte apoptosis by the interruption of mitochondrial apoptotic signaling through Snitrosylation of caspase-8. Hepatology 32, 770–778. Knudson, C. M., and Korsmeyer, S. J. (1997). Bcl-2 and Bax function independently to regulate cell death. Nat. Genet. 16, 358–363. Kontush, A., Chancharme, L., Escargueil-Blanc, I., Therond, P., Salvayre, R., NegreSalvayre, A., and Chapman, M. J. (2003). Mildly oxidized LDL particle subspecies are distinct in their capacity to induce apoptosis in endothelial cells: role of lipid hydroperoxides. FASEB J. 17, 88–90 Epub 2002 Nov 2015. Kröncke, K. D., Suschek, C. V., and Kolb-Bachofen, V. (2000). Implications of inducible nitric oxide synthase expression and enzyme activity. Antioxid. Redox. Signal. 2, 585–605. Lander, H. M., Hajjar, D. P., Hempstead, B. L., Mirza, U. A., Chait, B. T., Campbell, S., and Quilliam, L. A. (1997). A molecular redox switch on p21(ras). Structural basis for the nitric oxide-p21(ras) interaction. J. Biol. Chem. 272, 4323–4326. Lander, H. M., Ogiste, J. S., Pearce, S. F., Levi, R., and Novogrodsky, A. (1995). Nitric oxide-stimulated guanine nucleotide exchange on p21ras. J. Biol. Chem. 270, 7017–7020. Li, J., Billiar, T. R., Talanian, R. V., and Kim, Y. M. (1997). Nitric oxide reversibly inhibits seven members of the caspase family via S-nitrosylation. Biochem. Biophys. Res. Commun. 240, 419–424.

242

Nitric Oxide, Cell Signaling, and Gene Expression

Lipton, A. J., Johnson, M. A., Macdonald, T., Lieberman, M. W., Gozal, D., and Gaston, B. (2001). S-nitrosothiols signal the ventilatory response to hypoxia. Nature, 413, 171–174. Loke, K. E., Laycock, S. K., Mital, S., Wolin, M. S., Bernstein, R., Oz, M., Addonizio, L., Kaley, G., and Hintze, T. H. (1999). Nitric oxide modulates mitochondrial respiration in failing human heart. Circulation 100, 1291–1297. Mannick, J. B., Hausladen, A., Liu, L., Hess, D. T., Zeng, M., Miao, Q. X., Kane, L. S., Gow, A. J., and Stamler, J. S. (1999). Fas-induced caspase denitrosylation. Science, 284, 651–654. Messmer, U. K., Ankarcrona, M., Nicotera, P., and Brüne, B. (1994). p53 expression in nitric oxide-induced apoptosis. FEBS Lett. 355, 23–26. Miyashita, T., Krajewski, S., Krajewska, M., Wang, H. G., Lin, H. K., Liebermann, D. A., Hoffman, B., and Reed, J. C. (1994). Tumor suppressor p53 is a regulator of bcl2 and bax gene expression in vitro and in vivo. Oncogene 9, 1799–1805. Moroi, M., Zhang, L., Yasuda, T., Virmani, R., Gold, H. K., Fishman, M. C., and Huang, P. L. (1998). Interaction of genetic deficiency of endothelial nitric oxide, gender, and pregnancy in vascular responses to injury in mice. J. Clin. Invest. 101, 1225–1232. Murohara, T., Asahara, T., Silver, M., Bauters, C., Masuda, H., Kalka, C., Kearney, M., Chen, D., Symes, J. F., Fishman, M. C., et al. (1998). Nitric oxide synthase modulates angiogenesis in response to tissue ischemia. J. Clin. Invest. 101, 2567–2578. Nathan, C., and Xie, Q. W. (1994). Regulation of biosynthesis of nitric oxide. J. Biol. Chem. 269, 13725–13728. Nicholson, D. W. (1999). Caspase structure, proteolytic substrates, and function during apoptotic cell death. Cell Death Differ. 6, 1028–1042. Park, C., Kang, M., Kim-Kwon, Y., Kim, J., Ahn, H., and Huh, Y. (2002). Inhibition of neuronal nitric oxide synthase increases adrenalectomy-induced granule cell death in the rat dentate gyrus. Brain Res. 933, 81–84. Park, H. S., Huh, S. H., Kim, M. S., Lee, S. H., and Choi, E. J. (2000). Nitric oxide negatively regulates c-Jun N-terminal kinase/stress-activated protein kinase by means of S-nitrosylation. Proc. Natl. Acad. Sci. USA 97, 14022–14024. Pawloski, J. R., Hess, D. T., and Stamler, J. S. (2001). Export by red blood cells of nitric oxide bioactivity. Nature 409, 622–626. Poderoso, J. J., Lisdero, C., Schopfer, F., Riobo, N., Carreras, M. C., Cadenas, E., and Boveris, A. (1999). The regulation of mitochondrial oxygen uptake by redox reactions involving nitric oxide and ubiquinol. J. Biol. Chem. 274, 37709–37716. Rameau, G. A., Chiu, L. Y., and Ziff, E. B. (2003). NMDA receptor regulation of nNOS phosphorylation and induction of neuron death. Neurobiol. Aging 24, 11231133. Rössig, L., Fichtlscherer, B., Breitschopf, K., Haendeler, J., Zeiher, A. M., Mülsch, A., and Dimmeler, S. (1999). Nitric oxide inhibits caspase-3 by S-nitrosation in vivo. J. Biol. Chem. 274, 6823–6826. Sabourin, L. A., Tamai, K., Seale, P., Wagner, J., and Rudnicki, M. A. (2000). Caspase 3 cleavage of the Ste20-related kinase SLK releases and activates an apoptosisinducing kinase domain and an actin-disassembling region. Mol. Cell Biol. 20, 684–696. Sarti, P., Lendaro, E., Ippoliti, R., Bellelli, A., Benedetti, P. A., and Brunori, M. (1999). Modulation of mitochondrial respiration by nitric oxide: investigation by single cell fluorescence microscopy. FASEB J. 13, 191–197.

Nitric Oxide and Caspase Activation

243

Schachinger, V., Britten, M. B., and Zeiher, A. M. (2000). Prognostic impact of coronary vasodilator dysfunction on adverse long-term outcome of coronary heart disease. Circulation 101, 1899–1906. Shiva, S., Crawford, J. H., Ramachandran, A., Ceaser, E. K., Hillson, T., Brookes, P. S., Patel, R. P., and Darley-Usmar, V. M. (2004). Mechanisms of the interaction of nitroxyl with mitochondria. Biochem. J. 379, 359–366. Stamler, J. S., Lamas, S., and Fang, F. C. (2001). Nitrosylation. the prototypic redox-based signaling mechanism. Cell 106, 675–683. Stamler, J. S., Singel, D. J., and Loscalzo, J. (1992). Biochemistry of nitric oxide and its redox-activated forms [see comments]. Science 258, 1898–1902. Stumpe, T., Decking, U. K., and Schrader, J. (2001). Nitric oxide reduces energy supply by direct action on the respiratory chain in isolated cardiomyocytes. Am. J. Physiol. Heart Circ. Physiol. 280, H2350–2356. Sudoh, N., Toba, K., Akishita, M., Ako, J., Hashimoto, M., Iijima, K., Kim, S., Liang, Y. Q., Ohike, Y., Watanabe, T., et al. (2001). Estrogen prevents oxidative stressinduced endothelial cell apoptosis in rats. Circulation 103, 724–729. Sun, J., Xin, C., Eu, J. P., Stamler, J. S., and Meissner, G. (2001). Cysteine-3635 is responsible for skeletal muscle ryanodine receptor modulation by NO. Proc. Natl. Acad. Sci. USA 98, 11158–11162. Tschudi, M. R., Barton, M., Bersinger, N. A., Moreau, P., Cosentino, F., Noll, G., Malinski, T., and Luscher, T. F. (1996). Effect of age on kinetics of nitric oxide release in rat aorta and pulmonary artery. J. Clin. Invest. 98, 899–905. Valentino, K. L., Gutierrez, M., Sanchez, R., Winship, M. J., and Shapiro, D. A. (2003). First clinical trial of a novel caspase inhibitor: anti-apoptotic caspase inhibitor, IDN-6556, improves liver enzymes. Int. J. Clin. Pharmacol. Ther. 41, 441–449. Virag, L., Szabo, E., Gergely, P., and Szabo, C. (2003). Peroxynitrite-induced cytotoxicity: mechanism and opportunities for intervention. Toxicol. Lett. 140–141, 113–124. Welter, R., Yu, L., and Yu, C. A. (1996). The effects of nitric oxide on electron transport complexes. Arch. Biochem. Biophys. 331, 9–14. Wink, D. A., Kasprzak, K. S., Maragos, C. M., Elespuru, R. K., Misra, M., Dunams, T. M., Cebula, T. A., Koch, W. H., Andrews, A. W., Allen, J. S., et al. (1991). DNA deaminating ability and genotoxicity of nitric oxide and its progenitors. Science 254, 1001–1003. Wolin, M. S., Xie, Y. W., and Hintze, T. H. (1999). Nitric oxide as a regulator of tissue oxygen consumption. Curr. Opin. Nephrol. Hypertens. 8, 97–103. Yoo, J., and Fukuto, J. M. (1995). Oxidation of N-hydroxyguanidine by nitric oxide and the possible generation of vasoactive species. Biochem. Pharmacol. 50, 1995–2000. Zech, B., Wilm, M., van Eldik, R., and Brune, B. (1999). Mass spectrometric analysis of nitric oxide-modified caspase-3. J. Biol. Chem. 274, 20931–20936. Zeiher, A. M., Drexler, H., Saurbier, B., and Just, H. (1993). Endothelium-mediated coronary blood flow modulation in humans. Effects of age, atherosclerosis, hypercholesterolemia, and hypertension. J. Clin. Invest. 92, 652–662. Zhang, J., Dawson, V. L., Dawson, T. M., and Snyder, S. H. (1994). Nitric oxide activation of poly(ADP-ribose) synthetase in neurotoxicity. Science 263, 687–689.

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

246

Nitric Oxide, Cell Signaling, and Gene Expression

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

247

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

Signaling Effects of Peroxynitrite in Mammalian Cells

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

Signaling Effects of Peroxynitrite in Mammalian Cells

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.

252

Nitric Oxide, Cell Signaling, and Gene Expression

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:

Signaling Effects of Peroxynitrite in Mammalian Cells

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

Nitric Oxide, Cell Signaling, and Gene Expression

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].

Signaling Effects of Peroxynitrite in Mammalian Cells

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).

256

Nitric Oxide, Cell Signaling, and Gene Expression

REFERENCES 1. Radi, R., Peluffo, G., Alvarez, M. N., Naviliat, M., and Cayota, A. (2001) Unraveling peroxynitrite formation in biological systems. Free Radic. Biol. Med. 30, 463–488. 2. Pryor, W. A. and Squadrito, G. L. (1995) The chemistry of peroxynitrite: a product from the reaction of nitric oxide with superoxide. Am. J. Physiol. 268, L699–L722. 3. Marla, S. S., Lee, J., and Groves, J. T. (1997) Peroxynitrite rapidly permeates phospholipid membranes. Proc. Natl. Acad. Sci. USA 94, 14243–14248. 4. Denicola, A., Souza, J. M., and Radi, R. (1998) Diffusion of peroxynitrite across erythrocyte membranes. Proc. Natl. Acad. Sci. USA 95, 3566–3571. 5. Macfadyen, A. J., Reiter, C., Zhuang, Y., and Beckman, J. S. (1999) A novel superoxide dismutase-based trap for peroxynitrite used to detect entry of peroxynitrite into erythrocyte ghosts. Chem. Res. Toxicol. 12, 223–229. 6. Koppenol, W. H., Moreno, J. J., Pryor, W. A., Ischiropoulos, H., and Beckman, J. S. (1992) Peroxynitrite, a cloaked oxidant formed by nitric oxide and superoxide. Chem. Res. Toxicol. 5, 834–842. 7. Arteel, G. E., Briviba, K., and Sies, H. (1999) Protection against peroxynitrite. FEBS Lett. 445, 226–230. 8. Sies, H., Sharov, V. S., Klotz, L. O., and Briviba, K. (1997) Glutathione peroxidase protects against peroxynitrite-mediated oxidations. A new function for selenoproteins as peroxynitrite reductase. J. Biol. Chem. 272, 27812–27817. 9. Briviba, K., Klotz, L. O., and Sies, H. (1999) Defenses against peroxynitrite. Methods Enzymol. 301, 301–311. 10. Greenacre, S. A. B. and Ischiropoulos, H. (2001) Tyrosine nitration: localisation, quantification, consequences for protein function and signal transduction. Free Rad. Res. 34, 541–581. 11. Eiserich, J. P., Cross, C. E., Jones, A. D., Halliwell, B., and van der Vliet, A. (1996) Formation of nitrating and chlorinating species by reaction of nitrite with hypochlorous acid. A novel mechanism for nitric oxide-mediated protein modification. J. Biol. Chem. 271, 19199–19208. 12. Panasenko, O. M., Briviba, K., Klotz, L. O., and Sies, H. (1997) Oxidative modification and nitration of human low-density lipoproteins by the reaction of hypochlorous acid with nitrite. Arch. Biochem. Biophys. 343, 254–259. 13. Pfeiffer, S., Lass, A., Schmidt, K., and Mayer, B. (2001) Protein tyrosine nitration in cytokine-activated murine macrophages. Involvement of a peroxidase/nitrite pathway rather than peroxynitrite. J. Biol. Chem. 276, 34051–34058. 14. Kilinc, K., Kilinc, A., Wolf, R. E., and Grisham, M. B. (2001) Myoglobincatalyzed tyrosine nitration: no need for peroxynitrite. Biochem. Biophys. Res. Commun. 285, 273–276. 15. Robinson, K. M., Morre, J. T., and Beckman, J. S. (2004) Triuret: a novel product of peroxynitrite-mediated oxidation of urate. Arch. Biochem. Biophys. 423, 213–217. 16. Schroeder, P., Klotz, L. O., Buchczyk, D. P., Sadik, C. D., Schewe, T., and Sies, H. (2001) Epicatechin selectively prevents nitration but not oxidation reactions of peroxynitrite. Biochem. Biophys. Res. Commun. 285, 782–787. 17. Schroeder, P., Zhang, H., Klotz, L. O., Kalyanaraman, B., and Sies, H. (2001) (−)Epicatechin inhibits nitration and dimerization of tyrosine in hydrophilic as well as hydrophobic environments. Biochem. Biophys. Res. Commun. 289, 1334–1338.

Signaling Effects of Peroxynitrite in Mammalian Cells

257

18. Tibi, S. and Koppenol, W. H. (2000) Reactions of peroxynitrite with phenolic and carbonyl compounds: flavonoids are not scavengers of peroxynitrite. Helv. Chim. Acta 83, 2412–2424. 19. Teng, R. J., Ye, Y. Z., Parks, D. A., and Beckman, J. S. (2002) Urate produced during hypoxia protects heart proteins from peroxynitrite-mediated protein nitration. Free Radic. Biol. Med. 33, 1243–1249. 20. Uchida, K. and Kumagai, T. (2003) 4-Hydroxy-2-nonenal as a COX-2 inducer. Mol. Aspects Med. 24, 213–218. 21. Awasthi, Y. C., Sharma, R., Cheng, J. Z., Yang, Y., Sharma, A., Singhal, S. S., and Awasthi, S. (2003) Role of 4-hydroxynonenal in stress-mediated apoptosis signaling. Mol. Aspects Med. 24, 219–230. 22. Folden, D. V., Gupta, A., Sharma, A. C., Li, S. Y., Saari, J. T., and Ren, J. (2003) Malondialdehyde inhibits cardiac contractile function in ventricular myocytes via a p38 mitogen-activated protein kinase-dependent mechanism. Br. J. Pharmacol. 139, 1310–1316. 23. Takeuchi, K., Kato, M., Suzuki, H., Akhand, A. A., Wu, J., Hossain, K., Miyata, T., Matsumoto, Y., Nimura, Y., and Nakashima, I. (2001) Acrolein induces activation of the epidermal growth factor receptor of human keratinocytes for cell death. J. Cell Biochem. 81, 679–688. 24. Aust, A. E. and Eveleigh, J. F. (1999) Mechanisms of DNA oxidation. Proc. Soc. Exp. Biol. Med. 222, 246–252. 25. Dedon, P. C. and Tannenbaum, S. R. (2004) Reactive nitrogen species in the chemical biology of inflammation. Arch. Biochem. Biophys. 423, 12–22. 26. Rouse, J. and Jackson, S. P. (2002) Interfaces between the detection, signaling, and repair of DNA damage. Science 297, 547–551. 27. Buchczyk, D. P., Grune, T., Sies, H., and Klotz, L. O. (2003) Modifications of glyceraldehyde-3-phosphate dehydrogenase induced by increasing concentrations of peroxynitrite: early recognition by 20S proteasome. Biol. Chem. 384, 237–241. 28. Grune, T., Klotz, L. O., Gieche, J., Rudeck, M., and Sies, H. (2001) Protein oxidation and proteolysis by the nonradical oxidants singlet oxygen or peroxynitrite. Free Radic. Biol. Med. 30, 1243–1253. 29. Beckman, J. S., Ye, Y. Z., Anderson, P. G., Chen, J., Accavitti, M. A., Tarpey, M. M., and White, C. R. (1994) Extensive nitration of protein tyrosines in human atherosclerosis detected by immunohistochemistry. Biol. Chem. 375, 81–88. 30. Kong, S. K., Yim, M. B., Stadtman, E. R., and Chock, P. B. (1996) Peroxynitrite disables the tyrosine phosphorylation regulatory mechanism: Lymphocyte-specific tyrosine kinase fails to phosphorylate nitrated cdc2(6-20)NH2 peptide. Proc. Natl. Acad. Sci. USA 93, 3377–3382. 31. Brito, C., Naviliat, M., Tiscornia, A. C., Vuillier, F., Gualco, G., Dighiero, G., Radi, R., and Cayota, A. M. (1999) Peroxynitrite inhibits T lymphocyte activation and proliferation by promoting impairment of tyrosine phosphorylation and peroxynitrite-driven apoptotic death. J. Immunol. 162, 3356–3366. 32. Mallozzi, C., Di Stasi, A. M., and Minetti, M. (1997) Peroxynitrite modulates tyrosine-dependent signal transduction pathway of human erythrocyte band 3. FASEB J. 11, 1281–1290. 33. Reinehr, R., Görg, B., Höngen, A., and Häussinger, D. (2004) CD95-tyrosine nitration inhibits hyperosmotic and CD95 ligand-induced CD95 activation in rat hepatocytes. J. Biol. Chem. 279, 10364–10373.

258

Nitric Oxide, Cell Signaling, and Gene Expression

34. MacMillan-Crow, L. A., Greendorfer, J. S., Vickers, S. M., and Thompson, J. A. (2000) Tyrosine nitration of c-SRC tyrosine kinase in human pancreatic ductal adenocarcinoma. Arch. Biochem. Biophys. 377, 350–356. 35. Mallozzi, C., Di Stasi, A. M., and Minetti, M. (2001) Nitrotyrosine mimics phosphotyrosine binding to the SH2 domain of the src family tyrosine kinase lyn. FEBS Lett. 503, 189–195. 36. Fauman, E. B. and Saper, M. A. (1996) Structure and function of the protein tyrosine phosphatases. Trends. Biochem. Sci. 21, 413–417. 37. Kolmodin, K. and Aqvist, J. (2001) The catalytic mechanism of protein tyrosine phosphatases revisited. FEBS Lett. 498, 208–213. 38. Abdelmohsen, K., Gerber, P. A., von Montfort, C., Sies, H., and Klotz, L. O. (2003) Epidermal growth factor receptor is a common mediator of quinoneinduced signaling leading to phosphorylation of connexin-43: role of glutathione and tyrosine phosphatases. J. Biol. Chem. 278, 38360–38367. 39. Abdelmohsen, K., Patak, P., von Montfort, C., Melchheier, I., Sies, H., and Klotz, L. O. (2004) Signaling effects of menadione: from tyrosine phosphatase inactivation to connexin phosphorylation. Methods Enzymol. 378, 258–272. 40. Quijano, C., Alvarez, B., Gatti, R. M., Augusto, O., and Radi, R. (1997) Pathways of peroxynitrite oxidation of thiol groups. Biochem. J. 322, 167–173. 41. Takakura, K., Beckman, J. S., MacMillan-Crow, L. A., and Crow, J. P. (1999) Rapid and irreversible inactivation of protein tyrosine phosphatases PTP1B, CD45, and LAR by peroxynitrite. Arch. Biochem. Biophys. 369, 197–207. 42. Salmeen, A., Andersen, J. N., Myers, M. P., Meng, T. C., Hinks, J. A., Tonks, N. K., and Barford, D. (2003) Redox regulation of protein tyrosine phosphatase 1B involves a sulphenyl-amide intermediate. Nature 423, 769–773. 43. van Montfort, R. L., Congreve, M., Tisi, D., Carr, R., and Jhoti, H. (2003) Oxidation state of the active-site cysteine in protein tyrosine phosphatase 1B. Nature 423, 773–777. 44. Klotz, L. O., Schieke, S. M., Sies, H., and Holbrook, N. J. (2000) Peroxynitrite activates the phosphoinositide 3-kinase/Akt pathway in human skin primary fibroblasts. Biochem. J. 352, 219–225. 45. Mallozzi, C., Di Stasi, M. A., and Minetti, M. (2001) Peroxynitrite-dependent activation of src tyrosine kinases lyn and hck in erythrocytes is under mechanistically different pathways of redox control. Free Radic. Biol. Med. 30, 1108–1117. 46. Antunes, F. and Cadenas, E. (2001) 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. 47. Karin, M., Liu, Z., and Zandi, E. (1997) AP-1 function and regulation. Curr. Opin. Cell Biol. 9, 240–246. 48. Whitmarsh, A. J. and Davis, R. J. (1996) Transcription factor AP-1 regulation by mitogen-activated protein kinase signal transduction pathways. J. Mol. Med. 74, 589–607. 49. Klotz, L. O., Buchczyk, D., and Sies, H. (2001) MAP kinase activation and environmental stress. In: Micronutrients and Health: Molecular Biological Mechanisms (Nesaretnam, K. and Packer, L., Eds.), pp. 184–193, AOCS Press, Champaign, IL.

Signaling Effects of Peroxynitrite in Mammalian Cells

259

50. Chen, Z., Gibson, T. B., Robinson, F., Silvestro, L., Pearson, G., Xu, B., Wright, A., Vanderbilt, C., and Cobb, M. H. (2001) MAP kinases. Chem. Rev. 101, 2449–2476. 51. Ono, K. and Han, J. (2000) The p38 signal transduction pathway: activation and function. Cell Signal. 12, 1–13. 52. Gupta, S., Barrett, T., Whitmarsh, A. J., Cavanagh, J., Sluss, H. K., Derijard, B., and Davis, R. J. (1996) Selective interaction of JNK protein kinase isoforms with transcription factors. EMBO J. 15, 2760–2770. 53. Schieke, S. M., Briviba, K., Klotz, L. O., and Sies, H. (1999) Activation pattern of mitogen-activated protein kinases elicited by peroxynitrite: attenuation by selenite supplementation. FEBS Lett. 448, 301–303. 54. Bapat, S., Verkleij, A., and Post, J. A. (2001) Peroxynitrite activates mitogenactivated protein kinase (MAPK) via a MEK-independent pathway: a role for protein kinase C. FEBS Lett. 499, 21–26. 55. Go, Y. M., Patel, R. P., Maland, M. C., Park, H., Beckman, J. S., Darley-Usmar, V. M., and Jo, H. (1999) Evidence for peroxynitrite as a signaling molecule in flow-dependent activation of c-Jun NH2-terminal kinase. Am. J. Physiol. 277, H1647–H1653. 56. Oh-hashi, K., Maruyama, W., Yi, H., Takahashi, T., Naoi, M., and Isobe, K. (1999) Mitogen-activated protein kinase pathway mediates peroxynitrite-induced apoptosis in human dopaminergic neuroblastoma SH-SY5Y cells. Biochem. Biophys. Res. Commun. 263, 504–509. 57. Lee, C., Miura, K., Liu, X., and Zweier, J. L. (2000) Biphasic regulation of leukocyte superoxide generation by nitric oxide and peroxynitrite. J. Biol. Chem. 275, 38965–38972. 58. Zouki, C., Zhang, S. L., Chan, J. S., and Filep, J. G. (2001) Peroxynitrite induces integrin-dependent adhesion of human neutrophils to endothelial cells via activation of the Raf-1/MEK/Erk pathway. FASEB J. 15, 25–27. 59. Müller, T., Haussmann, H. J., and Schepers, G. (1997) Evidence for peroxynitrite as an oxidative stress-inducing compound of aqueous cigarette smoke fractions. Carcinogenesis 18, 295–301. 60. Foresti, R., Sarathchandra, P., Clark, J. E., Green, C. J., and Motterlini, R. (1999) Peroxynitrite induces haem oxygenase-1 in vascular endothelial cells: a link to apoptosis. Biochem. J. 339, 729–736. 61. Oh-hashi, K., Maruyama, W., and Isobe, K. (2001) Peroxynitrite induces GADD34, 45, and 153 via p38 MAPK in human neuroblastoma SH-SY5Y cells. Free Radic. Biol. Med. 30, 213–221. 62. Zhang, P., Wang, Y. Z., Kagan, E., and Bonner, J. C. (2000) Peroxynitrite targets the epidermal growth factor receptor, Raf-1, and MEK independently to activate MAPK. J. Biol. Chem. 275, 22479–22486. 63. Jope, R. S., Zhang, L., and Song, L. (2000) Peroxynitrite modulates the activation of p38 and extracellular regulated kinases in PC12 cells. Arch. Biochem. Biophys. 376, 365–370. 64. van der Vliet, A., Hristova, M., Cross, C. E., Eiserich, J. P., and Goldkorn, T. (1998) Peroxynitrite induces covalent dimerization of epidermal growth factor receptors in A431 epidermoid carcinoma cells. J. Biol. Chem. 273, 31860–31866.

260

Nitric Oxide, Cell Signaling, and Gene Expression

65. Dewas, C., Fay, M., Gougerot-Pocidalo, M.A., and El Benna, J. (2000) The mitogen-activated protein kinase extracellular signal-regulated kinase 1/2 pathway is involved in formyl-methionyl-leucyl-phenylalanine-induced p47phox phosphorylation in human neutrophils. J. Immunol. 165, 5238–5244. 66. Lampe, P. D. and Lau, A. F. (2000) Regulation of gap junctions by phosphorylation of connexins. Arch. Biochem. Biophys. 384, 205–215. 67. Sharov, V. S., Briviba, K., and Sies, H. (1999) Peroxynitrite diminishes gap junctional communication: protection by selenite supplementation. IUBMB Life 48, 379–384. 68. Datta, S. R., Brunet, A., and Greenberg, M. E. (1999) Cellular survival: a play in three Akts. Genes Dev. 13, 2905–2927. 69. Scheid, M. P. and Woodgett, J. R. (2001) Pkb/akt: functional insights from genetic models. Nat. Rev. Mol. Cell Biol. 2, 760–768. 70. Zwick, E., Hackel, P. O., Prenzel, N., and Ullrich, A. (1999) The EGF receptor as central transducer of heterologous signalling systems. Trends Pharmacol. Sci. 20, 408–412. 71. Chen, K., Vita, J. A., Berk, B. C., and Keaney, J. F., Jr. (2001) c-Jun N-terminal kinase activation by hydrogen peroxide in endothelial cells involves SRC-dependent epidermal growth factor receptor transactivation. J. Biol. Chem. 276, 16045–16050. 72. Wang, X., McCullough, K. D., Franke, T. F., and Holbrook, N. J. (2000) Epidermal growth factor receptor-dependent Akt activation by oxidative stress enhances cell survival. J. Biol. Chem. 275, 14624–14631. 73. Ostrakhovitch, E. A., Lordnejad, M. R., Schliess, F., Sies, H., and Klotz, L. O. (2002) Copper ions strongly activate the phosphoinositide-3-kinase/Akt pathway independent of the generation of reactive oxygen species. Arch. Biochem. Biophys. 397, 232–239. 74. Mallozzi, C., Di Stasi, A. M., and Minetti, M. (1999) Activation of src tyrosine kinases by peroxynitrite. FEBS Lett. 456, 201–206. 75. Zou, M. H., Hou, X. Y., Shi, C. M., Nagata, D., Walsh, K., and Cohen, R. A. (2002) Modulation by peroxynitrite of Akt- and AMP-activated kinase-dependent Ser1179 phosphorylation of endothelial nitric oxide synthase. J. Biol. Chem. 277, 32552–32557. 76. Zou, M. H., Hou, X. Y., Shi, C. M., Kirkpatick, S., Liu, F., Goldman, M. H., and Cohen, R. A. (2003) Activation of 5'-AMP-activated kinase is mediated through c-Src and phosphoinositide 3-kinase activity during hypoxia-reoxygenation of bovine aortic endothelial cells. Role of peroxynitrite. J. Biol. Chem. 278, 34003–34010. 77. Carling, D. (2004) The AMP-activated protein kinase cascade—a unifying system for energy control. Trends Biochem. Sci. 29, 18–24. 78. Buckley, B. J. and Whorton, A. R. (2000) Adaptive responses to peroxynitrite: increased glutathione levels and cystine uptake in vascular cells. Am. J. Physiol. Cell Physiol. 279, C1168–C1176. 79. Vallette, G., Tenaud, I., Branka, J. E., Jarry, A., Sainte-Marie, I., Dreno, B., and Laboisse, C. L. (1998) Control of growth and differentiation of normal human epithelial cells through the manipulation of reactive nitrogen species. Biochem. J. 331, 713–717.

Signaling Effects of Peroxynitrite in Mammalian Cells

261

80. Cobbs, C. S., Samanta, M., Harkins, L. E., Gillespie, G. Y., Merrick, B. A., and MacMillan-Crow, L. A. (2001) Evidence for peroxynitrite-mediated modifications to p53 in human gliomas: possible functional consequences. Arch. Biochem. Biophys. 394, 167–172. 81. Cobbs, C. S., Whisenhunt, T. R., Wesemann, D. R., Harkins, L. E., Van Meir, E. G., and Samanta, M. (2003) Inactivation of wild-type p53 protein function by reactive oxygen and nitrogen species in malignant glioma cells. Cancer Res. 63, 8670–8673. 82. Kröncke, K. D., Klotz, L. O., Suschek, C. V., and Sies, H. (2002) Comparing nitrosative versus oxidative stress toward zinc finger-dependent transcription. Unique role for NO. J. Biol. Chem. 277, 13294–13301. 83. Virag, L., Scott, G. S., Cuzzocrea, S., Marmer, D., Salzman, A. L., and Szabo, C. (1998) Peroxynitrite-induced thymocyte apoptosis: the role of caspases and poly (ADP-ribose) synthetase (PARS) activation. Immunology 94, 345–355. 84. Lin, K. T., Xue, J. Y., and Wong, P. Y. (1999) Mechanisms of peroxynitrite-induced apoptosis in HL-60 cells. Adv. Exp. Med. Biol. 469, 569–575. 85. Zhuang, S. and Simon, G. (2000) Peroxynitrite-induced apoptosis involves activation of multiple caspases in HL-60 cells. Am. J. Physiol. Cell Physiol. 279, C341–C351. 86. Xia, Z., Dickens, M., Raingeaud, J., Davis, R. J., and Greenberg, M. E. (1995) Opposing effects of ERK and JNK-p38 MAP kinases on apoptosis. Science 270, 1326–1331. 87. Assefa, Z., Vantieghem, A., Declercq, W., Vandenabeele, P., Vandenheede, J. R., Merlevede, W., de Witte, P., and Agostinis, P. (1999) The activation of the c-Jun N-terminal kinase and p38 mitogen-activated protein kinase signaling pathways protects HeLa cells from apoptosis following photodynamic therapy with hypericin. J. Biol. Chem. 274, 8788–8796. 88. Wang, X., Martindale, J. L., and Holbrook, N. J. (2000) Requirement for ERK activation in cisplatin-induced apoptosis. J. Biol. Chem. 275, 39435–39443. 89. Klotz, L. O. (2002) Oxidant-induced signaling: effects of peroxynitrite and singlet oxygen. Biol. Chem. 383, 443–456.

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 267

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 271

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

Ras S-nitrosylation (mole)

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

Ras S-nitrosylation (mole)

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

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 275

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 (<0.05–0.20 ppm) (>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

Nitric Oxide, Cell Signaling, and Gene Expression

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).

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 279

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

Nitric Oxide, Cell Signaling, and Gene Expression

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• −-

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 281

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

Nitric Oxide, Cell Signaling, and Gene Expression

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.

284

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 285

REFERENCES 1. Oxford, G., and Theodorescu, D. (2003) Ras superfamily monomeric G proteins in carcinoma cell motility, Cancer Lett. 189, 117–128. 2. Paduch, M., Jelen, F., and Otlewski, J. (2001) Structure of small G proteins and their regulators, Acta Biochim. Pol. 48, 829–850. 3. Corbett, K. D., and Alber, T. (2001) The many faces of Ras: recognition of small GTP-binding proteins, Trends Biochem. Sci. 26, 710–716. 4. Reuther, G. W., and Der, C. J. (2000) The Ras branch of small GTPases: Ras family members don’t fall far from the tree, Curr. Opin. Cell Biol. 12, 157–165. 5. Malliri, A., and Collard, J. G. (2003) Role of Rho-family proteins in cell adhesion and cancer, Curr. Opin. Cell Biol. 15, 583–589. 6. Takai, Y., Sasaki, T., and Matozaki, T. (2001) Small GTP-binding proteins, Physiol. Rev. 81, 153–208. 7. Shields, J. M., Pruitt, K., McFall, A., Shaub, A., and Der, C. J. (2000) Understanding Ras: “it ain’t over ‘til it’s over”, Trends Cell Biol. 10, 147–154. 8. Campbell, S. L., Khosravi-Far, R., Rossman, K. L., Clark, G. J., and Der, C. J. (1998) Increasing complexity of Ras signaling, Oncogene 17, 1395–1413. 9. Karnoub, A. E., Symons, M., Campbell, S. L., and Der, C. J. (2004) Molecular basis for Rho GTPase signaling specificity, Breast Cancer Res. Treat. 84, 61–71. 10. Sprang, S. (2001) GEFs: master regulators of G-protein activation, Trends Biochem. Sci. 26, 266–267. 11. Sprang, S. R. (1997) G proteins, effectors and GAPs: structure and mechanism, Curr. Opin. Struct. Biol. 7, 849–856. 12. Geyer, M., and Wittinghofer, A. (1997) GEFs, GAPs, GDIs and effectors: taking a closer (3D) look at the regulation of Ras-related GTP-binding proteins, Curr. Opin. Struct. Biol. 7, 786–792. 13. Lander, H. M. (1997) An essential role for free radicals and derived species in signal transduction, FASEB J. 11, 118–124. 14. Irani, K., and Goldschmidt-Clermont, P. (1998) Ras, superoxide and signal transduction, Biochem. Pharmacol. 55, 1339–1346. 15. Moon, S. Y., and Zheng, Y. (2003) Rho GTPase-activating proteins in cell regulation, Trends Cell Biol. 13, 13–22. 16. Pfeffer, S. R. (2001) Rab GTPases: specifying and deciphering organelle identity and function, Trends Cell Biol. 11, 487–491. 17. Olofsson, B. (1999) Rho guanine dissociation inhibitors: pivotal molecules in cellular signalling, Cell Signal. 11, 545–554. 18. Heo, J., and Campbell, S. L. (2004) Mechanism of p21Ras S-nitrosylation and kinetics of nitric oxide-mediated guanine nucleotide exchange, Biochemistry 43, 2314–2322. 19. Schrammel, A., Gorren, A., Schmidt, K., Pfeiffer, S., and Mayer, B. (2003) Snitrosation of glutathione by nitric oxide, peroxynitrite, and ΝO/O2 Free Radic. Biol. Med. 34, 1078–88. 20. Jourd’heuil, D., Jourd’heuil, F., and Feelisch, M. (2003) Oxidation and nitrosation of thiols at low micromolar exposure to nitric oxide. Evidence for a free radical mechanism, J. Biol. Chem. 278, 15720–6. 21. Ford, E., Hughes, M., and Wardman, P. (2002) Kinetics of the reactions of nitrogen dioxide with glutathione, cysteine, and uric acid at physiological pH, Free Radic. Biol. Med. 32, 1314–23.

286

Nitric Oxide, Cell Signaling, and Gene Expression

22. Heo, J., Prutzman, K. C., Mocanu, V., and Campbell, S. L. (2005) Mechanism of free radical nitric oxide-mediated Ras guanine nucleotide dissociation, J. Mol. Biol., 346, 1423–1440. 23. Forstermann, U., and Kleinert, H. (1995) Nitric oxide synthase: expression and expressional control of the three isoforms, Naunyn Schmiedebergs Arch. Pharmacol. 352, 351–364. 24. Khan, A. U., and Kasha, M. (1994) Singlet molecular oxygen in the Harber–Weiss reaction, Proc. Natl. Acad. Sci. USA 91, 12365–12367. 25. Stubbe, J., and van der Donk, W. A. (1998) Protein radicals in enzyme catalysis, Chem. Rev. 98, 705–762. 26. Weast, R. C. (1982) CRC Handbook of Chemistry and Physics, 63rd ed., CRC Press, Boca Raton, FL. 27. Quimby, B. B., and Dasso, M. (2003) The small GTPase Ran: interpreting the signs, Curr. Opin. Cell. Biol. 15, 338–344. 28. Nie, Z., Hirsch, D. S., and Randazzo, P. A. (2003) Arf and its many interactors, Curr. Opin. Cell Biol. 15, 396–404. 29. Spang, A. (2002) ARF1 regulatory factors and COPI vesicle formation, Curr. Opin. Cell Biol. 14, 423–427. 30. Yamada, M., Tachibana, T., Imamoto, N., and Yoneda, Y. (1998) Nuclear transport factor p10/NTF2 functions as a Ran-GDP dissociation inhibitor (Ran-GDI), Curr. Biol. 8, 1339–1342. 31. Dawson, T. M., Sasaki, M., Gonzalez-Zulueta, M., and Dawson, V. L. (1998) Regulation of neuronal nitric oxide synthase and identification of novel nitric oxide signaling pathways, Prog. Brain Res. 118, 3–11. 32. Lander, H. M., Hajjar, D. P., Hempstead, B. L., Mirza, U. A., Chait, B. T., Campbell, S., and Quilliam, L. A. (1997) A molecular redox switch on p21(ras). Structural basis for the nitric oxide-p21(ras) interaction, J. Biol. Chem. 272, 4323–4326. 33. Lander, H. M., Ogiste, J. S., Teng, K. K., and Novogrodsky, A. (1995) p21ras as a common signaling target of reactive free radicals and cellular redox stress, J. Biol. Chem. 270, 21195–21198. 34. Lander, H. M., Sehajpal, P. K., and Novogrodsky, A. (1993) Nitric oxide signaling: a possible role for g proteins, Am. Assoc. Immunol. 151, 7182–7187. 35. Kinbara, K., Goldfinger, L. E., Hansen, M., Chou, F. L., and Ginsberg, M. H. (2003) Ras GTPases: integrins’ friends or foes? Nat. Rev. Mol. Cell Biol. 4, 767–776. 36. Nassar, N., Horn, G., Herrmann, C., Block, C., Janknecht, R., and Wittinghofer, A. (1996) Ras/Rap effector specificity determined by charge reversal, Nat. Struct. Biol. 3, 723–729. 37. Cox, A. D., and Der, C. J. (2003) The dark side of Ras: regulation of apoptosis, Oncogene 22, 8999–9006. 38. Stork, P. J. (2003) Does Rap1 deserve a bad Rap? Trends Biochem. Sci. 28, 267–275. 39. Stenmark, H., and Olkkonen, V. M. (2001) The Rab GTPase family, Genome. Biol. 2, reviews 3007.1–3007.7. 40. Olkkonen, V. M., and Stenmark, H. (1997) Role of Rab GTPases in membrane traffic, Int. Rev. Cytol. 176, 1–85. 41. Darchen, F., and Goud, B. (2000) Multiple aspects of Rab protein action in the secretory pathway: focus on Rab3 and Rab6, Biochimie 82, 375–384.

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 287 42. Lipschutz, J. H., and Mostov, K. E. (2002) Exocytosis: the many masters of the exocyst, Curr. Biol. 12, R212– R214. 43. Adachi, T., Pimentel, D. R., Heibeck, T., Hou, X., Lee, Y. J., Jiang, B., Ido, Y., and Cohen, R. A. (2004) S-glutathiolation of Ras mediates redox-sensitive signaling by angiotensin II in vascular smooth muscle cells, J. Biol. Chem. 279, 29857– 29862. 44. Baker, T., Booden, M., and Buss, J. (2000) S-Nitrosocysteine increases palmitate turnover on Ha-Ras in NIH 3T3 cells, J. Biol. Chem. 275, 22037–22047. 45. Lander, H. M., Hajjar, D. P., Hempstead, B. L., Mirza, U. A., Chait, B. T., Campbell, S. L., and Quilliam, L. A. (1997) A molecular redox switch on p21ras. Structural basis for the nitric oxide-p21ras interaction, J. Biol. Chem. 272, 4323–4326. 46. Lander, H. M., Milbank, A. J., Tauras, J. M., Hajjar, D. P., Hempstead, B. L., Schwartz, G. D., Kraemer, R. T., Mirza, U. A., Chait, B. T., Burk, S. C., and Quilliam, L. A. (1996) Redox regulation of cell signalling, Nature 381, 380–381. 47. Lander, H. M., Ogiste, J. S., Pearce, S. F., Levi, R., and Novogrodsky, A. (1995) Nitric oxide-stimulated guanine nucleotide exchange on p21ras, J. Biol. Chem. 270, 7017–7020. 48. Ridley, A. J. (2004) Rho proteins and cancer, Breast Cancer Res. Treat. 84, 13–19. 49. Burridge, K., and Wennerberg, K. (2004) Rho and Rac take center stage, Cell 116, 167–179. 50. Aznar, S., and Lacal, J. C. (2001) Rho signals to cell growth and apoptosis, Cancer Lett. 165, 1–10. 51. Kaibuchi, K., Kuroda, S., and Amano, M. (1999) Regulation of the cytoskeleton and cell adhesion by the Rho family GTPases in mammalian cells, Annu. Rev. Biochem. 68, 459–486. 52. Takeya, R., and Sumimoto, H. (2003) Molecular mechanism for activation of superoxide-producing NADPH oxidases, Mol. Cells 16, 271– 277. 53. Bokoch, G. M., and Knaus, U. G. (2003) NADPH oxidases: not just for leukocytes anymore! Trends Biochem. Sci. 28, 502–508. 54. Bokoch, G. M. (2000) Regulation of cell function by Rho family GTPases, Immunol. Res. 21, 139–148. 55. Evers, E. E., Zondag, G. C., Malliri, A., Price, L. S., ten Klooster, J. P., van der Kammen, R. A., and Collard, J. G. (2000) Rho family proteins in cell adhesion and cell migration, Eur. J. Cancer 36, 1269–1274. 56. Scita, G., Tenca, P., Frittoli, E., Tocchetti, A., Innocenti, M., Giardina, G., and Di Fiore, P. P. (2000) Signaling from Ras to Rac and beyond: not just a matter of GEFs, EMBO. J. 19, 2393–2398. 57. Raftopoulou, M., and Hall, A. (2004) Cell migration: Rho GTPases lead the way, Developmental Biology 265, 23–32. 58. Stamler, J. S., Jaraki, O., Osborne, J., Simon, D. I., Keaney, J., Vita, J., Singel, D., Valeri, C. R., and Loscalzo, J. (1992) Nitric oxide circulates in mammalian plasma primarily as an S-nitroso adduct of serum albumin, Proc. Natl. Acad. Sci. USA 89, 7674–7677. 59. Sheu, F., Zhu, W., and Fung, P. (2000) Direct observation of trapping and release of nitric oxide by glutathione and cysteine with electron paramagnetic resonance spectroscopy, Biophys. J. 78, 1216–1226. 60. Gow, A. J., Buerk, D. G., and Ischiropoulos, H. (1997) A novel reaction mechanism for the formation of S-nitrosothiol in vivo, J. Biol. Chem. 272, 2841–2845.

288

Nitric Oxide, Cell Signaling, and Gene Expression

61. Heo, J., Gao, G., and Campbell, S. (2004) pH-dependent perturbation of Ras guanine-nucleotide interactions and Ras-guanine-nucleotide exchange, Biochemistry 43, 10102–10111. 62. Reinstein, J., Schlichting, I., Frech, M., Goody, R. S., and Wittinghofer, A. (1991) p21 with a phenylalanine 28—leucine mutation reacts normally with the GTPase activating protein GAP but nevertheless has transforming properties, J. Biol. Chem. 266, 17700–17706. 63. Schlichting, I., John, J., Frech, M., Chardin, P., Wittinghofer, A., Zimmermann, H., and Rosch, P. (1990) Proton NMR studies of transforming and nontransforming H-ras p21 mutants, Biochemistry 29, 504–511. 64. Page, C. C., Moser, C. C., Chen, X., and Dutton, P. L. (1999) Natural engineering principles of electron tunnelling in biological oxidation-reduction, Nature 402, 47–52. 65. Arkin, M. R., Stemp, E. D., Pulver, S. C., and Barton, J. K. (1997) Long-range oxidation of guanine by Ru(III) in duplex DNA, Chem. Biol. 4, 389–400. 66. Niles, J. C., Wishnok, J. S., and Tannenbaum, S. R. (2001) A novel nitroimidazole compound formed during the reaction of peroxynitrite with 2′,3′,5′-tri-O-acetylguanosine, J. Am. Chem. Soc. 123, 12147–12151. 67. Wagenknecht, H. A., Rajski, S. R., Pascaly, M., Stemp, E. D., and Barton, J. K. (2001) Direct observation of radical intermediates in protein-dependent DNA charge transport, J. Am. Chem. Soc. 123, 4400–4407. 68. Gu, F., Stillwell, W. G., Wishnok, J. S., Shallop, A. J., Jones, R. A., and Tannenbaum, S. R. (2002) Peroxynitrite-induced reactions of synthetic oligo 2′-deoxynucleotides and DNA containing guanine: formation and stability of a 5-guanidino-4-nitroimidazole lesion, Biochemistry 41, 7508–7518. 69. Joffe, A., Mock, S., Yun, B. H., Kolbanovskiy, A., Geacintov, N. E., and Shafirovich, V. (2003) Oxidative generation of guanine radicals by carbonate radicals and their reactions with nitrogen dioxide to form site specific 5-guanidino-4-nitroimidazole lesions in oligodeoxynucleotides, Chem. Res. Toxicol. 16, 966–973. 70. Kobayashi, K., and Tagawa, S. (2003) Direct observation of guanine radical cation deprotonation in duplex DNA using pulse radiolysis, J. Am. Chem. Soc. 125, 10213–10218. 71. Heo, J., and Campbell, S.L. (2005) Superoxide anion radical modulates the activity of ras and ras-related GTPases by a radical-base mechanism similar to that of nitric oxide, J. Biol. Chem. 280, 12438–12445. 72. Harrison, R. (2002) Structure and function of xanthine oxidoreductase: where are we now? Free Radic. Biol. Med. 33, 774–797. 73. Hirshberg, M., Stockley, R. W., Dodson, G., and Webb, M. R. (1997) The crystal structure of human rac1, a member of the rho-family complexed with a GTP analogue, Nat. Struct. Biol. 4, 147–152. 74. Ihara, K., Muraguchi, S., Kato, M., Shimizu, T., Shirakawa, M., Kuroda, S., Kaibuchi, K., and Hakoshima, T. (1998) Crystal structure of human RhoA in a dominantly active form complexed with a GTP analogue, J. Biol. Chem. 273, 9656–9666. 75. Longenecker, K., Read, P., Derewenda, U., Dauter, Z., Liu, X., Garrard, S., Walker, L., Somlyo, A. V., Nakamoto, R. K., Somlyo, A. P., and Derewenda, Z. S. (1999) How RhoGDI binds Rho, Acta Crystallogr. D. Biol. Crystallogr. 55 (Pt 9), 1503–1515.

Nitric Oxide and Cell Signaling: Redox Regulation of Ras Superfamily GTPases 289 76. Abdul-Manan, N., Aghazadeh, B., Liu, G. A., Majumdar, A., Ouerfelli, O., Siminovitch, K. A., and Rosen, M. K. (1999) Structure of Cdc42 in complex with the GTPase-binding domain of the “Wiskott–Aldrich syndrome” protein, Nature 399, 379–383. 77. Scheffzek, K., Stephan, I., Jensen, O. N., Illenberger, D., and Gierschik, P. (2000) The Rac-RhoGDI complex and the structural basis for the regulation of Rho proteins by RhoGDI, Nat. Struct. Biol. 7, 122–126. 78. Worthylake, D. K., Rossman, K. L., and Sondek, J. (2000) Crystal structure of Rac1 in complex with the guanine nucleotide exchange region of Tiam1, Nature 408, 682–688. 79. Hodgkiss, R. J. (1998) Use of 2-nitroimidazoles as bioreductive markers for tumour hypoxia, Anticancer Drug Des. 13, 687–702. 80. Denny, W. A. (2000) The role of hypoxia-activated prodrugs in cancer therapy, Lancet Oncol. 1, 25–29. 81. Evans, S. M., and Koch, C. J. (2003) Prognostic significance of tumor oxygenation in humans, Cancer Lett. 195, 1–16. 82. Bussink, J., Kaanders, J. H., and van der Kogel, A. J. (2003) Tumor hypoxia at the micro-regional level: clinical relevance and predictive value of exogenous and endogenous hypoxic cell markers, Radiother. Oncol. 67, 3–15. 83. Cairns, R. A., and Hill, R. P. (2004) Acute hypoxia enhances spontaneous lymph node metastasis in an orthotopic murine model of human cervical carcinoma, Cancer Res. 64, 2054–2061. 84. Cherfils, J., and Chardin, P. (1999) GEFs: structural basis for their activation of small GTP-binding proteins, Trends Biochem. Sci. 24, 306–311. 85. Boriack-Sjodin, P. A., Margarit, S. M., Bar-Sagi, D., and Kuriyan, J. (1998) The structural basis of the activation of Ras by Sos, Nature 394, 337–343. 86. Sayle, R. A., and Milner-White, E. J. (1995) RASMOL: biomolecular graphics for all, Trends Biochem. Sci. 20, 374. 87. Saville, B. (1958) A scheme for the colorimetric determination of microgram amounts of thiols, Analyst 83, 670–672. 88. Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding, Anal. Biochem. 72, 248–254. 89. Koradi, R., Billeter, M., and Wührich, K. (1996) A program for display and analysis of macromolecular structures, J. Mol. Graphics. 14, 51–55.

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

292

Nitric Oxide, Cell Signaling, and Gene Expression

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

293

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,

294

Nitric Oxide, Cell Signaling, and Gene Expression

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-

Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System

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,

296

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System

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.

298

Nitric Oxide, Cell Signaling, and Gene Expression

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,

Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System

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.

Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System

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

Nitric Oxide, Cell Signaling, and Gene Expression

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.

Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System

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).

REFERENCES 1. Nathan, C. (1992) Nitric oxide as a secretory product of mammalian cells. FASEB J. 6, 3051–3064 2. Schmidt, H. H., and Walter, U. (1994) NO at work. Cell 78, 919–925. 3. Moncada, S., and Higgs, E. A. (1995) Molecular mechanisms and therapeutic strategies related to nitric oxide. FASEB J. 9, 1319–1330. 4. Alderton, W. K., Cooper, C. E., and Knowles, R. G. (2001) Nitric oxide synthases: structure, function and inhibition. Biochem. J. 357, 593–615. 5. Kleinert, H., Schwarz, P. M., and Forstermann, U. (2003) Regulation of the expression of inducible nitric oxide synthase. Biol. Chem. 384, 1343–1364. 6. Stamler, J. S. (1994) Redox signaling: nitrosylation and related target interactions of nitric oxide. Cell 78, 931–936.

306

Nitric Oxide, Cell Signaling, and Gene Expression

7. Grisham, M. B., Jourd’Heuil, D., and Wink, D. A. (1999) Nitric oxide. I. Physiological chemistry of nitric oxide and its metabolites: implications in inflammation. Am. J. Physiol. 276, G315–321. 8. Furchgott, R. F., and Zawadzki, J. V. (1980) The obligatory role of endothelial cells in the relaxation of arterial smooth muscle by acetylcholine. Nature 288, 373–376. 9. Butler, A. R., Flitney, F. W., and Williams, D. L. (1995) NO, nitrosonium ions, nitroxide ions, nitrosothiols and iron-nitrosyls in biology: a chemist’s perspective. Trends Pharmacol. Sci. 16, 18–22. 10. Koppenol, W. H. (2002) NO nomenclature? Nitric Oxide 6, 96–98. 11. Marshall, H. E., Merchant, K., and Stamler, J. S. (2000) Nitrosation and oxidation in the regulation of gene expression. FASEB J. 14, 1889–1900. 12. Bogdan, C. (2001) Nitric oxide and the regulation of gene expression. Trends. Cell Biol. 11, 66–75. 13. Kroncke, K. D. (2003) Nitrosative stress and transcription. Biol. Chem. 384, 1365–1377. 14. von Knethen, A., and Brune, B. (2003) PPARgamma—an important regulator of monocyte/macrophage function. Arch. Immunol. Ther. Exp. (Warsz) 51, 219–226. 15. Nathan, C., and Shiloh, M. U. (2000) Reactive oxygen and nitrogen intermediates in the relationship between mammalian hosts and microbial pathogens. Proc. Natl. Acad. Sci. USA 97, 8841–8848. 16. Semenza, G. L. (2002) HIF-1 and tumor progression: pathophysiology and therapeutics. Trends Mol. Med. 8, S62–67. 17. Pugh, C. W., and Ratcliffe, P. J. (2003) Regulation of angiogenesis by hypoxia: role of the HIF system. Nat. Med. 9, 677–684. 18. Brüne, B., and Zhou, J. (2003) The role of nitric oxide (NO) in stability regulation of hypoxia inducible factor-1alpha (HIF-1alpha). Curr. Med. Chem. 10, 845–855. 19. Huang, L. E., and Bunn, H. F. (2003) Hypoxia-inducible factor and its biomedical relevance. J. Biol. Chem. 278, 19575–19578. 20. Semenza, G. L., and Wang, G. L. (1992) A nuclear factor induced by hypoxia via de novo protein synthesis binds to the human erythropoietin gene enhancer at a site required for transcriptional activation. Mol. Cell. Biol. 12, 5447–5454. 21. Maxwell, P. H., Pugh, C. W., and Ratcliffe, P. J. (2001) The pVHL-HIF-1 system. A key mediator of oxygen homeostasis. Adv. Exp. Med. Biol. 502, 365–376. 22. Wenger, R. H. (2002) Cellular adaptation to hypoxia: O2-sensing protein hydroxylases, hypoxia-inducible transcription factors, and O2-regulated gene expression. FASEB J. 16, 1151–1162. 23. Kaelin, W. G., Jr. (2002) How oxygen makes its presence felt. Genes Dev. 16, 1441–1445. 24. Wang, G. L., and Semenza, G. L. (1995) Purification and characterization of hypoxia-inducible factor 1. J. Biol. Chem. 270, 1230–1237. 25. Salceda, S., and Caro, J. (1997) Hypoxia-inducible factor 1alpha (HIF-1alpha) protein is rapidly degraded by the ubiquitin-proteasome system under normoxic conditions. J. Biol. Chem. 272, 22642–22647. 26. Kallio, P. J., Wilson, W. J., O’Brien, S., Makino, Y., and Poellinger, L. (1999) Regulation of the hypoxia-inducible transcription factor 1alpha by the ubiquitin–proteasome pathway. J. Biol. Chem. 274, 6519–6525.

Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System

307

27. Huang, L. E., Gu, J., Schau, M., and Bunn, H. F. (1998) Regulation of hypoxiainducible factor 1alpha is mediated by an O2-dependent degradation domain via the ubiquitin-proteasome pathway. Proc. Natl. Acad. Sci. USA 95, 7987–7992. 28. Jaakkola, P., Mole, D. R., Tian, Y. M., Wilson, M. I., Gielbert, J., Gaskell, S. J., Kriegsheim, A. V., Hebestreit, H. F., Mukherji, M., Schofield, C. J., Maxwell, P. H., Pugh, C. W., and Ratcliffe, P. J. (2001) Targeting of HIF-alpha to the von Hippel–Lindau ubiquitylation complex by O2-regulated prolyl hydroxylation. Science 292, 468–472. 29. Ivan, M., Kondo, K., Yang, H., Kim, W., Valiando, J., Ohh, M., Salic, A., Asara, J. M., Lane, W. S., and Kaelin, W. G., Jr. (2001) HIFalpha targeted for VHLmediated destruction by proline hydroxylation: implications for O2 sensing. Science 292, 464–468. 30. Epstein, A. C., Gleadle, J. M., McNeill, L. A., Hewitson, K. S., O’Rourke, J., Mole, D. R., Mukherji, M., Metzen, E., Wilson, M. I., Dhanda, A., Tian, Y. M., Masson, N., Hamilton, D. L., Jaakkola, P., Barstead, R., Hodgkin, J., Maxwell, P. H., Pugh, C. W., Schofield, C. J., and Ratcliffe, P. J. (2001) C. elegans EGL-9 and mammalian homologs define a family of dioxygenases that regulate HIF by prolyl hydroxylation. Cell 107, 43–54. 31. Bruick, R. K., and McKnight, S. L. (2001) A conserved family of prolyl-4hydroxylases that modify HIF. Science 294, 1337–1340. 32. Oehme, F., Ellinghaus, P., Kolkhof, P., Smith, T. J., Ramakrishnan, S., Hutter, J., Schramm, M., and Flamme, I. (2002) Overexpression of PH-4, a novel putative proline 4-hydroxylase, modulates activity of hypoxia-inducible transcription factors. Biochem. Biophys. Res. Commun. 296, 343–349. 33. Maxwell, P. H., Wiesener, M. S., Chang, G.-W., Clifford, S. C., Vaux, E. C., Cockman, M. E., Wykoff, C. C., Pugh, C. W., Maher, E. R., and Ratcliffe, P. J. (1999) The tumour suppressor protein VHL targets hypoxia-inducible factors for oxygen-dependent proteolysis. Nature 399, 271–275. 34. Valegard, K., van Scheltinga, A. C., Lloyd, M. D., Hara, T., Ramaswamy, S., Perrakis, A., Thompson, A., Lee, H. J., Baldwin, J. E., Schofield, C. J., Hajdu, J., and Andersson, I. (1998) Structure of a cephalosporin synthase. Nature 394, 805–809. 35. Roach, P. L., Clifton, I. J., Fulop, V., Harlos, K., Barton, G. J., Hajdu, J., Andersson, I., Schofield, C. J., and Baldwin, J. E. (1995) Crystal structure of isopenicillin N synthase is the first from a new structural family of enzymes. Nature 375, 700–704. 36. Hirsila, M., Koivunen, P., Gunzler, V., Kivirikko, K. I., and Myllyharju, J. (2003) Characterization of the human prolyl 4-hydroxylases that modify the hypoxiainducible factor. J. Biol. Chem. 278, 30772–30780. 37. Knowles, H. J., Raval, R. R., Harris, A. L., and Ratcliffe, P. J. (2003) Effect of ascorbate on the activity of hypoxia-inducible factor in cancer cells. Cancer Res. 63, 1764–1768. 38. Wang, G. L., and Semenza, G. L. (1993) Desferrioxamine induces erythropoietin gene expression and hypoxia-inducible factor 1 DNA-binding activity: implications for models of hypoxia signal transduction. Blood 82, 3610–3615. 39. Lando, D., Peet, D. J., Whelan, D. A., Gorman, J. J., and Whitelaw, M. L. (2002) Asparagine hydroxylation of the HIF transactivation domain: a hypoxic switch. Science 295, 858–861.

308

Nitric Oxide, Cell Signaling, and Gene Expression

40. Lee, C., Kim, S. J., Jeong, D. G., Lee, S. M., and Ryu, S. E. (2003) Structure of human FIH-1 reveals a unique active site pocket and interaction sites for HIF-1 and von Hippel-Lindau. J. Biol. Chem. 278, 7558–7563. 41. Dann, C. E., 3rd, Bruick, R. K., and Deisenhofer, J. (2002) Structure of factorinhibiting hypoxia-inducible factor 1: an asparaginyl hydroxylase involved in the hypoxic response pathway. Proc. Natl. Acad. Sci. USA 99, 15351–15356. 42. Elkins, J. M., Hewitson, K. S., McNeill, L. A., Seibel, J. F., Schlemminger, I., Pugh, C. W., Ratcliffe, P. J., and Schofield, C. J. (2003) Structure of factorinhibiting hypoxia-inducible factor (HIF) reveals mechanism of oxidative modification of HIF-1 alpha. J. Biol. Chem. 278, 1802–1806. 43. Semenza, G. L. (2000) HIF-1 and human disease: one highly involved factor. Genes Dev. 14, 1983–1991. 44. Semenza, G. L. (2003) Targeting HIF-1 for cancer therapy. Nat. Rev. Cancer 3, 721–732. 45. del Peso, L., Castellanos, M. C., Temes, E., Martin-Puig, S., Cuevas, Y., Olmos, G., and Landazuri, M. O. (2003) The von Hippel–Lindau/hypoxia-inducible factor (HIF) pathway regulates the transcription of the HIF-proline hydroxylase genes in response to low oxygen. J. Biol. Chem. 278, 48690–48695. 46. Berra, E., Benizri, E., Ginouves, A., Volmat, V., Roux, D., and Pouyssegur, J. (2003) HIF prolyl-hydroxylase 2 is the key oxygen sensor setting low steady-state levels of HIF-1alpha in normoxia. EMBO J. 22, 4082–4090. 47. Cioffi, C. L., Liu, X. Q., Kosinski, P. A., Garay, M., and Bowen, B. R. (2003) Differential regulation of HIF-1 alpha prolyl-4-hydroxylase genes by hypoxia in human cardiovascular cells. Biochem. Biophys. Res. Commun. 303, 947–953. 48. D’Angelo, G., Duplan, E., Boyer, N., Vigne, P., and Frelin, C. (2003) Hypoxia up-regulates prolyl hydroxylase activity: a feedback mechanism that limits HIF1 responses during reoxygenation. J. Biol. Chem. 278, 38183–38187. 49. Berra, E., Richard, D. E., Gothie, E., and Pouyssegur, J. (2001) HIF-1-dependent transcriptional activity is required for oxygen-mediated HIF-1alpha degradation. FEBS Lett. 491, 85–90. 50. Sandau, K. B., Fandrey, J., and Brüne, B. (2001) Accumulation of HIF-1alpha under the influence of nitric oxide. Blood 97, 1009–1015. 51. Kimura, H., Weisz, A., Kurashima, Y., Hashimoto, K., Ogura, T., D’Acquisto, F., Addeo, R., Makuuchi, M., and Esumi, H. (2000) Hypoxia response element of the human vascular endothelial growth factor gene mediates transcriptional regulation by nitric oxide: control of hypoxia-inducible factor-1 activity by nitric oxide. Blood 95, 189–197. 52. Kimura, H., Weisz, A., Ogura, T., Hitomi, Y., Kurashima, Y., Hashimoto, K., D’Acquisto, F., Makuuchi, M., and Esumi, H. (2001) Identification of hypoxiainducible factor 1 ancillary sequence and its function in vascular endothelial growth factor gene induction by hypoxia and nitric oxide. J. Biol. Chem. 276, 2292–2298. 53. Palmer, L. A., Gaston, B., and Johns, R. A. (2000) Normoxic stabilization of hypoxia-inducible factor-1 expression and activity: Redox-dependent effect of nitrogen oxides. Mol. Pharm. 58, 1197–1203 54. Sumbayev, V. V., Budde, A., Zhou, J., and Brüne, B. (2003) HIF-1alpha protein as a target for S-nitrosation. FEBS Lett. 535, 106–112.

Nitric Oxide and the Hypoxia Inducible Factor-1 Transducing System

309

55. Zhou, J., Fandrey, J., Schumann, J., Tiegs, G., and Brüne, B. (2003) NO and TNFalpha released from activated macrophages stabilize HIF-1alpha in resting tubular LLC-PK1 cells. Am. J. Physiol. Cell Physiol. 284, C439–446. 56. Sheta, E. A., Trout, H., Gildea, J. J., Harding, M. A., and Theodorescu, D. (2001) Cell density mediated pericellular hypoxia leads to induction of HIF-1alpha via nitric oxide and Ras/MAP kinase mediated signaling pathways. Oncogene 20, 7624–7634. 57. Xiong, M., Elson, G., Legarda, D., and Leibovich, S. J. (1998) Production of vascular endothelial growth factor by murine macrophages: regulation by hypoxia, lactate, and the inducible nitric oxide synthase pathway. Am. J. Pathol. 153, 587–598. 58. Albina, J. E., Mastrofrancesco, B., Vessella, J. A., Louis, C. A., Henry, W. L., Jr., and Reichner, J. S. (2001) HIF-1 expression in healing wounds: HIF-1alpha induction in primary inflammatory cells by TNF-alpha. Am. J. Physiol. 281, C1971–1977. 59. Burke, B., Tang, N., Corke, K. P., Tazzyman, D., Ameri, K., Wells, M., and Lewis, C. E. (2002) Expression of HIF-1alpha by human macrophages: implications for the use of macrophages in hypoxia-regulated cancer gene therapy. J. Pathol. 196, 204–212. 60. Cramer, T., Yamanishi, Y., Clausen, B. E., Forster, I., Pawlinski, R., Mackman, N., Haase, V. H., Jaenisch, R., Corr, M., Nizet, V., Firestein, G. S., Gerber, H. P., Ferrara, N., and Johnson, R. S. (2003) HIF-1alpha is essential for myeloid cellmediated inflammation. Cell 112, 645–657. 61. Metzen, E., Zhou, J., Jelkmann, W., Fandrey, J., and Brüne, B. (2003) Nitric oxide impairs normoxic degradation of HIF-1alpha by inhibition of prolyl hydroxylases. Mol. Biol. Cell 14, 3470–3481. 62. Arciero, D. M., Orville, A. M., and Lipscomb, J. D. (1985) [17O]Water and nitric oxide binding by protocatechuate 4,5-dioxygenase and catechol 2,3-dioxygenase. Evidence for binding of exogenous ligands to the active site Fe2+ of extradiol dioxygenases. J. Biol. Chem. 260, 14035–14044. 63. Hegg, E. L., and Que, L., Jr. (1997) The 2-His-1-carboxylate facial triad—an emerging structural motif in mononuclear non-heme iron(II) enzymes. Eur. J. Biochem. 250, 625–629. 64. Kasuno, K., Takabuchi, S., Fukuda, K., Kizaka-Kondoh, S., Yodoi, J., Adachi, T., Semenza, G. L., and Hirota, K. (2004) Nitric oxide induces hypoxia-inducible factor 1 activation that is dependent on MAPK and phosphatidylinositol 3-kinase signaling. J. Biol. Chem. 279, 2550–2558. 65. Zhang, S. X., Gozal, D., Sachleben, L. R., Jr., Rane, M., Klein, J. B., and Gozal, E. (2003) Hypoxia induces an autocrine-paracrine survival pathway via plateletderived growth factor (PDGF)-B/PDGF-beta receptor/phosphatidylinositol 3kinase/Akt signaling in RN46A neuronal cells. FASEB J. 17, 1709–1711. 66. Sandau, K. B., Zhou, J., Kietzmann, T., and Brüne, B. (2001) Regulation of the hypoxia-inducible factor 1alpha by the inflammatory mediators nitric oxide and tumor necrosis factor-alpha in contrast to desferroxamine and phenylarsine oxide. J. Biol. Chem. 276, 39805–39811. 67. Wellman, T. L., Jenkins, J., Penar, P. L., Tranmer, B., Zahr, R., and Lounsbury, K. M. (2004) Nitric oxide and reactive oxygen species exert opposing effects on the stability of hypoxia inducible factor-1alpha (HIF-1alpha) in explants of human pial arteries. FASEB J. 18, 379–381.

310

Nitric Oxide, Cell Signaling, and Gene Expression

68. Liu, Y., Christou, H., Morita, T., Laughner, E., Semenza, G. L., and Kourembanas, S. (1998) Carbon monoxide and nitric oxide suppress the hypoxic induction of vascular endothelial growth factor gene via the 5′ enhancer. J. Biol. Chem. 273, 15257–15262. 69. Sogawa, K., Numayama-Tsuruta, K., Ema, M., Abe, M., Abe, H., and FujiiKuriyama, Y. (1998) Inhibition of hypoxia-inducible factor 1 activity by nitric oxide donors in hypoxia. Proc. Natl. Acad. Sci. USA 95, 7368–7373. 70. Huang, L. E., Willmore, W. G., Gu, J., Goldberg, M. A., and Bunn, H. F. (1999) Inhibition of hypoxia-inducible factor 1 activation by carbon monoxide and nitric oxide. J. Biol. Chem. 274, 9038–9044 71. Srinivas, V., Zhu, X., Salceda, S., Nakamura, R., and Caro, J. (1998) Hypoxiainducible factor 1alpha (HIF-1alpha) is a non-heme iron protein. Implications for oxygen sensing. J. Biol. Chem. 273, 18019–18022. 72. Hagen, T., Taylor, C. T., Lam, F., and Moncada, S. (2003) Redistribution of intracellular oxygen in hypoxia by nitric oxide: effect on HIF1alpha. Science 302, 1975–1978. 73. Mateo, J., Garcia-Lecea, M., Cadenas, S., Hernandez, C., and Moncada, S. (2003) Regulation of hypoxia-inducible factor-1alpha by nitric oxide through mitochondria-dependent and -independent pathways. Biochem. J. 376, 537–544. 74. Wang, F., Sekine, H., Kikuchi, Y., Takasaki, C., Miura, C., Heiwa, O., Shuin, T., Fujii-Kuriyama, Y., and Sogawa, K. (2002) HIF-1alpha-prolyl hydroxylase: molecular target of nitric oxide in the hypoxic signal transduction pathway. Biochem. Biophys. Res. Commun. 295, 657–662. 75. Ramachandran, A., Ceaser, E., and Darley-Usmar, V. M. (2004) Chronic exposure to nitric oxide alters the free iron pool in endothelial cells: role of mitochondrial respiratory complexes and heat shock proteins. Proc. Natl. Acad. Sci. USA 101, 384–389. 76. Petrat, F., de Groot, H., Sustmann, R., and Rauen, U. (2002) The chelatable iron pool in living cells: a methodically defined quantity. Biol. Chem. 383, 489–502. 77. Zhong, H., De Marzo, A. M., Laughner, E., Lim, M., Hilton, D. A., Zagzag, D., Buechler, P., Isaacs, W. B., Semenza, G. L., and Simons, J. W. (1999) Overexpression of hypoxia-inducible factor 1alpha in common human cancers and their metastases. Cancer Res. 59, 5830–5835. 78. Wink, D. A., Vodovotz, Y., Laval, J., Laval, F., Dewhirst, M. W., and Mitchell, J. B. (1998) The multifaceted roles of nitric oxide in cancer. Carcinogenesis 19, 711–721. 79. Morbidelli, L., Donnini, S., and Ziche, M. (2003) Role of nitric oxide in the modulation of angiogenesis. Curr. Pharm. Des. 9, 521–530. 80. Xie, K., Huang, S., Dong, Z., Juang, S. H., Wang, Y., and Fidler, I. J. (1997) Destruction of bystander cells by tumor cells transfected with inducible nitric oxide (NO) synthase gene. J. Natl. Cancer Inst. 89, 421–427. 81. Xu, W., Liu, L. Z., Loizidou, M., Ahmed, M., and Charles, I. G. (2002) The role of nitric oxide in cancer. Cell Res. 12, 311–320. 82. Xie, K., and Huang, S. (2003) Contribution of nitric oxide-mediated apoptosis to cancer metastasis inefficiency. Free Radic. Biol. Med. 34, 969–986. 83. Crowell, J. A., Steele, V. E., Sigman, C. C., and Fay, J. R. (2003) Is inducible nitric oxide synthase a target for chemoprevention? Mol. Cancer. Ther. 2, 815–823.

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

311

312

Nitric Oxide, Cell Signaling, and Gene Expression

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

313

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

Nitric Oxide, Cell Signaling, and Gene Expression

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

The Cross Talk between Nitric Oxide and Ceramide

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

Nitric Oxide, Cell Signaling, and Gene Expression

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.

REFERENCES 1. Huwiler, A. Kolter, T., Pfeilschifter, J., and Sandhoff, K. (2000) Physiology and pathophysiology of sphingolipid metabolism and signalling. Biochim. Biophys. Acta 1485, 63–99. 2. Brown, D. A. and London, E. (2000) Structure and function of sphingolipid- and cholesterol-rich membrane rafts. J. Biol. Chem. 275, 17221–17224.

The Cross Talk between Nitric Oxide and Ceramide

317

3. Hannun, A. and Obeid L. M. (2002) The ceramide-centric universe of lipidmediated cell regulation: stress encounters of the lipid kind. J. Biol. Chem. 277, 25847–25850. 4. Hla, T. (2003) Signaling and biological actions of sphingosine 1-phosphate. Pharmacol. Res. 47, 401–407. 5. Alderton, W. K., Cooper, C. E., and Knowles, R. G. (2001) Nitric oxide synthases: structure, function and inhibition. Biochem. J. 357, 593–615. 6. Brenman, J. E., Chao, D. S., Gee, S. H., McGee, A. W., Craven, S. E., Santillano, D. R., Wu, Z., Huang, F., Xia, H., Peters, M. F., Froehner, S. C., and Bredt, D. S. (1996) Interaction of nitric oxide synthase with the postsynaptic density protein PSD-95 and α 1-syntrophin mediated by PDZ domains. Cell 84, 757–767. 7. Fulton, D., Gratton, J. P., and Sessa, W. C. (2001) Post-translational control of endothelial nitric oxide synthase: why isn't calcium/calmodulin enough? J. Pharmacol. Exp. Ther. 299, 818–24. 8. Felley-Bosco, E., Bender, F. C., Courjault-Gautier, F., Bron, C., and Quest, A. F. (2000) Caveolin-1 down-regulates inducible nitric oxide synthase via the proteasome pathway in human colon carcinoma cells. Proc. Natl. Acad. Sci. USA 97, 14334–14339. 9. Clementi, E., Borgese, N., and Meldolesi, J. (2003) Interactions between nitric oxide and sphingolipids and the potential consequences in physiology and pathology. Trends Pharmacol. Sci. 24, 518–523. 10. Florio, T., Arena, S., Pattarozzi, A., Thellung, S., Corsaro, A., Villa, V., Massa, A., Diana, F., Spoto, G., Forcella, S., Damonte, G., Filocamo, M., Benatti, U., and Schettini, G. (2003) Basic fibroblast growth factor activates endothelial nitric oxide synthase in CHO-CK1 cells via the activation of ceramide synthesis. Mol. Pharmacol. 63, 297–310. 11. Bulotta, S. Barsacchi, R., Rotiroti, D., Borgese, N., and Clementi, E. (2001) Activation of the endothelial nitric-oxide synthase by tumor necrosis factor-α: a novel feedback mechanism regulating cell death. J. Biol. Chem. 276, 6659–6536. 12. Barsacchi, R. Perrotta, C., Bulotta, S., Moncada, S., Borgese, N., and Clementi, E. (2003) Activation of endothelial nitric oxide synthase by tumor necrosis factorα: a novel pathway involving sequential activation of neutral sphingomyelinase, phosphatidylinositol-3’ kinase and Akt. Mol. Pharmacol. 63, 886–895. 13. Igarashi, J. and Michel, T. (2000) Agonist-modulated targeting of the EDG-1 receptor to plasmalemmal caveolae. J. Biol. Chem. 275, 32363–32370. 14. Kwon, Y.-G., Min, J. K., Kim, K. M., Lee, D. J., Billiar, T. R., and Kim, Y. M. (2001) Sphingosine 1-phosphate protects human umbilical vein endothelial cells from serum-deprived apoptosis by nitric oxide production. J. Biol. Chem. 276, 10627–10633. 15. Igarashi, J., Bernier, S. G., and Michel, T. (2001) Sphingosine 1-phosphate and activation of endothelial nitric-oxide synthase. J. Biol. Chem. 276, 12420–12426. 16. Morales-Ruiz, M., Lee, M. J., Zollner, S., Gratton, J. P., Scotland, R., Shiojima, I., Walsh, K., Hla, T., and Sessa, W. C. (2001) Sphingosine 1-phosphate activates Akt, nitric oxide production, and chemotaxis through a Gi protein/phosphoinositide 3-kinase pathway in endothelial cells. J. Biol. Chem. 276, 19672–19677. 17. Rikitake, Y., Hirata, K., Kawashima, S., Ozaki, M., Takahashi, T., Ogawa, W., Inoue, N., and Yokoyama, M. (2002) Involvement of endothelial nitric oxide in sphingosine-1 phosphate-induced angiogenesis. Aterioscler. Thromb. Vasc. Biol. 22, 108–114.

318

Nitric Oxide, Cell Signaling, and Gene Expression

18. Liu, L. and Stamler J. S. (1999) NO: an inhibitor of cell death. Cell Death Differ. 6, 937–942. 19. Brune, B. (2003) Nitric oxide: NO apoptosis or turning it ON? Cell Death Differ. 10, 864–869. 20. Sciorati, C., Rovere. P., Ferrarini, M., Heltai, S., Manfredi, A. A., and Clementi, E. (1997) Autocrine nitric oxide modulates CD95-induced apoptosis in γδ T lymphocytes. J. Biol. Chem. 272, 23211–23215. 21. Sciorati, C., Rovere, P., Ferrarini, M., Paolucci, C., Heltai, S., Vaiani, R., Clementi, E., and Manfredi, A. A. (1999) Generation of nitric oxide by the inducible nitric oxide synthase protects γδ T cells from Mycobacterium tuberculosis-induced apoptosis. J. Immunol. 163, 1570–1576. 22. Barsacchi, R., Perrotta, C., Sestili, P., Cantoni, O., Moncada, S., and Clementi, E. (2002) Cyclic GMP-dependent inhibition of acid sphingomyelinase by nitric oxide: an early step in protection against apoptosis. Cell Death Differ. 9, 1248–1255. 23. Gulbins, E. (2003) Regulation of death receptor signaling and apoptosis by ceramide. Pharmacol. Res. 47, 393–399. 24. De Nadai, C., Sestili, P., Cantoni, O., Lievremont, J. P., Sciorati, C., Barsacchi, R., Moncada, S., Meldolesi, J., and Clementi, E. (2000) Nitric oxide inhibits tumor necrosis factor-α-induced apoptosis by reducing the generation of ceramide. Proc. Natl. Acad. Sci. USA 97, 5480–5485. 25. Huwiler, A., Pfeilschifter, J., and van den Bosch, H. (1999) Nitric oxide donors induce stress signaling via ceramide formation in rat renal mesangial cells. J. Biol. Chem. 274, 7190–7195. 26. Takeda, Y., Tashima, M., Takahashi, A., Uchiyama, T., and Okazaki, T. (1999) Ceramide generation in nitric oxide-induced apoptosis. Activation of magnesiumdependent neutral sphingomyelinase via caspase-3. J. Biol. Chem. 274, 10654–10660. 27. Baud, V. and Karin, M. (2001) Signal transduction by tumor necrosis factor and its relatives. Trends Cell Biol. 11, 372–377. 28. Bogdan, C. (2001) Nitric oxide and the immune response. Nat. Immunol. 2, 907–916. 29. Paolucci, C., Rovere, P., De Nadai, C., Manfredi, A. A., and Clementi, E. (2000) Nitric oxide inhibits the tumor necrosis factor- α-regulated endocytosis of human dendritic cells in a cyclic GMP-dependent way. J. Biol. Chem. 275, 19638–19644. 30. Vann, L. R., Payne, S. G., Edsall, L. C., Twitty, S., Spiegel, S., and Milstien, S. (2002) Involvement of sphingosine kinase in TNF-α-stimulated tetrahydrobiopterin biosynthesis in C6 glioma cells. J. Biol. Chem. 277, 12649–12656. 31. Garcìa-Cardeña, G., Oh, P., Liu, J., Schnitzer, J. E., and Sessa, W.C. (1996) Targeting of nitric oxide synthase to endothelial cell caveolae via palmitoylation: implications for nitric oxide signaling. Proc. Natl. Acad. Sci. USA 93, 6448–6453. 32. Ko, Y. G., Lee, J. S., Kang, Y. S., Ahn, J. H., and Seo, J. S. (1999) TNF-αmediated apoptosis is initiated in caveolae-like domains. J. Immunol. 162, 7217–7223. 33. Clementi, E. and Meldolesi, J. (1997) The cross-talk between nitric oxide and Ca2+: a story with a complex past and a promising future. Trends Pharmacol. Sci. 18, 266–269. 34. Kester, M., and Kolesnick, R. (2003) Sphingolipids as therapeutics. Pharmacol. Res. 47, 365–371.

The Cross Talk between Nitric Oxide and Ceramide

319

35. Burgaud, J. L., Ongini, E., and Del Soldato, P. (2002) Nitric oxide-releasing drugs: a novel class of effective and safe therapeutic agents. Ann. NY Acad. Sci. 962, 360–371. 36. Adam-Klages S., Adam, D., Wiegmann, K., Struve, S., Kolanus, W., SchneiderMergener, J., and Kronke, M. (1996) FAN, a novel WD-repeat protein, couples the p55 TNF-receptor to neutral sphingomyelinase. Cell 86, 937–947.

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

Nitric Oxide, Cell Signaling, and Gene Expression

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 <10 mmHg, PHDs no longer target HIF 1α, HIF 1α is no longer degraded, and hypoxia-dependent transcription increases (23). Of note, NO itself inhibits this hypoxia-driven up-regulation of HIF 1α, possibly by affecting iron binding to the oxygen-dependent degradation domain (22). Remarkably S-Nitrosoglutathione (GSNO), which is endogenously present in the brainstem and elsewhere (24, 25), prevents HIF 1α degradation and increases HIF 1α-dependent transcription under conditions of normoxia (7) (Figure 14.1).

S-Nitrosothiol Signaling and Gene Regulation in Pulmonary Pathophysiology

323

TABLE 14.1 Proposed Criteria to Establish that a Specific Bioactivity Is Associated with S-Nitrosation or Denitrosation of a Specific Protein 1.The altered bioactivity of the target protein is associated with increased (or decreased) activity of a nitric oxide synthase (NOS) isoform. 2.S-nitrosation of the target protein isolated from cells (or in situ) following NOS activation can be demonstrated by more than one independent assay (ideally, three assays). In addition, the extent of nitrosation from endogenous nitric oxide formation is sufficient in magnitude to affect the activity of the protein, and nitrosation/denitrosation occurs rapidly enough to account for regulated changes in activity. 3.Mutation of a specific cysteine in the target protein results in: loss of the NOS-responsive bioactivity and inability to identify the S-nitrosated protein following NOS activation. 4.In association with termination of the bioactivity, loss of the cellular protein S-nitrosation is demonstrated. 5.Alteration of the function of the purified protein can be demonstrated in association with S-nitrosation under conditions relevant to the protein’s cellular environment (i.e., in the presence of a nitrogen oxide at concentrations measured in the specifically relevant tissue or cell compartment at the appropriate PO2, pH, etc.). 6.Pharmacological experiments demonstrate that cGMP is not exclusively involved in mediating the bioactivity. 7.Pharmacological experiments suggest that a thiol modification is involved (i.e., the altered bioactivity is blocked by pretreatment with N-ethylmaleimide or reversed by excess DTT). 8.Pharmacological modifications of specific SNO metabolic enzymes relevant to the putative signaling process, such as -γ glutamyl transpeptidase or glutathione-dependent formaldehyde dehydrogenase), appropriately alter the bioactivity, or the bioactivity is caused by S-nitrosoL-cysteine-containing compounds but not S-nitroso-D-cysteine-containing compounds.

This effect appears to involve cysteine S-nitrosylation reactions: It is independent of NO or cyclic GMP and reversed by the reducing agent, dithiothreitol (DDT). Moreover, it is inhibited acivicin, an inhibitor of γ glutamyl transpeptidase (GGT), suggesting that intact transport of S-nitroso-cysteinyl glycine across the cell membrane is of importance (7, 19). There are several possible S-nitrosylation targets that could prevent HIF 1α degradation; these include pvHL, PHDs, the oxygendependent domain of HIF 1α, the proteosome, and a variety by the chaperone proteins. Additionally, GSNO may target components of the inositol triphosphate (PI3)/Akt pathway (26). Inhibitors of this pathway attenuate or ablate the effect of GSNO on HIF 1α stability. This is particularly true in experiments in which higher concentrations of GSNO are used. Based on these data, it appears clear that GSNO can activate HIF 1 via more than one pathway. It has been appreciated for some time at the whole organism level that there should be a mechanism by which HIF 1α can be stabilized—and HIF 1 activated—at PO2s that are substantially higher than those (1% oxygen) required to activate HIF 1 in cell cultures and cell systems in vitro. This paradox might be resolved by considering the problem in the context of GSNO signaling. Researchers agree that hemoglobin (Hb) undergoes S-nitrosylation at the β 93

324 A

Nitric Oxide, Cell Signaling, and Gene Expression 0

+24 NOC-18 H

C

0

100 500

H

HIF-1

B

0

500

1000

H

HIF-1

D HIF-1

0

100 500

H HIF-1

FIGURE 14.1 The effect of NOC-18 is mediated by reactions of the nitrosonium ion. Nuclear proteins isolated from bovine pulmonary artery endothelial cells treated with 500 mM NOC-18 that was allowed to decay for 24 h (124) (A); 8-Br cGMP (B); Angeli’s salt (C); or GSNO (D) for a period of 4 h. HIF-1 DNA-binding activity was determined by EMSA as described in Figure 14.1. C = constitutive protein-DNA complexes; H = hypoxia.

cysteine, forming SNOHb. Deoxygenation with Hb transition to tense (T) state, exposes the β 93 SNO bond to transnitrosation reactions (11, 15, 27). The NO+ equivalent can be transferred to anion exchange protein 1 or to erythrocytic GSH (11, 15, 27, 28). GSNO formed by Hb deoxygenation can be transferred outside of the red cell to signal bioactivities, such as increased minute ventilation and relaxation of systemic arterioles (11, 28). Thus, GSNO can signal a regulatory response to oxyHb desaturation—as opposed to profoundly low pO2 (7). Indeed, levels of pO2 (<10 mmHg) that increase HIF 1α expression and up-regulate HIF 1 binding are almost never seen in the pulmonary vascular endothelium; we hypothesize the regulation of HIF 1-dependent genes in pulmonary hypertension occurs through this oxy Hb-desaturation/SNOmediated mechanism, as opposed to the purely hypoxia-dependent mechanism. Recently, Hildebrandt and co-workers reported that systemic administration of N-acetyl-cysteine (NAC) is hypoxia-mimetic in humans, dramatically increasing hypoxic ventilatory response as well as circulating erythropoietin levels (29). They proposed that the SNO-transnitrosation mechanism from deoxyHb may be augmented by NAC; that is, that transfer of NO+ from the cys β 93 of Hb to NAC occurs, forming S-nitroso-N-acetyl cysteine (SNOAC)—a low-mass, cell-permeable SNO that can increase minute ventilation and HIF 1-dependent transcription (in this case, of erythropoietin). Consistent with these observations, we have found that NAC causes murine pulmonary hypertension that exactly recapitulates—in terms of pulmonary artery pressure, right to left heart weight ratio, and pulmonary vascular remodeling—hypoxia-induced murine pulmonary hypertension (5). It is increasingly apparent that NO/HIF 1 interactions may also be relevant to airways diseases. In asthma, for example, inducible NOS expression is upregulated (30) and airway pH appears to be low (31), particularly in the context of acute asthma exacerbations. Both of these conditions would favor SNO formation, though SNO catabolism also appears to be accelerated (32). This increased SNO flux (33) is of interest in light of evidence that HIF 1-dependent genes, including vascular endothelial growth factor (VEGF), are substantially up-

S-Nitrosothiol Signaling and Gene Regulation in Pulmonary Pathophysiology

325

regulated in the asthmatic airway, despite the presence of atmospheric oxygen tensions (34). This pattern of HIF-dependent gene expression is believed to play a role in airway “remodeling” in asthma. Paradoxically, the opposite extreme may be relevant in the CF airway. Oxygen metabolism is increased in the cystic fibrosis (CF) airway epithelium to the extent that epithelial surface pO2 concentrations in mucus approach zero (35). Moreover, virtually no iNOS is available to produce NO-mediated inhibition of classical hypoxic HIF 1 induction (34). Ironically, then, this environment also favors HIF 1 expression and VEGF transcription.

14.3 SPECIFICITY FACTORS 1 AND 3 Specificity factors bind to a GC rich motif in the promoter region of many genes, particularly those coding for constitutively expressed proteins, such as CF transmembrane regulatory protein (CFTR) (37). Four Sps (Sp1, Sp2, Sp3, and Sp4) have been characterized that can serve as gene activators or repressors. Indeed, competitive binding by one Sp can repress transcription by preventing binding of another. Remarkably, GSNO has substantial effects on Sp 1 and Sp 3 expression and binding. GSNO at concentrations up to 10 µM increases Sp 3 expression and binding, whereas at concentrations over 10 µM inhibit Sp 3 and increase Sp 1 expression and binding (37) (Figure 14.2). Of note, these effects are not exclusively mediated through changes in Sp expression; there appears also to be an effect on Sp-DNA interaction that may be mediated through nitrosylation of cysteine on the protein’s zinc finger. The net effect is that, when cells are exposed to physiological concentrations of GSNO, Sp 3-dependent transcription is favored; whereas conditions of nitrosative stress (iNOS up-regulation, airway acidification, and other conditions favoring increased SNO formation) lead to increased Sp 1 expression and binding and inhibit Sp 3 binding (37). These gene regulatory effects appear to have relevance to CF, and perhaps to other airways diseases in which CFTR may play a role. GSNO levels in the CF airway are lower than those in normal control airways (38). This may be the result of decreased airway epithelial iNOS expression or glutathione transport. Additionally, increased GSNO catabolism may be relevant, as it is in asthma (38–40). GSNO has several potential beneficial effects, including bronchodilatation, ventilation/perfusion matching, augmented ciliary motility, inhibition of amiloridesensitive sodium transport, and antimicrobial effects (39). It has also recently been discovered that physiological concentrations of GSNO increase expression, maturation and function of CFTR, both the wild type protein and the common mutant form, ∆F508 (38, 41, 42). Though this effect is primarily post-translational, a transcriptional component also exists (37, 41). Therefore, substantial interest exists regarding GSNO replacement as a therapy for CF. However, the effects of GSNO to enhance Sp 1 expression and binding at high concentrations (37), in addition to its potential to serve as an NO donor, augmenting CFTR tyrosine nitration and

326

Nitric Oxide, Cell Signaling, and Gene Expression GSNO (µm) C

0.5

1

10

100

500

SP1/SP3

SP3

Free probe

FIGURE 14.2 Effect of GSNO on Sp3 and Sp1 DNA-binding activity in A549 cells. Electrophoretic mobility shift assay using a consensus Sp3/Sp1 oligonucleotide as probe and nuclear extracts from control (lane 1) and GSNO-treated cells at concentration 0.5 to 500 µM (lane 2 to 6). Physiological GSNO concentrations increased Sp3 and Sp1 DNAbinding. Supraphysiological GSNO concentrations inhibited Sp3 binding but augmented Sp1 binding.

degradation (43), mean that it is important to achieve and maintain physiological GSNO levels in the airway and not to “overshoot” the dose. These S-nitrosylation effects may also be relevant to pulmonary hypertension. A variety of genes relevant to the development of pulmonary vascular remodeling have Sp consensus sites. Indeed, three weeks of systemic NAC treatment—as noted previously—mimics 3 weeks of hypoxia exposure in causing pulmonary hypertension and, in parallel, decreases whole lung levels of Sp 3, while increasing Sp 1 expression and binding.

14.4 NFκB NFκB is a transcription factor that has a central role in regulating both inflammation and cell survival (44). Its activity is modulated through its association with inhibitory κB (IκB); dissociation of NFκB from IκB allows nuclear localization of NFκB and initiation of transcription. The interaction of these proteins, in turn, is partly regulated by IκB phosphorylation catalyzed by IκB kinase. IκB kinase has three subunits (IKKα; IKKβ, and IKKγ). IKKβ is primarily responsible for phosphorylating IκB. Cysteine S-nitrosylation reactions appear to target several elements of this NFκB activation pathway, preventing NFκB–DNA binding. These mechanisms appear, at least in part, to serve as a counter-regulatory mechanism to decrease iNOS gene expression. Mechanisms have been described by which NOS-derived SNOs can: 1. Directly stabilize IκB, preventing its degradation (45) 2. Inhibit IKKβ (preventing IκB phosphorylation-induced degradation) (46) 3. Directly inhibit the p50 subunit and of NFκB, preventing DNA binding (47)

S-Nitrosothiol Signaling and Gene Regulation in Pulmonary Pathophysiology

327

Indeed, inhibition of NOS in lymphocytes (Jurkat cells) leads to increased NFκB binding, in part through activation of IKK. These observations are relevant to murine alveolar type 2 epithelial cells and to activation of leukocytes (46, 47). Further, NO-mediated inhibition of NFκB may be relevant to CF, where baseline decreased expression of iNOS may prevent this counter-regulatory mechanism in airway epithelial cells, allowing up-regulation of other NFκB dependent genes such as interleukin 8. NFκB is also critical to the patho-physiology of ARDS, pneumonia, and a variety of other inflammatory lung diseases.

14.5 SUMMARY S-Nitrosothiols are endogenous, metabolically regulated compounds formed in association with activation of all three isoforms of NOS. In general, they mediate cellular effects through transnitrosation reactions. This SNO/transnitrosation chemistry appears to be relevant to a variety of cGMP-independent cell signaling events, one of which is gene regulation. Several of these gene regulatory effects are relevant to pulmonary patho-physiology. In particular, GSNO derived from hemoglobin deoxygenation increases HIF 1α stability and HIF 1 activity in conditions of relative normoxia, such as those to which the pulmonary vascular endothelium is exposed during the development of pulmonary hypertension. Further, this GSNO-mediated HIF 1 activation may be involved in the patho-physiology of other pulmonary diseases, such as normoxic HIF 1-mediated VEGF up-regulation in asthmatic airway remodeling. Sp 1 and Sp 3 expression and binding have opposing regulatory effects under physiological conditions and conditions of nitrosative stress. These effects appear to be designed to maintain expression of housekeeping proteins in health, and to shift resources to stress response proteins in disease. These observations appear to be particularly relevant to CF because of the potential therapeutic effects of SNOs on CFTR. Finally, down-regulation of NFκB activity by a variety of S-nitrosylation mechanisms may be important to attenuation of inflammation in CF and other airways diseases. Each of these mechanisms, in one way or the other, can oppose a classic or cGMP-mediated NO effect. SNO synthesis, bioactivation, storage, and catabolism appear to be highly regulated to accomplish this level of specificity; however, a great deal of work remains to be done to clarify these SNO-mediated gene regulatory pathways in the lung and elsewhere.

14.6 ACKNOWLEDGMENTS The authors are supported by NIH grant no. 2 RO1 HL 59337; NIH Asthma Center Grant no. RO1 HL69170, CF Foundation grant no. Zaman04GO, and NIH/NHLBI grant no. HL68173.

328

Nitric Oxide, Cell Signaling, and Gene Expression

REFERENCES 1. Gaston BM, Carver J, Doctor A, Palmer LA. S-nitrosylation signaling in cell biology, Mol. Interv. 3:253–263, 2003. 2. Arnelle D, Stamler JS. NO+, NO• and NO− donation by S-Nitrosothiols: implications for regulation of physiological functions by S-Nitrosylation and acceleration of disulfide formation, Arch. Biochem. Biophys. 318:279–285, 1995. 3. Vieira H, Kroemer G. Mitochondria as targets of apoptosis regulation by nitric oxide. IUBMB Life. 55:613–616, 2003. 4. Mannick J, Hausladen A, Liu L, et al. Fas-induced caspase denitrosylation, Science 284:651–654, 1999. 5. Palmer L. N-acetyl cysteine induces pulmonary hypertension: role of S-nitrosothiols. (abstract) American Thoracic Society, 2005. 6. Hoehm T, Preston AA, McPhaden, AR, Stiller B, Vogel M, Buhrer C, Wadsworth RM. Endothelial nitric oxide synthase (NOS) is up regulated in rapid progressive pulmonary hypertension of the newborn. Int. Care Med. 29:1757–1762, 2003. 7. Palmer L, Gaston B, Johns R. Normoxic stabilization of hypoxia-inducible factor 1 expression and activity: redox-dependent effect of nitrogen oxides. Molec. Pharmacol. 58:1197–1203, 2000. 8. Gow AJ, Chen Q, Hess DT, et al. Basal and stimulated protein S-nitrosylation in multiple cell types and tissues, J. Biol. Chem. 277:9637–9640, 2002. 9. Gow A, Buerk D, Ischiropoulos H. A novel reaction mechanism for the formation of S-nitrosothiol in vivo, J. Biol. Chem. 272:2841–2845, 1997. 10. Rafikova O, Rafikov R, Nudler E. Catalysis of S-nitrosothiols formation by serum albumin: the mechanism and implication in vascular control, Proc. Nat. Acad. Sci. USA 99:5913–5918, 2002. 11. Jia L, Bonaventura C, Bonaventura J, et al. S-Nitrosohaemoglobin: a dynamic activity of blood involved in vascular control, Nature 380:221–226, 1996. 12. Inoue K, Akaike T, Iyamoto Y, et al. Nitrosothiol formation catalyzed by ceruloplasmin. Implication for cytoprotective mechanism in vivo, J. Biol. Chem. 274:27069–27075, 1999. 13. Lancaster JR, Gaston B. NO and nitrosothiols: spatial confinement and free diffusion, Am. J. Physiol. Lung Cell Mol. Physiol. 287(3):465–456, 2004. 14. Vanin A, Malenkova I, Serezhenkov V. Iron catalyzes both decomposition and synthesis of S-nitrosothiols: optical and electron paramagnetic resonance studies, Nitric Oxide: Biol. Chem. 1:191–203, 1997. 15. Pawloski J, Hess D, Stamler JS. Export by red blood cells of nitric oxide bioactivity, Nature 409:622–626, 2001. 16. Ramachandran N, Root P, Jiang SM, Hogg PJ, Mutus B. Mechanism of transfer of NO from extracellular S-nitrosothiols into the cytosol by cell-surface protein disulfide isomerase. Proc. Nat. Acad. Sci. USA 98:9539–9544, 2001. 17. Zhang Y, Hogg N. The mechanism of transmembrane S-nitrosothiol transport, PNAS 101:7891–7896, 2004. 18. Liu L, Yan Y, Zeng M, Zhang J, Hanes MA, Ahearn G, McMahon T, Dickfeld T, Marshall H, Que L, Stamler JS. Essential roles of S-nitrosothiols in vascular homeostasis and endotoxic shock. Cell 116:617–618, 2004. 19. Liu L, Hausladen A, Zeng M, et al. A metabolic enzyme for S-nitrosothiol conserved from bacteria to humans, Nature 410:490, 2001.

S-Nitrosothiol Signaling and Gene Regulation in Pulmonary Pathophysiology

329

20. Hogg N, Singh RJ, Konorev E, Joseph J, Kalyanaraman B. Biochem. J. 323:477–481, 1997. 21. Meyer D, Kramer H, Özer N, et al. Kinetics and equilibria of S-nitrosothiol-thiol exchange between glutaminase, cysteine, penicillamines and serum albumin, FEBS Lett. 345:177–180, 1994. 22. Scharfstein J, Jeaney, Jr., J, Slivka A, et al. In vitro transfer of nitric oxide between a plasma protein-bound reservoir and low molecular weight thiols, J. Clin. Invest. 94:1432–1439, 1994. 23. McQuillin LP, Leund GK, Marsden PA, Kostyl SK, Kourembanas S. Hypoxia inhibits expression of eNOS via transcriptional and post-transcriptional mechanisms. Am. J. Physiol. 267:1921–1927, 1994. 24. Semenza GL. O2-regulated gene expression: transcriptional control of cardiorespiratory physiology by HIF-1 J. App. Physiol. 96, 1173–1177, 2004. 25. Gaston B, Reilly J, Drazen JM, et al. Endogenous nitrogen oxides and bronchodilator S-nitrosothiols in human airways, Proc. Natl. Acad. Sci. USA 90:10957–10961, 1993. 26. Kluge I, Gutteck-Amsler U, Zollinger M, et al. S-nitrosoglutathione in rat cerebellum: identification and quantification by liquid chromatography-mass spectrometry, J. Neurochem. 69:2599–2607, 1997. 27. Sandau KB, Faus HG, Brune B. Biochem. Biophys. Res. Commun. 278:263–267, 2000. 28. Gow A, Luchsinger B, Pawloski J, et a;. The oxyhemoglobin reaction of nitric oxide, PNAS USA 96:9027–9032, 1999. 29. Lipton A, Johnson M, Macdonald T, et al. S-nitrosothiols signal the ventilatory response to hypoxia, Nature 413:171–174, 2001. 30. Hildebrandt W, Alexander S, Bartsch P, Droge W. Blood 99:1551–1555, 2002. 31. Howarth PH, Redington AE, Springall DR, Martin U, Bloom SR, Polak JM, Holgate ST. Epithelially derived endothelin and nitric oxide in asthma. Int. Arch. Allerg. Immunol. 107:228–230, 1995. 32. Hunt J, Fang K, Malik R, Snyder A, Malhotra N, et al. Endogenous airway acidification: implications for asthma pathophysiology, Am. J. Respir. Crit. Care Med. 161:694–699, 2000. 33. Gaston B, Sears S, Woods J, Hunt J, Ponaman J, McMahon T, Stamler J. Bronchodilator S-nitrosothiol deficiency in asthmatic respiratory failure. Lancet 351:1317–1319, 1998. 34. Dweik R, Comhair S, Gaston B, Thunnissen F, Farver C, Thomassen M, Kavuru M, Hammel J, Abu-Soud H, Erzurum S. NO chemical events in the human airway during the immediate and late antigen-induced asthmatic response. Proc. Natl. Acad. Sci. USA 98:2622–2627, 2001. 35. Asai K, Kanazawa H, Kamoi H, Shiraishi S, Hirata K, Yoshikawa J. Increased levels of vascular endothelial growth factor in induced sputum in asthmatic patients. Clin. Exp. Allerg. 33:595–599, 2003. 36. Worlitzsch D, Tarran R, Ulrich M, Schwab U, Cekici A, Meyer KC, Birrer P, Bellon G, Berger J, Weiss T, Botzenhart K, Yankaskas JR, Randell S, Boucher RC, Doring G. Effects of reduced mucus oxygen concentration in airway Pseudomonas infections of cystic fibrosis patients. J. Clin. Investig. 109:317–325, 2002. 37. Wooldridge JL, Deutsch GH, Sontag MK, Osberg I, Chase DR, Silkoff PE, Wagener JS, Abman SH, Accurso FJ. NO pathway in CF and non-CF children. Ped. Pulm. 37:338–350, 2004.

330

Nitric Oxide, Cell Signaling, and Gene Expression

38. Zaman K, Palmer LA, Doctor A, et al. Concentration-dependent effects of endogenous S-nitrosoglutathione on gene regulation by specificity proteins Sp3 and Sp1, Biochem. J. 380(Pt 1):67–74, 2004. 39. Grasemann H, Gaston B, Fang K, Ratjen F. Decreased levels of nitrosothiols in the lower airways of patients with cystic fibrosis and normal pulmonary function. J. Pediatr. 135:770–772, 1999. 40. Snyder A, McPherson ME, Hunt JF, Johnson M, Stamler JS, Gaston B. Acute effects of aerosolized S-nitrosoglutathione in cystic fibrosis. Am. J. Respir. Crit. Care Med. 165:922–926, 2002. 41. Gaston B, Sears S, Woods J, Hunt J, Ponaman M, McMahon T, Stamler J. Bronchodilator S-nitrosothiol deficiency in asthmatic respiratory failure. Lancet 351:1317–1319, 1998. 42. Zaman K, McPherson M, Vaughan J, Hunt J, Mendes F, Gaston B, Palmer L. Snitrosoglutathione increases cystic fibrosis transmembrane regulator maturation. Biochem. Biophys. Res. Commun. 284:65–70, 2001. 43. Howard M, Fischer H, Roux J, Santos B, Gullans S, Yancey P, Welch W. Mammalian osmolytes and S-nitrosoglutathione promote ∆F508 cystic fibrosis transmembrane conductance regulator (CFTR) protein maturation and function. J. Biol. Chem. 278:35159–35167, 2003. 44. Jilling T, Haddad IY, Cheng SH, Matalon S. Nitric oxide inhibits heterologous CFTR expression in polarized epithelial cells. Am. J. Physiol. 277:89–96, 1999. 45. Karin M, Ben-Neriah Y. Phosphorylation meets ubiquitination: the control of NFκB activity. Annu. Rev. Immunol. 18:621-663, 2000. 46. Peng H, Libby P, Liao J. Induction and Stabilization of IkBa by Nitric Oxide Mediates Inhibition of NF-κB. J. Biol. Chem 270:14214–14219, 1995. 47. Reynaert N, Ckless K, Korn S, Vos N, Guala A, Wouters E, van der Vliet A, Janssen-Heininger YMW. Nitric oxide represses inhibitory κB kinase through Snitrosylation. PNAS 101:8945–8950, 2004.1

Oxide and Gene 15 Nitric Expression Josef Pfeilschifter and Karl-Friedrich Beck Pharmazentrum Frantfurt/ZAFES, Klinikum der Johann Wolfgang Goethe-Universität, Frankfurt, Germany

CONTENTS 15.1 Introduction ............................................................................................331 15.2 The Role of Guanylyl Cyclase in NO-Mediated Gene Expression .....332 15.2.1 Transcription Factors That Are Directly Activated by cGMP ..334 15.2.2 “Immediate Early Gene” Transcription Factors That Are Induced by cGMP ......................................................................337 15.2.3 Interference of the cGMP/PKG Pathway with Other Signaling Cascades That Lead to Changes in the Gene Expression Pattern ..........................................................................338 15.2.3.1 Interference with the cAMP/PKA Pathway ................338 15.2.3.2 Interference with the MAP Kinase Mediated Pathways ......................................................................338 15.2.3.3 Cross Talk between cGMP and the Calcineurin NF/AT Pathway ............................................................339 15.2.3.4 Effects of cGMP Signaling on the Expression of Pro-Inflammatory Genes .........................................339 15.2.3.5 Effects of cGMP on the Expression of Genes Involved in cGMP Signaling .......................................339 15.2.3.6 Effects of cGMP on mRNA Stability .........................340 15.3 The Role of NO in Hypoxia-Mediated Gene Expression ....................341 15.4 NO: A Redox Signaling Paradigm ........................................................342 15.4 Conclusion .............................................................................................344 References ...............................................................................................345

15.1 INTRODUCTION In the late 1980s, nitric oxide (NO) was identified as a signaling molecule that triggers cGMP-dependent smooth muscle relaxation and as a cytotoxic agent that potently kills invading microorganisms. In the last decade, however, it has 331

332

Nitric Oxide, Cell Signaling, and Gene Expression

become more evident that NO is a versatile mediator that not only triggers the protective activation of the soluble guanylyl cyclase in the picomolar range or exerts cytotoxic effects when synthesized in nanomolar quantities (e.g., in the course of inflammatory processes), but it is able to initiate well-coordinated signaling pathways that are followed by changes in the gene expression pattern of a target cell. The immense progress in messenger ribonucleic acid (mRNA) or protein-based techniques to analyze differential gene expression enabled us to identify a series of genes that are under expressional control of NO. But the aim of these studies, namely to define a more or less unique and ubiquitous signaling machinery that triggers NO-mediated gene expression (1) has been, so far, not attained. Moreover, the analysis of NO-modulated mRNA and protein expression revealed that nearly all signaling cascades known to date are involved in NO-evoked cell responses. Nevertheless, our knowledge of NO signaling enables us to identify three different pathways that are involved in triggering NO-driven responses directly to the transcriptional machinery of the cell or in mediating post-transcriptional or post-translational mechanisms and, subsequently, changes in the gene expression pattern. These pathways include: 1. The cGMP signaling pathway 2. Interference with hypoxia-mediated signaling 3. Interference with redox signaling (e.g., post-translational changes of proteins by nitration of tyrosine residues or nitrosation of cysteine residues) (2). A compilation of NO-regulated genes and the main mechanisms that may account for the NO-mediated effects are presented in Table 15.1 and Figures 15.1–15.3, respectively.

15.2 THE ROLE OF GUANYLYL CYCLASE IN NOMEDIATED GENE EXPRESSION The physiologically most relevant action of NO is the activation of the soluble guanylyl cyclase by binding to the enzyme’s heme moiety. The subsequent increase in cGMP levels alters the activity of three main target proteins (3, 4): 1. cGMP-dependent protein kinases 2. cGMP-regulated phosphodiesterases 3. cGMP-regulated ion channels Importantly, high concentrations of cGMP can also activate cAMP-dependent protein kinases. Therefore, NO is also, at least in part, able to use the cAMP signaling cascade for mediating gene expression (5). To date, most actions of cGMP on gene expression are considered to be mediated via cGMP-dependent

Nitric Oxide and Gene Expression

333

RGC

NO

Natriuretic peptides

PDE’s

sGC cGMP

ion channels PKG 1 PKG 2

PKG 1 CREB Gene transcription

FIGURE 15.1 Nitric oxide (NO) elevates cGMP levels by activation of the soluble guanylyl cyclase (sGC) and natriuretic peptides do so by activation of the membrane-bound receptor guanylyl cyclase (RGC). Both pathways affect phosphodiesterases (PDEs), ion channels, and protein kinases G (PKG 1, PKG 2). PKG 1 is able to enter the nucleus and phosphorylates transcription factors (e.g., CREB).

NO

Prolyl hydroxylases O2

Protein nitrosation HIF-1α destabilisation N2O3

HIF-1α Gene transcription

Cytochrome c oxidase

FIGURE 15.2 Nitric oxide (NO) is able to mimic hypoxia signaling by affecting HIF-1α stability due to the inhibition of prolyl hydroxylases. By contrast, NO is able to antagonize hypoxia by inhibiting the mitochondrial cytochrome c oxidase resulting in an increase in cellular O2 availability and a subsequent decrease in HIF-1α stability. Elevated O2 levels facilitate the reaction with NO to form N2O3, a potent nitrosating agent.

334

Nitric Oxide, Cell Signaling, and Gene Expression

NFκB, AP-1

NO ONOO− O2−

Gene transcription NFκB, AP-1

FIGURE 15.3 Nitric oxide (NO) and superoxide anion (O2−) neutralize each other and react to form peroxynitrite (ONOO−) that potently causes protein nitration.

protein kinase 1 (PKG 1) instead of PKG 2 or by cGMP-regulated ion channels. Nevertheless, effects of cGMP on different phosphodiesterases (PDEs) may account for an additional level of cross talk between cGMP and cAMP signaling events. Natriuretic peptides, such as ANP, BNP, and CNP, elevate cGMP levels independently from NO via activation of the particular guanylyl cyclase and a possible cross talk between these peptides and NO has additionally to be considered.

15.2.1 TRANSCRIPTION FACTORS THAT ARE DIRECTLY ACTIVATED BY CGMP Many genes that are regulated by cyclic nucleotides bear one or more so-called cyclic AMP responsive elements (CRE) in their promoter region. This element has originally been reported to bind the transcription factor CRE binding protein (CREB) in response to elevated cAMP levels after phosphorylation of Ser133 (6, 7). As indicated for the induction (e.g., of the c-fos promoter in vascular smooth muscle cells [VSMC]), however, CREB is also phosphorylated at Ser133 after translocation of PKG I to the nucleus (8–10). Interestingly, in neuronal and osteogenic cells that lack PKG I but express PKG 2, cGMP acts synergistically with calcium to induce the c-fos promoter. This effect is not directly mediated by PKG II because this kinase, in contrast to PKG I, is obviously not able to enter the nucleus. Chen et al. (11) have reported that a synergism between calcium and cGMP leads to a cooperation of CREB and CAAT/enhancer binding protein β (C/EBP-β) that mediates c-fos expression. Another transcription factor that is directly phosphorylated by elevated cGMP levels is the CRE-binding protein ATF-1. This has been reported in VSMC for the induction of the Rho A promoter (9). A further transcription factor that can be affected by the cGMP/PKG signaling cascade is nuclear factor κB (NFκB). This important mediator of gene transcription is activated by the phosphorylation and degradation of its naturally

Nitric Oxide and Gene Expression

335

TABLE 15.1 Compilation of NO-Regulated Gene Products Transcription Cross Cross Factor Talk with Talk with (Mechanism) ROS Hypoxia Probably n.d. n.d. yes n.d. n.d. n.d.

Gene Adrenomodullin

Up/ Down ↑

cGMP Signaling No

ANP

↓

Yes

AT-1 receptor

↓

No

BNP

↓

Yes

Probably yes n.d.

C-fos

↑

Yes

Collagen

↓

n.d

COX-2

↑↓

CuZn/SOD

Ref. 94 38

n.d.

Transcriptional

95

n.d.

NF-AT

29

n.d.

n.d.

CREB, TF-II-I

8, 18

n.d.

n.d.

n.d.

96

Yes

n.d.

Yes

32, 37

↑

No

n.d.

n.d.

NFκB (early) NFκB (late) Transcriptional

Cyclin A

↓

Yes

n.d.

n.d.

Transcriptional

97

ecSOD

↓

Yes

n.d.

n.d.

98

egr-1

↑↓

No

n.d.

n.d.

p38MAP kinasedependent Transcriptional

Erythropoetin

↓

n.d

n.d.

n.d.

99

E-selectin

↓

n.d

Probably yes n.d.

n.d.

n.d.

100

ET-1

↓

n.d

Fibronectin

↓

FLT-1 FMR1

76

24, 25

n.d.

Transcriptional

101

n.d

Probably yes n.d.

n.d.

n.d.

96

↓

Yes

n.d.

n.d.

n.d.

102

↓

n.d

n.d.

n.d.

CpG methylation

92

↑ Gammaglutamylcysteine synthetase Heme oxigenase 1 ↑

n.d

n.d.

n.d.

Transcriptional

103

No

n.d.

HIF-1

104

HPRT

↓

n.d

Probably yes n.d.

n.d.

CpG meth.

92

HuR

↓

Yes

n.d.

n.d.

n.d.

42, 46

ICAM–1

↓

No

n.d.

n.d.

105

IκB

↑

n.d

n.d.

Probably not n.d.

n.d.

85

IL-12 (p40)

↑

n.d

n.d.

n.d.

n.d.

106

IL-8

↑

n.d

n.d.

n.d.

NFκB, NF-IL-6

107

ILK

↓

No

n.d.

n.d.

n.d.

108

336

Nitric Oxide, Cell Signaling, and Gene Expression

TABLE 15.1 Compilation of NO-Regulated Gene Products (continued) Transcription Cross Cross Factor Talk with Talk with (Mechanism) ROS Hypoxia Probably Probably yes yes n.d. n.d. Post-transcriptional

Ref. 31, 36, 67,109 47

n.d

n.d.

n.d.

n.d.

96

No

n.d.

n.d.

110

↓

No

n.d.

n.d.

MHC class II

↓

n.d.

n.d.

n.d.

Inhibition of NFκB activity Inhibition of NFκB activity n.d.

112

MIP-1α

↑

n.d.

n.d.

n.d.

n.d.

113

MIP-2

↑

No

n.d.

NFκB

65, 66

MKP-1

↑

Yes

Probably yes n.d.

n.d.

n.d.

114

MMP-2

↓

n.d.

n.d.

n.d.

ATF-3

115

MMP-9

↓

Yes

n.d.

n.d.

mRNA stability (HuR)

34. 46

MnSOD

↑

Yes

n.d.

n.d.

PAI-1

↓

n.d

n.d.

n.d.

n.d.

33

PDGF-B

↓

n.d

n.d.

PKG-1α

↓

Yes

Probably yes n.d.

n.d.

rGC-A

↓

Yes

n.d.

n.d.

RhoA

↑

Yes

n.d.

n.d.

Glucocorticoid receptor sGC

↑

n.d.

n.d.

No?

↓

Yes

n.d.

n.d.

SPARC

↓

No

n.d.

n.d.

117

sPLA2

↑

No

n.d.

n.d.

118

TGF-β

↓

No

n.d.

n.d.

119

TIMP-1

↓

No

n.d.

n.d.

TNF-α

↑

Yes

n.d.

n.d.

t-PA

↓

No

n.d.

n.d.

Up/ Down ↑↓

cGMP Signaling Yes/No

KCl-cotransporter 3 Laminin

↑

Yes

↑

MCP-1

↓

M-CSF

Gene iNOS

111

35

101 Sp1 (PKAdependent) n.d. ATF-1, cAMPresponsive element GR (nuclear localization) mRNA stability (HuR)

26 39, 40 9 116 41, 42

120 Sp1

14, 30, 80, 121 122

Nitric Oxide and Gene Expression

337

TABLE 15.1 Compilation of NO-Regulated Gene Products (continued)

Gene VCAM-1

Up/ Down ↓

cGMP Signaling No

VEGF

↑

No

Xanthine oxidase

↓

No

Transcription Cross Cross Factor Talk with Talk with (Mechanism) ROS Hypoxia n.d. n.d. NFκB Probably yes n.d.

n.d.

HIF, NFκB

Yes

Ref. 85, 123, 124 64, 102 125

Note: n.d., not determined; AT-1, angiotensin II type 1 receptor; EcSOD, extracellular SOD; FLT1, fms-like-tyrosine kinase; ICAM-1, endothelial intercellular adhesion molecule-1; ILK, integrinlinked kinase; MCP-1, monocyte chemoattractant protein-1; M-CSF, macrophage colony-stimulating factor; MIP, macrophage inflammatory protein; PDGF-B, platelet-derived growth factor (B chain); SPARC, secreted protein acidic and rich in cysteine; sPLA2, secretory phospholipase A2; TGF-β, transforming growth factor β; t-PA, tissue plasminogen activator; VCAM-1, vascular adhesion molecule-1; VEGF, vascular endothelial growth factor.

occurring inhibitor IκB. Degradation of IκB then allows NFκB to enter the nucleus and to activate the transcription of several genes, most of them involved in inflammatory processes (12). PKG 1 directly phosphorylates and thereby activates the p50 and p65 subunits of NFκB (13); however, most of the actions of cGMP on NFκB-mediated gene expression are more indirect and finally lead to an activation or inhibition of NFκB signaling. In cardiac myocytes, PKG phosphorylates IκB leading to NFκB-driven TNF-α expression (14). By contrast, a stabilization of IκB by cGMP-elevating reagents was also observed in cytokine-stimulated human endothelial cells and in a rat model of hepatic ischemia/reperfusion injury (15, 16). Remarkably, the cGMP-elicited PKG I activity phosphorylates the transcriptional regulator TFII-I. This factor lacks a functional DNA binding domain but it co-activates a series of transcription factors (17). This factor also plays a role in co-activating cGMP-induced c-fos expression (18).

15.2.2 “IMMEDIATE EARLY GENE” TRANSCRIPTION FACTORS THAT ARE INDUCED BY CGMP Activator protein 1 (AP-1) is a transcription factor that mediates not only the processes of proliferation and differentiation, but also tumor progression (19). The transcription factor consists of two subunits denoted as c-Jun and c-Fos that are dramatically up-regulated by different stimuli, in particular by mitogens. CFos and JunB dimerization and subsequently binding of AP-1 to the respective promoter elements are enhanced when cells are exposed to elevated cGMP levels (20, 21). Moreover, the lack of the NO/cGMP signaling pathway as observed

338

Nitric Oxide, Cell Signaling, and Gene Expression

(e.g., in endothelial NO synthase [eNOS] knock-out mice) results in a diminished c-fos mRNA expression in cardiac tissue (22). In some cell types, additional costimuli, such as angiotensin II (AngII) or tumor necrosis factor-α (TNF-α), are required to enhance cGMP-mediated AP-1 activity (23). Another important transcription factor that is mainly involved in cell proliferation is Egr-1. As described for c-Jun and c-Fos, binding of this zinc-finger transcription factor to its respective responsive element strictly depends on newly synthesized Egr-1 protein. In pheocytochroma cells, cGMP elevating agents induce Egr-1 activity (24). Interestingly, the NO-releasing compound S-nitrosoglutathione (GSNO) attenuates proliferation and inhibits Egr-1 expression and activity in rat renal mesangial cells (25). This NO-mediated effect is probably independent of cGMP and highlights the often different effects of NO and cGMP-elevating agents on gene transcription.

15.2.3 INTERFERENCE OF THE CGMP/PKG PATHWAY WITH OTHER SIGNALING CASCADES THAT LEAD TO CHANGES IN THE GENE EXPRESSION PATTERN 15.2.3.1 Interference with the cAMP/PKA Pathway Because elevated cGMP levels act on the activity of phosphodiesterases, which in turn alter cellular cAMP levels, it is not surprising that the cGMP signaling pathway partially overlaps with the cAMP/PKA cascade. Since cGMP differentially affects different phosphodiesterases by stimulation or inhibition, a cGMP-dependent alteration of cAMP levels into both directions is possible. As mentioned previously, PKG I is able to phosphorylate CREB although with a lower potency when compared with PKA. Therefore, we cannot exclude that cGMP also mediates CREB phosphorylation by the cAMP/PKA pathway. Moreover, elevated cAMP levels can influence the cGMP signaling pathway by affecting another important mediator of this cascade. Sellak et al. (26) observed a down-regulation of the PKG Iα form by cyclic nucleotides and NO in bovine VSMC, and in embryonic VSMC that is mediated by decreased binding of the transcription factor SP1 to a tandem repeat of SP1 responsive elements within the core promoter region of the PKG Iα gene. They further demonstrated that inhibition of PKG Iα promoter activity is reversed by the PKA inhibitor KT5823 indicating a role for cAMP in cGMP-mediated signaling. Interestingly, NO mediates down-regulation of PKG Iα promoter activity independently of cGMP, thus providing a further example of NO acting both in a cGMP-dependent and independent manner on the expression of the same gene. 15.2.3.2 Interference with the MAP Kinase Mediated Pathways All three classical MAP kinase pathways, namely the p38, the JNK, and the Erk1/Erk-2 cascade, are affected by changes in cGMP levels. Because these pathways directly or indirectly phosphorylate transcription factors (27), it is obvious that

Nitric Oxide and Gene Expression

339

cGMP crosstalks with the MAPK pathways that in turn lead to changes in the gene expression patterns. For example, elevated cGMP levels induce the expression of MAP kinase phosphatase in glomerular mesangial cells leading to a reduced activity of Erk-1 and Erk-2 (28). Further examples for this topic are reviewed in Reference 2. 15.2.3.3 Cross Talk between cGMP and the Calcineurin NF/AT Pathway Recently, Fiedler et al. reported that the cGMP/PKG 1 pathway inhibits hypertrophy in cardiac myocytes by affecting calcineurin-NFAT signaling (29). They observed that the brain natriuretic peptide (BNP) expression is suppressed by NO via a PKG I-dependent inhibition of NFAT activation. They further demonstrated that this effect is due to a reduced open probability of single L-type Ca2+ channels. Taken together, this indicates a valuable role for NO and other cGMP elevating agents in the reduction of cardiac hypertrophy. 15.2.3.4 Effects of cGMP Signaling on the Expression of ProInflammatory Genes cGMP elevating agents act potently on the expression of genes involved in inflammatory processes. For example, the potent inflammatory cytokine tumor necrosis factor α (TNF-α), the inducible NO-synthase (iNOS), the cyclooxygenase 2 (COX-2), the plasminogen activator inhibitor-1 (PAI-1) the matrix metalloproteinase 9 (MMP-9), and the manganese superoxide dismutase (MnSOD, SOD-2) are under the control of the cGMP signaling pathway in different cell types (14, 30–35). In most cases, cGMP acts not at all or only weakly on the basal expression of these genes, but it potently amplifies their expression induced by inflammatory cytokines such as interleukin 1β (IL-1β). Interestingly, the effects of cGMP on the expression of inflammatory genes often occur in a biphasic manner. As exemplified for iNOS, COX, and MMP-9, some of these genes are co-induced by inflammatory cytokines and NO/cGMP at early time-points at the transcriptional level. After longer incubation periods, however, elevated cGMP levels decrease the stability of the respective mRNA species (34, 36, 37), indicating a post-transcriptional mechanism involved in cGMP signaling (see below). 15.2.3.5 Effects of cGMP on the Expression of Genes Involved in cGMP Signaling Elevated cGMP levels control phosphodiesterases that regulate the bioavailability of cyclic nucleotides in a well-coordinated manner; however, the cGMP mediated signaling pathways are also able to change the expression of important genes that are related to the generation or the action of cGMP. As mentioned previously, cGMP triggers the down-regulation of BNP expression, a potent activator of particulate GC, via suppression of the transcription factor NFAT

340

Nitric Oxide, Cell Signaling, and Gene Expression

activity, indicating that cGMP has the potency to antagonize its own synthesis (29). Moreover, a negative effect of elevated cGMP levels on ANP expression has also been described (38). As also discussed previously, NO and cyclic nucleotides suppress PKG Iα expression by targeting the transcription factor Sp1, indicating that cGMP also is able to regulate an important element of its downstream signaling cascade, which may serve to attenuate cGMP- triggered signals. Interestingly, both genes that code for cGMP synthesizing enzymes are affected by the cGMP signaling pathway. It has been demonstrated in vitro and in vivo that the expression of the receptor for natriuretic peptides rGC-A is transcriptionally repressed by cGMP and that knockout mice lacking a functional gene for the rGC-A ligand ANP express higher levels of the rGC-A receptor compared with non-genetically manipulated controls (39, 40). Furthermore, the α1 and β1 subunits of the soluble guanylyl cyclase (sGC) are also expressionally affected by cGMP. In contrast to the cGMP-dependent regulation of rGC-A, the sGC subunits are down-regulated by a post-transcriptional mechanism (41, 42). 15.2.3.6 Effects of cGMP on mRNA Stability As previously mentioned, elevated cGMP levels affect a series of genes by a destabilization of their respective mRNAs. The genes regulated post-transcriptionally by elevated cGMP levels, such as MMP-9, iNOS, COX2, sGC, TNFα, and TGF-β, possess AU-rich motifs in their 3′-untranslated regions that could be targeted by mRNA stabilizing or destabilizing protein factors (34, 36, 37, 41–44). As observed recently, a major role in the cGMP-induced destabilization of these mRNAs plays the mRNA binding protein HuR that belongs to the ELAV family of mRNA binding proteins (45). In rat mesangial cells NO and 8-Bromo-cGMP cause a down-regulation of HuR mRNA and protein levels and this subsequently leads to a decreased binding of HuR to the 3′UTR of the MMP-9 mRNA (46). This suggests that the down-regulation of HuR might be a ubiquitous mechanism for cGMP-mediated gene expression. As indicated for the potassium chloride cotransporter-3, cGMP-mediated signals can also stabilize mRNA, but the molecular mechanisms are, so far, not completely understood (47). In summary, cGMP is not only an acutely acting vasodilating agent, but it has also a sustained and long-lasting impact on gene expression. cGMP acts directly or indirectly on transcription factors, affects mRNA stabilization or interferes with signaling pathways to modulate transcription of genes involved in physiological and pathophysiological processes such as vascular tone, differentiation, cell proliferation, inflammation, apoptosis, and cardiac hypertrophy. Importantly, elevated cGMP levels regulate genes that are involved in cGMP formation or cGMP-mediated signal transduction, indicating that cGMP signaling is a fine-tuned process that includes a series of feedback mechanisms that guarantee physiological cGMP levels in a healthy cell.

Nitric Oxide and Gene Expression

341

15.3 THE ROLE OF NO IN HYPOXIA-MEDIATED GENE EXPRESSION Nearly three decades ago, long before the important function of NO was known, it became evident that NO binds to cytochrome c oxidase. This mitochondrial protein represents the terminal member of the respiratory chain proteins (48–50); however, a physiological role for this reversible process was demonstrated only 10 years ago (51). Inhibition of cytochrome c by NO leads to an insufficient mitochondrial oxygen consumption and therefore provides elevated oxygen availability in the cytosol. The consequence of this pathway was recently reported in a very impressive article suggesting that the elevated oxygen levels resulting from an inhibition of the respiratory chain by NO are responsible for the bioluminescent flashing of the firefly (52). More recently, Hagen et al. (53) demonstrated that a NO-induced redistribution of intracellular oxygen during hypoxia might interfere with the hypoxiainduced signaling cascade. Meanwhile, great efforts have been made to elucidate both the hypoxia induced and the NO induced signaling pathways, and it became clear that these signaling cascades extensively cross-communicate with each other. The terminal target of hypoxia is the transcription factor HIF-1α that is rapidly degraded under normoxic conditions. Inhibited degradation of HIF-1α as observed under hypoxic conditions leads to a dimerization with HIF-1α’s binding partner HIF-1β that drives the transcription factor into the nucleus where it controls the expression of genes, that in particular are involved in energy metabolism, angiogenesis, and erythropoiesis. The von Hippel–Lindau protein (pVHL) plays a crucial role in the process of HIF-1α degradation. Binding of HIF-1α to pVHL is necessary to initiate its degradation by the ubiquitin-proteasome system and HIF-1α has to be hydroxylated by prolyl hydroxylases, which function as the oxygen sensors of a cell to be bound by pVHL (54, 55). Under hypoxic conditions, the activity of prolyl hydroxylases is decreased, and consequently, HIF-1α is stabilized and enabled to form transcriptionally active heterodimers with HIF-1β. The complex signaling mechanism of oxygen sensing is affected at three important switches by NO, namely the cytochrome c oxidase (50, 53), the HIF-1α subunit (56, 57), and the prolyl hydroxylases (58). The activity of cytochrome c oxidase is inhibited by NO simply by competition with oxygen (59). The effects of NO on the HIF-1α subunit are much more complex. Because different reactive nitrogen species derived from different NO donors exert in part adverse effects on target molecules, Mateo et al. (60) performed experiments with stably transfected cells that bear the iNOS gene under the control of a tetracycline inducible promoter. They found a biphasic effect of NO on HIF-1α accumulation and destabilization. NO concentrations below 400 nM led to a decreased stability of HIF-1α. By contrast, concentrations of NO above 1 µM stabilize the HIF-1α protein. Moreover, the effects of low concentrations of NO are dependent on the respiratory chain, whereas high amounts of NO appear to directly affect HIF-1α (e.g., by S-nitrosation) (61).

342

Nitric Oxide, Cell Signaling, and Gene Expression

The third currently known group of targets includes the prolyl hydroxylases that are inhibited in their activity by the NO donor S-nitrosoglutathione, which in turn stabilizes HIF-1α. Taken together, NO acts differentially and on different switches of the oxygen sensing system, and thus alters hypoxia-induced gene expression. Not surprisingly, a series of genes are regulated by NO and hypoxia (2). One of the first genes found to be up-regulated by hypoxia was VEGF (62) and this effect of hypoxia can be mimicked by NO (63, 64). Other well-studied examples for NO and hypoxia-mediated gene expression are the chemokine MIP-2 (65, 66) or iNOS (67, 68). Most of the genes coregulated by oxygen and NO bear a HIF-1 binding site in their 5′−flanking but, as reported for MIP-2, hypoxia can also affect the NFκB pathway (68). The upregulation of iNOS via a HIF-1 site (65) further documents the cross talk between NO and oxygen sensing pathways. In summary, NO is able to partly mimic hypoxia and therefore to drive gene expression, mostly by stabilizing HIF1α. Interestingly, aside from HIF-1α, other transcription factors, such as AP-1, Egr-1, or NFκB, are regulated by both hypoxia and NO (reviewed by Semenza [69]). Moreover, NO acts through mitochondria-dependent and -independent mechanisms on hypoxia-mediated gene transcription and the inhibition of cytochrome c by NO may also cause an elevated availability of oxygen in the cytosol that may counteract hypoxia-driven gene expression (53, 70). This possibility has to be considered by the analysis of hypoxia and NO-mediated gene expression. Moreover, increased levels of molecular oxygen facilitate the formation of N2O3, which is a potent nitrosating agent, that is able to modulate gene expression by direct modification of transcription factors (see below).

15.4 NO: A REDOX SIGNALING PARADIGM The role of NO in gene transcription involves redox signaling pathways and is rather complex because NO can act both as an antioxidant as well as an oxidant dependent on the presence of reactive oxygen species (71). Similar to hypoxia, the action of NO is strictly determined by the microenvironmental conditions surrounding a target cell. Under physiological conditions, NO production is restricted to the activity of the constitutive isoforms of NOS (eNOS, bNOS) that produce only physiologic concentrations of NO. As discussed previously, these low amounts of NO mediate in particular the cGMP-dependent signaling cascades. In this setting, only low amounts of ROS are produced that are rapidly degraded by superoxide dismutases and catalase, and therefore, NO is only marginally neutralized by ROS (72, 73). In the state of inflammation, invading monocytes/macrophages and resident cells, such as renal mesangial cells, vascular smooth muscle cells, and pancreatic islet cells, are forced by cytokines to produce high amounts of NO and ROS. This shifts the balance from the mainly protective NO/cGMP signaling pathway to a redox-driven setting, which may result in cell death by necrosis or, in a more coordinated manner, by apoptosis. Cells that are able to cope with this toxic environment respond with dramatic changes in their gene expression pattern.

Nitric Oxide and Gene Expression

343

Importantly, ROS and NO are, in large part, not produced simultaneously during inflammation. ROS generation is mediated generally by a rapid activation of ROS producing enzymes, such as NADPH oxidases or xanthine oxidases that precedes the generation of NO, which in turn requires the de novo protein synthesis by a transcriptional induction of the iNOS gene by cytokines. Therefore, in the course of inflammation an early ROS-dominated state that affects gene transcription independently from NO (74) precedes a subsequent increase of NO following the induction of iNOS and results in the formation of the potent oxidant peroxynitrite. Peroxynitrite is generated by the reaction of NO with O2− and is able to nitrate proteins on tyrosine residues. Nitrosation of proteins on thiol groups of cysteine catalyzed by N2O3, another oxidation product of NO, often alters the function of target proteins and abolishes the activity of targeted enzymes (reviewed in Reference 75). Interestingly, high levels of iNOS nitrated on tyrosine residues were found in patients with sepsis; however, whether tyrosine-nitration targets specific proteins and constitutes a relevant signaling pathway that leads to changes in gene expression is to date not clear. In the further course of inflammation, ROS formation decreases and this coincides with the up-regulation of the ROS decomposing enzymes CuZn-SOD and Mn-SOD (SOD1 and SOD2, respectively (35, 76), whereas NO is further produced and is no longer scavenged by O2−. In this setting, NO is again able to exert its mainly protective effects by cGMP-signaling; however, in contrast to the physiological state where NO is exclusively produced by the constitutive NOS isoforms (eNOS and nNOS), the huge amounts of NO generated by iNOS are able to trigger cGMP-independent effects. S-nitrosation of cysteine residues can occur in the presence of molecular oxygen by the generation of nitrogen oxides (77), but more importantly, transition metals are facilitating nitrosation of proteins on their cysteine residues. In contrast to protein nitration, this reaction is reversible and a physiological role is well documented, most importantly in modulating the activity of transcription factors. To date a series of transcription factors are directly affected by NO. Transcription factors of the zinc finger type are inhibited in their DNA-binding activity by the displacement of zinc. This was first described for the yeast transcription factor LAC9 (78). Meanwhile a similar mechanism has been also reported for other transcription factors in higher eucaryotes such as Egr-1 and SP-1 (reviewed in Reference 79). Interestingly, when of SP-1 acts as a repressor, inhibition of SP-1 can lead to an increased gene transcription as indicated for the NO-stimulated TNF-α gene expression (80). Nuclear receptors of the zinc finger type that are inactivated by NO are the vitamin D3 receptor (VDR9) and the peroxisome proliferator-activated receptor γ (PPARγ) (81, 82). Another important group of NO regulated transcription factors are proteins that bear on ore more cysteine groups near to the DNA-binding domain. In particular, the effects of NO on NFκB, a transcription factor that is mainly involved in inflammatory processes has been intensively investigated. NO is able to Snitrosate the p50 subunit of NFκB on cysteine 62 that diminishes its activity (83, 84); however, as exemplified for the expression of VCAM-1 expression (85), NO is also able to inhibit activation by an increased expression of its natural inhibitor IκB. By contrast, activation of NFκB-driven gene expression has also been

344

Nitric Oxide, Cell Signaling, and Gene Expression

described. For example, in a mouse hemorrhagic shock model NFκB activity is enhanced by endogenous NO formation (86) and NO-mediated signaling events that lead to an activation of NFκB have been reported (87–89). Another important transcription factor that is modulated in its activity is AP-1. Interestingly, AP-1 can be directly affected by NO via S-nitrosation of the c-Jun and c-Fos subunits (90) as well as changes in the expression of c-Fos or JunB mRNA. In particular, c-Fos mRNA expression can be up- or down-regulated by NO depending on the cell type or the stimulation protocol used for the experiments (reviewed in Reference 91). Processes that alter the structure of the chromatin and therefore precede the specific binding of transcription factors have also been reported to be modulated by NO. NO is able to activate DNA methyltransferase. The NOinduced methylation of so-called CpG islands in the genomic DNA flanking the genes for fragile X mental retardation gene (FMR1) and hypoxanthine phosphoribosyltransferase (HPRT) is responsible for an enhanced chromatin condensation in this genomic region and therefore, these genes are silenced (92). Another example for the transcription factor independent suppression of gene expression by NO was reported for the regulation of the murine κ opioid receptor (KOR). A down-regulation of c-Myc by NO leads to a diminished recruitment of histone acetyltransferases, leading to a hypoacetylation of the KOR promoter and, therefore, to a decreased transcriptional activity (93).

15.4 CONCLUSION In summary, NO is a highly versatile signaling molecule and its ability to mediate gene expression is not restricted to the well-coordinated cGMP signaling cascade. The unpaired electron allows covalent modification of a series of biomolecules and this further potentiates the spectrum of actions of NO. Importantly, the effects of NO are strictly dependent from the microenvironmental conditions, such as the availability of oxygen and reactive oxygen species that enable NO to affect hypoxia sensing and the redox state of a cell, respectively. Therefore, it is not surprising that the huge pool of data published within the last decade that deals with the action of NO on gene expression appears often contradictory. Actually, this reflects the ability of NO to activate simultaneously or, as exemplified for the regulation of the cox-2 gene and other genes in a temporally well coordinated manner, different signaling cascades that antagonize each other and allow a finetuned action of NO on gene expression to maintain homeostasis of a cell. In a pathological setting, these self-regulatory mechanisms run out of control and this may result in apoptotic and necrotic processes that affect the proper function of the affected tissue. By contrast, reduced availability of NO as observed in cardiovascular diseases may also lead to a loss of function (e.g., of vascular contractility). The detailed knowledge of the mechanisms that trigger gene expression may help to develop therapeutic strategies to treat a number of diseases that are characterized by an inappropriate NO formation.

Nitric Oxide and Gene Expression

345

REFERENCES 1. Beck KF, Eberhardt W, Frank S, Huwiler A, Messmer UK, Mühl H, Pfeilschifter J. Inducible NO synthase: role in cellular signalling. J. Exp. Biol. 1999, 202:645–653. 2. Pfeilschifter J, Eberhardt W, Beck KF. Regulation of gene expression by nitric oxide. Pflugers Arch. 2001, 442:479–486. 3. Schmidt HH, Lohmann SM, Walter U. The nitric oxide and cGMP signal transduction system: regulation and mechanism of action. Biochim. Biophys. Acta. 1993, 1178:153–175. 4. Pilz RB, Casteel DE. Regulation of gene expression by cyclic GMP. Circ Res. 2003, 93:1034–1046. 5. Lohmann SM, Vaandrager AB, Smolenski A, Walter U, De Jonge HR. Distinct and specific functions of cGMP-dependent protein kinases. Trends Biochem. Sci. 1997, 22:307–312. 6. Montminy MR, Bilezikjian LM. Binding of a nuclear protein to the cyclic-AMP response element of the somatostatin gene. Nature. 1987, 328:175–178. 7. Shaywitz AJ, Greenberg ME. CREB: a stimulus-induced transcription factor activated by a diverse array of extracellular signals. Annu. Rev. Biochem. 1999, 68:821–861. 8. Gudi T, Casteel DE, Vinson C, Boss GR, Pilz RB. NO activation of fos promoter elements requires nuclear translocation of G-kinase I and CREB phosphorylation but is independent of MAP kinase activation. Oncogene. 2000, 19:6324–6333. 9. Sauzeau V, Rolli-Derkinderen M, Marionneau C, Loirand G, Pacaud P. RhoA expression is controlled by nitric oxide through cGMP-dependent protein kinase activation. J. Biol. Chem. 2003, 278:9472–9480. 10. Gudi T, Lohmann SM, Pilz RB. Regulation of gene expression by cyclic GMPdependent protein kinase requires nuclear translocation of the kinase: identification of a nuclear localization signal. Mol. Cell Biol. 1997, 17:5244–5254. 11. Chen Y, Zhuang S, Cassenaer S, Casteel DE, Gudi T, Boss GR, Pilz RB. Synergism between calcium and cyclic GMP in cyclic AMP response element-dependent transcriptional regulation requires cooperation between CREB and C/EBP-beta. Mol. Cell Biol. 2003, 23:4066–4082. 12. Janssen-Heininger YM, Poynter ME, Baeuerle PA. Recent advances towards understanding redox mechanisms in the activation of nuclear factor kappaB. Free Radic. Biol. Med. 2000, 28:1317–1327. 13. He B, Weber GF. Phosphorylation of NF-kappaB proteins by cyclic GMP-dependent kinase. A noncanonical pathway to NF-kappaB activation. Eur. J. Biochem. 2003, 270:2174–2185. 14. Kalra D, Baumgarten G, Dibbs Z, Seta Y, Sivasubramanian N, Mann DL. Nitric oxide provokes tumor necrosis factor-alpha expression in adult feline myocardium through a cGMP-dependent pathway. Circulation. 2000, 102:1302–1307. 15. Tsao PS, Buitrago R, Chan JR, Cooke JP. Fluid flow inhibits endothelial adhesiveness. Nitric oxide and transcriptional regulation of VCAM-1. Circulation. 1996, 94:1682–1689. 16. Kiemer AK, Gerbes AL, Bilzer M, Vollmar AM. The atrial natriuretic peptide and cGMP: novel activators of the heat shock response in rat livers. Hepatology. 2002, 35:88–94.

346

Nitric Oxide, Cell Signaling, and Gene Expression

17. Roy AL. Biochemistry and biology of the inducible multifunctional transcription factor TFII-I. Gene. 2001, 274:1–13. 18. Casteel DE, Zhuang S, Gudi T, Tang J, Vuica M, Desiderio S, Pilz RB. cGMPdependent protein kinase I beta physically and functionally interacts with the transcriptional regulator TFII-I. J. Biol. Chem. 2002, 277:32003–32014. 19. Shaulian E, Karin M. AP-1 as a regulator of cell life and death. Nat. Cell Biol. 2002, 4:E131– E136. 20. Pilz RB, Suhasini M, Idriss S, Meinkoth JL, Boss GR. Nitric oxide and cGMP analogs activate transcription from AP-1-responsive promoters in mammalian cells. FASEB J. 1995, 9:552–558. 21. Wu J, Fang L, Lin Q, Willis WD. Fos expression is induced by increased nitric oxide release in rat spinal cord dorsal horn. Neuroscience. 2000; 96:351–357. 22. Cappola TP, Cope L, Cernetich A, Barouch LA, Minhas K, Irizarry RA, Parmigiani G, Durrani S, Lavoie T, Hoffman EP, Ye SQ, Garcia JG, Hare JM. Deficiency of different nitric oxide synthase isoforms activates divergent transcriptional programs in cardiac hypertrophy. Physiol. Genomics. 2003, 14:25–34. 23. Gertzberg N, Clements R, Jaspers I, Ferro TJ, Neumann P, Flescher E, Johnson A. Tumor necrosis factor-alpha-induced activating protein-1 activity is modulated by nitric oxide-mediated protein kinase G activation. Am. J. Respir. Cell Mol. Biol. 2000, 22:105–115. 24. Esteve L, Lutz P, Thiriet N, Revel M, Aunis D, Zwiller J. Cyclic GMP-dependent protein kinase potentiates serotonin-induced Egr-1 binding activity in PC12 cells. Cell Signal. 2001, 13:425–432. 25. Rupprecht HD, Akagi Y, Keil A, Hofer G. Nitric oxide inhibits growth of glomerular mesangial cells: role of the transcription factor EGR-1. Kidney Int. 2000, 57:70–82. 26. Sellak H, Yang X, Cao X, Cornwell T, Soff GA, Lincoln T. Sp1 transcription factor as a molecular target for nitric oxide- and cyclic nucleotide-mediated suppression of cGMP-dependent protein kinase-Iα expression in vascular smooth muscle cells. Circ. Res. 2002, 90:405–412. 27. Hazzalin CA, Mahadevan LC. MAPK-regulated transcription: a continuously variable gene switch? Nat. Rev. Mol. Cell Biol. 2002, 3:30–40. 28. Sugimoto T, Haneda M, Togawa M, Isono M, Shikano T, Araki S, Nakagawa T, Kashiwagi A, Guan KL, Kikkawa R. Atrial natriuretic peptide induces the expression of MKP-1, a mitogen-activated protein kinase phosphatase, in glomerular mesangial cells. J. Biol. Chem. 1996, 271:544–547. 29. Fiedler B, Lohmann SM, Smolenski A, Linnemuller S, Pieske B, Schröder F, Molkentin JD, Drexler H, Wollert KC. Inhibition of calcineurin-NFAT hypertrophy signaling by cGMP-dependent protein kinase type I in cardiac myocytes. Proc. Natl. Acad. Sci. USA. 2002, 99:11363–11368. 30. Gong JH, Renz H, Sprenger H, Nain M, Gemsa D. Enhancement of tumor necrosis factor-alpha gene expression by low doses of prostaglandin E2 and cyclic GMP. Immunobiology. 1990, 182:44–55. 31. Inoue T, Fukuo K, Nakahashi T, Hata S, Morimoto S, Ogihara T. cGMP upregulates nitric oxide synthase expression in vascular smooth muscle cells. Hypertension. 1995, 4:711–714. 32. Tetsuka T, Daphna-Iken D, Miller BW, Guan Z, Baier LD, Morrison AR. Nitric oxide amplifies interleukin 1-induced cyclooxygenase-2 expression in rat mesangial cells. J. Clin. Invest. 1996, 97:2051–2056.

Nitric Oxide and Gene Expression

347

33. Bouchie JL, Hansen H, Feener EP. Natriuretic factors and nitric oxide suppress plasminogen activator inhibitor-1 expression in vascular smooth muscle cells. Role of cGMP in the regulation of the plasminogen system. Arterioscler. Thromb. Vasc. Biol. 1998, 18:1771–1779. 34. Eberhardt W, Akool el-S, Rebhan J, Frank S, Beck KF, Franzen R, Hamada FM, Pfeilschifter J. Inhibition of cytokine-induced matrix metalloproteinase 9 expression by peroxisome proliferator-activated receptor alpha agonists is indirect and due to a NO-mediated reduction of mRNA stability. J. Biol. Chem. 2002, 277:33518–33528. 35. Keller T, Pleskova M, McDonald MC, Thiemermann C, Pfeilschifter J, Beck KF. Identification of manganese superoxide dismutase as a NO-regulated gene in rat glomerular mesangial cells by 2D gel electrophoresis. Nitric Oxide. 2003, 9:183–193. 36. Perez-Sala D, Cernuda-Morollon E, Diaz-Cazorla M, Rodriguez-Pascual F, Lamas S. Posttranscriptional regulation of human iNOS by the NO/cGMP pathway. Am. J. Physiol. Renal Physiol. 2001, 280:F466– F473. 37. Diaz-Cazorla M, Perez-Sala D, Lamas S. Dual effect of nitric oxide donors on cyclooxygenase-2 expression in human mesangial cells. J. Am. Soc. Nephrol. 1999, 10:943–952. 38. Lee J, Lee SH, Kang DG, Kim SW. Effects of nitric oxide synthesis inhibition on the atrial natriuretic peptide system. Pharmacol. Res. 2000, 42:275–279. 39. Cao L, Wu J, Gardner DG. Atrial natriuretic peptide suppresses the transcription of its guanylyl cyclase-linked receptor. J. Biol. Chem. 1995, 270:24891–24897. 40. Vera N, Tse MY, Watson JD, Sarda S, Steinhelper ME, John SW, Flynn TG, Pang SC. Altered expression of natriuretic peptide receptors in proANP gene disrupted mice. Cardiovasc. Res. 2000, 46:595–603. 41. Filippov G, Bloch DB, Bloch KD. Nitric oxide decreases stability of mRNAs encoding soluble guanylate cyclase subunits in rat pulmonary artery smooth muscle cells. J. Clin. Invest. 1997, 100:942–948. 42. Klöss S, Furneaux H, Mülsch A. Post-transcriptional regulation of soluble guanylyl cyclase expression in rat aorta. J. Biol. Chem. 2003, 278:2377–2383. 43. Sinha B, Eigler A, Baumann KH, Greten TF, Moeller J, Endres S. Nitric oxide downregulates tumour necrosis factor in mRNA in RAW 264.7 cells. Res. Immunol. 1998, 149:139–150. 44. Abdelaziz N, Colombo F, Mercier I, Calderone A. Nitric oxide attenuates the expression of transforming growth factor-beta(3) mRNA in rat cardiac fibroblasts via destabilization. Hypertension. 2001, 38:261–266. 45. Brennan CM, Steitz JA. HuR and mRNA stability. Cell Mol. Life. Sci. 2000, 58:266–277. 46. Akool el-S, Kleinert H, Hamada FM, Abdelwahab MH, Förstermann U, Pfeilschifter J, Eberhardt W. Nitric oxide increases the decay of matrix metalloproteinase 9 mRNA by inhibiting the expression of mRNA-stabilizing factor HuR. Mol. Cell Biol. 2003, 23:4901–4916. 47. Di Fulvio M, Lincoln TM, Lauf PK, Adragna NC. Protein kinase G regulates potassium chloride cotransporter-3 expression in primary cultures of rat vascular smooth muscle cells. J. Biol. Chem. 2001, 276:21046–21052. 48. Tiesjema RH, van Gelder BF. Biochemical and biophysical studies on cytochrome c oxidase. XVI. Circular dichroic study of cytochrome c oxidase and its ligand complexes. Biochim. Biophys. Acta. 1974, 347:202–214.

348

Nitric Oxide, Cell Signaling, and Gene Expression

49. Brudvig GW, Stevens TH, Chan SI. Reactions of nitric oxide with cytochrome c oxidase. Biochemistry. 1980, 19:5275–5285. 50. Lizasoain I, Moro MA, Knowles RG, Darley-Usmar V, Moncada S. Nitric oxide and peroxynitrite exert distinct effects on mitochondrial respiration which are differentially blocked by glutathione or glucose. Biochem. J. 1996, 314:877–880. 51. Cleeter MW, Cooper JM, Darley-Usmar VM, Moncada S, Schapira AH. Reversible inhibition of cytochrome c oxidase, the terminal enzyme of the mitochondrial respiratory chain, by nitric oxide. Implications for neurodegenerative diseases. FEBS Lett. 1994, 345:50–54. 52. Trimmer BA, Aprille JR, Dudzinski DM, Lagace CJ, Lewis SM, Michel T, Qazi S, Zayas RM. Nitric oxide and the control of firefly flashing. Science. 2001, 292:2486–2488. 53. Hagen T, Taylor CT, Lam F, Moncada S. Redistribution of intracellular oxygen in hypoxia by nitric oxide: effect on HIF1alpha. Science. 2003, 302:1975–1978. 54. Ohh M, Park CW, Ivan M, Hoffman MA, Kim TY, Huang LE, Pavletich N, Chau V, Kaelin WG. Ubiquitination of hypoxia-inducible factor requires direct binding to the beta-domain of the von Hippel-Lindau protein. Nat. Cell Biol. 2000, 2:423–427. 55. Semenza GL. HIF-1, O(2), and the 3 PHDs: how animal cells signal hypoxia to the nucleus. Cell. 2001, 107:1–3. 56. Sogawa K, Numayama-Tsuruta K, Ema M, Abe M, Abe H, Fujii-Kuriyama Y. Inhibition of hypoxia-inducible factor 1 activity by nitric oxide donors in hypoxia. Proc. Natl. Acad. Sci. USA. 1998, 95:7368–7373. 57. Brüne B, Zhou J. The role of nitric oxide (NO) in stability regulation of hypoxia inducible factor-1alpha (HIF-1alpha). Curr. Med. Chem. 2003, 10:845–855. 58. Metzen E, Zhou J, Jelkmann W, Fandrey J, Brüne B. Nitric oxide impairs normoxic degradation of HIF-1alpha by inhibition of prolyl hydroxylases. Mol. Biol. Cell. 2003, 14:3470–3481. 59. Brown GC, Cooper CE. Nanomolar concentrations of nitric oxide reversibly inhibit synaptosomal respiration by competing with oxygen at cytochrome oxidase. FEBS Lett. 1994, 356:295–298. 60. Mateo J, Garcia-Lecea M, Cadenas S, Hernandez C, Moncada S. Regulation of hypoxia-inducible factor-1alpha by nitric oxide through mitochondria-dependent and -independent pathways. Biochem. J. 2003, 376:537–544. 61. Sumbayev VV, Budde A, Zhou J, Brüne B. HIF-1 alpha protein as a target for Snitrosation. FEBS Lett. 2003, 535:106–112. 62. Levy AP, Levy NS, Wegner S, Goldberg MA. Transcriptional regulation of the rat vascular endothelial growth factor gene by hypoxia. J. Biol. Chem. 1995, 270:13333–13340. 63. Tuder RM, Flook BE, Voelkel NF. Increased gene expression for VEGF and the VEGF receptors KDR/Flk and Flt in lungs exposed to acute or to chronic hypoxia. Modulation of gene expression by nitric oxide. J. Clin. Invest. 1995, 95:1798–1807. 64. Chin K, Kurashima Y, Ogura T, Tajiri H, Yoshida S, Esumi H. Induction of vascular endothelial growth factor by nitric oxide in human glioblastoma and hepatocellular carcinoma cells. Oncogene. 1997, 15:437–442. 65. Walpen S, Beck KF, Schaefer L, Raslik I, Eberhardt W, Schaefer RM, Pfeilschifter J. Nitric oxide induces MIP-2 transcription in rat renal mesangial cells and in a rat model of glomerulonephritis. FASEB J. 2001, 15:571–573.

Nitric Oxide and Gene Expression

349

66. Zampetaki A, Mitsialis SA, Pfeilschifter J, Kourembanas S Hypoxia induces macrophage inflammatory protein-2 (MIP-2) gene expression in murine macrophages via NF-κB. FASEB J. 2004, 18:1090–1092. 67. Mühl H, Pfeilschifter J. Amplification of nitric oxide synthase expression by nitric oxide in interleukin 1 beta-stimulated rat mesangial cells. J. Clin. Invest. 1995, 95:1941–1946. 68. Melillo G, Musso T, Sica A, Taylor LS, Cox GW, Varesio L. A hypoxia-responsive element mediates a novel pathway of activation of the inducible nitric oxide synthase promoter. J. Exp. Med. 1995, 182:1683–1693. 69. Semenza GL. Oxygen-regulated transcription factors and their role in pulmonary disease. Respir. Res. 2000, 1:159–162. 70. Xu W, Charles IG, Moncada S. Nitric oxide: orchestrating hypoxia regulation through mitochondrial respiration and the endoplasmic reticulum stress response. Cell Res. 2005, 15:63–65. 71. Pfeilschifter J, Eberhardt W, Huwiler A. Nitric oxide and mechanisms of redox signaling. J. Am. Soc. Nephrol. 2003, 14:S237– S240. 72. Grisham MB, Jourd’Heuil D, Wink DA. Nitric oxide. I. Physiological chemistry of nitric oxide and its metabolites: implications in inflammation. Am. J. Physiol. 1999, 276:G315– G321. 73. Pfeilschifter J, Beck KF, Eberhardt W, Huwiler A. Changing gears in the course of glomerulonephritis by shifting superoxide to nitric oxide-dominated chemistry. Kidney Int. 2002, 61:809–815. 74. Sen CK, Packer L. Antioxidant and redox regulation of gene transcription. FASEB J. 1996, 10:709–720. 75. Schopfer FJ, Baker PR, Freeman BA. NO-dependent protein nitration: a cell signaling event or an oxidative inflammatory response? Trends Biochem. Sci. 2003, 28:646–654. 76. Frank S, Zacharowski K, Wray GM, Thiemermann C, Pfeilschifter J. Identification of copper/zinc superoxide dismutase as a novel nitric oxide-regulated gene in rat glomerular mesangial cells and kidneys of endotoxemic rats. FASEB J. 1999, 13:869–882. 77. Shinyashiki M, Chiang KT, Switzer CH, Gralla EB, Valentine JS, Thiele DJ, Fukuto JM. The interaction of nitric oxide (NO) with the yeast transcription factor Ace1: a model system for NO-protein thiol interactions with implications to metal metabolism. Proc. Natl. Acad. Sci. USA. 2000; 97:2491–2496. 78. Kröncke KD, Fehsel K, Schmidt T, Zenke FT, Dasting I, Wesener JR, Bettermann H, Breunig KD, Kolb-Bachofen V. Nitric oxide destroys zinc-sulfur clusters inducing zinc release from metallothionein and inhibition of the zinc finger-type yeast transcription activator LAC9. Biochem. Biophys. Res. Commun. 1994, 200:1105–1110. 79. Kröncke KD. Zinc finger proteins as molecular targets for nitric oxide-mediated gene regulation. Antioxid. Redox. Signal. 2001, 3:565–575. 80. Wang S, Wang W, Wesley RA, Danner RL. A Sp1 binding site of the tumor necrosis factor alpha promoter functions as a nitric oxide response element. J. Biol. Chem. 1999, 274:33190–33193. 81. Kröncke KD, Carlberg C. Inactivation of zinc finger transcription factors provides a mechanism for a gene regulatory role of nitric oxide. FASEB J. 2000, 14:166–173.

350

Nitric Oxide, Cell Signaling, and Gene Expression

82. Von Knethen A, Brüne B. Activation of peroxisome proliferator-activated receptor gamma by nitric oxide in monocytes/macrophages down-regulates p47phox and attenuates the respiratory burst. J. Immunol. 2002, 169:2619–2626. 83. Matthews JR, Botting CH, Panico M, Morris HR, Hay RT. Inhibition of NFkappaB DNA binding by nitric oxide. Nucleic Acids Res. 1996, 24:2236–2242. 84. delaTorre A, Schroeder RA, Punzalan C, Kuo PC. Endotoxin-mediated S-nitrosylation of p50 alters NF-kappaB-dependent gene transcription in ANA-1 murine macrophages. J. Immunol. 1999, 162:4101–4108. 85. Spiecker M, Peng HB, Liao JK. Inhibition of endothelial vascular cell adhesion molecule-1 expression by nitric oxide involves the induction and nuclear translocation of IkappaBalpha. J. Biol. Chem. 1997, 272:30969–30974. 86. Hierholzer C, Harbrecht B, Menezes JM, Kane J, MacMicking J, Nathan CF, Peitzman AB, Billiar TR, Tweardy DJ. Essential role of induced nitric oxide in the initiation of the inflammatory response after hemorrhagic shock. J. Exp. Med. 1998, 187:917–928. 87. Lander HM. An essential role for free radicals and derived species in signal transduction. FASEB J. 1997, 11:118–124. 88. Deora AA, Win T, Vanhaesebroeck B, Lander HM. A redox-triggered ras-effector interaction. Recruitment of phosphatidylinositol 3'-kinase to Ras by redox stress. J. Biol. Chem. 1998, 273:29923–29928. 89. Aga M, Watters JJ, Pfeiffer ZA, Wiepz GJ, Sommer JA, Bertics PJ. Evidence for nucleotide receptor modulation of cross talk between MAP kinase and NF-kappa B signaling pathways in murine RAW 264.7 macrophages. Am. J. Physiol. Cell Physiol. 2004, 286:C923– C930. 90. Nikitovic D, Holmgren A, Spyrou G. Inhibition of AP-1 DNA binding by nitric oxide involving conserved cysteine residues in Jun and Fos. Biochem. Biophys. Res. Commun. 1998, 242:109–112. 91. Bogdan C. Nitric oxide and the regulation of gene expression.Trends Cell Biol. 2001, 11:66–75. 92. Hmadcha A, Bedoya FJ, Sobrino F, Pintado E. Methylation-dependent gene silencing induced by interleukin 1beta via nitric oxide production. J. Exp. Med. 1999, 190:1595–1604. 93. Park SW, Li J, Loh HH, Wei LN. A novel signaling pathway of nitric oxide on transcriptional regulation of mouse kappa opioid receptor gene. J. Neurosci. 2002, 22:7941–7947. 94. Hofbauer KH, Schoof E, Kurtz A, Sandner P. Inflammatory cytokines stimulate adrenomedullin expression through nitric oxide-dependent and -independent pathways. Hypertension. 2002, 39:161–167. 95. Ichiki T, Usui M, Kato M, Funakoshi Y, Ito K, Egashira K, Takeshita A. Downregulation of angiotensin II type 1 receptor gene transcription by nitric oxide. Hypertension. 1998, 31:342–348. 96. Trachtman H, Futterweit S, Singhal P. Nitric oxide modulates the synthesis of extracellular matrix proteins in cultured rat mesangial cells. Biochem. Biophys. Res. Commun. 1995, 207:120–125. 97. Guo K, Andres V, Walsh K. Nitric oxide-induced downregulation of Cdk2 activity and cyclin A gene transcription in vascular smooth muscle cells. Circulation. 1998, 97:2066–2072.

Nitric Oxide and Gene Expression

351

98. Fukai T, Siegfried MR, Ushio-Fukai M, Cheng Y, Kojda G, Harrison DG. Regulation of the vascular extracellular superoxide dismutase by nitric oxide and exercise training. J. Clin. Invest. 2000, 105:1631–1639. 99. Todorov V, Gess B, Godecke A, Wagner C, Schrader J, Kurtz A. Endogenous nitric oxide attenuates erythropoietin gene expression in vivo. Pflugers Arch. 2000, 439:445–448. 100. Kosonen O, Kankaanranta H, Uotila J, Moilanen E. Inhibition by nitric oxidereleasing compounds of E-selectin expression in and neutrophil adhesion to human endothelial cells. Eur. J. Pharmacol. 2000, 394:149–156. 101. Kourembanas S, McQuillan LP, Leung GK, Faller DV. Nitric oxide regulates the expression of vasoconstrictors and growth factors by vascular endothelium under both normoxia and hypoxia. J. Clin. Invest. 1993, 92:99–104. 102. Frank S, Stallmeyer B, Kämpfer H, Schaffner C, Pfeilschifter J. Differential regulation of vascular endothelial growth factor and its receptor fms-like-tyrosine kinase is mediated by nitric oxide in rat renal mesangial cells. Biochem. J. 1999, 338:367–374. 103. Kuo PC, Abe KY, Schroeder RA. Interleukin-1-induced nitric oxide production modulates glutathione synthesis in cultured rat hepatocytes. Am. J. Physiol. 1996, 271:C851– C862. 104. Durante W, Kroll MH, Christodoulides N, Peyton KJ, Schafer AI. Nitric oxide induces heme oxygenase-1 gene expression and carbon monoxide production in vascular smooth muscle cells. Circ. Res. 1997, 80:557–564. 105. Ikeda M, Ikeda U, Takahashi M, Shimada K, Minota S, Kano S. Nitric oxide inhibits intracellular adhesion molecule-1 expression in rat mesangial cells. J. Am. Soc. Nephrol. 1996, 7:2213–2218. 106. Rothe H, Hartmann B, Geerlings P, Kolb H. Interleukin-12 gene-expression of macrophages is regulated by nitric oxide. Biochem. Biophys. Res. Commun. 1996, 224:159–163. 107. Andrew PJ, Harant H, Lindley IJ. Nitric oxide regulates IL-8 expression in melanoma cells at the transcriptional level. Biochem. Biophys. Res. Commun. 1995, 214:949–956. 108. Beck KF, Walpen S, Eberhardt W, Pfeilschifter J. Downregulation of integrinlinked kinase mRNA expression by nitric oxide in rat glomerular mesangial cells. Life Sci. 2001, 69:2945–2955. 109. Taylor BS, Kim YM, Wang Q, Shapiro RA, Billiar TR, Geller DA. Nitric oxide down-regulates hepatocyte-inducible nitric oxide synthase gene expression. Arch. Surg. 1997, 132:1177–1183. 110. Zeiher AM, Fisslthaler B, Schray-Utz B, Busse R. Nitric oxide modulates the expression of monocyte chemoattractant protein 1 in cultured human endothelial cells. Circ. Res. 1995, 76:980–986. 111. Peng HB, Rajavashisth TB, Libby P, Liao JK. Nitric oxide inhibits macrophagecolony stimulating factor gene transcription in vascular endothelial cells. J. Biol. Chem. 1995, 270:17050–17055. 112. Sicher SC, Chung GW, Vazquez MA, Lu CY. Augmentation or inhibition of IFNgamma-induced MHC class II expression by lipopolysaccharides. The roles of TNF-alpha and nitric oxide, and the importance of the sequence of signaling. J. Immunol. 1995, 155:5826–5834.

352

Nitric Oxide, Cell Signaling, and Gene Expression

113. Mühl H, Dinarello CA. Macrophage inflammatory protein-1 alpha production in lipopolysaccharide-stimulated human adherent blood mononuclear cells is inhibited by the nitric oxide synthase inhibitor N(G)-monomethyl-L-arginine. J. Immunol. 1997, 159:5063–5069. 114. Sugimoto T, Haneda M, Togawa M, Isono M, Shikano T, Araki S, Nakagawa T, Kashiwagi A, Guan KL, Kikkawa R. Atrial natriuretic peptide induces the expression of MKP-1, a mitogen-activated protein kinase phosphatase, in glomerular mesangial cells. J. Biol. Chem. 1996, 271:544–547. 115. Chen HH, Wang DL. Nitric oxide inhibits matrix metalloproteinase-2 expression via the induction of activating transcription factor 3 in endothelial cells. Mol. Pharmacol. 2004, 65:1130–1140. 116. Ji JY, Diamond SL. Exogenous nitric oxide activates the endothelial glucocorticoid receptor. Biochem. Biophys. Res. Commun. 2004, 318:192–197. 117. Walpen S, Beck KF, Eberhardt W, Apel M, Chatterjee PK, Wray GM, Thiemermann C, Pfeilschifter J. Downregulation of SPARC expression is mediated by nitric oxide in rat mesangial cells and during endotoxemia in the rat. J. Am. Soc. Nephrol. 2000, 11:468–476. 118. Rupprecht G, Scholz K, Beck KF, Geiger H, Pfeilschifter J, Kaszkin M. Crosstalk between group IIA-phospholipase A2 and inducible NO-synthase in rat renal mesangial cells. Br. J. Pharmacol. 1999, 127:51–56. 119. Craven PA, Studer RK, Felder J, Phillips S, DeRubertis FR. Nitric oxide inhibition of transforming growth factor-beta and collagen synthesis in mesangial cells. Diabetes. 1997, 46:671–681. 120. Eberhardt W, Beeg T, Beck KF, Walpen S, Gauer S, Böhles H, Pfeilschifter J. Nitric oxide modulates expression of matrix metalloproteinase-9 in rat mesangial cells. Kidney Int. 2000, 57:59–69. 121. Van Dervort AL, Yan L, Madara PJ, Cobb JP, Wesley RA, Corriveau CC, Tropea MM, Danner RL. Nitric oxide regulates endotoxin-induced TNF-alpha production by human neutrophils. J. Immunol. 1994, 152:4102–4109. 122. Eberhardt W, Beck KF, Pfeilschifter J. Cytokine-induced expression of tPA is differentially modulated by NO and ROS in rat mesangial cells. Kidney Int. 2002, 61:20–30. 123. Spiecker M, Peng HB, Liao JK. Inhibition of endothelial vascular cell adhesion molecule-1 expression by nitric oxide involves the induction and nuclear translocation of IkappaBalpha. J. Biol. Chem. 1997, 272:30969–30974. 124. De Caterina R, Libby P, Peng HB, Thannickal VJ, Rajavashisth TB, Gimbrone MA, Jr., Shin WS, Liao JK. Nitric oxide decreases cytokine-induced endothelial activation. Nitric oxide selectively reduces endothelial expression of adhesion molecules and proinflammatory cytokines. J. Clin. Invest. 1995, 96:60–68. 125. Lee CI, Liu X, Zweier JL. Regulation of xanthine oxidase by nitric oxide and peroxynitrite. J. Biol. Chem. 2000, 275:9369–9376.

Oxide as a Modifier 16 Nitric of Gene Expression Santiago Lamas, Antonio Martínez-Ruiz, and Carlos Zaragoza Fundación Centro Nacional de Investigaciones Cardiovasculares (CNIC), Centro de Investigaciones Biológicas (CIB, CSIC), Instituto Reina Sofía de Investigaciones Nefrológicas, Madrid, Spain

CONTENTS 16.1 16.2 16.3 16.4

Introduction ............................................................................................353 Signaling Pathways Controlled by NO .................................................354 Biochemical Mechanisms ......................................................................356 Conclusion .............................................................................................360 References ...............................................................................................360

16.1 INTRODUCTION Nitric oxide (NO) is a free radical gas molecule produced by the activity of NO synthase (NOS). Three NOS isoforms have been described: Two constitutive isoforms are expressed in endothelial cells and central nervous system (eNOS and nNOS, respectively) [1, 2] among other tissues, and on the other hand, an inducible isoform is synthesized by several cell types (iNOS) [3, 4]. Although eNOS and nNOS are calcium dependent and produce low levels of NO in a very tightly controlled way [5], iNOS is calcium independent, and in response to proinflammatory stimuli, it generates NO very rapidly and at higher levels than the constitutive isoforms. NO plays different roles depending on the cell type in which it is produced. In general, it is a signaling molecule, playing neurotransmission properties in the central nervous system [6–9]. Immune cells generate NO during inflammation [10–17], and it is an immune effector against several microorganisms [14, 18–22]. In the vascular endothelium, it is synthesized by the constitutive calcium dependent NOS isoform (eNOS). It acts as a potent vasodilator and thus is key for the maintenance of vascular tone [23]. The main target of NO produced by the endothelial cells resides in the smooth muscle cell layer, namely the soluble guanylate cyclase, which produces cyclic GMP (cGMP) and induces vasorelaxation [24]. 353

354

Nitric Oxide, Cell Signaling, and Gene Expression

In recent years, it has been demonstrated that NO regulates gene expression. This may occur through its natural effector, cGMP, or by post-translational modification of proteins, including S-nitrosylation [25], nitration [26], or glutathionylation [27, 28]. As described in Section 16.2 in more detail, NO has been reported to interact with several signaling cascades that in turn modulate gene expression.

16.2 SIGNALING PATHWAYS CONTROLLED BY NO Cyclic GMP-induced gene expression by NO is mediated by the activation of the natural NO receptor soluble guanylate cyclase [29–31], which synthesizes cGMP, the activator of the cGMP-dependent protein kinase (PKG). Two forms of PKG have been described with different roles in physiology [32–41]. Although PKG II has been implicated in intestinal secretion and bone development [42–45], PKG I deals with the majority of the NO-mediated gene expression effects [46, 47]. PKG I mediates the effects of NO in the regulation of gene expression under different pathophysiological situations. NO affects the mRNA of several genes involved in cell proliferation, cell migration, cell adhesion, and apoptosis, all processes related with angiogenesis and cell injury responses to external stimuli [48–59]. In the cardiovascular system, NO also controls vascular tone [60, 61], smooth muscle cell differenciation [33], and is actively participating by controlling the gene expression of genes related to inflammatory events [62–64]. Energy supply as well as cellular responses to changes of the redox state balance are also controlled by NO, due to the capacity to regulate the expression of genes involved in the biosynthesis of mitochondria, cell survival, and cell death [65–71]. The effects of NO on cell proliferation are closely related with the cross talk between cGMP/PKG and other kinase signaling cascades. Depending on the tissue and cell type, NO induces cell proliferation [72] or inhibits cell proliferation [73–77]. In this regard NO modulates the activity of the Mitogen Activating Associated Protein Kinase (MAPK) family of proteins ERK, p38, and JNK, regulating the expression and activity of different transcription factors, and therefore controlling gene expression. ERK1,2 stimulation induces the activation of two main transcription factors involved in proliferation: Elk and AP-1 [63, 78–82]. In the case of AP-1 the activation of this transcription factor is mediated by PKG phosphorylation, inducing the expression of jun and fos members of AP-1, the effect mediated in some cases by the activation of Elk at the promoter region of these genes [46, 83, 84]. The anti-proliferative effects of NO related to the MAPK pathway are mediated by the effect of cGMP in the regulation of MAPK protein phosphatases (MKP). In particular, MKP-1 expression is increased via PKG [85, 86]. NFkB is another transcription factor regulated by NO, in which PKG phosphorylates the two components p50 and p65 [87–89], increasing the DNA-binding and trans-activation activity. Other factors modulated by NO are NFAT [90, 91], ATF-1 [92], CREB [93], SRF [94], or Egr [95, 96]. Another transcription factor controlled by NO through cGMP/PKG is TFII-I, which regulates the expression of many genes, including AP-1, and NF-κB [97]. A detailed list of genes controlled transcriptionally by NO is presented in Table 16.1.

Nitric Oxide as a Modifier of Gene Expression

355

TABLE 16.1 Gene Expression Regulated by NO/cGMP/PKG Gene

mRNA Levels

Cell Types

PKG Dependent Yes

Refs.

NOS-2

Increased

SMCs, CMs, MCs

[163, 164]

COX-2

Increased

MCs, HNCs RCs

Yes

[165–167]

ET-1

Decreased

ECs, fibroblasts

ND

[168]

PKG

Decreased

SMCs

TNF-α

Increased

SMCs, CMs

ANP receptor

Decreased

ANP

Decreased

BNP

[169–171] Yes

[89]

Yes

[172]

CMs

Yes

[173]

Decreased

CMs

Yes

[91]

VEGF

Increased

SMCs, macrophages, cancer, ECs

ND

[174, 175]

MKP-1

Increased

SMCs, ECs, MCs, ND

ND

[85, 176]

Rho

Increased

SMCs

Yes

[92]

p11

Increased

BEC

Yes

[177]

Cyclins

Decreased

SMCs, MCs, CCs

ND

[49, 51, 178]

p16

Increased

SMCs

ND

[179]

p21

Increased

Fibroblasts

ND

[180–182]

Bcl2

Increased

CNS, DCs

Yes

[56, 183]

PGC1

Increased

HeLa, fibroblasts

ND

[184]

MMP-2

Decreased

ECs

ND

Increased

Fibroblasts Cancer

MMP-9

Increased

Bone marrow

[185] [186] [187]

ND

[188]

Increased

SMCs

[189]

Decreased

SMCs

[190]

MMP-13

Increased

ECs

Yes

[63]

TGF-β

Decreased

Fibroblasts

ND

[191]

AP-1

Increased

Fibroblasts, ECs

Yes

[83, 84, 177]

IL8

Increased

LECs

No cGMP, no PKG

[192]

Caveolin-1

Decreased

ECs

ND

[193]

IP-10

Decreased

CCs

ND

[174]

MIG

Decreased

CCs

ND

[174]

P-selectin

Decreased

ECs

ND

[194]

MCP-1

Decreased

ECs

ND

[195]

Note: SMCs: smooth muscle cells; CMs: cardiomiocytes; DCs: dendritic cells; MCs: mesangial cells; HNCs; renal cortical cells; ECs: endothelial cells; CCs: colon cancer cells; LECs: lung epithelial cells; ND: no data.

356

Nitric Oxide, Cell Signaling, and Gene Expression

NO also regulates gene expression at the post-transcriptional level by means of two different ways: 1. NO alters the mRNA stability. 2. NO induces post-translational modifications. The mechanism exerted by NO concerning mRNA stability resides at the AU-rich regions that several genes harbor at the 3′-UTR end. In most cases, the effect is cGMP-dependent and is related with the expression of proteins that regulate mRNA stability at the AU-rich regions. In particular, cGMP regulates several genes at the post-translational level by controlling HuR expression, an AU-rich-binding protein, which confers mRNA stability. Genes affected by this way are soluble guanylate cyclase [98], MMP-9 [99, 100], iNOS [101], COX-2 [102], VEGF [103, 104], IL-8 [105], TGF-β, and TNF-α [106–108].

16.3 BIOCHEMICAL MECHANISMS NO was first postulated as a second messenger when it was reported that it could activate soluble guanylate cyclase (sGC) to produce cGMP from GTP [109, 110]. NO interacts with the heme group of sGC forming a coordinate bond. NO binding promotes the release of a histidine residue from the coordination structure and triggers the catalytic activity [111, 112]. sGC is characterized by a high affinity and dissociation rates for NO. These two features make this interaction especially suitable for signal transduction because small quantities of NO are perceived by sGC performing a rapid response [113, 114]. Activation of sGC to produce cGMP leads to the activation of different signalling cascades that influence gene expression in many ways, as explained before. Another relevant interaction between NO and a metal center is the one with cytochrome-c-oxidase complex IV from the mitochondrial respiration chain. It was demonstrated long ago that both NO and oxygen compete for the binding to its active center, in which a heme and a copper groups are localized. Although the precise mechanism of this competitive binding is still under discussion [115–117], it is known that the competition is established at physiological concentrations of the two gas molecules. This competition is able to inhibit the mitochondrial respiration chain at its last step, thus inducing the production of ROS (O2−, mainly as an excess of electrons are produced in the first steps of the chain), activating different signalling mechanisms. Another consequence of the competition between NO and O2 for the binding to cytochrome c oxidase has been established: When NO inhibits O2 binding, relocation of the oxygen molecule inside the cell occurs, influencing O2 sensing by the prolyl hidroxylases that act as oxygen sensors by signalling HIF-α degradation [118]. Hypoxia-inducible factors (HIF factors) are a family of transcription factors that induce the expression of a number of genes under hypoxic conditions, such as erythropoietin or VEGF [119], in which the α subunit is regulated by oxygen concentrations.

Nitric Oxide as a Modifier of Gene Expression

357

Indirect mechanisms by which NO production can alter gene expression rely on covalent modification of protein residues, in which the chemical species directly involved is not usually NO itself, but many of the so-called “reactive nitrogen species” (RNS) formed by the interaction of NO with other radicals and oxygen. Among these covalent modifications, S-nitrosylation (or S-nitrosation [120]) has been more studied in recent years. S-nitrosylation implies the incorporation of an NO group to cysteine thiol residues of proteins. NO itself is not a nitrosylating species, and the reaction is mediated by the formation of different RNS [121, 122]. NO reacts with O2 to form different higher nitrogen oxides, among which N2O3 has been proposed as the essential S-nitrosylating species [123, 124]: 2 NO + O2 → 2 NO2 NO + NO2 → N2O3 N2O3 + RSH → RSNO + NO2− + H+ The relevance of protein S-nitrosylation in physiological contexts is still controversial. Although some authors consider it a signalling mechanism, others claim that it is mainly a collateral effect when an overproduction of RNS takes place. Technical difficulties arising from nitrosothiol bond lability have hampered its detection in individual proteins from intact cells, although this modification has been described for over 100 proteins in studies with purified proteins [25, 120]. Thus, further studies will be needed, as well as development of methodologies with higher sensitivity. One way in which S-nitrosylation can alter gene expression is by modifying critical cysteines in transcription factors. This has been described for the p50 subunit of NF-κB. This factor has a complex regulation that can be influenced by oxidative and nitrosative stress in many ways [125], as it harbors a cysteine residue located within the DNA-binding domain, required to be reduced for the binding [126, 127]. S-nitrosylation of p50 has been described both in vitro [128, 129] and in cells [130, 131], and is correlated with an inactivation of DNA binding [132, 133]. Regarding NF-κB, a subunit of the kinase controlling NFκB activity, IKK, is also inactivated by S-nitrosylation, providing an additional mechanism of control for this transcription factor [134]. Another transcription factor modified b S-nitrosylation is AP-1 and more precisely the c-Jun subunit [129, 135]. Another set of transcription factors that can be inhibited by S-nitrosylation is that including zinc fingers with cysteines as zinc chelators [136]. Inhibition by S-nitrosylation of the DNA-binding activity has been reported for several members of this family [137–139], probably by disruption of the zinc finger structure. The effect of S-nitrosylation either increases or decreases transcription rates of the target genes, depending on the activator or repressor role of the transcription factor modified [137, 140].

358

Nitric Oxide, Cell Signaling, and Gene Expression

S-nitrosylation also affects zinc homeostasis in cells. Zn2+ is primarily stored by complexing it to cysteine centers in metallothionein, and is released in response to Zn2+ requirements [141]. NO releases Zn2+ from the thiolates in metallothionein by S-nitrosylation [142–145], although in certain situations it may promote a transient modification resulting in a disulfide bond formation [146]. S-nitrosylation has also been described to impair DNA binding of a repressor protein, heterogeneous nuclear ribonucleoprotein A/B, which activates transcription of the osteopontin gene [147]. S-nitrosylation is not the only mechanism by which NO and RNS modify protein thiols because they can also induce other oxidative modifications, providing a connection among oxidative and nitrosative chemistry. We must also consider S-thiolation, the formation of mixed disulfide bridges between two thiols. Because glutathione is the main free thiol reservoir in cells, S-glutathionylation is the S-thiolation that has been more thoroughly analyzed [148]. NO donors can induce S-glutathionylation in several proteins, including transcription factors such as the cJun subunit of AP-1 [28] and the p50 subunit of NF-κB [27]. A nitrosative agent could modify either of the two thiols, forming a nitrosothiol or it can create a disulfide bridge [148]. Besides, nitrosothiols react with a free thiol to form the disulfide bridge and HNO [149, 150]: RSNO + R′SH → RSSR′ + HNO In this regard, S-nitrosoglutathione (GSNO) has been used as a probe to bind proteins susceptible of being S-glutathionylated [148]. Thus, modification of a protein thiol by GSNO can lead to a functional alteration by at least two different mechanisms: S-nitrosylation or S-glutathionylation. Transnitrosation, the transfer of the NO moiety between two thiols, is also a mechanism that can be significant in the cell, because of the high concentration of intracellular low molecular mass thiols like glutathione [149, 151, 152]: RSNO + R′SH → RSH + R′SNO Tyrosine nitration, the incorporation of a nitro (−NO2) group in position 3 of the phenolic ring of tyrosine, is another mechanism of protein modification by RNS that can account for functional changes. Although 3-nitrotyrosine formation is considered to proceed mainly by radical reactions, involving species like the tyrosyl radical and nitrogen dioxide (NO2), two pathways are nowadays invoked as the main ones leading to formation of these nitrating species in biological systems [26, 153]. One is the formation of peroxynitrite (ONOO−), mainly by the fast reaction between NO and superoxide (O2−), when they are produced at the same time in the same place, as both are free radicals that can react with many other species. The other pathway is the formation of nitrating species from H2O2 and nitrite by heme peroxidase proteins, including myeloperoxidase and eosinophil peroxidase. Tyrosine nitration is considered irreversible in biological conditions because it is very much favored thermodynamically. In comparison

Nitric Oxide as a Modifier of Gene Expression

359

Other O2− sources

O2− Peroxidase pathway

ONOO−

NOS Prolyl hidroxilases NO

HIF-α Cyt. c ox.

P-Tyr-3-NO2 P-Men+-NO

N2O3

Tyrosine nitration

sGC

O2 Metal nitrosylation cGMP

GSNO P-SS-G

P-SNO

S-glutathionylation Signalling cascades

S-nitros(yl)ation

Signalling cascades

Transcription factors

FIGURE 16.1 Biochemical mechanisms of gene expression modification by nitric oxide. The mechanisms described in the text are schematized. NO, produced by nitric oxide synthases (NOS), can modify metal nitrosylation of soluble guaylate cyclase (sGC, which produces cGMP), cytocrome c oxidase (cyt. c ox., competing with O2, which is then liberated to the cell, inducing degradation of HIF-α by prolyl hydroxylases), and prolyl hydroxylases (when NO is in high amounts, inhibiting degradation of HIF-α). It can also produce N2O3, which induces S-nitrosylation. In turn, formation of S-nitrosoglutathione (or reaction of a protein S-nitrosothiol with glutathione) can induce S-glutathionylation. Two pathways have been described to favor tyrosine nitration, one being the formation of peroxynitrite (ONOO−), which can be formed by reaction of NO and superoxide (O2−). O2− can be produced by many sources, including NOS itself.

with phenylalanine, the presence of the phenol group in tyrosine stabilizes the incorporation of the nitro group in the ortho position with respect to it. There is a discussion about its biological relevance (reviewed in [26], [153], and [154]) because, although it has been detected in a number of situations that are mainly associated with pathological overproduction of ROS and RNS, quantitative measurements have indicated that a scarce number of tyrosine residues are actually modified. This can be related to the reported specificity of certain proteins, and certain residues among them, for being nitrated. In addition, although a substantial fraction of a residue has to be modified for losing its function, a gain of function would only require a small fraction of the protein to be modified for arising. Another issue to be considered is that many tyrosine residues can be phosphorylated and nitration of these residues would alter their signalling properties [155]. Gene expression affected by protein nitration has been reported by the alteration of transcription factors like the glucocorticoid receptor [156], retinoic acid receptor [157], PPARγ [158], or NF-κB through the inhibitory

360

Nitric Oxide, Cell Signaling, and Gene Expression

kinase IKK [159], as well as the alteration of different signalling pathways [160–162].

16.4 CONCLUSION It is increasingly evident that NO contributes to modify gene expression, not only through its natural effector cGMP, but also through precise post-translational modifications of proteins involved in this process. These modifications are progressively being recognized and their occurrence depends on the presence of NO or related species as well as on the interaction with molecular oxygen. Profound knowledge of stoichiometry, cellular spatio-temporal relationships, and in vivo relevance of these modifications will contribute to establish a definitive role for NO as a second messenger and key molecule in signal transduction.

REFERENCES 1. Bredt, D.S., et al., Cloned and expressed nitric oxide synthase structurally resembles cytochrome P-450 reductase. Nature, 1991. 351(6329): 714–8. 2. Lamas, S., et al., Endothelial nitric oxide synthase: molecular cloning and characterization of a distinct constitutive enzyme isoform. Proc. Natl. Acad. Sci. USA, 1992. 89(14): 6348–52. 3. Lowenstein, C.J. and E. Padalko, iNOS (NOS2) at a glance. J. Cell Sci., 2004. 117(Pt 14): 2865–7. 4. Cooke, J.P. and V.J. Dzau, Nitric oxide synthase: role in the genesis of vascular disease. Annu. Rev. Med., 1997. 48: 489–509. 5. Marletta, M.A., Another activation switch for endothelial nitric oxide synthase: why does it have to be so complicated? Trends Biochem. Sci., 2001. 26(9): 519–21. 6. Yun, H.Y., V.L. Dawson, and T.M. Dawson, Nitric oxide in health and disease of the nervous system. Mol. Psychiatry, 1997. 2(4): 300–10. 7. Karson, C.N., et al., Nitric oxide synthase (NOS) in schizophrenia: increases in cerebellar vermis. Mol. Chem. Neuropathol., 1996. 27(3): 275–84. 8. Nelson, R.J., et al., Behavioural abnormalities in male mice lacking neuronal nitric oxide synthase. Nature, 1995. 378(6555): 383–6. 9. Huang, P.L., et al., Targeted disruption of the neuronal nitric oxide synthase gene. Cell, 1993. 75(7): 1273–86. 10. Karupiah, G., et al., Rapid interferon gamma-dependent clearance of influenza A virus and protection from consolidating pneumonitis in nitric oxide synthase 2deficient mice. J. Exp. Med., 1998. 188(8): 1541–6. 11. Hickey, M.J., et al., Inducible nitric oxide synthase-deficient mice have enhanced leukocyte-endothelium interactions in endotoxemia. FASEB J., 1997. 11(12): 955–64. 12. Vodovotz, Y., et al., Vesicle membrane association of nitric oxide synthase in primary mouse macrophages. J. Immunol., 1995. 154(6): 2914–25. 13. Zaragoza, C., et al., The role of inducible nitric oxide synthase in the host response to Coxsackievirus myocarditis. Proc. Natl. Acad. Sci. USA, 1998. 95(5): 2469–74.

Nitric Oxide as a Modifier of Gene Expression

361

14. Zaragoza, C., et al., Inducible nitric oxide synthase protection against coxsackievirus pancreatitis. J. Immunol., 1999. 163(10): 5497–504. 15. Wahl, S.M., et al., Nitric oxide in experimental joint inflammation. Benefit or detriment? Cells Tissues Organs, 2003. 174(1-2): 26–33. 16. Kolios, G., et al., Expression of inducible nitric oxide synthase activity in human colon epithelial cells: modulation by T lymphocyte derived cytokines. Gut, 1998. 43(1): 56–63. 17. Kabashima, H., et al., Interferon-gamma-producing cells and inducible nitric oxide synthase-producing cells in periapical granulomas. J. Oral Pathol. Med., 1998. 27(3): 95–100. 18. Saura, M., et al., An antiviral mechanism of nitric oxide: inhibition of a viral protease. Immunity, 1999. 10(1): 21–8. 19. Zaragoza, C., et al., Nitric oxide inhibition of coxsackievirus replication in vitro. J. Clin. Invest., 1997. 100(7): 1760–7. 20. De Groote, M.A. and F.C. Fang, NO inhibitions: antimicrobial properties of nitric oxide. Clin. Infect. Dis., 1995. 21 (Suppl 2): S162–5. 21. Mannick, J.B., The antiviral role of nitric oxide. Res. Immunol., 1995. 146(9): 693–7. 22. Powell, K.L. and S.A. Baylis, The antiviral effects of nitric oxide. Trends Microbiol., 1995. 3(3): 81–2. 23. Shesely, E.G., et al., Elevated blood pressures in mice lacking endothelial nitric oxide synthase. Proc. Natl. Acad. Sci. USA, 1996. 93(23): 13176–81. 24. Murad, F., Cyclic guanosine monophosphate as a mediator of vasodilation. J. Clin. Invest., 1986. 78(1): 1–5. 25. Stamler, J.S., S. Lamas, and F.C. Fang, Nitrosylation: the prototypic redox-based signaling mechanism. Cell, 2001. 106(6): 675–83. 26. Radi, R., Nitric oxide, oxidants, and protein tyrosine nitration. Proc. Natl. Acad. Sci. USA, 2004: 0307446101. 27. Pineda-Molina, E., et al., Glutathionylation of the p50 subunit of NF-kappaB: a mechanism for redox-induced inhibition of DNA binding. Biochemistry, 2001. 40(47): 14134–42. 28. Klatt, P., E. Pineda Molina, and S. Lamas, Nitric oxide inhibits c-Jun DNA binding by specifically targeted S-glutathionylation. J. Biol. Chem., 1999. 274(22): 15857–64. 29. Klein, C., Nitric oxide and the other cyclic nucleotide. Cell Signal., 2002. 14(6): 493–8. 30. Andreopoulos, S. and A. Papapetropoulos, Molecular aspects of soluble guanylyl cyclase regulation. Gen. Pharmacol., 2000. 34(3): 147–57. 31. Denninger, J.W. and M.A. Marletta, Guanylate cyclase and the •NO/cGMP signaling pathway. Biochim. Biophys. Acta, 1999. 1411(2-3): 334–50. 32. Doerig, C., Protein kinases as targets for anti-parasitic chemotherapy. Biochim. Biophys. Acta, 2004. 1697(1-2): 155–68. 33. Lincoln, T.M., N. Dey, and H. Sellak, Invited review: cGMP-dependent protein kinase signaling mechanisms in smooth muscle: from the regulation of tone to gene expression. J. Appl. Physiol., 2001. 91(3): 1421–30. 34. Wang, X. and P.J. Robinson, Cyclic GMP-dependent protein kinase and cellular signaling in the nervous system. J. Neurochem., 1997. 68(2): 443–56. 35. Scott, J.D., Cyclic nucleotide-dependent protein kinases. Pharmacol. Ther., 1991. 50(1): 123–45.

362

Nitric Oxide, Cell Signaling, and Gene Expression

36. Hofmann, F., A. Ammendola, and J. Schlossmann, Rising behind NO: cGMPdependent protein kinases. J. Cell Sci., 2000. 113 (Pt 10): 1671–6. 37. Francis, S.H. and J.D. Corbin, Cyclic nucleotide-dependent protein kinases: intracellular receptors for cAMP and cGMP action. Crit. Rev. Clin. Lab. Sci., 1999. 36(4): 275–328. 38. Ruth, P., Cyclic GMP-dependent protein kinases: understanding in vivo functions by gene targeting. Pharmacol. Ther., 1999. 82(2-3): 355–72. 39. Pfeifer, A., et al., Structure and function of cGMP-dependent protein kinases. Rev. Physiol. Biochem. Pharmacol., 1999. 135: 105–49. 40. Smolenski, A., et al., Functional analysis of cGMP-dependent protein kinases I and II as mediators of NO/cGMP effects. Naunyn Schmiedebergs Arch. Pharmacol., 1998. 358(1): 134–9. 41. Lohmann, S.M., et al., Distinct and specific functions of cGMP-dependent protein kinases. Trends Biochem. Sci., 1997. 22(8): 307–12. 42. Selvaraj, N.G., et al., Evidence for the presence of cGMP-dependent protein kinase-II in human distal colon and in T84, the colonic cell line. Biochim. Biophys. Acta, 2000. 1498(1): 32–43. 43. Vaandrager, A.B., et al., Differential role of cyclic GMP-dependent protein kinase II in ion transport in murine small intestine and colon. Gastroenterology, 2000. 118(1): 108–14. 44. Wagner, C., et al., Role of cGMP-kinase II in the control of renin secretion and renin expression. J. Clin. Invest., 1998. 102(8): 1576–82. 45. Pfeifer, A., et al., Intestinal secretory defects and dwarfism in mice lacking cGMPdependent protein kinase II. Science, 1996. 274(5295): 2082–6. 46. Gudi, T., et al., Regulation of gene expression by cGMP-dependent protein kinase. Transactivation of the c-fos promoter. J. Biol. Chem., 1996. 271(9): 4597–600. 47. Pilz, R.B. and D.E. Casteel, Regulation of gene expression by cyclic GMP. Circ. Res., 2003. 93(11): 1034–46. 48. Stotz, W.H., D. Li, and R.A. Johns, Exogenous nitric oxide upregulates p21(waf1/cip1) in pulmonary microvascular smooth muscle cells. J. Vasc. Res., 2004. 41(3): 211–9. 49. Hanada, S., et al., Overexpression of protein kinase G using adenovirus inhibits cyclin E transcription and mesangial cell cycle. Am. J. Physiol. Renal Physiol., 2001. 280(5): F851–9. 50. Fischer, T.A., et al., Activation of cGMP-dependent protein kinase Ibeta inhibits interleukin 2 release and proliferation of T cell receptor-stimulated human peripheral T cells. J. Biol. Chem., 2001. 276(8): 5967–74. 51. Tanner, F.C., et al., Nitric oxide modulates expression of cell cycle regulatory proteins: a cytostatic strategy for inhibition of human vascular smooth muscle cell proliferation. Circulation, 2000. 101(16): 1982–9. 52. Komalavilas, P., et al., Activation of mitogen-activated protein kinase pathways by cyclic GMP and cyclic GMP-dependent protein kinase in contractile vascular smooth muscle cells. J. Biol. Chem., 1999. 274(48): 34301–9. 53. Firestein, B.L. and D.S. Bredt, Regulation of sensory neuron precursor proliferation by cyclic GMP-dependent protein kinase. J. Neurochem., 1998. 71(5): 1846–53. 54. Kawasaki, K., et al., Activation of the phosphatidylinositol 3-kinase/protein kinase Akt pathway mediates nitric oxide-induced endothelial cell migration and angiogenesis. Mol. Cell Biol., 2003. 23(16): 5726–37.

Nitric Oxide as a Modifier of Gene Expression

363

55. Jadeski, L.C., C. Chakraborty, and P.K. Lala, Nitric oxide-mediated promotion of mammary tumour cell migration requires sequential activation of nitric oxide synthase, guanylate cyclase and mitogen-activated protein kinase. Int. J. Cancer, 2003. 106(4): 496–504. 56. Ciani, E., M. Virgili, and A. Contestabile, Akt pathway mediates a cGMP-dependent survival role of nitric oxide in cerebellar granule neurones. J. Neurochem., 2002. 81(2): 218–28. 57. Fiscus, R.R., Involvement of cyclic GMP and protein kinase G in the regulation of apoptosis and survival in neural cells. Neurosignals, 2002. 11(4): 175–90. 58. Suenobu, N., et al., Natriuretic peptides and nitric oxide induce endothelial apoptosis via a cGMP-dependent mechanism. Arterioscler. Thromb. Vasc. Biol., 1999. 19(1): 140–6. 59. Smolenski, A., et al., Regulation of human endothelial cell focal adhesion sites and migration by cGMP-dependent protein kinase I. J. Biol. Chem., 2000. 275(33): 25723–32. 60. Taddei, S., et al., Endothelial dysfunction in hypertension. J. Nephrol., 2000. 13(3): 205–10. 61. Buchwalow, I.B., et al., Vascular smooth muscle and nitric oxide synthase. FASEB J., 2002. 16(6): 500–8. 62. Aizawa, T., et al., Role of phosphodiesterase 3 in NO/cGMP-mediated antiinflammatory effects in vascular smooth muscle cells. Circ. Res., 2003. 93(5): 406–13. 63. Zaragoza, C., et al., Activation of the mitogen activated protein kinase extracellular signal-regulated kinase 1 and 2 by the nitric oxide-cGMP-cGMP-dependent protein kinase axis regulates the expression of matrix metalloproteinase 13 in vascular endothelial cells. Mol. Pharmacol., 2002. 62(4): 927–35. 64. Tomita, H., et al., Inhibition of NO synthesis induces inflammatory changes and monocyte chemoattractant protein-1 expression in rat hearts and vessels. Arterioscler. Thromb. Vasc. Biol., 1998. 18(9): 1456–64. 65. Oddis, C.V. and M.S. Finkel, Cytokine-stimulated nitric oxide production inhibits mitochondrial activity in cardiac myocytes. Biochem. Biophys. Res. Commun., 1995. 213(3): 1002–9. 66. Takehara, Y., et al., Oxygen-dependent regulation of mitochondrial energy metabolism by nitric oxide. Arch. Biochem. Biophys., 1995. 323(1): 27–32. 67. Brookes, P.S., J.P. Bolanos, and S.J. Heales, The assumption that nitric oxide inhibits mitochondrial ATP synthesis is correct. FEBS Lett., 1999. 446(2-3): 261–3. 68. Brown, G.C., Nitric oxide and mitochondrial respiration. Biochim. Biophys. Acta, 1999. 1411(2-3): 351–69. 69. Murphy, M.P., Nitric oxide and cell death. Biochim. Biophys. Acta, 1999. 1411(2-3): 401–14. 70. Boczkowski, J., et al., Endogenous peroxynitrite mediates mitochondrial dysfunction in rat diaphragm during endotoxemia. FASEB J., 1999. 13(12): 1637–46. 71. Pervin, S., R. Singh, and G. Chaudhuri, Nitric-oxide-induced Bax integration into the mitochondrial membrane commits MDA-MB-468 cells to apoptosis: essential role of Akt. Cancer Res., 2003. 63(17): 5470–9. 72. Cooke, J.P., NO and angiogenesis. Atheroscler. Suppl., 2003. 4(4): 53–60. 73. Somoza, B., et al., Chronic l-arginine treatment reduces vascular smooth muscle cell hypertrophy through cell cycle modifications in spontaneously hypertensive rats. J. Hypertens., 2004. 22(4): 751–8.

364

Nitric Oxide, Cell Signaling, and Gene Expression

74. Kibbe, M.R., et al., Inducible nitric oxide synthase (iNOS) expression upregulates p21 and inhibits vascular smooth muscle cell proliferation through p42/44 mitogen-activated protein kinase activation and independent of p53 and cyclic guanosine monophosphate. J. Vasc. Surg., 2000. 31(6): 1214–28. 75. Yu, S.M., L.M. Hung, and C.C. Lin, cGMP-elevating agents suppress proliferation of vascular smooth muscle cells by inhibiting the activation of epidermal growth factor signaling pathway. Circulation, 1997. 95(5): 1269–77. 76. Shi, H.P., et al., The role of iNOS in wound healing. Surgery, 2001. 130(2): 225–9. 77. Romanska, H.M., et al., iNOS gene upregulation is associated with the early proliferative response of human lung fibroblasts to cytokine stimulation. J. Pathol., 2002. 197(3): 372–9. 78. Cibelli, G., et al., Nitric oxide-induced programmed cell death in human neuroblastoma cells is accompanied by the synthesis of Egr-1, a zinc finger transcription factor. J. Neurosci. Res., 2002. 67(4): 450–60. 79. Chiu, J.J., et al., Nitric oxide regulates shear stress-induced early growth response1. Expression via the extracellular signal-regulated kinase pathway in endothelial cells. Circ. Res., 1999. 85(3): 238–46. 80. Larsen, C.M., et al., Interleukin-1beta-induced rat pancreatic islet nitric oxide synthesis requires both the p38 and extracellular signal-regulated kinase 1/2 mitogen-activated protein kinases. J. Biol. Chem., 1998. 273(24): 15294–300. 81. Ishii, Y., et al., Induction of matrix metalloproteinase gene transcription by nitric oxide and mechanisms of MMP-1 gene induction in human melanoma cell lines. Int. J. Cancer, 2003. 103(2): 161–8. 82. Taimor, G., A. Rakow, and H.M. Piper, Transcription activator protein 1 (AP-1) mediates NO-induced apoptosis of adult cardiomyocytes. FASEB J., 2001. 15(13): 2518–20. 83. Pilz, R.B., et al., Nitric oxide and cGMP analogs activate transcription from AP1-responsive promoters in mammalian cells. FASEB J., 1995. 9(7): 552–8. 84. Wu, J., et al., Fos expression is induced by increased nitric oxide release in rat spinal cord dorsal horn. Neuroscience, 2000. 96(2): 351–7. 85. Jacob, A., et al., Insulin inhibits PDGF-directed VSMC migration via NO/ cGMP increase of MKP-1 and its inactivation of MAPKs. Am. J. Physiol. Cell Physiol., 2002. 283(3): C704–13. 86. Suhasini, M., et al., Cyclic-GMP-dependent protein kinase inhibits the Ras/Mitogen-activated protein kinase pathway. Mol. Cell Biol., 1998. 18(12): 6983–94. 87. He, B. and G.F. Weber, Phosphorylation of NF-kappaB proteins by cyclic GMPdependent kinase. A noncanonical pathway to NF-kappaB activation. Eur. J. Biochem., 2003. 270(10): 2174–85. 88. Wang, X. and P.J. Robinson, Cyclic GMP-dependent protein kinase substrates in rat brain. J. Neurochem., 1995. 65(2): 595–604. 89. Kalra, D., et al., Nitric oxide provokes tumor necrosis factor-alpha expression in adult feline myocardium through a cGMP-dependent pathway. Circulation, 2000. 102(11): 1302–7. 90. Gonzalez Bosc, L.V., et al., Intraluminal pressure is a stimulus for NFATc3 nuclear accumulation: role of calcium, endothelium-derived nitric oxide, and cGMPdependent protein kinase. J. Biol. Chem., 2004. 279(11): 10702–9. 91. Fiedler, B., et al., Inhibition of calcineurin-NFAT hypertrophy signaling by cGMPdependent protein kinase type I in cardiac myocytes. Proc. Natl. Acad. Sci. USA, 2002. 99(17): 11363–8.

Nitric Oxide as a Modifier of Gene Expression

365

92. Sauzeau, V., et al., RhoA expression is controlled by nitric oxide through cGMPdependent protein kinase activation. J. Biol. Chem., 2003. 278(11): 9472–80. 93. Chan, S.H., et al., Nitric oxide regulates c-fos expression in nucleus tractus solitarii induced by baroreceptor activation via cGMP-dependent protein kinase and cAMP response element-binding protein phosphorylation. Mol. Pharmacol., 2004. 65(2): 319–25. 94. Gudi, T., et al., cGMP-dependent protein kinase inhibits serum-response elementdependent transcription by inhibiting rho activation and functions. J. Biol. Chem., 2002. 277(40): 37382–93. 95. Subbaramaiah, K., et al., Microsomal prostaglandin E synthase-1 is overexpressed in inflammatory bowel disease. Evidence for involvement of the transcription factor Egr-1. J. Biol. Chem., 2004. 279(13): 12647–58. 96. Esteve, L., et al., Cyclic GMP-dependent protein kinase potentiates serotonininduced Egr-1 binding activity in PC12 cells. Cell Signal., 2001. 13(6): 425–32. 97. Casteel, D.E., et al., cGMP-dependent protein kinase I beta physically and functionally interacts with the transcriptional regulator TFII-I. J. Biol. Chem., 2002. 277(35): 32003–14. 98. Kloss, S., R. Srivastava, and A. Mulsch, Down-regulation of soluble guanylyl cyclase expression by cyclic AMP is mediated by mRNA-stabilizing protein HuR. Mol. Pharmacol., 2004. 65(6): 1440–51. 99. Akool el, S., et al., Nitric oxide increases the decay of matrix metalloproteinase 9 mRNA by inhibiting the expression of mRNA-stabilizing factor HuR. Mol. Cell Biol., 2003. 23(14): 4901–16. 100. Huwiler, A., et al., ATP potentiates interleukin-1 beta-induced MMP-9 expression in mesangial cells via recruitment of the ELAV protein HuR. J. Biol. Chem., 2003. 278(51): 51758–69. 101. Rodríguez-Pascual, F., et al., Complex contribution of the 3′-untranslated region to the expressional regulation of the human inducible nitric-oxide synthase gene. Involvement of the RNA-binding protein HuR. J. Biol. Chem., 2000. 275(34): 26040–9. 102. Denkert, C., et al., Overexpression of the embryonic-lethal abnormal vision-like protein HuR in ovarian carcinoma is a prognostic factor and is associated with increased cyclooxygenase 2 expression. Cancer Res., 2004. 64(1): 189–95. 103. Levy, A.P., Hypoxic Regulation of VEGF mRNA Stability by RNA-binding Proteins. Trends Cardiovasc. Med., 1998. 8(6): 246–50. 104. Tang, K., E.C. Breen, and P.D. Wagner, Hu protein R-mediated posttranscriptional regulation of VEGF expression in rat gastrocnemius muscle. Am. J. Physiol. Heart Circ. Physiol., 2002. 283(4): H1497–504. 105. Nabors, L.B., et al., Tumor necrosis factor alpha induces angiogenic factor upregulation in malignant glioma cells: a role for RNA stabilization and HuR. Cancer Res., 2003. 63(14): 4181–7. 106. Nabors, L.B., et al., HuR, a RNA stability factor, is expressed in malignant brain tumors and binds to adenine- and uridine-rich elements within the 3' untranslated regions of cytokine and angiogenic factor mRNAs. Cancer Res., 2001. 61(5): 2154–61. 107. Dixon, D.A., et al., Altered expression of the mRNA stability factor HuR promotes cyclooxygenase-2 expression in colon cancer cells. J. Clin. Invest., 2001. 108(11): 1657–65.

366

Nitric Oxide, Cell Signaling, and Gene Expression

108. Sengupta, S., et al., The RNA-binding protein HuR regulates the expression of cyclooxygenase-2. J. Biol. Chem., 2003. 278(27): 25227–33. 109. Arnold, W.P., et al., Nitric oxide activates guanylate cyclase and increases guanosine 3′:5′-cyclic monophosphate levels in various tissue preparations. Proc. Natl. Acad. Sci. USA, 1977. 74(8): 3203–7. 110. Gruetter, C.A., et al., Relaxation of bovine coronary artery and activation of coronary arterial guanylate cyclase by nitric oxide, nitroprusside and a carcinogenic nitrosoamine. J. Cyclic. Nucleotide Res., 1979. 5(3): 211–24. 111. Ballou, D.P., et al., eds. Revisiting the kinetics of nitric oxide (NO) binding to soluble guanylate cyclase: the simple NO-binding model is incorrect. Proc. Natl. Acad. Sci. USA. Vol. 99. 2002. 12097–101. 112. Zhao, Y., et al., A molecular basis for nitric oxide sensing by soluble guanylate cyclase. Proc. Natl. Acad. Sci. USA, 1999. 96(26): 14753–8. 113. Cooper, C.E., Nitric oxide and iron proteins. Biochim. Biophys. Acta, 1999. 1411(2-3): 290–309. 114. Forman, H.J., M. Torres, and J. Fukuto, Redox signaling. Mol. Cell Biochem., 2002. 234-235(1-2): 49–62. 115. Cooper, C.E. and N.A. Davies, Effects of nitric oxide and peroxynitrite on the cytochrome oxidase K(m) for oxygen: implications for mitochondrial pathology. Biochim. Biophys. Acta, 2000. 1459(2-3): 390–6. 116. Brown, G.C., Regulation of mitochondrial respiration by nitric oxide inhibition of cytochrome c oxidase. Biochim. Biophys. Acta, 2001. 1504(1): 46–57. 117. Moncada, S. and J.D. Erusalimsky, Does nitric oxide modulate mitochondrial energy generation and apoptosis? Nat. Rev. Mol. Cell Biol., 2002. 3(3): 214–20. 118. Hagen, T., et al., Redistribution of intracellular oxygen in hypoxia by nitric oxide: effect on HIF1α. Science, 2003. 302(5652): 1975–8. 119. Semenza, G.L., HIF-1: mediator of physiological and pathophysiological responses to hypoxia. J. Appl. Physiol., 2000. 88: 1474–80. 120. Martínez-Ruiz, A. and S. Lamas, S-nitrosylation: a potential new paradigm in signal transduction. Cardiovasc. Res., 2004. 62(1): 43–52. 121. Pryor, W.A., et al., Oxidation of thiols by nitric oxide and nitrogen dioxide: synthetic utility and toxicological implications. J. Org. Chem., 1982. 47: 156–9). 122. Wink, D.A., et al., Reaction kinetics for nitrosation of cysteine and glutathione in aerobic nitric oxide solutions at neutral pH. Insights into the fate and physiological effects of intermediates generated in the NO/O2 reaction. Chem. Res. Toxicol., 1994. 7(4): 519–25. 123. Wink, D.A., et al., Reactions of the bioregulatory agent nitric oxide in oxygenated aqueous media: determination of the kinetics for oxidation and nitrosation by intermediates generated in the NO/O2 reaction. Chem. Res. Toxicol., 1993. 6(1): 23–7. 124. Hogg, N., The biochemistry and physiology of S-nitrosothiols. Annu. Rev. Pharmacol. Toxicol., 2002. 42(1): 585–600. 125. Bogdan, C., Nitric oxide and the regulation of gene expression. Trends Cell Biol., 2001. 11(2): 66–75. 126. Matthews, J.R., et al., Thioredoxin regulates the DNA binding activity of NFkappa B by reduction of a disulphide bond involving cysteine 62. Nucleic Acids Res., 1992. 20(15): 3821–30.

Nitric Oxide as a Modifier of Gene Expression

367

127. Hayashi, T., Y. Ueno, and T. Okamoto, Oxidoreductive regulation of nuclear factor kappa B. Involvement of a cellular reducing catalyst thioredoxin. J. Biol. Chem., 1993. 268(15): 11380–8. 128. DelaTorre, A., R.A. Schroeder, and P.C. Kuo, Alteration of NF-kappa B p50 DNA binding kinetics by S-nitrosylation. Biochem. Biophys. Res. Commun., 1997. 238(3): 703–6. 129. DelaTorre, A., et al., Differential effects of nitric oxide-mediated S-nitrosylation on p50 and c-jun DNA binding. Surgery, 1998. 124(2): 137–42. 130. Marshall, H.E. and J.S. Stamler, Inhibition of NF-kappa B by S-nitrosylation. Biochemistry, 2001. 40(6): 1688–93. 131. DelaTorre, A., et al., Endotoxin-mediated S-nitrosylation of p50 alters NF-kappaB-dependent gene transcription in ANA-1 murine macrophages. J. Immunol., 1999. 162(7): 4101–8. 132. Park, S.K., H.L. Lin, and S. Murphy, Nitric oxide regulates nitric oxide synthase2 gene expression by inhibiting NF-kappaB binding to DNA. Biochem. J., 1997. 322(Pt 2): 609–13. 133. Marshall, H.E. and J.S. Stamler, Nitrosative stress-induced apoptosis through inhibition of NF-kappaB. J. Biol. Chem., 2002. 277(37): 34223–8. 134. Reynaert, N.L., et al., Nitric oxide represses inhibitory κB kinase through Snitrosylation. Proc. Natl. Acad. Sci. USA, 2004. 101(24): 8945–50. 135. Melino, G., et al., Nitric oxide inhibits apoptosis via AP-1-dependent CD95L transactivation. Cancer Res., 2000. 60(9): 2377–83. 136. Kröncke, K.-D., Zinc finger proteins as molecular targets for nitric oxide-mediated gene regulation. Antioxid. Redox. Signal., 2001. 3(4): 565–575. 137. Berendji, D., et al., Zinc finger transcription factors as molecular targets for nitric oxide-mediated immunosuppression: inhibition of IL-2 gene expression in murine lymphocytes. Mol. Med., 1999. 5(11): 721–30. 138. Kröncke, K.D. and C. Carlberg, Inactivation of zinc finger transcription factors provides a mechanism for a gene regulatory role of nitric oxide. FASEB J., 2000. 14(1): 166–73. 139. Galigniana, M.D., G. Piwien-Pilipuk, and J. Assreuy, Inhibition of glucocorticoid receptor binding by nitric oxide. Mol. Pharmacol., 1999. 55(2): 317–23. 140. Wang, S., et al., A Sp1 binding site of the tumor necrosis factor alpha promoter functions as a nitric oxide response element. J. Biol. Chem., 1999. 274(47): 33190–3. 141. Maret, W., The function of zinc metallothionein: a link between cellular zinc and redox state. J. Nutr., 2000. 130(5S Suppl): 1455S–8S. 142. Berendji, D., et al., Nitric oxide mediates intracytoplasmic and intranuclear zinc release. FEBS Lett., 1997. 405(1): 37–41. 143. Pearce, L.L., et al., Role of metallothionein in nitric oxide signaling as revealed by a green fluorescent fusion protein. Proc. Natl. Acad. Sci. USA, 2000. 97(1): 477–82. 144. St. Croix, C.M., et al., Nitric oxide-induced changes in intracellular zinc homeostasis are mediated by metallothionein/thionein. Am. J. Physiol. Lung Cell Mol. Physiol., 2002. 282(2): L185–92. 145. Chen, Y., et al., S-nitrosothiols react preferentially with zinc thiolate clusters of metallothionein III through transnitrosation. Biochemistry, 2002. 41(26): 8360–7.

368

Nitric Oxide, Cell Signaling, and Gene Expression

146. Aravindakumar, C.T., J. Ceulemans, and M. De Ley, Nitric oxide induces Zn2+ release from metallothionein by destroying zinc-sulphur clusters without concomitant formation of S-nitrosothiol. Biochem. J., 1999. 344 Pt 1: 253–8. 147. Gao, C., et al., S-nitrosylation of heterogeneous nuclear ribonucleoprotein A/B regulates osteopontin transcription in endotoxin-stimulated murine macrophages. J. Biol. Chem., 2004. 279(12): 11236–43. 148. Klatt, and S. Lamas, Regulation of protein function by S-glutathiolation in response to oxidative and nitrosative stress. Eur. J. Biochem., 2000. 267(16): 4928–44. 149. Arnelle, D.R. and J.S. Stamler, NO+, NO, and NO− donation by S-nitrosothiols: implications for regulation of physiological functions by S-nitrosylation and acceleration of disulfide formation. Arch. Biochem. Biophys., 1995. 318(2): 279–85. 150. Wong, P.S., et al., Reaction between S-nitrosothiols and thiols: generation of nitroxyl (HNO) and subsequent chemistry. Biochemistry, 1998. 37(16): 5362–71. 151. Hogg, N., The kinetics of S-transnitrosation—a reversible second-order reaction. Anal. Biochem., 1999. 272(2): 257–62. 152. Liu, Z., et al., S-transnitrosation reactions are involved in the metabolic fate and biological actions of nitric oxide. J. Pharmacol. Exp. Ther., 1998. 284(2): 526–34. 153. Turko, I.V. and F. Murad, Protein nitration in cardiovascular diseases. Pharmacol. Rev., 2002. 54(4): 619–34. 154. Ischiropoulos, H., Biological selectivity and functional aspects of protein tyrosine nitration. Biochem. Biophys. Res. Commun., 2003. 305(3): 776–83. 155. Monteiro, H.P., Signal transduction by protein tyrosine nitration: competition or cooperation with tyrosine phosphorylation-dependent signaling events? Free Radic. Biol. Med., 2002. 33(6): 765–73. 156. Paul-Clark, M.J., et al., Glucocorticoid receptor nitration leads to enhanced antiinflammatory effects of novel steroid ligands. J. Immunol., 2003. 171(6): 3245–52. 157. Mailfait, S., et al., Critical role of tyrosine 277 in the ligand-binding and transactivating properties of retinoic acid receptor alpha. Biochemistry, 2000. 39(9): 2183–92. 158. Shibuya, A., et al., Nitration of PPARgamma inhibits ligand-dependent translocation into the nucleus in a macrophage-like cell line, RAW 264. FEBS Lett., 2002. 525(1-3): 43–7. 159. Matata, B.M. and M. Galiñanes, Peroxynitrite is an essential component of cytokines production mechanism in human monocytes through modulation of nuclear factor-kappa B DNA binding activity. J. Biol. Chem., 2002. 277(3): 2330–5. 160. Klotz, L.-O., Schroeder, and H. Sies, Peroxynitrite signaling: receptor tyrosine kinases and activation of stress-responsive pathways, Free Radic. Biol. Med., 2002. 33(6): 737–43. 161. Minetti, M., C. Mallozzi, and A.M.M. Di Stasi, Peroxynitrite activates kinases of the src family and upregulates tyrosine phosphorylation signaling,. Free Radic. Biol. Med., 2002. 33(6): 744–54. 162. Llovera, M., et al., Impaired response to interferon-gamma in activated macrophages due to tyrosine nitration of STAT1 by endogenous nitric oxide. Br. J. Pharmacol., 2001. 132(2): 419–26. 163. Inoue, T., et al., cGMP upregulates nitric oxide synthase expression in vascular smooth muscle cells. Hypertension, 1995. 25(4 Pt 2): 711–4.

Nitric Oxide as a Modifier of Gene Expression

369

164. Choi, S.H., et al., Zaprinast, an inhibitor of cGMP-selective phosphodiesterases, enhances the secretion of TNF-alpha and IL-1beta and the expression of iNOS and MHC class II molecules in rat microglial cells. J. Neurosci. Res., 2002. 67(3): 411–21. 165. Park, S.W., et al., The effect of nitric oxide on cyclooxygenase-2 (COX-2) overexpression in head and neck cancer cell lines. Int. J. Cancer, 2003. 107(5): 729–38. 166. Chang, M.S., et al., YC-1-induced cyclooxygenase-2 expression is mediated by cGMP-dependent activations of Ras, phosphoinositide-3-OH-kinase, Akt, and nuclear factor-kappaB in human pulmonary epithelial cells. Mol. Pharmacol., 2004. 66(3): 561–71. 167. Cheng, H.F., et al., Nitric oxide regulates renal cortical cyclooxygenase-2 expression. Am. J. Physiol. Renal Physiol., 2000. 279(1): F122–9. 168. Kelly, L.K., et al., Nitric oxide decreases endothelin-1 secretion through the activation of soluble guanylate cyclase. Am. J. Physiol. Lung Cell Mol. Physiol., 2004. 286(5): L984–91. 169. Soff, G.A., et al., Smooth muscle cell expression of type I cyclic GMP-dependent protein kinase is suppressed by continuous exposure to nitrovasodilators, theophylline, cyclic GMP, and cyclic AMP. J. Clin. Invest., 1997. 100(10): 2580–7. 170. Sellak, H., et al., Sp1 transcription factor as a molecular target for nitric oxideand cyclic nucleotide-mediated suppression of cGMP-dependent protein kinaseIalpha expression in vascular smooth muscle cells. Circ. Res., 2002. 90(4): 405–12. 171. Browner, N.C., et al., Regulation of cGMP-dependent protein kinase expression by soluble guanylyl cyclase in vascular smooth muscle cells. J. Biol. Chem., 2004. 279(45): 46631–6. 172. Hum, D., et al., Characterization of a cGMP-response element in the guanylyl cyclase/natriuretic peptide receptor A gene promoter. Hypertension, 2004. 43(6): 1270–8. 173. Wollert, K.C., et al., Gene transfer of cGMP-dependent protein kinase I enhances the antihypertrophic effects of nitric oxide in cardiomyocytes. Hypertension, 2002. 39(1): 87–92. 174. Hellmuth, M., et al., Nitric oxide differentially regulates pro- and anti-angiogenic markers in DLD-1 colon carcinoma cells. FEBS Lett., 2004. 563(1-3): 98–102. 175. Ramanathan, M., A. Giladi, and S.J. Leibovich, Regulation of vascular endothelial growth factor gene expression in murine macrophages by nitric oxide and hypoxia. Exp. Biol. Med. (Maywood), 2003. 228(6): 697–705. 176. Kiemer, A.K., et al., Inhibition of p38 MAPK activation via induction of MKP1: atrial natriuretic peptide reduces TNF-alpha-induced actin polymerization and endothelial permeability. Circ. Res., 2002. 90(8): 874–81. 177. Pawliczak, R., et al., p11 expression in human bronchial epithelial cells is increased by nitric oxide in a cGMP-dependent pathway involving protein kinase G activation. J. Biol. Chem., 2001. 276(48): 44613–21. 178. Deguchi, A., W.J. Thompson, and I.B. Weinstein, Activation of protein kinase G is sufficient to induce apoptosis and inhibit cell migration in colon cancer cells. Cancer Res., 2004. 64(11): 3966–73. 179. Doi, K., et al., C-type natriuretic peptide induces redifferentiation of vascular smooth muscle cells with accelerated reendothelialization. Arterioscler. Thromb. Vasc. Biol., 2001. 21(6): 930–6.

370

Nitric Oxide, Cell Signaling, and Gene Expression

180. Gu, M., J. Lynch, and Brecher, Nitric oxide increases p21(Waf1/Cip1) expression by a cGMP-dependent pathway that includes activation of extracellular signalregulated kinase and p70(S6k). J. Biol. Chem., 2000. 275(15): 11389–96. 181. Sato, J., et al., eNOS gene transfer to vascular smooth muscle cells inhibits cell proliferation via upregulation of p27 and p21 and not apoptosis. Cardiovasc. Res., 2000. 47(4): 697–706. 182. Bauer, P.M., G.M. Buga, and L.J. Ignarro, Role of p42/p44 mitogen-activatedprotein kinase and p21waf1/cip1 in the regulation of vascular smooth muscle cell proliferation by nitric oxide. Proc. Natl. Acad. Sci. USA, 2001. 98(22): 12802–7. 183. Andoh, T., C.C. Chiueh, and P.B. Chock, Cyclic GMP-dependent protein kinase regulates the expression of thioredoxin and thioredoxin peroxidase-1 during hormesis in response to oxidative stress-induced apoptosis. J. Biol. Chem., 2003. 278(2): 885–90. 184. Nisoli, E., et al., Mitochondrial biogenesis in mammals: the role of endogenous nitric oxide. Science, 2003. 299(5608): 896–9. 185. Chen, H.H. and D.L. Wang, Nitric oxide inhibits matrix metalloproteinase-2 expression via the induction of activating transcription factor 3 in endothelial cells. Mol. Pharmacol., 2004. 65(5): 1130–40. 186. Choe, T., et al., Inhibition of matrix metalloproteinase-1 and -2 expression using nitric oxide synthase inhibitors in UV-irradiated human dermal fibroblasts. J. Cosmet. Sci., 2003. 54(3): 229–38. 187. Orucevic, A., et al., Nitric-oxide production by murine mammary adenocarcinoma cells promotes tumor-cell invasiveness. Int. J. Cancer, 1999. 81(6): 889–96. 188. Aicher, A., et al., Essential role of endothelial nitric oxide synthase for mobilization of stem and progenitor cells. Nat. Med., 2003. 9(11): 1370–6. 189. Marcet-Palacios, M., et al., Nitric oxide and cyclic GMP increase the expression of matrix metalloproteinase-9 in vascular smooth muscle. J. Pharmacol. Exp. Ther., 2003. 307(1): 429–36. 190. Knipp, B.S., et al., Increased MMP-9 expression and activity by aortic smooth muscle cells after nitric oxide synthase inhibition is associated with increased nuclear factor-kappaB and activator protein-1 activity. J. Surg. Res., 2004. 116(1): 70–80. 191. Abdelaziz, N., et al., Nitric oxide attenuates the expression of transforming growth factor-beta(3) mRNA in rat cardiac fibroblasts via destabilization. Hypertension, 2001. 38(2): 261–6. 192. Sparkman, L. and V. Boggaram, Nitric oxide increases IL-8 gene transcription and mRNA stability to enhance IL-8 gene expression in lung epithelial cells. Am. J. Physiol. Lung Cell Mol. Physiol., 2004. 287(4): L764–73. 193. Phillips, P.G. and L.M. Birnby, Nitric oxide modulates caveolin-1 and matrix metalloproteinase-9 expression and distribution at the endothelial cell/tumor cell interface. Am. J. Physiol. Lung Cell Mol. Physiol., 2004. 286(5): L1055–65. 194. dAhluwalia, A., et al., Antiinflammatory activity of soluble guanylate cyclase: cGMP-dependent down-regulation of P-selectin expression and leukocyte recruitment. Proc. Natl. Acad. Sci. USA, 2004. 101(5): 1386–91. 195. Desai, A., et al., Nitric oxide modulates MCP-1 expression in endothelial cells: implications for the pathogenesis of pulmonary granulomatous vasculitis. Inflammation, 2003. 27(4): 213–23.

Oxide and Post17 Nitric Transcriptional Control of Gene Expression by the IRE/IRP System Leonor Oliveira, Cécile Bouton, and Jean-Claude Drapier Institut de Chimie des Substances Naturelles, CNRS, Gif-sur-Yvette, France

CONTENTS 17.1 The Post-Transcriptional Regulation of Iron Metabolism in Mammalian Cells ...................................................................................372 17.1.1 Coordination of Cellular Iron Metabolism by Post-Transcriptional Regulation of Ferritin and Transferrin Receptor Expression ...................................................................372 17.1.2 The Iron-Responsive Elements (IREs) ......................................373 17.1.3 The Iron Regulatory Proteins as Major Regulators of Intracellular Iron Homeostasis ...................................................374 17.1.4 Implication of IRPs in Iron-Associated Disorders and Lessons from IRP Mutant Mice ................................................377 17.2 Interplay between Nitric Oxide (NO) and IRPs ...................................379 17.2.1 NO as a Modulator of [Fe-S] Cluster-Containing Proteins ......379 17.2.2 NO and the Post-Translational Regulation of IRP1 Activities ......380 17.2.3 Mechanism Underlying NO Modulation of IRP1 RNA-Binding Activity ...............................................................381 17.2.4 Regulation of IRP1 Protein Levels by NO ................................383 17.2.5 Modulation of IRP2 by NO .......................................................384 17.2.6 Functional Impact of the NO/IRP Regulatory System on Ferritin and TfR Expression ......................................................385 17.3 A Look Ahead .......................................................................................386 References ...............................................................................................387

371

372

Nitric Oxide, Cell Signaling, and Gene Expression

17.1 THE POST-TRANSCRIPTIONAL REGULATION OF IRON METABOLISM IN MAMMALIAN CELLS 17.1.1 COORDINATION OF CELLULAR IRON METABOLISM BY POST-TRANSCRIPTIONAL REGULATION OF FERRITIN AND TRANSFERRIN RECEPTOR EXPRESSION Many proteins containing iron or requiring iron for their activity play determinant roles in crucial biological functions. They include hemoproteins involved in oxygen binding and transport (e.g., hemoglobin and myoglobin), oxygen metabolism (e.g., oxidases, peroxidase, and catalase) and electron transfer (e.g., cytochromes). Other non-heme iron-containing proteins catalyze key reactions involved in energy metabolism, such as [Fe-S] proteins of the electron transport chain, and DNA synthesis, such as ribonucleotide reductase (reviewed in Griffiths, 1987; Cammack et al., 1990). The characteristics that make iron an excellent catalyst also make it a potentially hazardous metal, however. In the presence of oxygen, the one-electron redox reactions catalyzed by iron can lead to the formation of hydroxyl radicals which may cause lipid peroxidation, DNA strand breaks and damage to proteins (Halliwell and Gutteridge, 1984). To avoid both iron toxicity and starvation, cells must therefore keep free iron levels as low as possible, while maintaining an adequate concentration for metabolic requirements. This has been achieved during evolution by the development of systems specialized to deliver iron to the cells, transport it and store it in a soluble, nontoxic form. In mammals, there is a strict regulation of the proteins involved in the uptake, utilization, and storage of iron. This is mainly achieved through a highly regulated system in which iron availability coordinately regulates the expression of transferrin receptor (TfR*) and ferritin through a regulatory feed-back loop. Thus, when iron supply is high, the number of TfR at the cell surface decreases to reduce the incorporation of iron into the cell, and the levels of ferritin rise to store more iron. Conversely, in cells that are iron-deprived, the levels of TfR increase, thus allowing cells to take more iron and the levels of ferritin fall, keeping iron available for metabolic utilization. This fine regulation of ferritin and TfR by iron occurs mainly at the post-transcriptional level and is under the control of a tightly regulated genetic regulatory system (reviewed in Klausner et al., 1993 and in Hentze et al., 2004). It is important, however, to mention that ferritin and TfR expression is also subject to transcriptional regulation and may also be modulated by iron-independent regulatory mechanisms (reviewed in Ponka and Lok, 1999).

* Another TfR (TfR2) was recently discovered. It is restricted to certain tissues, such as hepatocytes and erythroid cells, and its messenger ribonucleic acid (mRNA) does not possess an iron-responsive element (IRE). Thus, TfR in this review will be referred to as TfR1, which is ubiquitously expressed.

Nitric Oxide and Post-Transcriptional Control of Gene Expression

373

17.1.2 THE IRON-RESPONSIVE ELEMENTS (IRES) Intensive research aimed at characterizing the molecular mechanisms by which iron regulates ferritin and TfR expression led to the discovery that the irondependent post-transcriptional regulation of both ferritin and TfR was mediated by conserved cis-acting elements located in the untranslated regions of their messenger ribonucleic acids (mRNAs). These elements were first identified in ferritin H- and L-chain mRNA by sequence analysis (Aziz and Munro, 1987; Hentze et al., 1987), and later referred to as the iron-responsive element or IRE (Figure 17.1). Ferritin mRNA contains one IRE in the 5′ end of its UTR, close to the cap site where translational factors initially bind. Five similar IRE sequences were also identified in the 3′UTR of TfR mRNA (Casey et al., 1988) of which three have a critical role in regulation of TfR mRNA stability (Müllner and Kühn, 1988; Casey et al., 1989). IREs present a high degree of phylogenetic conservation among animals. They appear to be absent in plant ferritin mRNA, the regulation of which occurs either transcriptionally or post-translationally (reviewed in Theil, 1994).

B

A

G

U G

A

G

U

C

C AU AU CG UG

G

A Loop

C

N NN NN NN NN

Upper stem

UA C G

NN C NN NN NN NN NN 5′ 3′

Bulge

Lower stem of variable length

U U

C CG CG UA UA GC GC GC 5′ 3′

FIGURE 17.1 The IRE motifs. (A) IREs are RNA stem-loop structures composed of a highly conserved loop sequence, 5′-CAGUGN-3′, where N can be any base but G, an upper stem of 5-base-pair nucleotides not conserved between species, an unpaired C 5′ in the stem and a lower stem that may vary in size depending on the IRE examined. (B) The IRE of human H-chain ferritin is depicted.

374

Nitric Oxide, Cell Signaling, and Gene Expression

IREs have also been found in the 5′UTR of the mRNAs encoding the erythroid 5-aminolevulinate synthase (eALAS) (Cox et al., 1991; Dandekar et al., 1991), and mitochondrial aconitase (m-aconitase) (Dandekar et al., 1991; Zheng et al., 1992) and of mitochondrial succinate dehydrogenase subunit b of Drosophila melanogaster (Kohler et al., 1995, Gray et al., 1996 reviewed in Hentze and Kühn, 1996). The functional impact of the IRE-dependent inhibition of m-aconitase translation has recently been discussed by Eisenstein and Ross (2003). One IRE has also been identified in the 3'UTR of one of the two alternatively spliced transcripts of the iron-transporter DMT1, which is involved both in intestinal absorption and in endosomal release of iron (Fleming et al., 1998; Fleming et al., 1998; Gunshin et al., 1997). An IRE-like motif is also located in the 5'UTR of the mRNA of Ireg1 (for iron-regulated mRNA) (McKie et al., 2000), a duodenal protein involved in iron transport from the basolateral membrane of epithelial cells to the circulation, which is also referred to as ferroportin (Donovan et al., 2000). Whether IRE of DMT1 and Ireg-1 mRNA is functionally involved in irondependent control of these proteins still needs to be confirmed. As depicted in Figure 17.2, IREs are found mainly in the untranslated region of mRNA encoding proteins involved in uptake, storage, transport, utilization, and export of iron.

17.1.3 THE IRON REGULATORY PROTEINS AS MAJOR REGULATORS OF INTRACELLULAR IRON HOMEOSTASIS The identification and characterization of cellular factors that interact with IREs was the key to understanding the mechanisms involved in the post-transcriptional regulation of ferritin and TfR by iron (Leibold and Munro, 1988; Müllner et al., 1989). The IREs are recognized by cytosolic RNA-binding proteins, known as iron regulatory proteins (IRPs). Two related IRPs, IRP1 and IRP2, have been identified to date. Binding of IRPs to the IREs in the 5′UTR of ferritin mRNA blocks translation initiation by preventing the association of the 43S translation pre-initiation complex (Gray and Hentze, 1994). Then, it was reported that the IRP-IRE complex prevents the recruitment of the small ribosomal subunit by the cap-binding complex eIF4F, probably by steric hindrance (Muckenthaler et al., 1998). Binding of IRP to TfR mRNA IREs is thought to protect TfR mRNA against degradation, by preventing access of an endonuclease with a cleavage site that is close to the IREs (Binder et al., 1994) (Figure 17.3). The mechanisms by which iron regulates IRP1 IRE-binding have been the subject of intensive investigation. Importantly, it was found that IRP1 IRE-binding activity lost by treatment of cells with iron sources could be recovered following exposure of cell lysates to reducing agents like 2-mercaptoethanol and dithiothreitol (Hentze et al., 1989). These results strongly suggested that iron levels regulate IRP1 by a post-translational mechanism, without changes in the total amount of protein. A fundamental clue to the regulation of IRP1 by iron was revealed in 1991, when computer-based analysis of data banks indicated a striking level of sequence conservation between IRP1 and mitochondrial aconitase from pig heart and Saccharomyces cerevisiae (Rouault et al., 1991; Hentze and Argos,

Nitric Oxide and Post-Transcriptional Control of Gene Expression

375

Tf-Fe

TfR

Ferritin DMT1 “Labile iron pool”

heme

ALAS2

TCA cycle

m-aco

Fe

Endosome

Mitochondrion

lreg1

Fe

FIGURE 17.2 Schematic representation of mammalian cell iron metabolism with the IRE/IRP system-dependent checkpoint control. This simplified scheme depicts the sites (marked by a ) where IRPs may function as translation regulators by binding to IRE(s) of mRNA of several proteins involved in iron homeostasis and energy metabolism. ALAS2 is one of the 2 genes that encodes 5-aminolevulinate synthase, the first enzyme involved in heme synthesis. ALAS2 is expressed only in erythrocytic cells.

IRPs IRE Ferritin mRNA

3′

5′

Translation inhibition

Ferritin

3′ transferrin receptor mRNA

5′

Stabilization

Transferrin receptor

FIGURE 17.3 The trans-regulatory activity of IRPs. Formation of IRE/IRP complexes on the 5′UTR of ferritin mRNA inhibits early steps of translation (Muckenthaler et al., 1998). IREs located in the 3′ UTR of TfR mRNA allow stabilization of the mRNA upon binding of IRPs.

376

Nitric Oxide, Cell Signaling, and Gene Expression

1991). Thus, by the end of 1992, evidence was strong enough to conclude that IRP1 and cytosolic aconitase were the same protein. Gel retardation assays and measurement of aconitase activity in cell extracts indicated that there was a reciprocal relationship between aconitase activity and IRP1 RNA-binding activity, which was related to the iron status of the cell. These results indicated that the RNA-binding state of IRP1 and its enzymatic activity were mutually exclusive, or, to put it another way, an individual molecule cannot have both activities at once. Studies were also undertaken to determine the status of the [Fe-S] cluster in the RNA-binding form of IRP1. It was reported that the removal of the labile fourth iron atom (Fea) results in loss of aconitase activity, but is not sufficient to induce IRP1 IRE-binding activity, suggesting that a more extensive cluster alteration is required to reproduce the high affinity of IRP1 IRE-binding seen in cells following iron depletion. A direct proof for the crucial role of the [Fe-S] cluster in regulating IRP1 functions was provided by site-directed mutagenesis experiments. Mutation of any of the three cysteines predicted to coordinate the [Fe-S] cluster in IRP1 (cys437, cys503, and cys506) to serine residues resulted in complete loss of aconitase activity (Philpott et al., 1993; Hirling et al., 1994). IRP2 differs from IRP1 by the insertion of a cysteine-rich 73-amino-acid domain near its amino terminus (Guo et al., 1995), and by the absence of a [Fe-S] cluster and, in turn, of aconitase activity (Guo et al., 1994). IRP2 has long been considered as weakly expressed in tissues and cell lines. It has recently been reported, however, that IRP2 protein (and activity) may have been underestimated due to its sensitivity to oxygen, which is more pronounced under bench-scale conditions than in tissues (Bourdon et al., 2003). As depicted in Figure 17.4, in contrast to IRP1, changes in the IRE-binding activity of IRP2 resulting from alterations in iron availability do not result from a post-translational modification but from degradation by the proteasome (Guo et al., 1995; Iwai et al., 1995). It was claimed that the 73-amino-acid domain unique to IRP2 is required for degradation of IRP2 in iron-replete cells because deletion of this sequence abolishes the iron-induced rapid turnover of IRP2. More recently, it was finally demonstrated by a mass spectrometry study that this domain forms an “iron binding site” with one cysteine residue oxidized to dehydrocysteine, which renders the protein susceptible to ubiquitination and targets IRP2 for degradation (Kang et al., 2003) (Figure 17.5). According to Yamanaka et al. (2003), upon binding heme, this "iron degradation domain" would be recognized by the RINGfinger protein HOIL-1 with E3 ubiquitin ligase activity and degraded by the proteasome. This well-established model was recently challenged by two reports, however, indicating that the 73-amino-acid domain is not required for iron or oxygen-mediated IRP2 degradation but, instead, that a 2-oxoglutarate-dependent oxygenase is involved in this process (Hanson, et al., 2003, Wang et al., 2004, Pantopoulos, 2004). The molecular mechanism explaining IRP2 regulation thus appears less clear than expected.

Nitric Oxide and Post-Transcriptional Control of Gene Expression

377

FIGURE 17.4 Quintessential difference between IRP1 and IRP2 with regard to induction of IRE-binding activity. Top: conversion of the aconitase form of IRP1 into a transregulatory factor depends on complete [4Fe-4S] cluster disassembly. Bottom: IRP2 mRNAbinding activity depends on protein stability.

17.1.4 IMPLICATION OF IRPS IN IRON-ASSOCIATED DISORDERS AND LESSONS FROM IRP MUTANT MICE Mutations in the IRE sequence affecting IRP-IRE interaction and the regulation of ferritin synthesis have been reported in the hereditary hyperferritinemia-cataract syndrome (Beaumont et al., 1995). This autosomal dominant disorder is associated with elevated serum ferritin in the absence of iron overload and bilateral cataract. A mutation in the IRE of the L-ferritin mRNA has been found in members of families with dominantly inherited hyperferritinemia and cataract, consisting of a single- or double-point mutation in the IRE loop, lower stem, or bulge (reviewed in Fodinger and Sunder-Plassmann, 1999). A dysregulation of IRP activity may also occur in hemochromatosis. Hereditary hemochromatosis is an autosomal recessive iron-overload disease characterized by inappropriately high intestinal iron absorption. In the prototypic disease, the hemochromatosis gene (HFE gene) encodes a nonclassic major histocompatibility complex (MHC) class I-like protein. A single-point mutation, C282Y, which accounts for a great majority of cases of hereditary hemochromatosis, prevents the association

378

Nitric Oxide, Cell Signaling, and Gene Expression de novo synthesis

−Fe 73-aa domain 5′

Stabilization and accumulation

3′ Ferritin untranslated iron storage

Ferritin mRNA

+ Fe

Ubiquitination

Modified IRP-2

Oxidation carbonylation ?

Degradation by the proteasome

5′

3′

Ferritin translated iron storage

FIGURE 17.5 Iron-dependent expression and activity of IRP2, and regulation of ferritin translation. See text for details.

of the HFE protein with β2-microglobulin, thus abolishing the transport of HFE to the cell surface. HFE modulates iron uptake by interacting with TfR, thus decreasing its affinity for iron-loaded transferrin. An interrelation between HFE and IRP activity was demonstrated in vitro by using HeLa cells stably transfected with HFE (Riedel et al., 1999). The authors reported that induction of HFE resulted in IRP activity with down-regulation of ferritin and up-regulation of TfR. It is of interest that in hemochromatosis patients, an increase in IRP activity has been found in monocytes (Cairo et al., 1997), which is in line with the idea that macrophages are iron-deprived in these patients (reviewed in Kühn, 1999). Development of targeted mutant mice can provide valuable information about the origin of human pathologies. Comparison of the effects of genetic disruption of IRP1 and IRP2 gene in mice recently revealed that IRP1−/− mice do not display obvious phenotypes whereas IRP2 −/− mice misregulate iron metabolism, particularly in specific areas of brain (LaVaute et al., 2001; Meyron-Holtz et al., 2004). In IRP2−/− mice, iron overload is associated with progressive neurodegenerative disorders with dysregulation of TfR and ferritin. This is reminiscent of an earlier report indicating that IRP2 co-localizes with redox-iron in the brain of

Nitric Oxide and Post-Transcriptional Control of Gene Expression

379

Alzheimer’s patients (Smith et al., 1998). Altogether, these observations indicate that IRP2 appears to dominate the post-transcriptional regulation of iron metabolism in response to dietary iron level even though IRP1 contributes to iron homeostasis because IRP1+/− IRP2−/− mice display aggravated neurological disorders (Smith et al., 2004). These results are in keeping with previous data indicating that IRP2 can be fully responsible for iron-dependent regulation of iron homeostasis in a murine pro-B lymphocyte cell line lacking IRP1 (Schalinske et al., 1997). At this stage, it should be recalled that IRP1 is a bifunctional protein, and operates mostly as an aconitase (or displays no function) under resting conditions. It is therefore not surprising that suppressing cytosolic aconitase activity does not markedly alter iron metabolism of mice kept under the sanitary conditions found in animal facilities. It is tempting to propose that the aconitase form of IRP1, with its perceptive [4Fe-4S] cluster, is prone to sense chemical signals like NO (see below). To tune iron homeostasis, IRP2 would prevail in the case of low-iron diet, but IRP1 may well be predominant in inflammatory settings or immune reaction involving NOS2-inducing stimuli including Th1 cytokines or bacterial products. Further studies with animal models for inflammatory diseases are required to clarify this point.

17.2 INTERPLAY BETWEEN NITRIC OXIDE (NO) AND IRPs Dissecting the mechanisms underlying the regulation of IRP RNA-binding activity was crucial to our understanding of the role that IRPs play in physiology and pathophysiology. The field was booming, and it rapidly became clear that stimuli other than changes in iron levels can regulate activity of either IRP, including oxidative stress (Pantopoulos and Hentze, 1998), hypoxia/reoxygenation conditions (Hanson and Leibold, 1999, Richardson, 2003) phosphorylation (Eisenstein, 2000) and NO synthesis, on which we will now focus.

17.2.1 NO AS A MODULATOR OF [FE-S] CLUSTER-CONTAINING PROTEINS Iron-sulfur clusters are polymetallic structures with iron atoms that are linked to inorganic sulfur and are usually liganded to proteins by cysteine thiolates. It is clear that [Fe-S] clusters are not only involved in electron transfer reactions but also play structural roles and are involved in enzymatic catalysis, sensing, signaling, and regulation of gene expression (reviewed in Beinert and Kiley, 1999). [Fe-S] proteins were originally designated as targets of NO by the work of Hibbs and colleagues who reported that the L-arginine-dependent mechanism of cytotoxicity of activated macrophages toward target cells involved mainly inhibition of DNA synthesis and of mitochondrial respiration (Granger and Lehninger, 1982; Drapier and Hibbs, 1986; Hibbs et al., 1987). In particular, it was stressed that the activity of mitochondrial iron-dependent enzymes, including the heme-containing cytochrome oxidase, the iron-sulfur enzymes NADH-ubiquinone

380

Nitric Oxide, Cell Signaling, and Gene Expression

oxidoreductase (complex I) and succinate-ubiquinone oxidoreductase (complex II) of the respiratory chain, as well as the citric acid cycle enzyme m-aconitase, was inhibited by this L-arginine-dependent pathway (Drapier and Hibbs, 1986; Drapier and Hibbs, 1988). The effector molecule of this pathway was then identified as NO (Hibbs et al., 1988; Stuehr and Nathan, 1989). The correlation between NO production, inhibition of m-aconitase activity and formation of the dinitrosyl-iron-dithiol complexes, together with the potential of NO to yield complexes with [Fe-S] clusters, strongly suggested that NO or an NO-derived molecule interacted directly with the [4Fe-4S] cluster of m-aconitase, resulting in loss of enzyme activity (Drapier et al., 1991). Other [Fe-S] protein are targets of NO include mammalian ferrochelatase, the last enzyme in heme biosynthesis (Sellers et al., 1996), as well as the bacterial regulators SoxR, which is involved in resistance to oxidative stress and antibiotics (reviewed in Hidalgo et al., 1997), and FNR which represses the flavohemoglobin Hmp (Cruz-Ramos et al., 2002). Altogether, these studies pointed to the interplay of NO with [Fe-S] clusters as a determinant factor in the regulation of protein activity (Drapier and Bouton, 1996, Drapier, 1997).

17.2.2 NO AND THE POST-TRANSLATIONAL REGULATION IRP1 ACTIVITIES

OF

Early studies on the previously mentioned interaction between NO and mitochondrial aconitase suggested that the [4Fe-4S] cluster containing IRP1 (the cytosolic aconitase) could be, similar to its mitochondrial counterpart, a potential target of NO action. In 1993, two independent groups reported that the stimulation of macrophages with IFN-γ or LPS, which induces NOS2, was accompanied by a loss of cytosolic aconitase activity and concomitant increase in IRP1 RNA-binding activity (Drapier et al., 1993, Weiss et al., 1993). Suppression of NO synthesis by N-mono-methyl-L-arginine or N-nitro-L-arginine, two specific NOS inhibitors, abolished the reciprocal modulation of IRP1 activities in stimulated cells, thus pointing to NO or an NO-derived species as the effective agent in the conversion of IRP1 from aconitase to RNA-binding protein. These two reports were the first demonstration of a post-translational regulation of IRP1 activities at the cellular level, by a physiological stimulus other than changes in intracellular iron availability. Furthermore, they revealed a novel connection between the cytokine-induced NOS and the IRP/IRE pathway. These initial findings were followed 1 year later by the demonstration that rat brain cells expressing constitutive NOS, as well as producing NO because of N-methyl-D-aspartate stimulation and calcium influx, also exhibited reciprocal modulation of IRP1 activities (Jaffrey et al., 1994). NO-induced IRP1 RNAbinding activity has also been reported in other cell types that are either stimulated to produce NO or exposed to NO-releasing chemicals. Although data concerning IRP regulation have in large part been obtained with rodent cell lines or primary macrophages, evidence exists that cytokine-driven NO synthesis may also be a crucial factor in the modulation of the IRP/IRE regulatory

Nitric Oxide and Post-Transcriptional Control of Gene Expression

381

system in human cells (Cairo et al., 1997). However, no discrimination was made between human IRP1 and IRP2, which co-migrate in the electrophoretic mobility shift assay. A similar increase in IRP IRE-binding activity induced by IFN-γ/LPS treatment was observed in the human monocytic cell line THP-1, which was abolished in the presence of the NOS2 inhibitor L-NMMA (Pietrangelo et al., 1998). NO-mediated IRP regulation has also been reported in the human erythroid cell line K562 (Weiss et al., 1993; Richardson et al., 1995; Oria et al., 1995) and in human umbilical vein endothelial cells (Juckett et al., 1996).

17.2.3 MECHANISM UNDERLYING NO MODULATION RNA-BINDING ACTIVITY

OF

IRP1

Although it is clear that NO biosynthesis induces a switch between the aconitase and the RNA-binding form of IRP1, the mechanism by which this regulation occurs has remained a matter of controversy. Indeed, different hypotheses have been proposed concerning the mechanism of action of NO on IRP1. Based on observations that NO would exert a slow effect on IRP1, with kinetics close to those of iron deprivation, it was proposed that NO may modulate IRP1 activities indirectly, by depleting cellular iron pools (Pantopoulos et al., 1996); however, several lines of evidence favor the hypothesis that NO modulates IRP1 activities by directly interacting with this protein. Evidence pointing to a direct effect of NO on IRP1 was first provided by Drapier and colleagues, who reported that in vitro exposure of purified recombinant human IRP1 to NO gas or to NO-releasing drugs resulted in a dose-dependent decrease in aconitase activity, which was not due to protein degradation. Aconitase activity was completely lost following a 15-min exposure to NO gas, which was accompanied by a rise in RNA-binding activity (Drapier et al., 1993). Furthermore, incubation of cytosols with an excess of aconitase substrates, which bind to the [4Fe-4S] cluster, prevented both the inhibition of aconitase activity and the induction of IRP1 IRE-binding activity by NO (Bouton et al., 1996). Additional evidence in favor of a direct effect of NO on IRP1 was provided by the demonstration that NO released from activated macrophages can rapidly induce the IRE-binding activity of IRP1 from target cells, without requirement of de novo protein synthesis (Bouton et al., 1998). Although these studies pointed to the capacity of NO to interact directly with IRP1, particularly with its [4Fe-4S] cluster, some questions have remained concerning the mechanism by which NO can induce the RNA-binding form of this protein. As stated previously, exposure of recombinant IRP1 to NO results in total loss of aconitase activity; however, the IRP1 IRE-binding activity of recombinant IRP1 treated with NO was not complete in contrast with what happens in the whole cell (Drapier et al., 1993). In brief, it appeared that simplification of the experimental system, from the whole cell to the purified protein, was accompanied by a gradual difficulty in achieving IRP1 IRE-binding capacity. Keeping in mind that IRP1 is a redox-sensitive protein (Hentze et al., 1989), the loss of activity upon purification suggested a lack of a cellular redox compound. Therefore, the role of the physiological reducer thioredoxin (Trx), which catalyzes

382

Nitric Oxide, Cell Signaling, and Gene Expression

dithiol-disulfide exchange reactions, was addressed. It was demonstrated that reduced Trx, at physiological concentrations, strongly increased the IRE-binding activity of IRP1 in cytosolic extracts previously exposed to NO. Conversely, neutralization of endogenous Trx with an anti-Trx monoclonal antibody abolished the increase in RNA-binding activity of IRP1 in cytosolic extracts exposed to NO (Oliveira et al., 1999). These results strongly suggested that reduced Trx exerts its action on an oxidized apoprotein form of IRP1 generated following exposure to NO. In vitro, it was confirmed by EPR spectroscopy that NO reacts directly with IRP1 [4Fe-4S] cluster (Kennedy et al., 1997) and that purified recombinant human IRP1 exposed to NO released 4 iron atoms per molecule (Soum et al., 2002), suggesting that the cluster is completely disassembled by NO or an NO-derived molecule. In view of these results, it is likely that disassembly of the Fe-S cluster induced by NO is accompanied by the formation of an oxidized apo-IRP1, which needs to be reduced to allow access to the IREs. Further, the Lewis acid character of iron would be expected to withdraw electrons from NO to produce an intermediate with nitrosonium (NO+)-like reactivity, which would then be able to interact with the proximal nucleophilic sulfurs. Thus, it is possible that NO liganded in the dinitrosyl-iron complex (DNIC) of IRP1 will promote the nitrosation of the thiolate group of one (or more) of the cysteines that hold the [Fe-S] cluster (Figure 17.6). It has been claimed that peroxynitrite instead of NO inhibits aconitases (Hausladen et al., 1994). Actually, exogenously added peroxynitrite inhibits aconitase activity of IRP1 but hardly activates its RNA-binding activity. Using a C437S IRP1 mutant, Bouton et al. (1997) reported that peroxynitrite promotes formation of a disulfide bridge involving C437 and probably C503 or C506, which may prevent binding to IRE. Moreover, resonance Raman experiments demonstrated that peroxynitrite could nitrate IRP1, which may also explain why peroxynitrite-exposed IRP1 has no activity (Soum et al., 2003). Nitration of IRP1 associated with loss of aconitase activity and low capacity to bind IRE has recently been confirmed in intact cells stimulated by IFN-γ/LPS and PMA (Gonzalez et al., 2004). To understand better the impact of the signaling molecule NO on IRP-1 functions, the fate of IRP-1 functions when NO flux stops was also investigated. Data indicated that upon NO removal, IRP-1 dissociates from IRE and, via a mitochondrial ATP-dependent process, rapidly recovers its aconitase activity, which implies [4Fe-4S] cluster repair (Bouton et al., 2002). These results demonstrate that fluctuation of NO levels allows the same IRP-1 molecule to commute quickly between its two activities through the disassembly/reassembly of its [4Fe-4S] cluster. In brief, NO determines IRP1 function by reversible modifications and does not alter (at least durably) the iron-sulfur cluster assembly machinery (Lill and Kispal, 2000; Tong and Rouault, 2000) involved in the recovery of initial aconitase activity. With IRP1 as accomplice, NO plays here its typical role of chemical signaling molecule: quick, efficient, and elusive (Bouton and Drapier, 2003).

Nitric Oxide and Post-Transcriptional Control of Gene Expression aconitase IRP-1

NO arg699

cys437S cys503 S

1 S

trans-regulator IRP-1

inactive IRP-1

NO cys437 S cys503 S

cys506

Fe

DNIC formation

NO 2

S cys506

cys437 S cys503 S

383

NO S NO

3

cys506

di-sulfide bridge cys437 S cys503 S

S cys506

4

cys437 S cys503 S

Thioredoxin

IRE S cys506

Transient S-nitrosation ?

(Fe-S) cluster disassembly (Fe-S) cluster repair

IRP-1.lRE dissociation

NO removal IRE

cys437 S cys S

Energized- 503 cysS 506 mitochondria-dependent process

FIGURE 17.6 Schematic representation of the post-translational regulation of IRP1 by NO. The most likely mechanism deduced from the data gathered in the literature points to a progressive disassembly of IRP1 [4Fe-4S] cluster by NO (1). An iron atom, from a remnant of the Fe-S cluster, would contribute to the formation of an EPR-detectable dinitrosyl iron complex (2) (Kennedy et al., 1997). Nitrosyl-iron complex, by promoting NO+ formation, can facilitate nitrosation of local cysteines (i.e., Cys437, Cys 503, or Cys 506). As S-nitrosation of vicinal thiols usually promotes disulfide bond, formation of an oxidized apo-IRP1 is likely (3). This null form would gain full IRE-binding protein after reduction by thioredoxin (4) (Oliveira et al., 1999). After NO removal, IRP1 recovers its initial aconitase activity by repairing its Fe-S cluster with the requirement of energized mitochondria (5) (Bouton et al., 2002).

17.2.4 REGULATION

OF

IRP1 PROTEIN LEVELS

BY

NO

Although intensive research has been conducted concerning the regulation of IRP1 activities, little is known with regard to how the expression of this protein is regulated. We have shown that stimulation of RAW 264.7 cells and of murine peritoneal macrophages with IFN-γ or LPS results in a decrease in IRP1 levels. Furthermore, experiments performed with macrophages from NOS2−/− mice demonstrated that NO was responsible for the down-regulation of IRP1 expression observed in IFN-γ/LPS stimulated cells and in cells adjacent to NO-producing cells (Oliveira and Drapier, 2000). It is also important to stress that, although the total amount of IRP1 molecules present in stimulated cells is reduced compared with control cells, the number of IRP1 molecules able to bind RNA is still higher in the former. In other words, the net result of IFN-γ/LPS stimulation as regards IRP1 is a marked increase in its IRE-binding activity. Thus, these data do not dispute the well-established fact that IFN-γ/LPS stimulation induces higher RNAbinding activity of IRP1, via NO. Instead, they reveal that NO exerts a dual effect on IRP1: In addition to promoting the conversion of IRP1 from aconitase to transregulator, it induces a parallel decrease in IRP1 levels.

384

Nitric Oxide, Cell Signaling, and Gene Expression

The opposite regulation of IRP1 IRE-binding activity and IRP1 expression in response to NO is titillating and raises some important questions. First, it is tempting to speculate that down-regulation of IRP1 expression may likewise represent a compensatory mechanism to mitigate NO-mediated IRP1 activation. NO-mediated down-regulation of IRP1 expression might thus be a novel mechanism to protect cells against an excessive iron-catalyzed formation of free radicals, in pathophysiological situations, such as immune and inflammatory responses, where NO may be produced in excessive amounts. At first glance, it is reasonable to consider that the down-regulation of IRP1 protein levels induced by NO could be due to an increased susceptibility of this protein to proteolytic degradation following NO exposure. In this regard, it is worth recalling that m-aconitase, upon exposure to the NO-derived oxidant peroxynitrite, is degraded by a proteasome-mediated pathway (Grüne et al., 1998). Moreover, IRP1 contains proteolytic cleavage sites near or within its putative cleft that are masked when the [4Fe-4S] cluster is present, and become accessible following cluster disruption (Schalinske et al., 1997, Gegout et al., 1999). Considering all these data, it is tempting to assume that NO, by disrupting the [4Fe-4S] cluster of IRP1, may also accelerate IRP1 degradation, either by simply exposing these proteolysis-susceptible sites, or by inducing some conformational modification that would predispose IRP1 to degradation. Alternatively, because the NO-mediated decrease in IRP1 levels in stimulated RAW 264.7 cells was associated with a reduction of the amount of IRP1 mRNA (Oliveira and Drapier, 2000), it is tempting to posit that NO could react with one or more redox-sensitive transcription factors required for IRP1 gene expression.

17.2.5 MODULATION

OF

IRP2

BY

NO

Although the role of NO as an inducer of IRP1 IRE-binding activity is well established, its effect on the modulation of IRP2 activity has been a matter of debate. Even though IRP2 does not possess an [Fe-S] cluster, it exhibits several redox-sensitive cysteine residues, which makes it a potential target of NO. In vitro, the RNA-binding activity of IRP2 was analyzed following treatment of cytosolic extracts of various cells with different classes of NO donors. NO (or an NO-derived species) caused a reduction of IRP2 activity, which was overcome by the physiological thioredoxin system (Oliveira et al., 1999). These data strongly suggest that NO is able to affect IRP2 activity by interacting with some critical cysteine residue(s), in a way that prevents access to the IRE sequences. Based on the model of iron-induced degradation of IRP2 (Iwai et al., 1998), it was tempting to consider that NO or some nitrosating species may also favor IRP2 proteolysis, by reacting with cysteine residues of the 73-amino-acid domain. Ponka and collaborators explored this line of research. They reported that in contrast to the nitrosothiol SNAP, sodium nitroprusside (SNP), by releasing the nitrosonium anion NO+, can nitrosate IRP2 (Kim and Ponka, 1999; Kim and Ponka, 2000). They described that exposure of RAW 264.7 macrophage-like cells to IFN-γ/LPS or to SNP mediated decrease in IRP2 activity and

Nitric Oxide and Post-Transcriptional Control of Gene Expression

385

protein level, associated with decrease in transferrin receptor mRNA level and increase in ferritin expression. Moreover, they reported that mutation of C178 in the 73-amino-acid domain prevented NO-mediated degradation of IRP2 and concluded that nitrosation of C178 is the crucial feature leading to IRP2 degradation by the ubiquitin-proteasome system (Kim et al., 2004). Another group, however, recently reported that HEK cells stably transfected with mutant IRP2 constructs with all cysteines of the 73-amino-acid domain (including C178) mutagenized to alanines also exhibited decrease in IRP2 level in response to SNP (Bourdon et al., 2003). These authors also claimed that desferal, an iron chelator, prevents the effect of SNP and suggested that SNP may actually mediate IRP2 degradation via release of iron instead of NO.

17.2.6 FUNCTIONAL IMPACT OF THE NO/IRP REGULATORY SYSTEM ON FERRITIN AND TFR EXPRESSION The question of whether NO affects the expression of ferritin and TfR through the IRP/IRE system was addressed as soon as the induction of IRP1 IRE-binding activity by NO was discovered. Several lines of evidence rapidly indicated that ferritin translation is repressed in response to NO-mediated IRP1 activation. A decrease in ferritin biosynthesis following NO production by the macrophage cell line J774 stimulated with IFN-γ and LPS was first reported without a reduction of ferritin mRNA levels, in agreement with a post-transcriptional regulation via IRP1 IRE-binding activity (Weiss et al., 1993). Moreover, NO-induced activation of IRP1 IRE-binding also caused repression of the translation of reporter mRNAs containing a ferritin IRE in their 5'UTR (Weiss et al., 1993). Interestingly, elimination of NOS2 induction in macrophages treated with the cytokines IL-4 and IL-13 was accompanied by a concomitant rise in ferritin translation without increase in ferritin mRNA levels (Weiss et al., 1997). A repression of ferritin translation following NO-induced IRP1 activation was also reproduced in B6 fibroblasts stably transfected with a cDNA encoding murine macrophage NOS2 (Pantopoulos and Hentze, 1995) and in the rat hepatoma cell line FTO2B following stimulation with IFN-γ/TNF-α/LPS (Phillips et al., 1996). The inhibition of ferritin biosynthesis observed in all these experiments was dependent on NO synthesis because it was abrogated by the NOS inhibitor L-NMMA (Weiss et al., 1993; Pantopoulos and Hentze, 1995; Phillips et al., 1996; Weiss et al., 1997). Induction of IRP1 IRE-binding activity and repression of ferritin synthesis was also reproduced by treatment of FTO2B cells and RAW 264.7 macrophages with the NO-donors SNAP and DETA/NO, respectively (Phillips et al., 1996; Bouton et al., 2002). Interestingly, IRP1 IRE dissociation after DETA/NO removal was accompanied by recovery of ferritin expression (Bouton et al., 2002). Finally, an NO-dependent reduction of ferritin expression was also reported in the erythroid cell line K562 stably transfected with a retroviral vector encoding murine macrophage NOS2 (Domachowske et al., 1996). Altogether, these data indicated that NO is sufficient to directly control ferritin translation in a variety of cell types.

386

Nitric Oxide, Cell Signaling, and Gene Expression

Not all data agree with an NO-induced decrease in ferritin expression, however. There was reported a rise in ferritin expression in macrophage cell lines following stimulation with a combination of IFN-γ plus LPS, which was reversed in the presence of NO synthase inhibitors (Recalcati et al., 1998; Kim and Ponka, 2000). The rise in ferritin expression induced by treatment with IFN-γ/LPS was attributed to the previously mentioned drastic decrease in IRP2 activity observed in these experiments (Kim and Ponka, 2000). In contrast to what was predicted (see Figure 17.3), activation of IRP1 following stimulation of macrophages with IFN-γ/LPS was not correlated with TfR mRNA stabilization (Drapier et al., 1993; Pantopoulos and Hentze, 1995). Indeed, treatment of the macrophages with these stimuli was accompanied by a decrease, instead of an increase, in TfR mRNA levels, which was not reversed by the NOS2 inhibitor L-NMMA (Pantopoulos and Hentze, 1995; Weiss et al., 1997). These results are in agreement with earlier studies demonstrating that stimulation of murine peritoneal macrophages with IFN-γ results in a reduction of TfR mRNA levels (Hamilton et al., 1984); however, in B6 fibroblasts transfected with NOS2, an up-regulation of TfR mRNA was associated with an increase in IRP1 IRE-binding activity (Pantopoulos and Hentze, 1995). In addition, the cell surface expression of TfR was increased in K562 cells stably transfected with NOS2, probably because of NO-mediated stabilization of TfR mRNA (Domachowske et al., 1996). These results were confirmed by Leibold and colleagues who analyzed TfR mRNA levels in the hepatoma cells FTO2B following treatment with NO-releasing agents to eliminate cytokine- and LPSmediated effects. IRP1 IRE-binding activity was increased and, in parallel, a twofold increase in TfR mRNA levels was found (Phillips et al., 1996). A longlasting increase in TfR mRNA levels induced by the NO donor DETA/NO was also associated with a rise in IRP1 IRE-binding activity in RAW 264.7 macrophages (Bouton et al., 2002). Overall, these results indicated that NO alone is able to stabilize TfR mRNA via IRP1 IRE interaction, and supported the hypothesis that the down-regulation of TfR expression induced by IFN-γ or LPS may be due to other pathways (e.g., transcriptional regulation) that might counteract NOmediated stabilization of TfR mRNA (discussed in Bouton and Drapier, 2003). Alternatively, as mentioned earlier, others explained the decrease in TfR expression observed in IFN-γ/LPS-stimulated RAW 264.7 macrophages by IRP2 degradation following S-nitrosation of a cysteine by the nitrosonium anion NO+ (Kim et al., 2004).

17.3 A LOOK AHEAD The two post-transcriptional regulators IRPs are major characters that play important parts in the adaptive response to iron dysregulation. The intrusion of NO obviously disturbs the play. The current view depicted by most studies points toward an activation of IRP1 RNA-binding capacity by NO, either produced endogenously in cells stimulated physiologically or exposed to NO-releasing chemicals. As a functional consequence, ferritin translation is repressed. With regard to TfR, the

Nitric Oxide and Post-Transcriptional Control of Gene Expression

387

scheme is more devious. In response to an induction of NOS2, by cytokines or LPS, most of the data agree with a decrease in TfR mRNA level. In contrast, in response to exogenous NO, TfR mRNA level is enhanced. Such a situation is likely to happen in a cell adjacent to NOS2 expressing cells and unresponsive to IFN-γ or LPS. Accordingly, iron uptake would be favored, but in the absence of storage in ferritin, excess iron could represent a potential hazard. To recapitulate, in the context of cell-mediated toxicity, a cell both responding to IFN-γ and LPS and producing NO (e.g., an activated macrophage) would be protected against noxious iron, whereas a target cell exposed to paracrine NO would not. Redundancy between both IRPs has been an enigma. According to data obtained with IRP mutant mice, IRP2 appears to be prevalent in dietary irondeficient animals. Yet, IRP1 remains the most important pool of latent IREbinding activity. The partnership between NO and IRP1 proved so efficient in cell culture that it may also be effective in vivo, particularly in pathological situations (e.g., endotoxemia, imbalance of the cytokine network, and chronic inflammation) that are known to induce NOS2 expression. Each IRP would specialize in responding to a particular signal: iron flux for IRP2 and environmental signals such as NO for IRP1. Further studies addressing this question, for instance, by challenging IRP mutant mice, are much awaited.

REFERENCES Aziz, N., and Munro, H. N. (1987). Iron regulates ferritin mRNA translation through a segment of its 5′ untranslated region. Proc. Natl. Acad. Sci. USA, 84:8478–82. Beaumont, C., Leneuve, P., Devaux, I., Scoazec, J. Y., Berthier, M., Loiseau, M. N., Grandchamp, B., and Bonneau, D. (1995). Mutation in the iron responsive element of the L ferritin mRNA in a family with dominant hyperferritinaemia and cataract. Nat. Genet., 11:444–6. Beinert, H., and Kiley, P. J. (1999). Fe-S proteins in sensing and regulatory functions. Curr. Opin. Chem. Biol., 3:152–7. Binder, R., Horowitz, J. A., Basilion, J. B., Koeller, D. M., Klausner, R. D., and Harford, J. B. (1994). Evidence that the pathway of transferrin receptor mRNA degradation involves an endonucleolytic cleavage within the 3′ UTR and does not involve poly(A) tail shortening. EMBO J, 13:1969–80. Bourdon, E., Kang, D. K., Ghosh, M. C., Drake, S. K., Wey, J., Levine, R. L., and Rouault TA. (2003). The role of endogenous heme synthesis and degradation domain cysteines in cellular iron-dependent degradation of IRP2. Blood Cells Mol. Dis., 31:247–55. Bouton, C., Hirling, H., and Drapier, J. C. (1997). Redox modulation of iron regulatory proteins by peroxynitrite. J. Biol. Chem., 272:19969–75. Bouton, C., Oliveira, L., and Drapier, J. C. (1998). Converse modulation of IRP1 and IRP2 by immunological stimuli in murine RAW 264.7 macrophages. J. Biol. Chem., 273:9403–8. Bouton, C., Raveau, M., and Drapier, J. C. (1996). Modulation of iron regulatory protein functions. Further insights into the role of nitrogen- and oxygen-derived reactive species. J. Biol. Chem., 271:2300–6.

388

Nitric Oxide, Cell Signaling, and Gene Expression

Bouton, C., Chauveau, M. J., Lazereg, S., and Drapier, J. C. (2002). Recycling of RNA binding iron regulatory protein 1 into an aconitase after nitric oxide removal depends on mitochondrial ATP. J. Biol. Chem., 277:31220–7. Bouton C., and Drapier J. C. (2003). Iron regulatory proteins as NO signal transducers. Sci. STKE, (182):pe17 Cairo, G., Recalcati, S., Montosi, G., Castrusini, E., Conte, D., and Pietrangelo, A. (1997). Inappropriately high iron regulatory protein activity in monocytes of patients with genetic hemochromatosis. Blood, 89:2546–53. Cammack R, Wrigglesworth J. M. and Baum, H. (1990). Iron-dependent enzymes in mammalian systems. In: Ponka, P., Schulman, H. M. and Woodworth R. C. (eds.), Iron transport and storage, pp. 17–39. Boca Raton: CRC Press. Casey, J. L., Hentze, M. W., Koeller, D. M., Caughman, S. W., Rouault, T. A., Klausner, R. D., and Harford, J. B. (1988). Iron-responsive elements: regulatory RNA sequences that control mRNA levels and translation. Science, 240:924–8. Casey, J. L., Koeller, D. M., Ramin, V. C., Klausner, R. D., and Harford, J. B. (1989). Iron regulation of transferrin receptor mRNA levels requires iron-responsive elements and a rapid turnover determinant in the 3' untranslated region of the mRNA. EMBO J., 8:3693–9. Cox, T. C., Bawden, M. J., Martin, A., and May, B. K. (1991). Human erythroid 5aminolevulinate synthase: promoter analysis and identification of an iron-responsive element in the mRNA. EMBO J., 10:1891–902. Cruz-Ramos, H., Crack, J., Wu, G., Hughes, M. N., Scott, C., Thomson, A.J., Green, J., and Poole, R.K. (2002) NO sensing by FNR: regulation of the Escherichia coli NO-detoxifying flavohaemoglobin, Hmp. EMBO J., 13:3235–44. Dandekar, T., Stripecke, R., Gray, N. K., Goossen, B., Constable, A., Johansson, H. E., and Hentze, M. W. (1991). Identification of a novel iron-responsive element in murine and human erythroid delta-aminolevulinic acid synthase mRNA. EMBO J., 10:1903–9. Domachowske, J. B., Rafferty, S. P., Singhania, N., Mardiney, M. R., and Malech, H. L. (1996). Nitric oxide alters the expression of gamma-globin, H-ferritin, and transferrin receptor in human K562 cells at the posttranscriptional level. Blood, 88:2980–8. Donovan, A., Brownlie, A., Zhou, Y., Shepard, J., Pratt, S. J., Moynihan, J., Paw, B. H., Drejer, A., Barut, B., Zapata, A., Law, T. C., Brugnara, C., Lux, S.E., Pinkus, G. S., Pinkus, J.L., Kingsley, P. D., Palis, J., Fleming, M.D., Andrews, N.C., Zon, L. I. (2000). Positional cloning of zebrafish ferroportin1 identifies a conserved vertebrate iron exporter. Nature, 403:776–81. Drapier, J. C. (1997). Interplay between NO and [Fe-S] clusters: relevance to biological systems. Methods, 11:319–29. Drapier, J. C., and Bouton, C. (1996). Modulation by nitric oxide of metalloprotein regulatory activities. Bioessays, 18:549–56. Drapier, J. C., and Hibbs, J. B., Jr. (1986). Murine cytotoxic activated macrophages inhibit aconitase in tumor cells. Inhibition involves the iron-sulfur prosthetic group and is reversible. J. Clin. Invest., 78:790–7. Drapier, J. C., and Hibbs, J. B., Jr. (1988). Differentiation of murine macrophages to express nonspecific cytotoxicity for tumor cells results in L-arginine-dependent inhibition of mitochondrial iron-sulfur enzymes in the macrophage effector cells. J. Immunol., 140:2829–38.

Nitric Oxide and Post-Transcriptional Control of Gene Expression

389

Drapier, J. C., Hirling, H., Wietzerbin, J., Kaldy, P., and Kühn, L. C. (1993). Biosynthesis of nitric oxide activates iron regulatory factor in macrophages. EMBO J., 12:3643–9. Drapier, J. C., Pellat, C., and Henry, Y. (1991). Generation of EPR-detectable nitrosyliron complexes in tumor target cells cocultured with activated macrophages. J. Biol. Chem., 266:10162–7. Eisenstein, R. S, and Ross, K. L. (2003) Novel roles for iron regulatory proteins in the adaptive response to iron deficiency. J. Nutr., 133:1510S–6S. Eisenstein, R. S. (2000). Iron regulatory proteins and the molecular control of mammalian iron metabolism. Annu. Rev. Nutr., 20:627–62. Fleming, M. D., Romano, M. A., Su, M. A., Garrick, L. M., Garrick, M. D., and Andrews, N. C. (1998). Nramp2 is mutated in the anemic Belgrade (b) rat: evidence of a role for Nramp2 in endosomal iron transport. Proc. Natl. Acad. Sci. USA, 95:1148–53. Fodinger, M., and Sunder-Plassmann, G. (1999). Inherited disorders of iron metabolism. Kidney Int., 69:S22–34. Gegout, V., Schlegl, J., Schlager, B., Hentze, M. W., Reinbolt, J., Ehresmann, B., Ehresmann, C., and Romby, P. (1999). Ligand-induced structural alterations in human iron regulatory protein-1 revealed by protein footprinting. J. Biol. Chem., 274:15052–8. Gonzalez, D, Drapier, J.C., and Bouton, C. (2004). Endogenous nitration of iron regulatory protein-1 in nitric oxide-producing murine macrophages. Further insights into the mechanism of nitration in vivo and its impact on IRP-1 functions. J. Biol. Chem., Jul. 2004 10.1074/jbc. Granger, D.L., and Lehninger, A. (1982). Sites of inhibition of mitochondrial electron transport in macrophage-injured neoplastic cells. J. Cell Biol., 95:527–35. Gray, N. K., and Hentze, M. W. (1994). Iron regulatory protein prevents binding of the 43S translation pre-initiation complex to ferritin and eALAS mRNAs. EMBO J., 13:3882–91. Gray, N. K., Pantopoulous, K., Dandekar, T., Ackrell, B. A., and Hentze, M. W. (1996). Translational regulation of mammalian and Drosophila citric acid cycle enzymes via iron-responsive elements. Proc. Natl. Acad. Sci. USA, 93:4925–30. Griffiths, E. (1987). Iron in biological systems. In: Bullen, J. J. and Griffiths (eds.), Iron and infection, pp. 1–25, John Wiley & Sons Ltd., New York. Grüne, T., Blasig, I. E., Sitte, N., Roloff, B., Haseloff, R., and Davies, K. J. A. (1998). Peroxynitrite increases the degradation of aconitase and other cellular proteins by proteasome. J. Biol. Chem., 273:10857–62. Gunshin, H., Mackenzie, B., Berger, U. V., Gunshin, Y., Romero, M. F., Boron, W. F., Nussberger, S., Gollan, J. L., and Hediger, M. A. (1997). Cloning and characterization of a mammalian proton-coupled metal-ion transporter. Nature, 388:482–8. Guo, B., Brown, F. M., Phillips, J. D., Yu, Y., and Leibold, E. A. (1995). Characterization and expression of iron regulatory protein 2 (IRP2). Presence of multiple IRP2 transcripts regulated by intracellular iron levels. J. Biol. Chem., 270:16529–35. Guo, B., Phillips, J. D., Yu, Y., and Leibold, E. A. (1995). Iron regulates the intracellular degradation of iron regulatory protein 2 by the proteasome. J. Biol. Chem., 270:21645–51. Guo, B., Yu, Y., and Leibold, E. A. (1994). Iron regulates cytoplasmic levels of a novel iron-responsive element-binding protein without aconitase activity. J. Biol. Chem., 269:24252–60.

390

Nitric Oxide, Cell Signaling, and Gene Expression

Halliwell, B. and Gutteridge, J. M. C. (1984). Oxygen toxicity, oxygen radicals, transition metals and disease. Biochem. J., 219:1–14. Hamilton, T. A., Gray, P. W., and Adams, D. O. (1984). Expression of the transferrin receptor on murine peritoneal macrophages is modulated by in vitro treatment with interferon gamma. Cell Immunol., 89:478–88. Hanson, E. S., and Leibold, E. A. (1998). Regulation of iron regulatory protein 1 during hypoxia and hypoxia/reoxygenation. J. Biol. Chem., 273:7588–93. Hanson, E. S., and Leibold, E. A. (1999). Regulation of the iron regulatory proteins by reactive nitrogen and oxygen species. Gene Expr., 7:367–76. Hanson, E. S., Rawlins, M. L., and Leibold, E. A. (2003). Oxygen and iron of iron regulation of iron regulatory protein 2. J. Biol. Chem., 278:40337–42. Hausladen, A., and Fridovich, I. (1994). Superoxide and peroxynitrite inactivate aconitases, but nitric oxide does not. J. Biol. Chem., 269:29405–8. Henderson, B. R. (1996). Iron regulatory proteins 1 and 2. Bioessays, 18:739–46. Hentze, M. W., and Kühn, L. C. (1996). Molecular control of vertebrate iron metabolism: mRNA-based regulatory circuits operated by iron, nitric oxide, and oxidative stress. Proc. Natl. Acad. Sci. USA, 93:8175–82. Hentze, M. W., Caughman, S. W., Rouault, T. A., Barriocanal, J. G., Dancis, A., Harford, J. B., and Klausner, R. D. (1987). Identification of the iron-responsive element for the translational regulation of human ferritin mRNA. Science, 238:1570–3. Hentze, M. W., Rouault, T. A., Harford, J. B., and Klausner, R. D. (1989). Oxidationreduction and the molecular mechanism of a regulatory RNA-protein interaction. Science, 244:357–9. Hentze, M.W., and Argos, P. (1991). Homology between IRE-BP, a regulatory RNAbinding protein, aconitase, and isopropylmalate isomerase. Nucleic Acids Res., 19; 1739–40. Hentze, M. W., Muckenthaler, M. U., and Andrews, N. C. (2004). Balancing acts: molecular control of mammalian iron metabolism. Cell, 117:285–97. Hibbs, J. B., Jr., Taintor, R. R., Vavrin, Z., and Rachlin, E. M. (1988). Nitric oxide: a cytotoxic activated macrophage effector molecule. Biochem. Biophys. Res. Commun., 157:87–94. Hibbs, J. B., Jr., Vavrin, Z., and Taintor, R. R. (1987). L-arginine is required for expression of the activated macrophage effector mechanism causing selective metabolic inhibition in target cells. J. Immunol., 138:550–65. Hidalgo, E., Ding, H., and Demple, B. (1997). Redox signal transduction via iron-sulfur clusters in the SoxR transcription activator. Trends Biochem. Sc.i, 22:207–10. Hirling, H., Henderson, B. R., and Kühn, L. C. (1994). Mutational analysis of the [4Fe4S]-cluster converting iron regulatory factor from its RNA-binding form to cytoplasmic aconitase. EMBO J., 13:453–61. Iwai, K., Drake, S. K., Wehr, N. B., Weissman, A. M., LaVaute, T., Minato, N., Klausner, R. D., Levine, R. L., and Rouault, T. A. (1998). Iron-dependent oxidation, ubiquitination, and degradation of iron regulatory protein 2: implications for degradation of oxidized proteins. Proc. Natl. Acad. Sci. USA, 95:4924–8. Jaffrey, S. R., Cohen, N. A., Rouault, T. A., Klausner, R. D., and Snyder, S. H. (1994). The iron-responsive element binding protein: a target for synaptic actions of nitric oxide. Proc. Natl. Acad. Sci. USA, 91:12994–8. Juckett, M. B., Weber, M., Balla, J., Jacob, H. S., and Vercellotti, G. M. (1996). Nitric oxide donors modulate ferritin and protect endothelium from oxidative injury. Free Radic. Biol. Med., 20:63–73.

Nitric Oxide and Post-Transcriptional Control of Gene Expression

391

Kang, D.K., Jeong, J., Drake, S.K., Wehr, N.B., Rouault, T.A., Levine, R.L. (2003) Iron regulatory protein 2 as iron sensor. Iron-dependent oxidative modification of cysteine. J. Biol. Chem., 278:14857–64. Kennedy, M. C., Antholine, W. E., and Beinert, H. (1997). An EPR investigation of the products of the reaction of cytosolic and mitochondrial aconitases with nitric oxide. J. Biol. Chem., 272:20340–7. Kim, S., and Ponka, P. (1999). Control of transferrin receptor expression via nitric oxidemediated modulation of iron-regulatory protein 2. J. Biol. Chem., 274:33035–42. Kim, S., and Ponka, P. (2000). Effects of interferon-gamma and lipopolysaccharide on macrophage iron metabolism are mediated by nitric oxide-induced degradation of iron regulatory protein 2. J. Biol. Chem., 275:6220–6. Kim, S., and Ponka, P. (2002) Nitrogen monoxide-mediated control of ferritin synthesis: implications for macrophage iron homeostasis. Proc. Natl. Acad. Sci. USA, 99:12214–9. Kim, S., Wing, S. S., and Ponka, P. (2004) S-nitrosylation of IRP2 regulates its stability via the ubiquitin-proteasome pathway. Mol. Cell Biol., 24:330–7. Klausner, R. D., Rouault, T. A., and Harford, J. B. (1993). Regulating the fate of mRNA: the control of cellular iron metabolism. Cell, 72:19–28. Kohler, S. A., Henderson, B. R., and Kühn, L. C. (1995). Succinate dehydrogenase b mRNA of Drosophila melanogaster has a functional iron-responsive element in its 5′-untranslated region. J. Biol. Chem., 270:30781–6. Kühn, L. C. (1999). Iron overload: molecular clues to its cause. Trends Biochem. Sci., 24:164–6. LaVaute, T., Smith, S., Cooperman, S., Iwai, K., Land, W., Meyron-Holtz, E., Drake, S. K., Miller, G., Abu-Asab, M., Tsokos, M., Switzer, R., Grinberg, A., Love, P., Tresser, N., and Rouault, T. (2001). Targeted deletion of the gene encoding iron regulatory protein-2 causes misregulation of iron metabolism and neurodegenerative disease in mice. Nat. Genet., 27:209–14. Leibold, E. A., and Munro, H. N. (1988). Cytoplasmic protein binds in vitro to a highly conserved sequence in the 5' untranslated region of ferritin heavy- and lightsubunit mRNAs. Proc. Natl. Acad. Sci. USA, 85:2171–75. Lill, R., and Kispal, G. (2000). Maturation of cellular Fe-S proteins: an essential function of mitochondria. Trends Biochem. Sci., 25:352–6. McKie, A. T., Marciani, P., Rolfs, A., Brennan, K., Wehr, K., Barrow, D., Miret, S., Bomford, A., Peters, T. J., Farzaneh, F., Hediger, M. A., Hentze, M. W., and Simpson, R. (2000). A novel duodenal iron-reguated transporter, IREG1, implicated in the basolateral transfer of iron to the circulation. Mol. Cell, 5:299–309. Meyron-Holtz, E. G., Ghosh, M. C., Iwai, K., LaVaute, T., Brazzolotto, X., Berger, U.V., Land, W., Ollivierre-Wilson, H., Grinberg, A., Love, P., and Rouault, T. A. (2004). Genetic ablations of iron regulatory proteins 1 and 2 reveal why iron regulatory protein 2 dominates iron homeostasis. EMBO J., 23:386–95. Muckenthaler, M., Gray, N. K., and Hentze, M. W. (1998). IRP-1 binding to ferritin mRNA prevents the recruitment of the small ribosomal subunit by the cap-binding complex eIF4F. Mol. Cell, 2:383–8. Müllner, E. W., and Kühn, L. C. (1988). A stem-loop in the 3′ untranslated region mediates iron-dependent regulation of transferrin receptor mRNA stability in the cytoplasm. Cell, 53:815–25.

392

Nitric Oxide, Cell Signaling, and Gene Expression

Müllner, E. W., Neupert, B., and Kühn, L. C. (1989). A specific mRNA binding factor regulates the iron-dependent stability of cytoplasmic transferrin receptor mRNA. Cell, 58:373–82. Oliveira L., Bouton C., and Drapier J. C. (1999). Thioredoxin activation of iron regulatory proteins. Redox regulation of RNA binding after exposure to nitric oxide. J. Biol. Chem., 274:516–21. Oliveira, L., and Drapier, J. C. (2000). Down-regulation of iron regulatory protein 1 gene expression by nitric oxide. Proc. Natl. Acad. Sci. USA, 97:6550–5. Oria, R., Sanchez, L., Houston, T., Hentze, M. W., Liew, F. Y., and Brock, J. H. (1995). Effect of nitric oxide on expression of transferrin receptor and ferritin and on cellular iron metabolism in K562 human erythroleukemia cells. Blood, 85:2962–6. Pantopoulos, K., and Hentze, M. W. (1995). Nitric oxide signaling to iron-regulatory protein: direct control of ferritin mRNA translation and transferrin receptor mRNA stability in transfected fibroblasts. Proc. Natl. Acad. Sci. USA, 92:1267–71. Pantopoulos, K., and Hentze, M. W. (1998). Activation of iron regulatory protein-1 by oxidative stress in vitro. Proc. Natl. Acad. Sci. USA, 95:10559–63. Pantopoulos, K., Weiss, G., and Hentze, M. W. (1996). Nitric oxide and oxidative stress (H2O2) control mammalian iron metabolism by different pathways. Mol. Cell Biol., 16:3781–8. Pantopoulos, K. Iron metabolism and the IRE/IRP regulatory system: an update (2004). Ann. NY Acad. Sci, 1012:1–13. Phillips, J. D., Kinikini, D. V., Yu, Y., Guo, B., and Leibold, E. A. (1996). Differential regulation of IRP1 and IRP2 by nitric oxide in rat hepatoma cells. Blood, 87:2983–92. Philpott, C. C., Haile, D., Rouault, T. A., and Klausner, R. D. (1993). Modification of a free Fe-S cluster cysteine residue in the active iron-responsive element-binding protein prevents RNA binding. J. Biol. Chem., 268:17655–8. Pietrangelo, A., Montosi, G., Recalcati, S., Garuti, C., and Cairo, G. (1998). Diacerhein blocks iron regulatory protein activation in inflamed human monocytes. Life Sci., 63:213–9. Ponka, P., and Lok, C. N. (1999). The transferrin receptor†: role in health and disease. Int. J. Biochem. Cell Biol., 31:1111–37. Recalcati, S., Taramelli, D., Conte, D., and Cairo, G. (1998). Nitric oxide-mediated induction of ferritin synthesis in J774 macrophages by inflammatory cytokines: role of selective iron regulatory protein-2 downregulation. Blood, 91:1059–66. Richardson, D. R., Neumannova, V., Nagy, E., and Ponka, P. (1995). The effect of redoxrelated species of nitrogen monoxide on transferrin and iron uptake and cellular proliferation of erythroleukemia (K562) cells. Blood, 86:3211–9. Richardson, D. R. (2003) The role of hypoxia and nitrogen monoxide in the regulation of cellular iron metabolism. J. Lab. Clin. Med., 141:289–91. Riedel, H.D., Muckenthaler, M.U., Gehrke, S.G., Mohr, I., Brennan, K., Herrmann, T., Fitscher, B.A., Hentze, M.W., and Stremmel, W. (1999). HFE downregulates iron uptake from transferrin and induces iron-regulatory protein activity in stably transfected cells. Blood, 94:3915–21. Rouault, T.A., Stout, C.D., Kaptain, S., Harford, J.B., and Klausner, R.D. (1991). Structural relationship between an iron-regulated RNA-binding protein (IRE-BP) and aconitase: functional implications. Cell 64:881–3.

Nitric Oxide and Post-Transcriptional Control of Gene Expression

393

Schalinske, K.L., Anderson, S.A., Tuazon, P.T., Chen, O.S., Kennedy, M.C., and Eisenstein, R. S. (1997). The iron-sulfur cluster of iron regulatory protein 1 modulates the accessibility of RNA binding and phosphorylation sites. Biochemistry, 36:3950–8. Sellers, V. M., Johnson, M. K., and Dailey, H. A. (1996). Function of the [2Fe-2S] cluster in mammalian ferrochelatase: a possible role as a nitric oxide sensor. Biochemistry, 35:2699–704. Smith, M. A., Wehr, K., Harris, P. L. R., Siedlak, S. L., Connor, J. R., and Perry, G. (1998). Abnormal localization of iron regulatory protein in Alzheimer's disease. Brain Res., 788:232–6. Smith, S.R., Cooperman, S., Lavaute, T., Tresser, N., Ghosh, M., Meyron-Holtz, E., Land, W., Ollivierre, H., Jortner, B., Switzer, R. 3rd, Messing, A., and Rouault, T.A. (2004). Severity of neurodegeneration correlates with compromise of iron metabolism in mice with iron regulatory protein deficiencies. Ann. NY Acad. Sci., :65–83. Soum, E., Brazzolotto, X., Goussias, C., Bouton, C., Moulis, J.M., Mattioli, T.A., Drapier, J.C. (2003) Peroxynitrite and nitric oxide differently target the iron-sulfur cluster and amino acid residues of human iron regulatory protein 1. Biochemistry, 42:7648–54. Soum E., and Drapier J. C. (2002) Nitric oxide and peroxynitrite promote complete disruption of the [4Fe-4S] cluster of recombinant human iron regulatory protein 1. J. Biol. Inorg. Chem., 8:226–32. Stuehr, D. J., and Nathan, C. F. (1989). Nitric oxide. A macrophage product responsible for cytostasis and respiratory inhibition in tumor target cells. J. Exp. Med., 169:1543–55. Theil, E. C. (1994). Iron regulatory elements (IREs): a family of mRNA non-coding sequences. Biochem. J., 304:1–11. Tong, W. H., and Rouault, T. (2000). Distinct iron-sulfur cluster assembly complexes exist in the cytosol and mitochondria of human cells. EMBO J., 19:5692–700. Wang, J., Chen, G., Muckenthaler, M., Galy, B., Hentze, M. W., and Pantopoulos K. (2004) Iron-mediated degradation of IRP2, an unexpected pathway involving a 2-oxoglutarate-dependent oxygenase activity. Mol. Cell Biol., 24:954–65. Weiss, G., Bogdan, C., and Hentze, M. W. (1997). Pathways for the regulation of macrophage iron metabolism by the anti-inflammatory cytokines IL-4 and IL-13. J. Immunol., 158:420–5. Weiss, G., Goossen, B., Doppler, W., Fuchs, D., Pantopoulos, K., Werner-Felmayer, G., Wachter, H., and Hentze, M. W. (1993). Translational regulation via iron-responsive elements by the nitric oxide/NO-synthase pathway. EMBO J., 12:3651–7. Yamanaka, K., Ishikawa, H., Megumi, Y., Tokunaga, F., Kanie, M., Rouault, T.A., Morishima, I., Minato, N., Ishimori, K., and Iwai, K. (2003) Identification of the ubiquitin-protein ligase that recognizes oxidized IRP2. Nat. Cell Biol., 5:336–40. Zheng, L., Kennedy, M. C., Blondin, G. A., Beinert, H., and Zalkin, H. (1992). Binding of cytosolic aconitase to the iron responsive element of porcine mitochondrial aconitase mRNA. Arch. Biochem. Biophys., 299:356–60.

Oxide and Tumor 18 Nitric Biology Pierre Sonveaux and Olivier Feron* UCL Medical School, Brussels, Belgium

CONTENTS 18.1 18.2 18.3 18.4

Introduction ............................................................................................395 Cancer and the Tumor Microenvironment ............................................396 NO and Tumorigenesis ..........................................................................398 NO and Tumor Angiogenesis ................................................................402 18.4.1 Regulation of eNOS Expression and Activation during Angiogenesis ..............................................................................402 18.4.2 NO as a Pro-Angiogenic Effector ..............................................405 18.5 NO and Tumor Vascular Reactivity ......................................................407 18.6 NO in Anti-Tumor Therapy ..................................................................409 18.6.1 NO Donors .................................................................................409 18.6.2 Modulation of Endogenous NO Production ..............................411 18.7 Conclusions ............................................................................................413 References ...............................................................................................414

18.1 INTRODUCTION Nitric oxide (NO) has dichotomous activities in many areas of biology. NO can promote cell survival but it also has pro-apoptotic effects. It can stimulate cell growth as well as cell death and necrosis. As a potent vasodilator, NO regulates the vasodynamic balance, but it can also promote angiogenesis, which is associated with the loss of vasoactivity. These apparent contradictory roles have been widely exemplified, leading to the conclusion that the net effect of NO depends on its available concentration in a determined environment (characterized by redox status, the level of reactive oxygen species (ROS), metal ions, specific proteins, and target cell type). In vivo, low levels of NO (picomolar to nanomolar range) are produced by the constitutively expressed NO-synthase (NOS) isoforms, endothelial and neuronal NOS (eNOS and nNOS, respectively) (Figure 18.1). Under calcium activation, these enzymes produce a transient increase in NO levels * Olivier Feron is a Fonds National de la Recherche Scientifique (FNRS) research associate. Pierre Sonveaux is a FNRS post-doctoral fellow.

395

396

Nitric Oxide, Cell Signaling, and Gene Expression

that control important physiological processes such as vasodilation (eNOS) and neuronal transmission (nNOS). Constitutive NOS activation can further be prolonged upon post-translational modifications. Changes in the eNOS phosphorylation status (see below) can, for instance, drive the enzyme toward pro-angiogenic capacities (sustained nanomolar levels of NO). In contrast to the constitutive isoforms, iNOS, the inducible NOS, is expressed under some pathological conditions and generates micromolar NO concentrations (Figure 18.1). It has been demonstrated that iNOS is induced by several agents involved in the inflammatory process, including lipopolysaccharides (LPS) and other bacterial products,1 cytokines, such as tumor necrosis factor-alpha (TNF-α) and interferongamma (IFN-γ),1–3 and viral proteins.4, 5 During host defense, NO generated by iNOS in macrophages behaves as a cytotoxic agent that alters protein integrity and function through S-nitrosylation and induces a variety of phenomena including caspase activation, lipid peroxidation, DNA damage and inhibition of mitochondrial respiration, all leading to cell damage and most often cell death. The genotoxic effects of NO have been attributed to its reaction with oxygen and superoxide to form peroxynitrites and other reactive nitrogen species (RNS) that produce either direct or indirect DNA damage.6, 7 iNOS itself can produce both NO and superoxide simultaneously, suggesting that their combination to form peroxynitrites could readily happen at the time of synthesis.8 Although iNOS is easily induced and expressed in macrophages, many other cell types express iNOS during inflammation or under comparable stresses (reviewed by Hofseth et al.9). Thus, based on the previously mentioned general observations, the role of endogenous NO in cancer should mainly be determined by the level of NO production, itself dependent on the nature of the NOS isoform activated. Accordingly, cell transformation would preferentially be fostered by iNOS expression through the selection of immunopreserved clones bearing RNS or ROS-induced DNA lesions, and eNOS would, instead, participate in the later phase of angiogenesis that supports exponential tumor growth (Figure 18.1). This chapter discusses the fact that although these general assumptions are somehow verified, NO bioactivity and its effects are further determined by the cellular nature and microenvironment of the tumor where it is produced.

18.2 CANCER AND THE TUMOR MICROENVIRONMENT Tumorigenesis is a multistep process that regulates the progressive transformation of normal cells into highly malignant derivatives. Observations of human cancers and animal models have led to the conclusion that tumor development proceeds via a process formally analogous to Darwinian evolution in which a succession of genetic changes confers growth advantages to cancer cells. Because tumor cells often occupy less than one half of the volume of a tumor, the elucidation of the roles played by NO during tumor initiation and development requires

Nitric Oxide and Tumor Biology

397 iNOS Necrosis

µM

Apoptosis

NO concentration

Mutagenesis

iNOS

nM

P-eNOS eNOS

Cell survival Angiogenesis

Vasodilation Minutes

Days Time

FIGURE 18.1 Proposed relationship between NO levels and tumor bioactivity. In a simplified view, the effects of endogenous NO in tumors depend on local concentrations over time. eNOS produces nanomolar concentrations of NO. Following calcium activation in endothelial cells, transient production of NO accounts for vasodilation. Further activation through phosphorylation on Ser1177 allows P-eNOS to produce sustained amounts of NO that promote cell survival and angiogenesis. Similarly, expression of eNOS by certain tumor cells would confer on them a survival advantage over neighbor cells (inhibition of apoptosis). By comparison, expression of iNOS by macrophages and inflamed cells produces higher micromolar amounts of NO. During inflammation, NO behaves as a cytotoxic agent that induces tissue cell apoptosis. If inflammation becomes chronic (in pre-cancerous and cancerous tissues), however, sub-lethal NO levels (dashed line) can promote the selection of apoptosis-resistant pre-neoplastic cells that can bear DNA lesions (mutagenesis). If local levels remain high, NO would eventually lethally damage these cells via direct reactions with DNA (single strand break, alkylation by nitrosamines, and deamination reactions), leading to senescence and necrosis. It requires conditions of oxidative stress where NO combines with superoxide to produce peroxinitrite and other reactive nitrogen species. Because cytokines and hypoxia synergistically induce iNOS expression, the tumor microenvironment may sustain high NO production, thereby further supporting the clonal selection of neoplastic cells and tumor growth.

considering tumors as highly integrated systems in which tumor cells reciprocally interact with the nearby nonmalignant host cells.10 Autopsies of individuals who died of other causes than cancer have revealed the frequent presence of microscopic dormant tumors (in which tumor cell division is equilibrated by apoptosis)11 (Figure 18.2). When compared with the statistics of clinical detection, these observations support the notion that only a very small subset of tumors enters the phase of exponential growth. It is therefore evident that the expression of oncogenic genes by tumor cells, although necessary, is not sufficient for progressive expansion: the acquisition of a specific tumor microenvironment is an absolute prerequisite for tumor growth. One of the main

398

Nitric Oxide, Cell Signaling, and Gene Expression

characteristics of the tumor microenvironment is the onset of hypoxia, which results from insufficient oxygen supply to meet the metabolic requirements (high oxygen consumption rate) for expansion12 (Figure 18.2). However, although hypoxia should eventually lead to cell death, low oxygen tensions in tumors are related to progression and poor treatment outcome.12, 13 Indeed, faced with the hypoxic microenvironment, tumor cells either adapt themselves to survive with a low proliferation rate under oxygen deprivation or adapt the host to their metabolic demand to increase the oxygen supply. The latter option involves the acquisition by the tumor of its own blood vascular network, a process known as “tumor angiogenesis”14 (Figure 18.2). During the multistep process of angiogenesis, the tumor starts to proliferate exponentially. NO is now widely recognized as a potent pro-angiogenic agent in tumors, especially as a mediator of the vascular endothelial growth factor (VEGF) cascade. As opposed to physiological angiogenesis (that occurs, for example, during wound healing), tumors fail to turn angiogenesis off,15 essentially because metabolic requirements of tumor cells perpetually increase with the energetic supply, thereby determining a continuous need for more blood vessels. Consequently, the continuous extension of the vascular network in tumors only leaves limited time for tumor capillaries to properly mature and become fully functional arterioles or veinules. Still, intratumor heterogeneity, especially in slow-growing human tumors (vs. fast-growing experimental animal tumors), enables sufficient maturation of capillaries to acquire a functional arteriolar reactivity necessary to control the tumor blood flow.15–20 Blood flow and flow-related microenvironmental parameters (tissue pO2, pH, distribution, bioenergetic status, and nutrient supply) are important factors known to modulate the sensitivity of cancer cells to ionizing radiations and anticancer agents. Radiotherapy, for example, takes advantage of molecular oxygen to stabilize DNA damage through oxidation of DNA radicals (oxygen effect). Tumor blood flow also serves as a vehicle for the delivery of systemically administered anticancer agents to tumor cells.21 In expanding tumors, blood flow is generally uneven due to the aberrant vascular architecture and low pH, and can even stop and resume.22–24 Various strategies exploiting the functional reactivity of mature tumor arterioles have been proposed to increase the efficiency of radio- and chemotherapies. These ‘pro-vascular’ approaches are aimed at transiently increasing tumor blood flow and O2 bioavailability (by modulating both vascular O2 delivery and local consumption) in tumors.25 As one of the most potent vasodilators and an inhibitor of mitochondrial respiration, NO could play an important role in regulating tumor perfusion and oxygenation. In consequence, NO donors have been tested as adjuvant anticancer strategies.

18.3 NO AND TUMORIGENESIS The relationship between chronic inflammation and tumorigenesis has been well documented in various tissues. Cancer is indeed a typical outcome of several inflammatory pathologies, including Crohn’s disease (colorectal cancer), Wilson’s

Nitric Oxide and Tumor Biology

1 Normal cell

2 Malignant cell

3 Neoplasm

399

H Dormant tumor

4

H Incipient angiogenesis

5

H

Invasive tumor

FIGURE 18.2 Tumor progression as a multistep process. During tumorigenesis (1), repeated mutagenic assaults (yellow arrow) can transform a normal cell into a malignant derivative thereof. This cell and its progeny grow a microscopic neoplasm (2) than can reach a size of a couple of millimeters. The resulting dormant tumor (3) is characterized by a dynamic equilibrium between cell survival and death. A gradient of tumor cell hypoxia (H) arises in regions distant from the closest blood supply. Hypoxia and other microenvironmental influences will eventually lead to the selection of a tumor cell population that produces pro-angiogenic factors (angiogenic switch) (4). Factors such as VEGF initiate the formation of new blood vessels, a process formally known as “tumor angiogenesis”. Angiogenesis is mandatory for further exponential tumor growth (5). Because tumor energetic needs continuously overcome its nutrient supply, event persistent tumor angiogenesis leaves behind zones of tumor hypoxia, and the tumor develops as a rim surrounding a central necrotic core. The preexisting blood vessels included in the tumor mass are known as “co-opted” arterioles and veinules. In reality, the situation is much more complex and regions of growth, dormancy, and regression are in close proximity in heterogenic tumors; their repartition varies over time.

disease (hepatoma and cholangiocarcinoma), ulcerative colitis (colon cancer), hemochromatosis (hepatocarcinoma), acid reflux in Barrett’s esophagus (oesophagal cancer), helicobacter pylori gastritis (gastric cancer), hepatitis B and C virus infections (liver cancer), human papillomavirus infection (cervical cancer), cholecystisis (gall bladder cancer), long-term bladder catheterization (bladder cancer), asbestos (meothelioma), smoking (lung cancer), and several parasitic infections (S. hematobium and bladder cancer, S. japonicum and colon cancer, liver flukes and cholangiocarcinoma).9 iNOS expression and nitrotyrosine accumulation is generally increased in the pre-cancerous mucosa of patients with chronic inflammation. It is also present in a large array of cancers of different types and grades, but its abundance appears to be regulated during tumor progression. In human gynecological and breast cancers, for example, the increased expression of iNOS is inversely associated with the differentiation grade of the tumor.26, 27 Similarly, in colon cancers, iNOS activity is highest in adenomas, then declines with advancing tumor stage, and is lowest in metastatic tumors.28, 29 In melanoma patients, low expression of iNOS is associated with higher risk of developing distant metastases.30 These observations collectively suggest that iNOS could play a particularly important role in the early phases of tumor development (initiation and promotion). Experimentally, tumor initiation is obtained in animal by the delivery of carcinogens, exposure to ionizing radiation, or genetic manipulations (spontaneous tumors in cancer-prone animals). These

400

Nitric Oxide, Cell Signaling, and Gene Expression

models have allowed a better understanding of the relationship between inflammation, iNOS activity, and tumorigenesis. N-nitrosomethylbenzamidine (NMBA) is a carcinogen present in the diet of high-risk populations for esophageal squamous cell carcinoma in China. Chen and Stoner31 reported that iNOS mRNA and protein expression increased progressively in the esophageal epithelium of rats after treatment with NMBA. iNOS expression increased as the tissues progressed from normal to preneoplastic to papilloma states. Macrophages in the involved tissues were also positively stained for iNOS expression, suggesting that NO produced by these cells could contribute to tumor progression. Importantly, the administration of a selective iNOS inhibitor (that decreased NO generation without affecting iNOS expression) significantly suppressed NMBA-induced tumor development.32 In another study, Takahashi et al.33 have examined iNOS expression in a model of rat colon initiated by azoxymethane (AOM). The presence of iNOS in dysplastic but not in hyperplastic epithelial cells suggested that iNOS played a role in the early stages of tumor formation. iNOS was also detected in infiltrating macrophages. In the corresponding model in mice, suppression of colon tumorigenesis by vitamin B6 was associated to suppression of iNOS expression.34 In another study, chemically induced skin tumorigenesis in mice was reduced by 75% upon topical application of nobiletin, an inhibitor of superoxide production (i.e., not a superoxide scavenger) and NO generation by iNOS.35 Chemoprevention by nobiletin was associated with reduced leukocyte infiltration and suppression of oxidative insults by these cells. A related compound, resveratrol, inhibited the growth of prostate cancer cells through inhibition of iNOS activation.36 This compound also suppressed the generation of superoxide and H2O2 by macrophages stimulated by LPS or phorbol esters (TPA).37 It is not yet known whether inhibition of superoxide production by nobiletin and resveratrol proceed directly via inhibition of iNOS or through interaction with other superoxide-producing enzymes. Nevertheless, these selected examples illustrate that, in various tissues, iNOS activation in association with inflammation plays an important role in the promotion of chemically induced tumors. As a consequence, a particularly efficient strategy for tumor chemoprevention in animals combines the suppression of iNOS-mediated NO generation and the inhibition of superoxide production. As in normal tissues, however, NO also possesses opposed functions in the course of chemically induced tumorigenesis. Indeed, it has been reported that nitroglycerin, a nitrovasodilator, can reduce skin tumorigenesis by TPA and the anthracene derivative DMBA in mice.38 Moreover, the nonselective NOS inhibitor L-NAME can promote AOM-induced preneoplastic changes in the colon of rats.39 Conflicting results are better interpreted by considering the dose-dependent bioactivity of NO (Figure 18.1). Excessive NO levels (endogenously produced by iNOS + exogenously delivered by NO donors) may reduce tumorigenesis by enhancing apoptosis and necrosis. Part of these effects may be due to higher DNA damage incompatible with the clonal selection of highly malignant cells. Exogenous NO can also quench free radicals (e.g., superoxide), and high levels of NO could lead to the formation of nontoxic

Nitric Oxide and Tumor Biology

401

metabolites like nitrites and nitrates instead of RNS. Local NO levels higher than physiologically produced by iNOS during inflammation could thus overcome the genotoxicity of ROS and RNS, major carcinogens associated with inflammation. Finally, NO donors, unlike RNS produced by iNOS during inflammation, could inhibit tumor cell proliferation via cGMP-mediated inhibition of PKC.38 The effects of nonselective NOS inhibitors, such as L-NAME, are complex as they will simultaneously influence tumor and non-tumor NOS activity, in ways that can either support or inhibit tumor promotion (Figure 18.1). Similar to chemically induced tumors, iNOS participates in radiationinduced tumorigenesis. Whole-body irradiation with a sub-lethal dose of γ-ray (as a tumor initiator) and administration of diethylstilbestrol (a tumor promoter) during pregnancy or lactation induce mammary tumors in rats.40 Using this model, Inano and Onodo41 have reported that both a nonselective NOS inhibitor and the selective iNOS inhibitor 1,4-PB-ITU prevented tumor initiation by γ-rays. Interestingly, NO formation is enhanced in various tissues of γ-irradiated mice,42 and iNOS is overexpressed in epithelial cells of alveoli and lactiferous ducts in the mammary glands of rats treated with the proinflammatory agent LPS.43 Based on in vitro experiments, it has been proposed that the enhancement of NO production by γ-irradiation is attributable to high levels of iNOS expression in infiltrating macrophages.44 Therefore, radiation-induced tumorigenesis may be partially promoted by excessive NO or RNS produced by radiation-induced iNOS. It is not surprising that both positive and negative effects for NO in tumorigenesis have been reported because the three NOS isoforms are generally present in preneoplastic tissues. Although some NOS inhibitors are relatively selective for iNOS, none are truly specific, making it imprudent to make definitive conclusions from experiments having used these pharmacological tools. In a model of multiple lung adenocarcinoma, Kisley et al.45 have studied urethane-induced lung tumorigenesis in iNOS−/− mice. Although iNOS deficiency did not affect tumor incidence (number of mice bearing cancer), iNOS−/− animals developed 80% fewer tumors than wild-type mice. The tumor growth rate remained unaffected by the absence of iNOS. In normal mice, iNOS expression was particularly apparent throughout the tumor epithelium and particularly intense in cells located at the tumor periphery. Conversely, iNOS was not detected in wild-type mouse lung tissues. The absence of iNOS did not affect the expression and distribution of constitutive NOS in tumor and normal tissues (thereby excluding any compensatory mechanism), and did not alter the recruitment of tumor macrophages. These data unequivocally demonstrate that chronic iNOS activation in lungs promotes tumorigenesis and that this effect is specific to iNOS vs. constitutive NOS. It may be associated with the cytotoxic activity of macrophages independent of their recruitment. Interestingly, tumors isolated from iNOS−/− mice contained twofold less VEGF than tumors derived from wild-type animals, suggesting that deficient neovascularization could further account for reduced lung tumor development in iNOS−/− mice. In another study, Ellies et al.46 have examined the initiation of spontaneous mammary tumors (polyomavirus middle T antigen

402

Nitric Oxide, Cell Signaling, and Gene Expression

under the control of the mouse MMTV-LTR promoter) in iNOS−/− mice. Whereas mammary gland development appeared normal in the absence of iNOS, progression to hyperplasia was delayed in iNOS−/− mice when compared with control mice. Tumor incidence, macrophage infiltration, and angiogenesis (microvascular density), however, were all unaltered in iNOS−/− mice vs. controls. Increased tumor latency in iNOS−/− mice suggested that iNOS action was early and sustained in the process of tumorigenesis. Altered tumor morphology and histologic profiles in iNOS−/− mice further indicated that iNOS might affect further tumor development. The concordance between the studies by Kisley et al.45 and Ellies et al.46 demonstrates that iNOS can stimulate early tumor induction and promotion in different tissues. They also provide compelling rationale for developing and evaluating iNOS-specific inhibitors as chemopreventive agents; however, the role of NO in tumorigenesis is not straightforward: both inhibition (determined as relative to aberrant crypt formation)47 and promotion (number of adenomas)48 of colon tumorigenesis have been reported in the absence of iNOS proteins. Furthermore, how and to which extent iNOS activity influences tumor development (i.e., exponential growth) remains obscure and deserves further characterization.

18.4 NO AND TUMOR ANGIOGENESIS 18.4.1 REGULATION OF ENOS EXPRESSION DURING ANGIOGENESIS

AND

ACTIVATION

Although iNOS plays a critical role in tumor induction, eNOS might be the main source of angiogenic NO in tumors. Several factors present in the tumor microenvironment have the intrinsic capacity to modulate eNOS expression. First, it has been consistently demonstrated that hypoxia down-regulates eNOS expression in cultured pulmonary human endothelial cells.49 In a corresponding animal model, it was attributed to both a decreased rate of transcription and a destabilization of eNOS mRNA.50 In non-pulmonary endothelial cells, the findings are more controversial: both increased51 and reduced52 eNOS expressions have been reported using similar models of endothelial cell hypoxia. Second, eNOS mRNA and eNOS protein are increased in proliferating vs. resting (i.e., confluent) endothelial cells. It was found to be the result of a greater stability of eNOS mRNA in proliferating cells, due to the down-regulation of a mRNA-destabilizing protein that interacts with the 3′-UTR of eNOS mRNA.53 Third, cytokines (e.g., TNF, IFN-γ, interleukin (IL)-12, and LPS-induced cytokines) can regulate eNOS expression in different ways depending on the cytokine combination, the animal species, and the cell type. For example, LPS injection in rats reduces eNOS mRNA expression in aorta54; by contrast, it increases mRNA abundance in cultured bovine endothelial cells.55 Finally, growth factors may up-regulate eNOS mRNA. Incubation of endothelial cells with transforming growth factor (TGF)β enhances the eNOS promoter activity.56 Similarly, basic fibroblast growth factor (bFGF) up-regulates eNOS mRNA, protein, and activity,57 and VEGF has been

Nitric Oxide and Tumor Biology

403

described to increase eNOS expression via the VEGF receptor-2 (VEGFR-2) pathway in cultured human endothelial cells.58 Brouet et al.59 have elucidated the succession of events leading to eNOS activation after treatment of endothelial cells with VEGF, a major pro-angiogenic factor present in tumors (Figure 18.3). The time-course study revealed that, chronologically, endothelial cell exposure to VEGF first leads to a transient increase in intracellular Ca2+ concentrations. This Ca2+ transient appears mandatory to promote the efficient binding of calmodulin (CaM) to eNOS, which induces eNOS dissociation from its inhibitory regulatory protein caveolin (cav)-1. This event is a hallmark of the CaM-mediated activation of eNOS. It then results in a shortterm burst of NO release independently of eNOS phosphorylation (Figure 18.1 and Figure 18.3). Importantly, the CaM/eNOS association is maintained even though intracellular Ca2+ levels return to baseline. This persistent interaction allows the subsequent recruitment of hsp90 to form the eNOS/CaM/hsp90 complex, which, in turn, recruits both the kinase Akt and the phosphatase calcineurin in the eNOS vicinity. The formation of this multi-protein complex fosters the phosphorylation of eNOS on the serine 1177, and its dephosphorylation on the threonine 495. Ser1177 phosphorylation leads to the long-term activation of eNOS, which produces sustained amounts of NO (Figure 18.1 and Figure 18.3). Importantly, the dissection of this multistep activation pathway revealed that the initial increase in intracellular Ca2+ leading to CaM recruitment (often referred to as “Ca2+-dependent eNOS activation”) is also critical for the late phase of eNOS phosphorylation (“Ca2+-independent eNOS activation”) even if Ca2+ levels are already back to baseline during this late step. Accordingly, intracellular delivery of a Ca2+ chelator not only prevented eNOS/CaM association (and, thereby, the early phase of eNOS activation), but also the late enzyme activation and prolonged NO production. Furthermore, geldanamycin, an inhibitor of hsp90, had no impact on the early phase, but completely blocked further Akt recruitment and consecutive eNOS phosphorylation and activation. Of note, the increase in Ca2+ levels upon VEGF stimulation has been documented to result from VEGFR2-mediated activation of PLC-γ,60 whereas Akt activation is known to occur following the VEGF-activation of PI3K61 (Figure 18.3). In addition, the chaperone role of hsp90 for Akt was recently extended to calcineurin that dephophorylates eNOS on the threonine 495.62 Similar to VEGF, other factors that are present in the tumor microenvironment are obviously capable of modulating eNOS activity at the post-translational level. For example, yet unidentified factors participate in decreasing eNOS activity by inhibiting Akt activity and Ser1177 phosphorylation during chronic hypoxia.63 In addition, bFGF induces eNOS activation,64 although unlike for VEGF, the underlying mechanisms appear to be strictly Ca2+-independent and to involve ceramide. Activation of eNOS by TNF-α in tumor cells utilizes both the PI3K/Akt and ceramide pathways.65 With respect to the VEGF pathway, the post-translational regulation of eNOS in tumors and, a fortiori, in the tumor vasculature is still a poorly explored field.

404

Nitric Oxide, Cell Signaling, and Gene Expression

VEGF

VEGFR-2 OUT

IN eNOS

eNOS P P

P P

(2)

(3)

cav-1

P Akt

(4) P Akt

CaM Ca2+

(2′)

hsp90

NO

(1)

PI3K

eNOS

P eNOS

hsp90

ER Myristoyl

Sphingosine

Palmitoyl

Calmodulin

FIGURE 18.3 Proposed model for eNOS activation through the VEGF pathway. Constitutive eNOS inactivation (basal activity) is maintained upon its inhibitory interaction with cav-1, a structural protein of caveolae. (1) The binding of VEGF to the tyrosinekinase VEGFR-2 expressed by endothelial cells results in the stimulation of agonist pathways. Important pathways controlling eNOS activity include the PI3K/Akt pathway and the phospho-inositoside cascade that promotes Ca2+ release from intracellular storage, including the endoplasmic reticulum (ER). (2) Increased Ca2+ availability activates calmodulin that can now interact with eNOS and releases the enzyme from the cav-1 interaction. It induces a burst in NO production (short-term activation). (2’) Akt is activated (phosphorylated) simultaneously to Ca2+ release, but remains away from eNOS in the cytosol. (3) The interaction of eNOS with CaM is a prerequisite for the further recruitment of hsp90 to the eNOS complex. This event is dependent on the earlier Ca2+ release but may occur when Ca2+ levels have already returned to basal. (4) Hsp90 functions as a chaperon protein, allowing the subsequent recruitment of activated Akt. Upon interaction, Akt phosphorylates eNOS on Ser1177, leading to sustained eNOS activation and long-term NO production. Once activated, eNOS is thought to be depalmitoylated, thereby abolishing its caveolar anchorage. Other phosphorylation sites in eNOS, including Thr495, function as control elements for the modulation of eNOS activity (not shown). As a freely diffusible radical, NO can cross membranes to exert its activity in the surrounding vascular smooth muscle cells. Other receptor-ligand interactions can activate the same intracellular cascades leading to short-term and, sometimes, long-term eNOS activation. P: phosphate residue.

Nitric Oxide and Tumor Biology

18.4.2 NO

AS A

405

PRO-ANGIOGENIC EFFECTOR

Tumor angiogenesis initiates with vasodilation, a process involving VEGFinduced endothelial NO release.66 Subsequently, vascular permeability increases in response to VEGF, which implicates the formation of vesiculo-vesicular organelles, partly through the coalescence of caveolae. Interestingly, VEGFdependent Akt-mediated increase in vascular permeability is inhibited by the NOS inhibitor L-NAME, indicating that NO could play a crucial role in the early steps of tumor angiogenesis.67 The implication of NO in physiological angiogenesis has now been well established. As first indirect evidence, end products of NO synthesis—nitrites and nitrates—are elevated early and transiently in fluids collected from sponges implanted in sub-cutaneous wounds.68 In vitro, NO donors, such as sodium nitroprusside, promote endothelial cell proliferation and migration; inhibitors of NOS suppress this response.69 Furthermore, systemic administration of L-arginine (the substrate of NOS) enhances wound healing.70 Topical application of an NO donor accelerates closure of excisional wounds, whereas NOS inhibitors given systemically or applied to the surface of wounds delay healing (see references in Lee et al.71). Wound healing occurs in response to the production of multiple growth factors, including VEGF, bFGF, platelet-derived growth factor (PDGF), insulin-like growth factor (IGF-1), and TGF-β. With this in mind, the direct evidence that NO induces angiogenesis was initially reported by two different groups who were investigating the mechanisms of VEGF-induced angiogenesis. Papapetropoulos et al.72 used an in vitro model in which 3-dimensional cultures of human venous endothelial cells (HUVEC) in type I collagen gels received exogenous VEGF. This model had been previously developed to demonstrate that VEGF, bFGF, TGF-β, and phorbol myristate acetate (PMA) stimulate the proliferation-independent formation of tube-like networks from unorganized endothelial cells. HUVEC exposed to VEGF appeared to engage in the formation of a vascular network within 24–48 hours. Vessel-like structures subsequently disassembled, a phenomenon attributable to the lack of factors necessary for network maintenance in this assay. The organization of HUVEC into cords and network structures was largely inhibited in the presence of the NOS inhibitor L-NAME. Using pharmacological tools, Papapetropoulos et al.72 further evidenced that VEGF-mediated NO production in cultured HUVEC required the activation of tyrosine-kinases, a transient raise in intracellular Ca2+ concentrations, the presence of bioavailable calmodulin, the activation of the PI3K pathway and downstream Akt, and cGMP production, all supporting the involvement of eNOS, but not iNOS or nNOS (not expressed in HUVEC) in this process. Long-term exposure to VEGF also increased eNOS expression in HUVEC. Ziche et al.73 utilized a model of angiogenesis in which slow-releasing VEGF pellets were implanted in the cornea of rabbits. VEGF-induced corneal angiogenesis was completely prevented when rabbits received L-NAME in the drinking water, demonstrating that NO is a key pro-angiogenic messenger in vivo, acting downstream VEGF. By contrast, angiogenesis elicited by bFGF appeared to be NO-independent. Due to

406

Nitric Oxide, Cell Signaling, and Gene Expression

the lack of selective inhibitors, the final demonstration that eNOS is the main source of pro-angiogenic NO further required the use of eNOS−/− mice. In a model of operatively induced hindlimb ischemia (in which the entire femoral and saphenous arteries and their major branches were dissected free), angiogenesis was significantly impaired in eNOS−/− vs. wild-type mice.74 Both eNOS−/− and eNOS+/+ mice expressed similar levels of VEGF gene and protein in the ischemic limb tissues. Thus, eNOS deletion had apparently no impact on VEGF expression. Moreover, angiogenesis was not restored by either recombinant VEGF protein administration or adenovirus-mediated VEGF gene transfer in eNOS−/− mice, demonstrating that eNOS, but not iNOS or nNOS, was required to transmit the VEGF pro-angiogenic signal. Importantly, systemic administration of an NO donor did not restore angiogenesis in vivo, suggesting that endogenous eNOS activity is critical to meet local or temporal requirements that are necessary for effective angiogenesis. These results were further confirmed in models where wound healing was reported to be almost completely absent in eNOS−/− mice and derived tissues. It was attributed to a reduced efficacy of VEGF signaling. Furthermore, VEGF-induced angiogenesis and vascular permeabilization were absent in eNOS−/− mice, but preserved in iNOS−/− mice.75 Of note, although iNOS plays only a minor role in VEGF-induced angiogenesis, it may be significantly more important in response to other pro-angiogenic factors. Indeed, wound closure was reported to be delayed by 31% in iNOS−/− mice vs. wild-type mice.76 In normal tissues, angiogenesis is strictly regulated by a reciprocal control of NO on VEGF expression: an excessive amount of NO, as can be produced upon longterm iNOS activation, acts negatively on VEGF synthesis, probably by limiting HIF-1α activity.77 Because tumors express high levels of VEGF to support angiogenesis in response to hypoxia, NO is likely to be an important mediator of tumor angiogenesis. Accordingly, NOS activity is elevated in various tumors, and NOS protein and activity have been positively correlated with the degree of malignancy for tumors of the human reproductive tract, breast, and central nervous system. Experimental tumors have provided more direct evidence of the contributory role of NO in tumor progression: systemic treatment of tumor-bearing mice with NOS inhibitors delayed tumor growth through eNOS inhibition, and prevented metastasis, suggesting a possible implication of endotheliumexpressed eNOS in metastasis.78 Interestingly, some studies reported an inverse correlation between NO and tumor progression and metastasis; when genetically transduced to overexpress iNOS, these tumors lost their tumorigenic and metastatic abilities as a result of NO-mediated apoptosis.79, 80 This suggests a dual role for NO in these processes: the susceptibility of tumor cells to NO-mediated injury may depend on levels of NO produced, which is determined by the nature and localization of the NOS isoform activated78 (Figure 18.1). The first evidence of the role of NO in tumor angiogenesis was obtained in a model of mice challenged for corneal angiogenesis after implantation of tumor cells that were engineered to express high amounts of VEGF.73 Mice rapidly developed extensive corneal angiogenesis upon cell transplantation, and it was inhibited by the

Nitric Oxide and Tumor Biology

407

systemic delivery of L-NAME. Consequently, the cells remain dormant in the cornea. Importantly, neovascularization induced by naive tumor cells appeared late and was insensitive to L-NAME, revealing that, beyond VEGF, other tumorderived pro-angiogenic factors, such as bFGF, can overcome the impairment or lack of VEGF signaling. Trying to elucidate which NOS isoform promoted tumor angiogenesis, Jadeski and Lala78 used a model of mammary adrenocarcinoma expressing eNOS but not iNOS, so that the major source of NO in the tumor was eNOS expressed in tumor and endothelial cells. Sustained systemic administration of L-NAME dramatically reduced angiogenesis in implanted matrix plugs containing the tumor cells, indicating that prolonged activation of eNOS is likely to play a major role in tumor neovascularization and, hypothetically, in tumor dissemination. This is consistent with the current view of angiogenesis-dependent metastasis. Progression of a syngeneic Lewis lung carcinoma was markedly reduced in eNOS−/− mice compared with wild-type mice, an effect associated with a twofold reduction in endothelium-specific staining in the tumor.81 Thus, tumor growth retardation in eNOS−/− mice can be attributed, at least in part, to a reduced angiogenic rate. Collectively, increasing evidence indicates that the eNOS pathway might be a promising therapeutic target to consider for anti-angiogenic strategies aimed at treating cancer. It will most certainly foster the development of specific eNOS inhibitors, as well as strategies targeting this pathway. Further support for that view comes from the observation that the irradiated tumor vasculature takes an active part in determining the overall efficiency of X-ray radiotherapy (XRT) through eNOS activation.82 Indeed, beside well-known cytotoxic effects, repeated irradiations select a population of endothelial cells that have acquired the capacity to undergo angiogenesis. Using in vitro, ex vivo, and in vivo assays, we provided the first evidence that XRT promotes angiogenesis by activating the eNOS pathway in the tumor vasculature.82 X-rays induce a dramatic increase in NO (released from eNOS) through both transcriptional and post-translational regulations of the enzyme expression and activity. These molecular events are likely to confer radioresistance to endothelial cells (e.g., through activation of the PI3K/Akt survival pathway),83 and to activate angiogenesis. Accordingly, we reported that irradiated endothelial cells acquire new migratory, networking, and sprouting capabilities, which all are dependent upon NO release. Dissection of the molecular and functional changes governing XRT-induced angiogenesis strongly suggests that such events occur in XRT-treated tumors in vivo. Indeed, irradiation induces a massive recruitment of endothelial cells at the margin of irradiated tumors, and these cells have the capacity to generate an extensive vascular network. The inhibitory effect of L-NAME on endothelial cell colonization further suggests that NO is not only involved in endothelial cell networking but also participates in their recruitment.

18.5 NO AND TUMOR VASCULAR REACTIVITY In non-tumoral vessels, the eNOS pathway of vasodilation is activated by receptor-dependent agonists (i.e., acetylcholine, bradykinin, substance P, vasopressin,

408

Nitric Oxide, Cell Signaling, and Gene Expression

adrenaline, noradrenaline, ATP, and platelet-derived products, such as serotonin, histamine, adenosine diphosphate [ADP], and thrombin) that increase intracellular Ca2+ levels.21, 84 It is further activated by shear forces exerted by the circulating blood cells, thereby causing flow-dependent vasodilation. Because the endothelial control of the vascular tone by NO is a sensitive and finely tuned process, peculiarities of the tumor microenvironment are likely to be dramatically important for the vascular function. The precise effects of hypoxia on eNOS expression require clarification, however. Discrepancies between different reports may well be related to differences in the extent and duration of hypoxia, as well as varying responses between tissues. Short-term hypoxia causes physiological and reversible modulation of vascular tone and blood flow, whereas chronic hypoxia results in irreversible remodeling of the vasculature and surrounding tissues, suggesting that gene expression may be different under the two conditions. Furthermore, NO production under hypoxia may be very low even if increased gene and protein expression of eNOS exists because of the requirement of O2 for enzymatic activity.12 More important, it is evident that the vasodynamic function of NO in tumor vessels should be restricted to sufficiently mature vessels having acquired a vascular smooth muscle (VSMC) coat. Despite those intrinsic limitations, accumulating evidence indicates that NO is a major factor controlling the tumor blood flow.12 For example, pharmacological inhibition of NOS, using systemic administration of L-NMMA and L-NAME, reduced blood flow in models of murine adenocarcinoma and melanoma.85 These effects were reversed by administration of L-arginine. In this study, the same inhibitors failed to modulate the blood flow in corresponding nonmalignant tissues. These results were confirmed using a model of mammary adenocarcinoma in which NOS inhibitors were locally infused: NOS inhibition reduced both tumor and control veinule perfusion, but the effect was blunted in the vicinity of tumors, possibly because of increased NOS levels.86 In another study, the effect of the NOS inhibitor L-NNA was selective for sarcomas vs. normal tissues, with a significant decrease in tumor blood flow and corresponding increase in vascular resistance in the tumor but no significant effect in any other normal tissue analyzed.87 Increasing the dose of the NOS inhibitor resulted in systemic vasoconstriction. The higher sensitivity of tumor vessels vs. normal vessels to NOS inhibition, which could be reversed by L-arginine, suggests that NO overproduction is important for maintaining a vasodilatory tone in some tumor types. In another study, the NOS inhibitor Nnitro-L-arginine, injected 30 minutes before X-rays, increased tumor cell survival 3–5-fold in an experimental squamous cell carcinoma and 50–200-fold in a murine radiation-induced fibrosarcoma.88 These effects were equivalent to those obtained from clamped tumors, indicating full radiobiological hypoxia (i.e., NO production sustained tumor oxygenation, and NOS inhibition aggravated tumor hypoxia). When combined with a bioreductive agent, NOS inhibition induced tumor necrosis88 in a way very similar to what was observed using anti-vascular strategies. Altogether, these experiments indirectly reveal that vasoactive NO is present in the tumor vasculature, and is functionally able to control tumor blood flow, oxygenation, and energetic supply.

Nitric Oxide and Tumor Biology

409

18.6 NO IN ANTI-TUMOR THERAPY 18.6.1 NO DONORS As a strong vasodilator, NO has the potency to induce vasodilation of mature tumor vessels, subsequently leading to tumor radiosensitization. Several groups of investigators have studied the effects of NO donors on the tumor vasculature. Jordan et al.89, 90 reported that a systemic delivery of the NO donor isosorbide dinitrate to fibrosarcoma (FSA)-II tumor-bearing mice was as efficient as carbogen breathing at inducing an increase in tumor blood flow, pO2, and response to XRT. Similarly, Wood et al.91 reported that the systemic administration of the NO donor 3-morpholinosydnonimine hydrochloride (SIN-1; 2 mg/kg) significantly decreased adenosine triphosphate (ATP) turnover in SCCVII/Ha tumor-bearing mice. They proposed that these changes reflected an increase in tumor blood flow and pO2. Such treatment administered immediately before irradiation increased tumor cell killing 2–4-fold over that obtained with 15 Gy X-rays alone.91, 92 The effects of SIN-1 on tumor metabolism, however, turned out to be highly variable in function of time and dose at which SIN-1 was delivered: the improvement in energy levels were maximal 5 minutes after injection of 2 mg/kg SIN-1 and returned to control levels by 20 minutes, whereas SIN-1 at 10 mg had no effect at 10 minutes but, instead, increased ATP metabolism 1 hour after delivery. In addition, SIN-1 (2 mg/kg) did not consistently alter energy levels in RIF-1 tumorbearing mice. Moreover, considering that only 10% of vessels in the SCCVII tumors possess smooth muscle, the effects of SIN-1 on the reactivity of the tumor microvasculature are questionable.21 These results instead argue for a steal effect (Figure 18.4) in which, on an individual tumor basis, the response of the host vasculature to increasing concentrations of SIN-1 shunts away the blood flow from the tumor to normal tissues. Song 21 investigated the effects of the NO donor diethylamine (DEA)/NO on the tumor blood flow in R3230 Ac mammary adenocarcinoma-tumor bearing rats. An intravenous injection of DEA/NO decreased tumor blood flow by 30%. Similarly, it decreased tumor pO2 for about 30 minutes in FSA-II tumor-bearing mice. It appeared that exogenous NO caused preferentially vasodilation in host tissues. Fukumura et al.16 observed that superfusion of the NO donor spermine/NO increased tumor vessel diameter and flow rate, whereas systemic injection had no significant effect on these parameters in tumorbearing mice. Similarly, Shan et al.93 documented a decrease in tumor oxygenation following an intravenous administration of DEA/NO. DEA/NO reduced mean arterial pressure (MAP) and tumor blood flow but did not affect muscle blood flow, except at higher doses. These findings suggest that DEA/NO decreases tumor blood flow in two ways: vascular steal occurs during the initial period of hypotension, and reduced perfusion pressure (induced by lowered MAP) further decreases flow to the tumor, probably because of relatively higher flow resistance. In tumor-bearing rats, Thews et al.94 did not report any reoxygenation associated to a systemic injection of the NO donor sodium nitroprusside. Instead, systemic application of the vasodilator drug resulted in a dose-dependent decrease in MAP

410

Nitric Oxide, Cell Signaling, and Gene Expression

paralleled by a decrease in tumor perfusion and pO2. Resistance to flow did not change during the infusion, indicating that sodium nitroprusside had no impact on tumor vessel diameter. As illustrated here, the benefits of NO donors in inducing tumor reoxygenation and, thereby, radiosensitization are quite questionable because modifications of tumor blood flow consecutive to the systemic administration of such drugs is mainly governed by vascular steal and variation in MAP. Thus, like NOS inhibitors but for different reasons, NO donors typically cause a reduction in tumor blood flow. In experimental tumors, the weak response of the tumor vasculature to exogenous NO can result either from a saturation of the NO dilatory pathway by endogenous NO or from the paucity in mature tumor vessels sensitive to NO and to poor NO delivery in the tumor microcirculation. The latter can be attributed to low microregional accessibility inherent to the nature of the tumor vasculature. It is also likely that, by contrast to endogenous NO, exogenous NO in tumors fails to reach the local subcellular threshold required to modulate vascular reactivity. (eNOS activity typically results in elevated concentrations of NO in the cellular compartment where eNOS is expressed, e.g., caveolae.) An additional level of variability is related to the intrinsic chemical activities of NO donors controlling the mechanisms and rate of NO generation. NO release from SIN-1, for example, requires the presence of oxygen.95 It is also conceivable that high systemic concentrations of NO could increase NO dependent S-nitrosylation of hemoglobin. Indeed, the binding of oxygen to haem irons in the lungs promotes the binding of NO derivatives to a specific cystein residue of hemoglobin, forming S-nitrosohemoglobin.96 Partial deoxygenation of S-nitrosohemoglobin in peripheral tissues is accompanied by an allosteric transition that further promotes oxygen delivery and unloading of NO-related vasodilators. In tumor-bearing patients, the transition to deoxyhemoglobin is believed to occur in moderately hypoxic tissues surrounding the tumor.97 By promoting hemoglobin S-nitrosylation in the host vasculature instead of local changes in the tumor microcirculation (for the reasons detailed previously), NO donors would facilitate blood deoxygenation at the tumor periphery where mature arterioles participate in the steal effect. In addition to its vasoactive properties, several NO donors have been reported to radiosensitize hypoxic cells in vitro, a process in which NO gas turned out to be as effective as oxygen.98, 99 Although it was proposed that such an effect was dependent on NO-mediated inhibition of mitochondrial respiration, the delivery of NO to hypoxic cells in culture still achieved radiosensitization. It is therefore reasonable to envisage that reactions between radiation-induced carbon-centered radicals on DNA and NO promote fixation of DNA damage in a way similar to the oxygen effect of radiosensitization; however, the use of NO gas in animals (or humans) to radiosensitize hypoxic cells is problematic because breathing high concentrations of NO gas can damage lung tissues. Moreover, their hypotensive effects limit the clinical use of NO donors. Recently, Jordan et al.100 have reported that stabilization of DNA damage by NO can be exploited in vivo using (e.g., insulin). Indeed, although insulin achieved a smaller increase in tumor pO2 than carbogen breathing, it radiosensitized tumors more efficiently. The

Nitric Oxide and Tumor Biology

411

Flow In series Flow Host tissue

Tumor

Flow

In parallel

Flow

FIGURE 18.4 Vascular steal effect. The changes in tumor blood flow caused by nontumor-selective vasoactive treatments are fundamentally influenced by the structural relationship between the tumor vascular bed and the vascular bed of the surrounding normal tissues. According to this model, an increase in the blood flow in normal tissues results in an increase in tumor perfusion if vascular beds are in series (top). Conversely, if vascular beds are in parallel (bottom), an increase in the blood flow in normal tissues reduces tumor perfusion (vascular steal). In most tumors, beds in parallel and in series are mixed, rendering unpredictable the overall effect of non-tumor-selective vasoactive drugs.

radiosensitizing properties of insulin were lost in eNOS−/− mice, demonstrating the implication of vascular NO in tumor radiosensitization. A key point for further preclinical studies is to selectively target hypoxic cells using this approach.

18.6.2 MODULATION

OF

ENDOGENOUS NO PRODUCTION

The lack of specificity for tumor vs. normal vessels blunts the therapeutic use of NO donors as pro-vascular agents (e.g., agents aiming at modifying the tumor vessel reactivity). One could alternatively envisage that the specific activation of the eNOS pathway in the tumor vasculature would tip the vasodynamic balance in favor of vasodilation. Interestingly, some lines of evidence suggest that XRT itself could selectively activate the eNOS pathway within the tumor vasculature. Indeed, irradiation is associated to the local production of reactive oxygen species (ROS),101 and independent reports indicate that some of these species, such as H2O2, activate the eNOS pathway when delivered to cultured endothelial cells.102 Vasomodulatory pathways are profoundly altered in mature tumor vessels. It was recently reported that in a some experimental tumor models, although the constrictive side of the vasodynamic balance is maintained aberrantly activated (through continuous activation of the endothelin ET-1/ETA pathway),103 NO-mediated vasodilation, which counterbalances ET-1 activity, is deficient in tumor vessels.104 Low-dose

412

Nitric Oxide, Cell Signaling, and Gene Expression

radiotherapy, delivered as a local treatment, can overcome one of the major limitations of NO donors-based approaches (i.e., vascular steal). Ionizing radiation constitutes a strong stimulus leading to the tumor-selective activation of the eNOS pathway in tumor microvessels.104 ROS are likely to mediate at least part of this response. It is important to stress out that, although irradiation produces large amounts of ROS known to react with NO and neutralize its action,105 the blocking effects of NOS inhibitors in functional read-outs demonstrate that NO production reaches sufficient levels or is compartmentalized to overcome the scavenging effects of ROS. eNOS activation accounts for the active participation of the functional tumor vasculature in the efficiency of fractionated radiotherapy (Figure 18.5). Unexpectedly, the tumoricidal effect does not arise from induced vascular cytotoxicity, but instead results from NO-mediated vasodilation (and probably permeabilization), leading to tumor reperfusion and reoxygenation. Tumor reoxygenation is blocked using the NOS inhibitor L-NAME, which excludes a mere decompression effect. Moreover, because irradiation-induced NO production by eNOS does not modify the respiration rate of tumor cells, tumor reoxygenation appears exclusively attributable to vascular responses to X-rays. Radiosensitization through vasomodulation constitutes a strong rationale for the use of XRT in its fractionated mode. It also suggests that the frequency of exposure to X-rays could be finely tuned to exploit the best window of tumor reoxygenation in treating a given tumor in a given patient. Tumor-selective reperfusion induced by ionizing radiations also improves tumor accessibility to circulating antitumor agents (Figure 18.5). Our own experiments demonstrate increased uptake of cationic liposome reporter cDNA plasmid complexes in irradiated tumors.104 Increased transgene delivery appears to be exquisitely NO-dependent because both L-NAME administration and the use of eNOS−/− mice almost completely prevented transgene expression in tumors. In slow-growing human tumors, an increased number of sufficiently mature vessels prone to autoregulation are likely to further improve tumor sensitivity to XRT-based pro-vascular strategies. Complementary studies are required to evaluate whether such an approach can be exploited in combination with other anti-tumor treatments (e.g., chemotherapeutic drugs). If so, it has the potential not only to increase the accessibility of circulating agents to the tumor, but also, consequently, to allow dose down-scheduling for an identical efficiency while decreasing collateral damages. Conversely, the key role of the tumor vasculature certainly deserves most attention in the design of therapeutic protocols or clinical trials in which XRT is combined with another anticancer treatment. The slow-growing nature of most human tumors, that allows time for post-angiogenic vessels to mature, certainly constitutes an appreciable advantage in favor of pro-vascular vs. anti-angiogenic approaches. The initiation of angiogenesis by ionizing radiations (as detailed previously), however, provides a strong rationale to combine radiotherapy to an antiangiogenic approach. The vasodynamic status and the effects of radiations on the eNOS pathway in the tumor vasculature are summarized in Figure 18.6.

Nitric Oxide and Tumor Biology

r Tumo

413

XRT TC pO2 VSMC eNOS blood

EC

X-rays Local irradiation

NO

NO eNOS Perfusion

Permeabilization ? Transient dilation

FIGURE 18.5 Low dose irradiation as a pro-vascular approach. Mature tumor vessels constitute an important barrier preventing the accessibility of circulating tumor agents (spots) to the tumor cell compartment (on left). The vascular pathway of NO production, which depends on eNOS activity in endothelial cells (EC) and controls the vasodilatory side of the vasodynamic balance, is defective. This functionality is restored upon local delivery of a small dose of X-rays that activates the eNOS pathway through expressional and post-translational modulations (on right). X-ray-induced NO release activates the cGMP-mediated pathway of vasodilation in vascular smooth muscle cells (VSMC). It is also likely to increase vascular permeability. Consequently, the irradiated tumor vasculature actively promotes a tumor-specific and transient increase in tumor perfusion and oxygenation, exploitable when combined with conventional anticancer treatments (chemo- and radiotherapies).

18.7 CONCLUSIONS Regarding the question of whether NO is a good or a bad thing in tumors, the answer is not more straightforward than in the heart or other tissues. Still, the roles of NO in tumorigenesis and tumor angiogenesis are now better understood and more generally, the tumor microenvironment appears as the key regulator of the influence of NO on the tumor’s fate. The use of NO donors or NOS inhibitors have revealed their limitations and today, additional research should focus on how to modulate endogenous NOS activity to exploit at best local NOS isoform expression. In other words, the major challenge is to determine in every set of circumstances peculiar to the development of a given tumor whether the blockade or the stimulation of NO-dependent processes should be preferred, depending on whether they support tumor survival or could help to eradicate the tumor, respectively.

414

Nitric Oxide, Cell Signaling, and Gene Expression

Myogenic tone

ET-1/ETA inhibition

Defective NO pathway

Low-dose irradiation

pO2

Angiogenesis NO

Vasodilation Flux Tumor accessibility Radiotherapy

Repeated irradiations (fractionated XRT)

Immunotherapy ? Chemotherapy

Anti-angiogenic therapy

FIGURE 18.6 The mature tumor vasculature as a key determinant of tumor response and resistance to treatments. Mature vessels are prone to blood flow regulation controlling energetic and oxygen supply. This functional reactivity is selectively altered in tumors. Indeed, the vasodynamic balance tips in favor of constriction because massive and local ET-1 production by tumor cells supports a basal constrictive myogenic tone (top left), and because the eNOS pathway of vasodilation is defective in tumor endothelial cells (top right). Importantly, both inhibition of the myogenic tone (using ETA inhibitors) and restoration of the NO pathway (using local low-dose irradiation) promote selective and transient tumor reperfusion and reoxygenation (bottom left). Such pro-vascular approaches increase the efficiency of conventional antitumor therapies. Conversely, the long-term activation of the eNOS pathway induced by repeated irradiations accounts for the activation of tumor angiogenesis (bottom right). This constitutes a strong rationale for strategies combining fractionated radiotherapy (XRT) to an anti-angiogenic approach to improve cancer cure.

REFERENCES 1. Stuehr, D.J., Cho, H.J., Kwon, N.S., Weise, M.F., and Nathan, C.F. Purification and characterization of the cytokine-induced macrophage nitric oxide synthase: an FAD- and FMN-containing flavoprotein. Proc. Natl. Acad. Sci. USA 88, 7773–7 (1991). 2. Markewitz, B.A., Michael, J.R., and Kohan, D.E. Cytokine-induced expression of a nitric oxide synthase in rat renal tubule cells. J. Clin. Invest. 91, 2138–43 (1993). 3. Koide, M., Kawahara, Y., Tsuda, T., and Yokoyama, M. Cytokine-induced expression of an inducible type of nitric oxide synthase gene in cultured vascular smooth muscle cells. FEBS Lett. 318, 213–7 (1993). 4. Amaro, M.J., Bartolome, J., and Carreno, V. Hepatitis B virus X protein transactivates the inducible nitric oxide synthase promoter. Hepatology 29, 915–23 (1999).

Nitric Oxide and Tumor Biology

415

5. Elmore, L.W. et al. Hepatitis B virus X protein and p53 tumor suppressor interactions in the modulation of apoptosis. Proc. Natl. Acad. Sci. U S A 94, 14707–12 (1997). 6. Szabo, C., Zingarelli, B., O'Connor, M., and Salzman, A.L. DNA strand breakage, activation of poly (ADP-ribose) synthetase, and cellular energy depletion are involved in the cytotoxicity of macrophages and smooth muscle cells exposed to peroxynitrite. Proc. Natl. Acad. Sci. USA 93, 1753–8 (1996). 7. Szabo, C. and Salzman, A.L. Inhibition of terminal calcium overload protects against peroxynitrite-induced cellular injury in macrophages. Immunol. Lett. 51, 163–7 (1996). 8. Xia, Y. and Zweier, J.L. Superoxide and peroxynitrite generation from inducible nitric oxide synthase in macrophages. Proc. Natl. Acad. Sci. USA 94, 6954–8 (1997). 9. Hofseth, L.J., Hussain, S.P., Wogan, G.N., and Harris, C.C. Nitric oxide in cancer and chemoprevention. Free Radic. Biol. Med. 34, 955–68 (2003). 10. Hanahan, D. and Weinberg, R.A. The hallmarks of cancer. Cell 100, 57–70 (2000). 11. Black, W.C. and Welch, H.G. Advances in diagnostic imaging and overestimations of disease prevalence and the benefits of therapy. N. Engl. J. Med. 328, 1237–43 (1993). 12. Dachs, G.U. and Tozer, G.M. Hypoxia modulated gene expression: angiogenesis, metastasis and therapeutic exploitation. Eur. J. Cancer 36, 1649–60 (2000). 13. Vaupel, P., Kelleher, D.K., and Hockel, M. Oxygen status of malignant tumors: pathogenesis of hypoxia and significance for tumor therapy. Semin. Oncol. 28, 29–35 (2001). 14. Folkman, J. Tumor angiogenesis: therapeutic implications. N. Engl. J. Med. 285, 1182–6 (1971). 15. Bergers, G. and Benjamin, L.E. Tumorigenesis and the angiogenic switch. Nat. Rev. Cancer 3, 401–10 (2003). 16. Fukumura, D., Yuan, F., Endo, M., and Jain, R.K. Role of nitric oxide in tumor microcirculation. Blood flow, vascular permeability, and leukocyte-endothelial interactions. Am. J. Pathol. 150, 713–25 (1997). 17. Lubbe, A.S. and Huhnt, W. Microvessel diameters of human colon adenocarcinoma during acute treatment with serotonin. Int. J. Microcirc. Clin. Exp. 14, 218–25 (1994). 18. Huhnt, W. and Lubbe, A.S. Venules and arterioles in xenotransplanted human colon adenocarcinoma critically constrict with hyperthermia and serotonin. J. Cancer Res. Clin. Oncol. 121, 267–74 (1995). 19. Thews, O., Kelleher, D.K., and Vaupel, P.W. Modulation of spatial O2 tension distribution in experimental tumors by increasing arterial O2 supply. Acta Oncol. 34, 291–5 (1995). 20. Emerich, D.F. et al. Bradykinin modulation of tumor vasculature: I. Activation of B2 receptors increases delivery of chemotherapeutic agents into solid peripheral tumors, enhancing their efficacy. J. Pharmacol. Exp. Ther. 296, 623–31 (2001). 21. Song, C.W. Blood perfusion and microenvironment of human tumors. (Molls, M. and Vaupel, P., eds.), pp. 193–208 (Springer-Verlag, Berlin, 1998). 22. Kiani, M.F., Pries, A.R., Hsu, L.L., Sarelius, I.H., and Cokelet, G.R. Fluctuations in microvascular blood flow parameters caused by hemodynamic mechanisms. Am. J. Physiol. 266, H1822–8 (1994). 23. Dewhirst, M.W. et al. Microvascular studies on the origins of perfusion-limited hypoxia. Br. J. Cancer Suppl. 27, S247–51 (1996).

416

Nitric Oxide, Cell Signaling, and Gene Expression

24. Kimura, H. et al. Fluctuations in red cell flux in tumor microvessels can lead to transient hypoxia and reoxygenation in tumor parenchyma. Cancer Res. 56, 5522–8 (1996). 25. Feron, O. Targeting the tumor vascular compartment to improve conventional cancer therapy. Trends Pharmacol. Sci. 25, 536–42 (2004). 26. Thomsen, L.L. et al. Nitric oxide synthase activity in human gynecological cancer. Cancer Res. 54, 1352–4 (1994). 27. Thomsen, L.L. et al. Nitric oxide synthase activity in human breast cancer. Br. J. Cancer 72, 41–4 (1995). 28. Ambs, S. et al. Frequent nitric oxide synthase-2 expression in human colon adenomas: implication for tumor angiogenesis and colon cancer progression. Cancer Res. 58, 334–41 (1998). 29. Ambs, S. et al. Relationship between p53 mutations and inducible nitric oxide synthase expression in human colorectal cancer. J. Natl. Cancer. Inst. 91, 86–8 (1999). 30. Tschugguel, W. et al. Inducible nitric oxide synthase (iNOS) expression may predict distant metastasis in human melanoma. Br. J. Cancer 79, 1609–12 (1999). 31. Chen, T. and Stoner, G.D. Inducible nitric oxide synthase expression in Nnitrosomethylbenzylamine (NMBA)-induced rat esophageal tumorigenesis. Mol. Carcinog. 40, 232–40 (2004). 32. Chen, T., Nines, R.G., Peschke, S.M., Kresty, L.A., and Stoner, G.D. Chemopreventive effects of a selective nitric oxide synthase inhibitor on carcinogen-induced rat esophageal tumorigenesis. Cancer Res. 64, 3714–7 (2004). 33. Takahashi, M., Mutoh, M., Kawamori, T., Sugimura, T., and Wakabayashi, K. Altered expression of beta-catenin, inducible nitric oxide synthase and cyclooxygenase-2 in azoxymethane-induced rat colon carcinogenesis. Carcinogenesis 21, 1319–27 (2000). 34. Komatsu, S., Watanabe, H., Oka, T., Tsuge, H., and Kat, N. Dietary vitamin B6 suppresses colon tumorigenesis, 8-hydroxyguanosine, 4-hydroxynonenal, and inducible nitric oxide synthase protein in azoxymethane-treated mice. J. Nutr. Sci Vitaminol (Tokyo) 48, 65–8 (2002). 35. Murakami, A. et al. Inhibitory effect of citrus nobiletin on phorbol ester-induced skin inflammation, oxidative stress, and tumor promotion in mice. Cancer Res. 60, 5059–66 (2000). 36. Kampa, M. et al. Wine antioxidant polyphenols inhibit the proliferation of human prostate cancer cell lines. Nutr. Cancer 37, 223–33 (2000). 37. Martinez, J. and Moreno, J.J. Effect of resveratrol, a natural polyphenolic compound, on reactive oxygen species and prostaglandin production. Biochem. Pharmacol. 59, 865–70 (2000). 38. Trikha, P., Sharma, N., and Athar, M. Nitroglycerin: an NO donor inhibits TPAmediated tumor promotion in murine skin. Carcinogenesis 22, 1207–11 (2001). 39. Schleiffer, R., Duranton, B., Gosse, F., Bergmann, C., and Raul, F. Nitric oxide synthase inhibition promotes carcinogen-induced preneoplastic changes in the colon of rats. Nitric Oxide 4, 583–9 (2000). 40. Inano, H., Suzuki, K., Ishii-Ohba, H., Ikeda, K., and Wakabayashi, K. Pregnancydependent initiation in tumorigenesis of Wistar rat mammary glands by 60Coirradiation. Carcinogenesis 12, 1085–90 (1991). 41. Inano, H. and Onoda, M. Role of nitric oxide in radiation-induced initiation of mammary tumorigenesis in rats. Nitric Oxide 8, 144–8 (2003).

Nitric Oxide and Tumor Biology

417

42. Voevodskaya, N.V. and Vanin, A.F. Gamma-irradiation potentiates L-argininedependent nitric oxide formation in mice. Biochem. Biophys. Res. Commun. 186, 1423–8 (1992). 43. Onoda, M. and Inano, H. Localization of nitric oxide synthases and nitric oxide production in the rat mammary gland. J. Histochem. Cytochem. 46, 1269–78 (1998). 44. Ibuki, Y. and Goto, R. Enhancement of NO production from resident peritoneal macrophages by in vitro gamma-irradiation and its relationship to reactive oxygen intermediates. Free Radic. Biol. Med. 22, 1029–35 (1997). 45. Kisley, L.R. et al. Genetic ablation of inducible nitric oxide synthase decreases mouse lung tumorigenesis. Cancer Res. 62, 6850–6 (2002). 46. Ellies, L.G. et al. Mammary tumor latency is increased in mice lacking the inducible nitric oxide synthase. Int. J. Cancer 106, 1–7 (2003). 47. Ahn, B. and Ohshima, H. Suppression of intestinal polyposis in Apc(Min/+) mice by inhibiting nitric oxide production. Cancer Res. 61, 8357–60 (2001). 48. Scott, D.J. et al. Lack of inducible nitric oxide synthase promotes intestinal tumorigenesis in the Apc(Min/+) mouse. Gastroenterology 121, 889–99 (2001). 49. Ziesche, R. et al. Lipopolysaccharide and interleukin 1 augment the effects of hypoxia and inflammation in human pulmonary arterial tissue. Proc. Natl. Acad. Sci. USA 93, 12478–83 (1996). 50. Liao, J.K. et al. Regulation of bovine endothelial constitutive nitric oxide synthase by oxygen. J. Clin. Invest. 96, 2661–6 (1995). 51. Arnet, U.A., McMillan, A., Dinerman, J.L., Ballermann, B., and Lowenstein, C.J. Regulation of endothelial nitric-oxide synthase during hypoxia. J. Biol. Chem. 271, 15069–73 (1996). 52. Phelan, M.W. and Faller, D.V. Hypoxia decreases constitutive nitric oxide synthase transcript and protein in cultured endothelial cells. J. Cell Physiol. 167, 469–76 (1996). 53. Searles, C.D., Miwa, Y., Harrison, D.G., and Ramasamy, S. Posttranscriptional regulation of endothelial nitric oxide synthase during cell growth. Circ Res. 85, 588–95 (1999). 54. Liu, S.F., Adcock, I.M., Old, R.W., Barnes, P.J., and Evans, T.W. Differential regulation of the constitutive and inducible nitric oxide synthase mRNA by lipopolysaccharide treatment in vivo in the rat. Crit. Care Med. 24, 1219–25 (1996). 55. Kaku, Y. et al. Differential induction of constitutive and inducible nitric oxide synthases by distinct inflammatory stimuli in bovine aortic endothelial cells. Biochim. Biophys. Acta 1356, 43–52 (1997). 56. Inoue, N. et al. Molecular regulation of the bovine endothelial cell nitric oxide synthase by transforming growth factor-beta 1. Arterioscler. Thromb. Vasc. Biol. 15, 1255–61 (1995). 57. Kostyk, S.K. et al. Basic fibroblast growth factor increases nitric oxide synthase production in bovine endothelial cells. Am. J. Physiol. 269, H1583–9 (1995). 58. Kroll, J. and Waltenberger, J. VEGF-A induces expression of eNOS and iNOS in endothelial cells via VEGF receptor-2 (KDR). Biochem. Biophys. Res. Commun. 252, 743–6 (1998). 59. Brouet, A., Sonveaux, P., Dessy, C., Balligand, J.L., and Feron, O. Hsp90 ensures the transition from the early Ca2+-dependent to the late phosphorylation-dependent activation of the endothelial nitric-oxide synthase in vascular endothelial growth factor-exposed endothelial cells. J. Biol. Chem. 276, 32663–9 (2001).

418

Nitric Oxide, Cell Signaling, and Gene Expression

60. He, H. et al. Vascular endothelial growth factor signals endothelial cell production of nitric oxide and prostacyclin through flk-1/KDR activation of c-Src. J. Biol. Chem. 274, 25130–5 (1999). 61. Michell, B.J. et al. The Akt kinase signals directly to endothelial nitric oxide synthase. Curr. Biol. 9, 845–8 (1999). 62. Kupatt, C., Dessy, C., Hinkel, R., Raake, P., Daneau, G., Bouzin, C., Boekstegers, P., and Feron O. Heat shock protein 90 transfection reduces ischemia-reperfusioninduced myocardial dysfunction via reciprocal endothelial NO synthase serine 1177 phosphorylation and threonine 495 dephosphorylation. Arterioscler. Thromb. Vasc. Biol. 24, 1435–41 (2004). 63. Murata, T., Sato, K., Hori, M., Ozaki, H., and Karaki, H. Decreased endothelial nitric-oxide synthase (eNOS) activity resulting from abnormal interaction between eNOS and its regulatory proteins in hypoxia-induced pulmonary hypertension. J. Biol. Chem. 277, 44085–92 (2002). 64. Arena, S. et al. Nitric oxide production stimulated by the basic fibroblast growth factor requires the synthesis of ceramide. Ann. NY Acad. Sci. 973, 94–104 (2002). 65. Barsacchi, R. et al. Activation of endothelial nitric-oxide synthase by tumor necrosis factor-alpha: a novel pathway involving sequential activation of neutral sphingomyelinase, phosphatidylinositol-3' kinase, and Akt. Mol. Pharmacol. 63, 886–95 (2003). 66. Carmeliet, P. Mechanisms of angiogenesis and arteriogenesis. Nat. Med. 6, 389–95 (2000). 67. Six, I., Kureishi, Y., Luo, Z., and Walsh, K. Akt signaling mediates VEGF/VPF vascular permeability in vivo. FEBS Lett. 532, 67–9 (2002). 68. Albina, J.E., Mills, C.D., Henry, W.L., Jr., and Caldwell, M.D. Temporal expression of different pathways of 1-arginine metabolism in healing wounds. J. Immunol. 144, 3877–80 (1990). 69. Ziche, M. et al. Nitric oxide mediates angiogenesis in vivo and endothelial cell growth and migration in vitro promoted by substance P. J. Clin. Invest. 94, 2036–44 (1994). 70. Kirk, S.J. et al. Arginine stimulates wound healing and immune function in elderly human beings. Surgery 114, 155–9; discussion 160 (1993). 71. Lee, P.C. et al. Impaired wound healing and angiogenesis in eNOS-deficient mice. Am. J. Physiol. 277, H1600–8 (1999). 72. Papapetropoulos, A., Garcia-Cardena, G., Madri, J.A., and Sessa, W.C. Nitric oxide production contributes to the angiogenic properties of vascular endothelial growth factor in human endothelial cells. J. Clin. Invest. 100, 3131–9 (1997). 73. Ziche, M. et al. Nitric oxide synthase lies downstream from vascular endothelial growth factor-induced but not basic fibroblast growth factor-induced angiogenesis. J. Clin. Invest. 99, 2625–34 (1997). 74. Murohara, T. et al. Nitric oxide synthase modulates angiogenesis in response to tissue ischemia. J. Clin. Invest. 101, 2567–78 (1998). 75. Fukumura, D. et al. Predominant role of endothelial nitric oxide synthase in vascular endothelial growth factor-induced angiogenesis and vascular permeability. Proc. Natl. Acad. Sci. USA 98, 2604–9 (2001). 76. Yamasaki, K. et al. Reversal of impaired wound repair in iNOS-deficient mice by topical adenoviral-mediated iNOS gene transfer. J. Clin. Invest. 101, 967–71 (1998).

Nitric Oxide and Tumor Biology

419

77. Kimura, H. et al. Hypoxia response element of the human vascular endothelial growth factor gene mediates transcriptional regulation by nitric oxide: control of hypoxia-inducible factor-1 activity by nitric oxide. Blood 95, 189–97 (2000). 78. Jadeski, L.C. and Lala, P.K. Nitric oxide synthase inhibition by N(G)-nitro-Larginine methyl ester inhibits tumor-induced angiogenesis in mammary tumors. Am. J. Pathol. 155, 1381–90 (1999). 79. Dong, Z., Staroselsky, A.H., Qi, X., Xie, K., and Fidler, I.J. Inverse correlation between expression of inducible nitric oxide synthase activity and production of metastasis in K-1735 murine melanoma cells. Cancer Res. 54, 789–93 (1994). 80. Juang, S.H. et al. Suppression of tumorigenicity and metastasis of human renal carcinoma cells by infection with retroviral vectors harboring the murine inducible nitric oxide synthase gene. Hum. Gene Ther. 9, 845–54 (1998). 81. Gratton, J.P. et al. Selective inhibition of tumor microvascular permeability by cavtratin blocks tumor progression in mice. Cancer Cell 4, 31–9 (2003). 82. Sonveaux, P. et al. Irradiation-induced angiogenesis through the up-regulation of the nitric oxide pathway: implications for tumor radiotherapy. Cancer Res. 63, 1012–9 (2003). 83. Folkman, J. and Camphausen, K. Cancer. What does radiotherapy do to endothelial cells? Science 293, 227–8 (2001). 84. d'Uscio, L.V. and Luscher, T.F. Nitric oxide: biology and pathobiology. (Ignarro, L.J., ed.), pp. 609–20 (Academic Press, San Diego, 2000). 85. Andrade, S.P., Hart, I.R., and Piper, P.J. Inhibitors of nitric oxide synthase selectively reduce flow in tumor-associated neovasculature. Br. J. Pharmacol. 107, 1092–5 (1992). 86. Meyer, R.E. et al. Nitric oxide synthase inhibition irreversibly decreases perfusion in the R3230Ac rat mammary adenocarcinoma. Br. J. Cancer 71, 1169–74 (1995). 87. Tozer, G.M., Prise, V.E., and Chaplin, D.J. Inhibition of nitric oxide synthase induces a selective reduction in tumor blood flow that is reversible with L-arginine. Cancer Res. 57, 948–55 (1997). 88. Wood, P.J. et al. Induction of hypoxia in experimental murine tumors by the nitric oxide synthase inhibitor, NG-nitro-L-arginine. Cancer Res. 54, 6458–63 (1994). 89. Jordan, B.F. et al. Changes in tumor oxygenation/perfusion induced by the NO donor, isosorbide dinitrate, in comparison with carbogen: monitoring by EPR and MRI. Int. J. Radiat. Oncol. Biol. Phys. 48, 565–70 (2000). 90. Jordan, B.F., Beghein, N., Aubry, M., Gregoire, V., and Gallez, B. Potentiation of radiation-induced regrowth delay by isosorbide dinitrate in FSaII murine tumors. Int. J. Cancer 103, 138–41 (2003). 91. Wood, P.J. et al. Modification of energy metabolism and radiation response of a murine tumour by changes in nitric oxide availability. Biochem. Biophys. Res. Commun. 192, 505–10 (1993). 92. Wood, P.J. et al. Changes in energy metabolism and X-ray sensitivity in murine tumours by the nitric oxide donor SIN-1. Br. J. Cancer Suppl. 27, S177–80 (1996). 93. Shan, S.Q. et al. Effects of diethylamine/nitric oxide on blood perfusion and oxygenation in the R3230Ac mammary carcinoma. Br. J. Cancer 76, 429–37 (1997). 94. Thews, O., Kelleher, D.K., and Vaupel, P. No improvement in perfusion and oxygenation of experimental tumors upon application of vasodilator drugs. Int. J. Oncol. 19, 1243–7 (2001).

420

Nitric Oxide, Cell Signaling, and Gene Expression

95. Feelisch, M., Ostrowski, J., and Noack, E. On the mechanism of NO release from sydnonimines. J. Cardiovasc. Pharmacol. 14 Suppl 11, S13–22 (1989). 96. Stamler, J.S. et al. Blood flow regulation by S-nitrosohemoglobin in the physiological oxygen gradient. Science 276, 2034–7 (1997). 97. Dewhirst, M.W. et al. Quantification of longitudinal tissue pO2 gradients in window chamber tumours: impact on tumour hypoxia. Br. J. Cancer 79, 1717–22 (1999). 98. Mitchell, J.B. et al. Hypoxic mammalian cell radiosensitization by nitric oxide. Cancer Res. 53, 5845–8 (1993). 99. Mitchell, J.B. et al. Radiation sensitisation by nitric oxide releasing agents. Br. J. Cancer Suppl. 27, S181–4 (1996). 100. Jordan, B.F. et al. Insulin increases the sensitivity of tumors to irradiation: involvement of an increase in tumor oxygenation mediated by a nitric oxide-dependent decrease of the tumor cells oxygen consumption. Cancer Res. 62, 3555–61 (2002). 101. Drummond, G.R., Cai, H., Davis, M.E., Ramasamy, S., and Harrison, D.G. Transcriptional and posttranscriptional regulation of endothelial nitric oxide synthase expression by hydrogen peroxide. Circ. Res. 86, 347–54 (2000). 102. Thomas, S.R., Chen, K., and Keaney, J.F., Jr. Hydrogen peroxide activates endothelial nitric-oxide synthase through coordinated phosphorylation and dephosphorylation via a phosphoinositide 3-kinase-dependent signaling pathway. J. Biol. Chem. 277, 6017–24 (2002). 103. Sonveaux, P. et al. Endothelin-1 is a critical mediator of myogenic tone in tumor arterioles: implications for cancer treatment. Cancer Res. 64, 3209–14 (2004). 104. Sonveaux, P. et al. Modulation of the tumor vasculature functionality by ionizing radiation accounts for tumor radiosensitization and promotes gene delivery. FASEB J. 16, 1979–81 (2002). 105. Cai, H. and Harrison, D.G. Endothelial dysfunction in cardiovascular diseases: the role of oxidant stress. Circ Res. 87, 840–4 (2000).

Index NUMERICS 2-deoxy-D-glucose-6-phosphate 149 2-deoxyglucose 149–151 2-oxoglutarate 296, 302 2,3-dimethoxy-1,4-naphthoquinone 301 3-morpholinosydnonimine 246, 409 3-nitrotyrosine 104, 247 4-hydroxynonenal 248 5’-AMP-activated protein kinase 150–151 5-aminoimidazole-4-carboxamide ribonucleoside 151 5-aminolevulinate synthase 374–375 6-phosphofructo-2-kinase 157–159 7-nitroindazole 3, 14 8-(2-chlorophenylthio)-cGMP 185 8-hydroxydeoxyguanosine 89 13-hydroxyoctadecanoic acid 185

A acetylcholine 189 aconitase 109, 111, 380 acrolein 248 activator protein 1 (AP-1) 337–338 acylation 2 adenine nucleotide translocase (ANT) 110 adenomas 402 adenylyl cyclases 172 adrenomodullin 335 aging 237–238 AhR nuclear translocator (ARNT 295 alanine 276 alanines 385 alpha-tocopherol acetate 89 Alzheimer’s disease 35, 155 aminoguanidine 59 5-aminoimidazole-4-carboxamide ribonucleoside 151 5-aminolevulinate synthase 374–375 5’-AMP-activated protein kinase (AMPK) 150–151 AMP/PKA pathway 338 amyotrophic lateral sclerosis (ALS) 119, 168 angiotensin II 31, 338 ANP receptor gene 355

antimycin 46 antioxidants 89, 110–114 antipsychotics 34 anti-tumor therapy 409–412 AP-1 gene 355 Apaf-1 adaptor protein 220 apoptosis 36–38 and bioavailability of nitric oxide (NO) 237–238 calcium ions in 86 definition of 232 Fas-induced 219–221 mediators of 110 modulation of caspases in 234–235 modulation of GTPases and kinases in 235–236 NO-cGMP signaling pathway 191–192 pathways 59–60 role of peroxynitrite in 109–110 apoptosis signal-regulating kinase 1 (ASK1) 60 apoptosome 220 arginine 2, 58, 101 aryl hydrocarbon receptor (AhR) 295 ascorbates 89, 114 asthma 324–325 astrocytes 148 glucose uptake by 149–151 glycolysis in 157–159 mitochondial network in 4 protection from cell death 156–157 ATP synthase 107–108 autoinhibition 182 autophosphorylation 182, 253 azide 171 azoxymethane 400

B basic fibroblast growth factor (bFGF) 312, 402 Bcl2 gene 355 Bcl-2 proteins 59–60 bimetallic copper 5 binuclear center 5 binuclear sites 7 bioactivity 323 bioenergetics 62–64, 83–84

421

422

Nitric Oxide, Cell Signaling, and Gene Expression

biogenesis 62–64 bladder cancer 58–59, 399 bone homeostasis 190 breast cancer 59

C C3635 cysteine residue 223 calcineurin-NFAT pathway 339 calcium ions 169–170 homeostasis 186 in mitochondrial apoptosis 86 reduction of 183–184 calcium overload 110 calmodulin 30, 169–170, 185 cAMP response elements (CREs) 191 cancer 396–398 carbon dioxide 102, 246 carbon monoxide 301 carbonate radical 103 carbonate radical anion 277 cardiac hypertrophy 339 cardiolipin 60 cardiomyopathy 108 carnitine palmitoyltransferase I 109, 112, 118 caspase-3 zymogens 220 caspases 219–220 activity of 237–238 inhibition of 110 modulation of 234–235 catabolite activator protein (CAP) 175 catalase 49–50 catalytic redox cycle 8 catechol 2,3-dioxygenase 299 caveolin 170 caveolin-1 355 CD95 cell line 249 cell cycle 53–54 cell death 156–157, 254–255 cell events 62–64 cell proliferation 51–53, 57–58 cell respiration 3–4 ceramide 312–316 activation of eNOS by 312–313 crosstalk 314–316 regulation of 313–314 cerebellum 35 ceruloplasmin 322 cervical cancer 399 C-fos gene 335 cGMP 332–334 crosstalk with calcineurin NF/AT pathways 339

effects on expression of pro-inflammatory genes 339 effects on mRNA stability 340 gene expression 339–340 gene transcription factor 337–338 interference with cAMP/PKA pathway 338 interference with MAP kinase pathways 338 transcription factors 334–337 See also NO-cGMP signaling pathway chaperone proteins 323 chemokine MIP-2 342 chemoprotective agents 56 chemotherapeutic drugs 412 Chernobyl accident 58 chloramphenicol 62 8-(2-chlorophenylthio)-cGMP 185 chlorpromazine 34 cholangiocarcinoma 399 citrulline 2, 82, 169, 293 C-linker 175 collagen 335 collagen prolyl hydroxylases 296 colon cancer 399 colorectal cancer 398 complementary deoxyribonucleic acid (cDNA) 51 COX-2 gene 335, 355 creatine kinase 111 CREB transcription factor 191 Crohn’s disease 398 C-terminal transactivation domain (CTAD) 296 cyclic nucleotide binding domain (CNBD) 175 cyclic nucleotide-gated (CNG) channels 174 expression and composition 174 functions 175–176 pathologies 176 regulation of 176–177 structure and regulation of 174–175 cyclic nucleotides 177–182 cyclin-dependent kinases 57 cyclins 57, 355 cyclooxygenase 185 cyclophylin D 110 cycloskeletal-associated proteins 187 cysteine 267, 277, 357 cystic fibrosis 325 cystic fibrosis transmembrane regulatory protein (CFTR) 325–326 cytochrome c oxidase (CcOX) 4–8 active site of 6 inhibition of 8–9, 341 inrteraction with nitric oxide (NO) 356 nitrite-inhibited 9–10 nitrosyl- and nitrite derivatives of 11

Index

423

nitrosyl-inhibited 10–12 reactions with nitric oxide (NO) 12–17 cytochrome oxidase 31 cytosol 117 cytosolic kinases 110 cytosolic nitric oxide (NO) 47

D dehydrocysteine 376 denitrosation 323 denitrosylation 219 2-deoxyglucose 149–151 deoxyhemoglobin 172 deoxymyoglobin 172 desferal 385 desferrioxamine 296 Dexras 222 diabetes 115–117 diacylglycerol 175 dichlorodihydrofluorescein 247 diethylstilbestrol 401 diffusion coefficient 47 2,3-dimethoxy-1,4-naphthoquinone 301 dimethyloxalglycine 302 dinitrosyl-iron complex (DNIC) 382 dithiothreitol 234, 323 Drosophila melanogaster 374 dystrophin 189

E egr-1 gene 335 electron transfer chain 45–46, 62 endoplasmic reticulum nitric oxide synthase (erNOS) 37 endothelial nitric oxide synthase (eNOS) 2 activation by ceramide 312–313 and tumor angionesis 402–403 See also nitric oxide synthases endothelium 3 endothelium-derived relaxation factor (EDRF) 78, 168, 292 endotoxemia 47, 117 eosinophil peroxidase 358 epicatechin 248 epidermal growth factor (EGF) 252 epidermal growth factor receptor (EGFR) 252–253 ER-1 cell line 53 erectile function 190

erythroid 5-aminolevulinate synthase (eALAS) 374–375 erythropoietin 296, 324, 335 Escherichia coli 5 esophageal squamous cell carcinoma 400 estrogen receptors 59 ET-1 gene 355 eukaryotes 4 eukaryotic cells 64 excitotoxicity 119 extracellular signal-related kinases (ERKs) 54–56, 252–253

F factor inhibiting HIF-1 (FIH) 296 Fas proteins 219–221 fenton chemistry 106 ferritin 106, 372, 385–386 ferrochelatase 380 [Fe-S] clusters 379–380 fibrinogen 186 fibroblasts 51 fibronectin 335 fibrosarcoma 409 flavanol (-)-epicatechin 247 flavine adenine dinucleotide (FAD) 169 flavohemoglobin 380 FLT-1 gene 335 fluorescence 3 fluorescence microscopy 14 FMR1 gene 335 formaldehyde dehydrogenase 219 fragile X mental retardation gene (FMR1) 344 free radicals 265 in guanine nucleotide exchange (GNE) 283 protein target sites of 265 redox potentials of 265 fructose-1,6-bisphosphate 157 fructose-2,6-bisphosphate 157–158 fructose-6-phosphate (F6P) 157

G gall bladder cancer 399 gamma radiation 401 gangliosides 312 gap-junctional intercellular communication (GJC) 253 gastric cancer 399 gene expression 331–344 biochemical mechanisms 356–360

424

Nitric Oxide, Cell Signaling, and Gene Expression

gene expression (continued) calcineurin-NFAT pathway 339 cAMP/PKA pathway 338 cGMP signaling genes 339–340 hypoxia-mediated 341–342 MAP kinase pathways 338 nitric oxide (NO)-mediated 332–340 post-transcriptional control of 372–379 pro-inflammatory genes 339 role of guanylyl cyclase in 332–340 role of nitric oxide (NO) in 341–342 signaling pathways 354–356 transcription factors 334–337 gene regulation 191 glial cells 147 glucocorticoid receptor 336 glucose metabolism 146 carriers in neural cells 148–149 glutamate-induced neurotoxicity 154–155 glycolysis in 155–156 nitric oxide (NO) regulation of 149–151 nitric oxide formation in neural cells 146–147 pentose-phosphate pathway 152 role of astrocyes in 148 role of glutathione in 153–154 glucose transporters 146, 148–149 glucose-6-phosphate dehydrogenase (G6PD) 152–153 glutamate dehydrogenase 109 glutamate-induced neurotoxicity 154–155 glutamate-oxaloacetate transaminase 112 glutamyl transpeptidase 323 glutathione 89, 113–114, 153–154, 358 glutathione disulfide 114 glutathione peroxidase 49–50, 114 glutathionyl radicals 114 glyceraldehyde 3-phosphate dehydrogenase (GAPDH) 155–156, 247, 249 glycogen 148 glycolysis 155–156 growth arrest 254–255 GTPases 235–236 activating proteins 266 NKCD-containing 279–280 NKSD-containing 280–281 5-guanidino-4-nitroimidazole diphosphate 277 guanidino group 101 guanine nucleotide exchange (GNE) 264 characterization of 273–274 free radical-mediated- 283 hydroxyl radical-mediated 280–281 NKCD-containing GTPases 279–280 NKSD-containing GTPases 280–281

Rab GTPases 281–283 Rho GTPases 281–283 structural and mechanistic basis of 275–279 guanine nucleotide exchange factors (GEFs) 264, 266, 284 guanine nucleotide inhibitors (GDIs) 266 guanylate cyclase (GC) 222 guanylyl cyclase 332–340

H half-reduced binuclear sites 7 haloperidol 34 Harber-Weiss reaction 265 heart mitochondrial nitric oxide synthases (mtNOS) 84–86 heat shock protein 20 (HSP20) 185 heat shock protein 27 (HSP27) 187 HeLa cell cloneS 312 heme 5, 172–173 heme domain 169 heme oxigenase 335 heme-copper oxidases 5, 18 hemochromatosis 377–378, 399 hemoglobin 323, 372, 410 hemoproteins 372 hepatocytes 51 hepatoma 399 heterodimers 171–172 hexokinases 149 hippocampal neurons 34 hippocampus 35, 176 homodimers 172 human venous endothelial cells (HUVEC) 405 HuR gene 335 hydrogen peroxide 49–50 and apoptosis 59–60 cell cycle arrest by 53–54 effects on cyclins 57 formation of 265 intracellular source of 100 proliferating effects of 51–53 See also oxygen active species 8-hydroxydeoxyguanosine 89 hydroxyl group 265 hydroxyl radicals 103, 265, 280–281 hydroxylamine 171 hydroxymethylglutaryl-CoA synthase 112 4-hydroxynonenal 248 13-hydroxyoctadecanoic acid 185 hyperglycemia 115–116 hypertension 89–91

Index

425

hypoxanthine phosphoribosyltransferase (HPRT) 344 hypoxia 294–297 and gene expression 341–342 role of mtNOS in 88–89 in tumor microenvironment 398 hypoxia inducible factor-1 322–325 activation of 298–300 destabilization of 301–303 redox sensitivity of 292 stabilization of 298–300 synthesis vs. stability regulation of 300 in tumor biology 303–304 hypoxia responsive element (HRE) 296 hypoxic-mimetics 296

I ICAM-1 gene 335 IDN-6556 pancaspase inhibitor 232 inducible nitric oxide synthases (iNOSs) 2 and cell proliferation 57–58 and tumors 58–59 See also nitric oxide synthases inositol triphosphate 323 insulin-like growth factor (IGF-1) 405 intestinal motility 192 ion channels 333 ionizing radiation 412 IP-10 gene 355 iron regulatory proteins (IRPs) 374–376 impact on ferritin and tranferrin receptor expression 385–386 in iron-associated disorders 377–379 modulation of [Fe-S] clusters by nitric oxide (NO) 379–380 modulation of IRP2 384–385 post-translational regulation of 380–381 regulation of IRP1 protein levels 383–384 RNA-binding activities of 381–382 iron responsive elements (IREs) 373–374 ischaemia 147–148 isocitrate dehydrogenase 109 isopenicillin N synthase 299

J JNK activation 56, 235–236 Jurkat T-cells 39, 327

K ketoglutarate dehydrogenase 109 kidney 189–190 kinases 235–236

L lactates 148 laminin 336 L-arginine 58 L-citrulline 169 leukemia 314 leukocytes 327 Lewis acid 103 lipid peroxyl radicals 114 lipoperoxidation 37, 89 lipopolysaccharides 147, 313, 396 lipoxygenase-1 247 liver cancer 399 L-nitrosoarginine 2 long-term depression (LTD) 188 long-term memory (LTM) 34 long-term potentiation (LTP) 34, 188 low-density lipoprotein (LDL) 237 lung cancer 399 lymphocytes 220, 313 lypooxygenase 185

M macrophages 3, 400 magnesium 85–86 malate dehydrogenase 111 malondialdehyde 248 mammary glands 401–402 mammary tumors 401 manganese porphyrins 121 MAP kinase pathways 338 matrix components 109 MCP-1 gene 336 M-CSF gene 336 MDA-MB-231 cell line 58 melanoma 399 meothelioma 399 messenger RNA (mRNA) 332 cis-acting elements 373 effects of cGMP on 340 and gene expression 354–356 metalloproteins 103 metallothionein 358 methionine adenosyltransferase (MAT) 218

426

Nitric Oxide, Cell Signaling, and Gene Expression

microglia 146 microglobulin 378 microsomes 49 mitochondria 45–46 bioenergetics 83–84 biogenesis 62–64 calcium ion homeostasis 84 effects of nitric oxide 30–34 formation of peroxynitrite in 101–102 functions of nitric oxide (NO) for 79–81 inhibition of cell respiration in 3–4 nitrated proteins 111–113 persistence of nitric oxide (NO) in 20–22 pharmacology 121–122 production of oxygen active species 46–48 structure and functions 80 tyrosine nitration in 103–107 mitochondrial creatine kinase (Mt-CK) 108–109 mitochondrial dysfunction 100 in diabetes 115–117 in neurodegenerative diseases 119–120 in sepsis 117–118 mitochondrial nitric oxide synthases (mtNOS) 81–82 calcium ion dependence of 82–83 effects on mitochondrial bioenergetics 83–84 existence of 2 heart 84–86 and hypoxia/reoxygenation 88–89 inhibition of 34 and life processes 61–62 protein expression 35 regulatory activity of 33 role in hypertension 89–91 See also nitric oxide synthases mitochondrial superoxide dismutase (Mn-SOD) 46 peroxynitrite reactions with 110–113 transfection of 62 mitogen-activated protein kinases (MAPKs) 39, 252, 354 MKP-1 gene 336 MMP-13 gene 355 MMP-2 gene 336, 355 MMP-9 gene 336, 355 3-morpholinosydnonimine 246, 409 multiple sclerosis 119 murine pulmonary hypertension 324 myeloperoxidase 247, 358 myocarditis 118 myoglobin 19, 372 myosin light chain kinase (MLCK) 184–185

myosin light chain phosphatase (MLCP) 184–185 myosin-binding subunit (MBS 184 myristoylation 170

N N-acetyl-cysteine (NAC) 324 naphthoquinone 250 natriuretic peptides 333 neuroblastoma cells 14 neurodegenerative diseases 119–120 neuronal nitric oxide synthases (nNOSs) 2 neuronal plasticity 34–35 neurons 3 mitochondrial network in 4 nitric oxide (NO) formation in 146–147 neuroprotection 147, 153–154 neurotoxicity 147, 154–155 neurotransmission 146 NO-cGMP signaling 187–189 perception of pain in 188 role of nitric oxide (NO) in 189 synaptic plasticity 188 NF-kB activation 56 NFkB transcription factor 326–327 nicotinamide adenine dinucleotide phosphate (NADPH) 169 NIH 3T3 cell line 53 nitrergic nerves 189 nitric oxide (NO) 2 and redox homeostasis 232–234 in anti-tumor therapy 409–412 apoptosis 36–38, 59–60 bioavailability of 237–238 crosstalk with ceramide 312–316 diffusion coefficient 47 donors 409–411 endogenous production of 411–412 formation in neural cells 146–147 functions for mitochondria 79–81 and gene expression 331–344 and glycolysis 155–156 inhibition of cell respiration by 3–4 and iron regulatory proteins (IRPs) 379–386 isoforms 29–30 light-induced dissociation of 12 mitochondrial production of 2, 30–34 modulation of caspases by 234–235 modulation of cell proliferation 57–58 neuronal plasticity 34–35 neurotoxic vs. neuroprotectieve roles 147 persistence in mitochondrion 20–22

Index as pro-angiogenic factor 405–407 pro-apoptopic effects of 236–237 reactions with cytochrome c oxidase (CcOX) 12–17 redox signaling 342–344 regulatory capacity of 33 role in neurotransmission 189 signaling pathways 354–356 steady-state concentration of oxygen active species 49–50 synthesis of 169 and tumor angiogenesis 402–407 and tumor vascular reactivity 407–408 and tumorigenesis 398–402 utilization pathways 46–48 nitric oxide synthases 2–3 calcium ion regulation of 169–170 functions of 169 inhibitors 117 isoforms 78–79 sub-cellular localization of 170–171 and tumors 58–59 nitrites 2, 105–106 nitrogen dioxide 105 nitroglycerin 171, 400 7-nitroindazole 3, 14 nitroprusside 171 nitrosation 323 nitrosoarginine 2 nitrosoglutathione 298, 322–323 nitrosohemoglobin. 410 nitrosothiols 89 nitrosylation 217–218 N-methyl-D-aspartate receptor signaling 221–222 redox-active GTPase 268–273 role in gene expression 357–358 ryanodine receptors 222–224 specificity of 218–219 thioredoxin signaling 224–225 nitrothiols 321–327 hypoxia inducible factor 1 (HIF 1) 322–325 NFkB transcriptin factor 326–327 regulation of 224–225 specificity factors 325–326 3-nitrotyrosine 118, 120, 104, 247 nitrovasodilator 400 nitroxyl 21 N-methyl-D-aspartate 380 N-methyl-D-aspartate (NMDA) 4, 221–222 N-nitrosomethylbenzamidine (NMBA) 400 nobiletin 400 NO-cGMP signaling pathway 169–183 and apoptosis 191–192

427 in bone homeostasis 190 cyclic nucleotide-gated (CNG) channels 174–177 cyclic nucleotides 177–182 and gene regulation 191 and intestinal motility 192 and kidney function 189–190 and neurotransmission 187–189 nitric oxide (NO) synthases 169–171 and platelet aggregation 185–187 in reproduction 190 self-regulatory interactions in 192–193 soluble guanylyl cyclase (sGC) 171–173 and vascular relaxation 183–185 non-steroidal anti-inflammatory agents (NSAIDs) 56 normoxia 322, 327 NOS-2 gene 355 Nox1 cell line 53 nuclear location sequence (NLS) 191

O oesophagal cancer 399 olfactory channels 175 olfactory sensory neurons (OSN) 177 oligodendrocytes 146 oligomycin 157 oncocytoma 62 osteoblasts 51 osteoclasts 190 oxidative stress 246 antioxidants 114 and cell proliferation 51–53 JNK activation 56 NF-kB activation 56 p38 MAPK cascade 54–56 oxo-ferryl species 106 2-oxoglutarate 296, 302 oxo-metals 103 oxygen active species 46–48 apoptosis 59–60 cell cycle arrest by 53–54 effects on cyclins 57 proliferating effects of 51–53 steady-state concentration 49–50 See also hydrogen peroxide oxygenase domain 169 oxygen-dependent degradation domain (ODD) 295

428

Nitric Oxide, Cell Signaling, and Gene Expression

P p11 gene 355 p16 gene 355 p21 gene 355 p38 MAPK 54–56 p53 protein 59 PAI-1 gene 336 pain, perception of 188 palmetoylation 170 pancaspase inhibitors 232, 235 Paracoccus denitrificans 5, 13 Parkinson’s disease 35, 120 PDE1 enzymes 178–179 PDE10 enzymes 181–182 PDE11A enzymes 182 PDE2 enzymes 179–180 PDE3 enzymes 180 PDE5 enzymes 180–181 PDE6 enzymes 181 PDE9 enzymes 181 penile erection 190 pentose-phosphate pathway 152 permeability transition pore (PTP) 91, 109 peroxidase 105–106 peroxides 105–106 peroxiredoxin 111, 121 peroxynitrite 101 ATP synthase 107–108 electron transport chain components 107–108 formation of 101–102 in diabetes 115–116 in neurodegenerative diseases 119–120 in sepsis 117–118 intermembrane components 108–109 matrix components 109 protein nitration with 7 reactions with apoptopic machinery 109–110 reactions with mitchondrial antioxidants 110–114 reactivity 102–103 signaling effects of 245–255 adaptive responses 252–254 cell death 254–255 cellullar reactions 251–255 growth arrest 254–255 intracellular targets 246–247 mechanisms 248–251 membrane permeability 246–247 observed biological effect 247 tyrosine phosphatases 250–251 tyrosine nitration 103–105

PGC1 gene 355 PH domain-containing enzymes (PHD) 296 pharmacology 121–122 phenyl N-tert-butylnitrone (PBN) 268 phenylalanine 277 phorbol esters 400 phorbol myristate acetate (PMA) 405 phosphatidyl inositol kinase (PI-kinase) 183 phosphatidyl inositol-4 phosphate 183 phosphatidylinositol 3’ kinase (PI3K)/Akt pathway 313 phosphatidylinositol 3-kinase (PI3-K) 266 phosphodiesterases (PDEs) 177–182 cGMP-regulated 333 PDE1 enzymes 178–179 PDE10 enzymes 181–182 PDE11 enzymes 182 PDE2 enzymes 179–180 PDE3 enzymes 180 PDE5 enzymes 180–181 PDE6 enzymes 181 PDE9 enzymes 181 6-phosphofructo-2-kinase 157–159 phosphoglycerate kinase (PGK) 155 phosphoinositides 253 phospholamban 184 phospholipase 186 phospholipid hydroperoxide glutathione peroxidas 110 phosphorylation 2, 57, 184–185, 249–250 photoreceptors 176 PKG gene 355 plasma 3-nitrotyrosine 116 plasminogen activator inhibitor 1 (PAI-1) 191 plasticity 34–35 platelet aggregation 185–187 calcium ion homeostasis 186 cycloskeletal-associated proteins 187 surface receptors 186–187 platelet-derived growth factor (PDGF) 253–254, 405 P-loop 175 poly(ADP-ribose) synthetase (PARS) 147 polyethylene glycol-SOD 117 porphyrins 121 post-synaptic density (PSD) protein 312 potassium channel 175, 184 pregnancy 190 pro-inflammatory genes 339 prostacyclin 185 protein kinase C (PKC) 186 protein tyrophosphatases (PTPases) 250–251 protocatechuate 4,5-dioxygenase 299 P-selectin 186

Index

429

Pseudomonas stutzeri 18 Purkinje cells 188

Q QM7 cell line 62 quinols 4

R Rab GTPases 267–268 radiation 401 radical spin-trap agents 268 radiotherapy 398 Raf kinase 266 rapamycin 300 Ras proteins 222 Ras superfamily GTPases 265–268 free radicals 265 guanine nucleotide exchange (GNE) 273–274 guanine nucleotide exchange factors (GEFs) 284 nitrosylation 268–273 overview 264 Rab GTPases 267–268 Rho GTPases 267–268 See also GTPases raxofelast 89 reactive nitrogen intermediates (RNIs) 293–294 transcription factors 295 in tumor biology 303–304 reactive nitrogen species (RNS) 357 reactive oxygen species (ROS) 265, 411–412 receptor guanylyl cyclase (RGC) 333 redox potentials 265 reductase domain 169 renal cell carcinoma 59 renal function] 189–190 Renilla luciferase 302 renin 189–190 reoxygenation 88–89 respirometry 14 resveratrol 400 retinitis pigmentosa 176 retinopathy 117 rho gene 355 Rho GTPases 267–268, 281–283 rhodamine 3, 14 Rhodobacter sphaeroides 5 rotenone 63 ryanodine receptors 222–224

S sarcoplasmic reticulum 184 selectin 335, 355 selenocysteine 114 semiquinone 48 sepsis 117–118 serine 184 serine-threonine kinases 57 serum response factor (SRF) 191 sexual arousal 190 shear stress 235 sildenafil 190 skeletal muscle 170 S-nitrosoglutathione {GSNO) 322–323 S-nitroso-N-acetyl cysteine (SNOAC) 324 S-nitroso-N-acetylpenicillamine (SNAP) 81 S-nitrothiols (SNOs) 321–327 hypoxia inducible factor 1 (HIF 1) 322–325 NFkB transcriptin factor 326–327 regulation of 224–225 specificity factors 325–326 sodium nitroprusside 300, 405 soluble guanylate cyclase (sGC) 356 soluble guanylyl cyclase (sGC) 171–173 activation of 333 role of heme in function of 172–173 structural functional organization of 171–172 sub-cellular localization of 173 SPARC gene 336 spatial learning 34–35 specificity factors 325–326 sphingolipids 312 sphingomyelinases (SMases] 313–314 sphingosine 1 phosphate (S1P) 313 sPLA2 gene 336 spontaneously hypertensive rats (SHR) 91 stroke 119–120 substantia nigra 35, 64 succinyl-CoA:3-oxoacid CoA-transferase (SCOT) 109 sulfenic acids 250 sulfinic acid 251 sulfonic acid 251 superoxide dismutase (SOD) 265 superoxides 265 formation of 101 in guanine nucleotide exchange (GNE) 279–280 reaction with nitric oxide (NO) 358 surface receptors 186–187 synaptic plasticity 188 synaptosomes 3, 13

430

Nitric Oxide, Cell Signaling, and Gene Expression

synthases 2–3 syntrophin 170, 312

T telokin 185 tetrahydrobiopterine 30, 169 TFII-I transcriptional regulator 191 thapsigargin 37–38 Thermus thermophilus 5, 18 thin filaments 185 thiolation 358 thiols 30, 358 thioperoxidase 49 thioredoxin reductase 110, 224–225 thioredoxin signaling 224–225 THP-1 cell line 381 thromboxane TXA2 receptor 186 thymocytes 37 thyroxine 31 tocopherols 114 tocopheroxyl radicals 114 transcription factors 295, 334–337 transducin 175 transferrin receptor 372, 385–386 transnitrosation 358 trifunctional protein 111 Trolox 89 Trypanosoma cruzi 108 tumor 396–413 angionesis 402–407 endothelial nitric oxide synthase (eNOS) expression 402–403 hypoxia inducible factor-1 303–304 microenvironment 396–398 pro-angiogenic factor 405–407 reactive nitrogen intermediates (RNIs) 303–304 treatment of 409–412 vascular reactivity 407–408 tumor necrosis factor (TNF) 225, 312–316 tumorigenesis 398–402 tumors 58–59 tyrosine nitration 115 as biological effect of peroxynitrite 247 fenton chemistry in 106 inhibitors of 248 mechanisms 358–359 nitrite-mediated 105–106 peroxidase-mediated 105–106 peroxide-mediated 105–106 peroxynitrite-mediated 103–105 radical termination reaction in 107

tyrosyl radicals in 106–107 vs. phosphorylation 249–250 tyrosine phosphatases 250–251 tyrosines 5 tyrosyl radicals 104–107

U ubiquinol 21, 32, 46, 48 ubiquinone 81, 114 ubisemiquinone 21, 46, 48, 114 UDP-glucosyltransferase (UDP-GT) 37 ulcerative colitis 399 urate 248 urothelial cell growth 58–59 uterus 190

V V79 cells 56 valine 276 vascular endothelial growth factor (VEGF) 296, 324, 405–406 vascular relaxation 183–185 calcium ion desensitization in 184–185 reduction of intracellular calcium ions 183–184 regulation of thin filament functions in 185 vascular steal 412 vasodilatory-stimulated phosphoprotein (VASP) 185, 187 VCAM-1 gene 337, 343 vitamin B6 400 voltage-dependent anion channel (VDAC) 110 von Hippel Lindau protein (pVHL) 295, 322, 341

W Wilson’s disease 398 working memory (WM) 34–35 wound healing 405

X–Z xanthine 301 xanthine oxidase 117, 265, 266, 283, 301, 337 X-ray radiotherapy (XRT) 407 zinc 358 zymogens 220, 234