Membrane Structure in Disease and Drug Therapy

Membrane Structure in Disease and Drug Therapy

Membrane Structure in Disease and Drug Therapy edited by

Guido Zirnrner Johann Wolfgang Goethe University Frankfurt, Germany

.M. .A. R . C.F I . ~

MARCELDEKKER, INC. D E K K E R

-

NEWYORK BASEL

ISBN: 0-8247-0361-8 This book is printed on acid-free paper. Headquarters Marcel Dekker, Inc. 270 Madison Avenue, New York, NY 10016 tel: 212-696-9000; fax: 212-685-4540 Eastern Hemisphere Distribution Marcel Dekker AG Hutgasse 4, Postfach 812, CH-4001 Basel, Switzerland tel: 41-61-261-8482; fax: 41-61-261-8896 World Wide Web http://www.dekker.com The publisher offers discounts on this book when ordered in bulk quantities. For more information, write to Special Sales/Professional Marketing at the headquarters address above. Copyright  2000 by Marcel Dekker, Inc. All Rights Reserved. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage and retrieval system, without permission in writing from the publisher. Current printing (last digit): 10 9 8 7 6 5 4 3 2 1 PRINTED IN THE UNITED STATES OF AMERICA

Preface

The first book on membranes and disease was published more than 20 years ago (1). Since that time, much has been learned about membrane structure and its wide influence on diseases as well as its potential for therapeutic use. Before turning our attention to pathophysiology (disease state), we first must learn about physiology (healthy state). Moreover, in the disease state, we must consider two issues: the multifold reactions that occur with changes in membrane structure and the complexity of membrane response from structure change due to therapy. In both cases, structural or conformational changes may have consequences or be a consequence of (1) interaction with membranous transport systems (opening or closing of ion or substrate channels); (2) reaction with receptors; (3) activation or inhibition of membrane enzymes; or (4) cytosolic membranous signaling and exchange. These consequences within the membrane influence intracellular wellbeing: life is possible only if a balance between intracellular and extracellular compartments is maintained. However, a new complexity, which was not suspected 20 years ago, has been introduced. It is becoming more and more obvious that comparisons of sequences of proteins and their primary and secondary structures result in a better understanding of correlations, homologies, and therefore multifunctionality (by domains or complexes). Thus, while the previously simple picture becomes more complex it also becomes more complete (e.g., insulin action, glucose-6-phosphatase, and cystic fibrosis). Hopefully, the reader will not view such complexity as a burden, but rather as a challenge. In reviewing the chapters of this book, it was rewarding to see how completely and effectively contributors covered their topics. To maximize this volume’s usefulness to the reader, the Introduction provides a brief overview that highlights important aspects of each chapter. I wish to express my sincere thanks to all contributors for their participation and enthusiasm in making this book a most valuable reference in the vast field of membrane structure. I am also grateful to the staff of Marcel Dekker, Inc.,— iii

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Preface

in particular, Sandra Beberman, Vice President, Dr. Linda Ogden-Wolgemuth, Assistant to Graham Garratt, and Elyce Misher, Production Editor—for their cooperation, and to many others who encouraged me in this endeavor. This book is dedicated to Graham Garratt, former Vice President and Publisher, who continually inspired me. Those who knew him will never forget an exceptional and noble friend. Guido Zimmer

REFERENCE 1. Bolis L, Hoffmann JF, Leaf A, eds. Membranes and Disease. New York: Raven Press, 1976.

Contents

Preface Contributors Introduction

iii ix xv

Part I. Antibacterial Actions 1. Antibacterial and Hemolytic Activity of Amphipathic Helical Peptides Margitta Dathe

1

2. Bacterial Membrane as a Target for a Novel Class of Diastereomers of Cytolytic Peptides Yechiel Shai

27

3. Bee Venom Toxicity: Synergistic Action of Melittin on Interfacial Hydrolysis by Phospholipase A2 Yolanda Cajal and Mahendra Kumar Jain

45

4. Polymyxins: Prototype for a New Class of Antibiotics Mahendra Kumar Jain and Martha Bruch

63

Part II. Dermatology 5. Environmental Effects on Skin Lipids and Impairment of Barrier Function Ju¨rgen Fuchs

77 v

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Contents

Part III. Drug Transporters and Oxidative Stress 6. Oxidative Stress and Loose Coupling/Uncoupling Guido Zimmer 7. Uncouplers of Oxidative Phosphorylation: Activities and Physiological Significance Yasuo Shinohara and Hiroshi Terada 8. Tetraether Lipid Liposomes Hans-Joachim Freisleben 9. Transfection of Eukaryotic Cells with Bipolar Cationic Derivatives of Tetraether Lipid Larissa A. Balakireva and Maxim Yu. Balakirev

95

107 127

153

Part IV. Inflammation and Infection 10. Cholera Toxin Conformational Changes Associated with Changes in Membrane Structure Jameson A. McCann and William D. Picking

167

11. Parasite Mitochondrial Membrane Functions as Targets for Chemotherapy Akhil B. Vaidya, Michael T. McIntosh, and Indresh K. Srivastava

183

12. Hepatic Cell Function in Liver Fluke Infection Linda M. Lenton and Carolyn A. Behm 13. Lipopolysaccharide: A Membrane-Forming and InflammationInducing Bacterial Macromolecule Ulrich Seydel, Artur J. Ulmer, Stefan Uhlig, and Ernst Theodor Rietschel 14. Effect of Cholestasis on Biomembranes Su¨krettin Gu¨ldu¨tuna and Ulrich Leuschner

201

217

253

Part V. Metabolism 15. 3,5-Diiodothyronine Binds to Subunit Va of Cytochrome c Oxidase: Possible Mechanism of Short-Term Effects of Thyroid Hormones Bernhard Kadenbach and Susanne Arnold

271

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16. Regulation of Glucose Transport by Insulin in Muscle and Fat Cells: Translocation and Activation of Glucose Transporters Romel Somwar, Gary Sweeney, Karen Yaworsky, Toolsie Ramlal, Peter Tong, Zayna Khayat, and Amira Klip 17. Glucose-6-Phosphatase: A Member of the Newly Identified HPP Superfamily That Consists of Histidine Phosphatases and Vanadium-Containing Peroxidases and Consequences for Membrane Topology, Active Site, and Reaction Mechanism Wieger Hemrika, Rokus Renirie, and Ron Wever

vii

283

303

Part VI. Neurology 18. Alzheimer’s Amyloid β-Peptide-Associated Oxidative Stress: Brain Membrane Lipid Peroxidation and Protein Oxidation D. Allan Butterfield 19. Membrane Orientation of the Alzheimer’s Disease–Associated Presenilins C. M. A. Boeve, P. Cupers, F. Van Leuven, W. Annaert, and Bart De Strooper

335

353

Part VII. Oncology 20. Membrane Structure Analysis in Apoptotic Processes of Leukemic Blasts and Leukemia-Derived Cell Lines Uwe Ebener, Sibylle Wehner, Christoph Rietschel, Hu¨lya Cakmak, Eckhard Niegemann, and Matthias Eishold 21. Use of Monoclonal Antibodies in Cloning and Identification of Membrane Antigens Marco Bestagno and Oscar R. Burrone

369

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Part VIII. Polycystic Diseases 22. Polycystins: Membrane-Associated Proteins Involved in Autosomal Dominant Polycystic Kidney Disease Katherine W. Klinger and Oxana Ibraghimov-Beskrovnaya

409

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Part IX. Pulmonary Physiology and Diseases 23. Action of β-Agonists Compared to Cromoglycate on Mononuclear Cell Membranes: Stabilizing or Destabilizing? Guido Zimmer, Markus Bernho¨rster, Patrizius Pilz, and Jutta Schuchmann-Fix

427

24. Cystic Fibrosis Transmembrane Conductance Regulator: A Chloride Channel Regulator of Ion Channels 439 Makoto Sugita and J. Kevin Foskett 25. Role of Ca2⫹-Independent Lysosomal Phospholipase A2 in Turnover of Lung Surfactant Phospholipids Aron B. Fisher

461

Part X. Transport Events 26. Role of Sodium Pump in Disease Zhimin (Tim) Cao and Roland Valdes, Jr.

479

Index

513

Contributors

W. Annaert Flanders Interuniversitary Institute, Gasthuisberg, and Catholic University Leuven, Leuven, Belgium Susanne Arnold, Ph.D. Children’s Hospital and Harvard Medical School, Boston, Massachusetts Maxim Yu. Balakirev, Ph.D. Institute of Chemical Kinetics and Combustion, Russian Academy of Science, Novosibirsk, Russia Larissa A. Balakireva, Ph.D. Institute of Cytology and Genetics, Russian Academy of Science, Novosibirsk, Russia Carolyn A. Behm, Ph.D. Division of Biochemistry and Molecular Biology, School of Life Sciences, Australian National University, Canberra, Australia Markus Bernho¨rster Membrane Structure Group, Center for Biological Chemistry, Johann Wolfgang Goethe University, Frankfurt, Germany Marco Bestagno, Ph.D. Department of Molecular Immunology, International Centre for Genetic Engineering and Biotechnology (ICGEB), Trieste, Italy C. M. A. Boeve Flanders Interuniversitary Institute, Gasthuisberg, and Catholic University Leuven, Leuven, Belgium Martha Bruch Department of Chemistry and Biochemistry, University of Delaware, Newark, Delaware Oscar R. Burrone, Ph.D. Department of Molecular Immunology, International Centre for Genetic Engineering and Biotechnology (ICGEB), Trieste, Italy ix

x

Contributors

D. Allan Butterfield, Ph.D. Department of Chemistry and Center of Membrane Sciences, University of Kentucky, Lexington, Kentucky Yolanda Cajal Department of Physiochemistry, University of Barcelona, Barcelona, Spain Hu¨lya Cakmak Department of Hematology and Oncology, Clinic of Pediatrics—III, Johann Wolfgang Goethe University, Frankfurt, Germany Zhimin (Tim) Cao* Department of Pathology and Laboratory Medicine, University of Louisville, Louisville, Kentucky P. Cupers Flanders Interuniversitary Institute, Gasthuisberg, and Catholic University Leuven, Leuven, Belgium Margitta Dathe, M.D., Ph.D. Department of Peptide Chemistry, Institute of Molecular Pharmacology, Berlin, Germany Bart De Strooper, M.D. , Ph.D. Center for Human Genetics, Flanders Interuniversitary Institute, Gasthuisberg, and Catholic University Leuven, Leuven, Belgium Uwe Ebener, Ph.D. Department of Hematology and Oncology, Clinic of Pediatrics—III, Johann Wolfgang Goethe University, Frankfurt, Germany Matthias Eishold, M.D. Center for Surgery, Department of Urology, Johann Wolfgang Goethe University, Frankfurt, Germany Aron B. Fisher, M.D. Institute for Environmental Medicine and Department of Physiology, University of Pennsylvania, Philadelphia, Pennsylvania J. Kevin Foskett, Ph.D. Department of Physiology, University of Pennsylvania, Philadelphia, Pennsylvania Hans-Joachim Freisleben, Dr.Med.Sci., Ph.D., M.Pharm. Faculty of Medicine, University of Indonesia, Jakarta, Indonesia

* Current address: Clinical Chemistry and Hematology, Wadsworth Center, New York State Department of Health, Albany, New York

Contributors

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Ju¨rgen Fuchs, M.D., Ph.D. Department of Dermatology, Johann Wolfgang Goethe University, Frankfurt, Germany Su¨krettin Gu¨ldu¨tuna, M.D., Ph.D. Medical Clinics II, Johann Wolfgang Goethe University, Frankfurt, Germany Wieger Hemrika, Ph.D. Faculty of Chemistry, E.C. Slater Institute, University of Amsterdam, Amsterdam, The Netherlands Oxana Ibraghimov-Beskrovnaya, Ph.D. Department of Genetics and Genomics, Genzyme Corporation, Framingham, Massachusetts Mahendra Kumar Jain, Ph.D. Department of Chemistry and Biochemistry, University of Delaware, Newark, Delaware Bernhard Kadenbach, Ph.D. Department of Chemistry, Philipps-University, Marburg, Germany Zayna Khayat The Hospital for Sick Children, Toronto, Ontario, Canada Katherine W. Klinger, Ph.D. Department of Genetics and Genomics, Genzyme Corporation, Framingham, Massachusetts Amira Klip, Ph.D. The Hospital for Sick Children, Toronto, Ontario, Canada Linda M. Lenton Division of Biochemistry and Molecular Biology, School of Life Sciences, Australian National University, Canberra, Australia Ulrich Leuschner, M.D. Medical Clinics II, Johann Wolfgang Goethe University, Frankfurt, Germany Jameson A. McCann Department of Biology, St. Louis University, St. Louis, Missouri Michael T. McIntosh, Ph.D. Department of Microbiology and Immunology, MCP Hahnemann University, Philadelphia, Pennsylvania Eckhard Niegemann, Ph.D. Department of Hematology and Oncology, Clinic of Pediatrics—III, Johann Wolfgang Goethe University, Frankfurt, Germany William D. Picking, Ph.D. Department of Molecular Biosciences, University of Kansas, Lawrence, Kansas

xii

Contributors

Patrizius Pilz Membrane Structure Group, Center of Internal Medicine, Johann Wolfgang Goethe University, Frankfurt, Germany Toolsie Ramlal The Hospital for Sick Children, Toronto, Ontario, Canada Rokus Renirie E. C. Slater Institute, University of Amsterdam, Amsterdam, The Netherlands Christoph Rietschel, M.D. Department of Hematology and Oncology, Clinic of Pediatrics—III, Johann Wolfgang Goethe University, Frankfurt, Germany Ernst Theodor Rietschel, Ph.D. Department of Immunochemistry and Biochemical Microbiology, Center for Medicine and Biosciences, Research Center Borstel, Borstel, Germany Jutta Schuchmann-Fix Johann Wolfgang Goethe University, Frankfurt, Germany Ulrich Seydel, Ph.D. Department of Immunochemistry and Biochemical Microbiology, Center for Medicine and Biosciences, Research Center Borstel, Borstel, Germany Yechiel Shai, Ph.D. Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Yasuo Shinohara, Ph.D. Faculty of Pharmaceutical Sciences, University of Tokushima, Tokushima, Japan Romel Somwar The Hospital for Sick Children and The University of Toronto, Toronto, Ontario, Canada Indresh K. Srivastava, Ph.D. Department of Microbiology and Immunology, MCP Hahnemann University, Philadelphia, Pennsylvania Makoto Sugita, Ph.D., D.D.S. Department of Oral Biology, Hiroshima University School of Medicine, Hiroshima, Japan Gary Sweeney The Hospital for Sick Children, Toronto, Ontario, Canada Hiroshi Terada, Ph.D. Faculty of Pharmaceutical Sciences, University of Tokushima, Tokushima, Japan

Contributors

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Peter Tong The Hospital for Sick Children, Toronto, Ontario, Canada Stefan Uhlig, Ph.D. Department of Immunochemistry and Biochemical Microbiology, Research Center Borstel, Borstel, Germany Artur J. Ulmer, Ph.D. Department of Immunology and Cell Biology, Research Center Borstel, Borstel, Germany Akhil B. Vaidya, Ph.D. Department of Microbiology and Immunology, MCP Hahnemann University, Philadelphia, Pennsylvania Roland Valdes, Jr., Ph.D. Department of Pathology and Laboratory Medicine, University of Louisville, Louisville, Kentucky F. Van Leuven Flanders Interuniversitary Institute, Gasthuisberg, and Catholic University Leuven, Leuven, Belgium Sibylle Wehner Department of Hematology and Oncology, Clinic of Pediatrics—III, Johann Wolfgang Goethe University, Frankfurt, Germany Ron Wever, Ph.D. Faculty of Chemistry, E. C. Slater Institute, University of Amsterdam, Amsterdam, The Netherlands Karen Yaworsky The Hospital for Sick Children and The University of Toronto, Toronto, Ontario, Canada Guido Zimmer, M.D., Ph.D. Center of Internal Medicine, Membrane Structure Group, Johann Wolfgang Goethe University, Frankfurt, Germany

Introduction

Part I of this book begins with chapters on antibacterial actions of peptides. Chapter 1 states that the permeabilizing effect on negatively charged membranes is promoted by cationic (⫹) charged peptides. With reduced membrane charge, the potential of structural parameters becomes more important. This pertains to the role of hydrophobic peptide–membrane lipid interactions. The findings agree with those in Chapter 2, which deal with a novel family of antibacterial peptides. Modulations of hydrophobicity and net positive charge are sufficient to increase antibacterial activity via a carpetlike mechanism. With regard to the long-known bee venom melittin, formation of pores at zwitter–ionic interfaces does not solely account for toxicity, but exchange of phospholipids and activation of phospholipase A2 appear to be highly important as well (Chapter 3). Another new class of antibiotics, described in Chapter 4, is the polymyxins (Pmx). Stable contacts between outer cytoplasmic membranes and inner phospholipid layers are mediated by Pmx, thus interfering with bacterial defense against osmotic stress. There may not be a genetically derived resistance against this action. Chapter 5, in Part II, describes the intercellular lipid barrier of skin epidermis as critical for first-line defense against microorganism infections, permeability changes, and consecutive fluid loss. This is critically dependent on the high percentage of unsaturated (linoleic acid) phospholipids, which are highly sensitive to oxidative (environmental) stress, resulting in free radical–induced lipid peroxidation. The role of drug transporters and oxidative stress is the focus of Part III. Chapter 6 takes us into the intracellular milieu. Mitochondrial membranes, containing about 80% protein, are very sensitive to protein structural change. The decrease of reactive -SH groups, frequently due to -SH-S-S- interchange and resulting in rigidizing disulfides, is most critical for oxidative phosphorylation. Uncoupling of this first line of generation of cellular energy from respiration is xv

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Introduction

described in Chapter 7, which clearly explains actions of different types of novel uncouplers. Low concentrations interact with mitochondrial membrane proteins; the type of uncoupling, however, also depends on membrane (lipid) fluidity. For the site of interaction on membrane proteins, a 30-kDa uncoupler binding protein has been described. Nowadays, we could compare this to the 28-kDa PVP protein in F0, which shows proton conduction and was characterized by Papa and coworkers. Under conditions of oxidative stress it may form a disulfide bridge to F1γ, thereby inhibiting ATP synthesis (Chapter 6). Interestingly, G-ketocholestanol has been found to act as a recoupling substance (see Chapter 14). Liposomes as drug transporters are considered in Chapters 8 and 9. Chapter 8 describes the development of the fascinating field of archebacteria, in particular those surviving at temperatures of 56 °C and at pH 1–2. Under these conditions, ultrastable tetraether lipids were developed that could be isolated and transformed into acid-stable and otherwise highly resistant liposomes as drug carriers. Such tetraether lipids could become derivatized and used for successful experiments in transfection into different cell lines (Chapter 9). A crucial role for lipofection appears to be played by the lipid with which the archefects were mixed: the hexagonal phase of dioleylphosphatidylethanolamine significantly augments the transfection efficiency. (Similar observations are described in Chapter 13.) Part IV deals with inflammation and infection. There is another intracellular membrane system, the endosomal environment, within which toxins (e.g., cholera toxin) are processed. Binding of the ‘‘B’’ part of the toxin is influenced by the in vivo existing endosomal pH of 5–6. Chapter 10 outlines the pH-dependent conformational changes of the toxin and correlated membrane structure changes. Chapter 11 deals with the problem of malaria infection. In the malaria parasite, the smallest mitochondrial genome encodes only three proteins (i.e., cytochrome b and subunits I and III of cytochrome c oxidase). Since there is only a functional bc1 complex and cytochrome c oxidase, NADH dehydrogenase, and citric acid cycle producing most of the NADH are absent. Reducing equivalents are delivered by dihydroorotate dehydrogenase, succinate dehydrogenase, and glycerol-3-phosphate dehydrogenase. A therapy has been developed, using a naphthoquinone derivative, atovaquone, in combination with an established antimalarial drug, proguanil. This combination allowed a 100% cure rate with no indication of recrudescence. Atovaquone inhibits parasite bc1 complexes with an EC50 of about 10⫺9 M, whereas mammalian bc1 is inhibited only at about 5 ⫻ 10⫺7 M. Combination with proguanil results in enhancement of ∆Ψm collapse, and thus resistance against atovaquone is avoided. The protein sequences of sensitivity as well as resistance to atovaquone can be localized in the cytochrome b molecule. Chapter 12 stresses the point that infection with liver fluke has different results in different species, which makes comparison of animal models difficult. However, much has been learned from rats—developing uncoupling of oxidative

Introduction

xvii

phosphorylation or changes in F0F1 interaction. Results on disrupted tissue in infected animals nevertheless appear not to be free of artifacts. Importantly, the frequently infected sheep behaved differently compared with the rat after being given the corticosteroid dexamethasone. Therefore, as far as bioenergetic metabolism is concerned, the rat is apparently not a suitable model for sheep. Progress in the field appears very difficult, and this chapter warns against generalizing findings from one species to others. Chapter 13 is devoted to structure/function considerations of lipid A, the endotoxic principal of lipopolysaccharide (LPS). The structure of lipid A is highly conserved throughout organisms containing six alkyl chains by amide or ester bonds, reminiscent of the structure of ceramide, which has been found to induce apoptosis of cells. Concerning the supramolecular structures it was found that lipid A is competely inactive when it occurs in lamellar structure; mixed lamellar/ cubic structures are intermediately active, whereas the QHII (hexagonal) structure is highly active. Similar observations were made in transfection studies (Chapter 9). Fewer than six alkyl chains means that the compounds become antagonistic to lipid A. Receptor binding of lipid A has been described, although it is only partially understood, but intercalation into the host (monocyte, macrophage) membrane via hydrophobic interactions is found. LPS in its active conformation brings about release of numerous cytokines like interleukins 1, 6, 8, 10, 12, and TNF-α. In addition, reactive oxygen species are formed. Chapter 14 describes basic research on cholestasis in plasma membranes of red cells, as well as on those of hepatocytes, and compares these findings to model membranes (liposomes). Addition of chenodeoxycholate results in leaky membranes up to full disintegration. Ursodeoxycholate, by contrast, protects plasma membranes in a cholesterol-like manner. Successful treatment of chronic biliary cholestatic diseases with ursodeoxycholate was initiated by this group. The different biophysical activities of the two compounds can now be considered in light of recent observations by Lamcharfi et al., who found that chenodeoxycholate undergoes stable but very rigid and bulky sandwich structure, whereas ursodeoxycholate reveals more flexible R-OH-OOC-R interactions. In Part V, Chapters 15 to 17 are concerned with metabolism. Chapter 15 tackles the old problem of mechanism of action of thyroid hormones. It has been recognized for a very long time that thyroid hormones should act by some sort of uncoupling of respiration from phosphorylation. The exact mechanism, however, remains obscure. Chapter 15 shows that binding of 3,5-diiodo-l-thyronine (3,5T2) to subunit Va of cytochrome c oxidase completely abolishes allosteric inhibition at high matrix ATP/ADP ratios. This also means that it abolishes the socalled second mechanism of respiratory control. 3,5-T2 also partially inhibits the first mechanism via proton motive force ∆p of the reconstituted enzyme. The increase of basal metabolic rate by thyroid hormones can be explained in this novel way.

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Introduction

The way in which the manifold metabolic interactions of insulin are brought about is definitely linked to membrane binding at receptor sites and a cascade of events stimulating glucose transport. Chapter 16 shows that activation of the insulin receptor tyrosine kinase appears to be the first biochemical consequence of a hormone binding to its receptor. Low-density microsomes represent just another intracellular fraction of membranes from which the intracellular pools of glucose transporters are derived. Apart from defects in signaling steps of insulin action in diabetes type 2, there may be additional impairment of glucose transport (e.g., defects of translocation, reduction in amount of transporters, or defect in final activation of the transporters in the plasma membrane). Moreover, the actin skeleton of the membrane appears changed—it has a ‘‘ruffled’’ appearance. Structural changes of cytoskeletal proteins have also been known to occur in oxidative stress (Chapter 18). The last chapter on metabolism (Chapter 17) concerns the site where glucose is produced from glycogen: glucose-6-phosphatase (G-6-Pase) is the only enzyme in the body that produces significant amounts of glucose. Moreover, it has been suggested that overexpression of G-6-Pase may play a role in some forms of diabetes. For this enzyme, which is located in the membrane of the endoplasmic reticulum, a substrate transport model of function has been favored. In this model, the hydrolytic enzyme is accompanied by accessory proteins responsible for substrate or product transport and for stabilization of the catalytic unit. Two of these transporters, T2 and T3, transport phosphate, whereas GLUT 7 transports glucose. Absence of G-6-Pase or of membrane transport activity was found to be the cause of glycogen-storage, or von Gierke’s, disease, which may cause hypoglycemia, growth retardation, hepatomegaly, kidney enlargement, hyperlipidemia, and lactic acidosis. The development of a model for G-6-Pase, including nine transmembrane helices, was based on the observation that the far more advanced structural analysis of haloperoxidase, activated by vanadium, aligns and shows close homology in many respects with G-6-Pase. Thus, through comparison within the superfamily, great progress (including correct positioning of the decisive amino acid residues at the luminal site of the endoplasmic reticulum as well as with the reaction mechanisms of G-6-Pases) has been made. New aspects of Alzheimer’s disease are discussed in Part VI. For some time, investigators have suspected that some age-related neurological diseases may develop along with increased oxidative stress. With oxidative stress, enzymes such as catalase superoxide dismutase and glutathione peroxidase were found to decline during aging. For unknown physiological reasons, some βAPP molecules are processed by a series of proteolytic cuts to generate small peptides of 40–42 amino acids known as amyloid β-peptides. Because of their β-pleated sheet structure, these peptides aggregate to form insoluble amyloid deposits that lead to neuronal degeneration (Alzheimer’s). Chapter 18 presents clear evidence for association of amyloid β-peptide

Introduction

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with oxidative stress, exemplified by EPR spin trapping results using the trapper PBN. In contrast to most other studies using spin trapping, there was no detectable signal in the extremely purified trap itself, which makes the results much more convincing. In addition, this chapter gives very thorough and substantial proof for oxidative stress on AD. There are indications of protein as well as lipid oxidation, and there are other spin label data on motional and reduction characteristics; therefore, antioxidant therapy is at hand and appears to be worthwhile. Information regarding the dangerous rise of amyloidogenic β-peptide in Alzheimer’s disease is delivered in Chapter 19. Point mutations mainly in the presenilin-1 gene (PS1) are a major cause. Most of the affected residues are located in the predicted transmembrane regions of the endoplasmic reticulum, probably arranged on one side of an α-helix. It has been proposed that presenilins form channels or transporters within the endoplasmic reticulum toward the Golgi apparatus, and mutations result in a selective increase in production of amyloidβ (1-42) peptide by proteolytic processing of the precursor protein (APP). Also, in this case, learning from homologs or closely related structures induces progress. Part VII focuses on oncology. Lymphoblast cells of peripheral blood as well as cell lines from sick children are analyzed in Chapter 20. In this chapter, biophysical EPR spin labeling methodology has been added to the conventional analytical arsenal. Some remarkable results were obtained using 16-doxyl-stearic acid. Compared to control lymphocytes, there was a much broader range of polarities as well as of (provisional) order of the plasma membranes, indicating diverse states of activities. Lymphoblast cells, by contrast, appeared much more uniform. Inducing apoptosis in the cell lines revealed a large increase in membrane polarity in the hydrophobic core, whereas necrosis made the spectrum more anisotropic, the apparent order parameter reaching values between 0.3 and 0.4. Chapter 21 reports on cDNA cloning and biochemical characterization of the antigen recognized by the monoclonal antibody (mAb) anti-breast-cancer GCG. The GCG mAb revealed a strong staining of the cell surface of the human breast-cancer-derived cell line MCF7, as well as of cell lines derived from other kinds of human tumors, including the erythroleukemia cell line K-562. GCG mAb recognized with high specificity a 28-kDa membrane protein from MCF7 as well as K-562 cells. In Part VIII, Chapter 22 decribes the molecular biological development of polycystins-1 and -2. Mutations within these two integral membrane proteins result in autosomal dominant polycystic kidney disease, which represents about 8 to 10% of all cases of end-stage renal disease. The genomic region of PKD1 codes for a large cell-surface glycoprotein (polycystin-1) of unknown function. Its predicted domain structure leads to the suggestion of involvement in protein–protein and protein–carbohydrate interaction. Approximately 30% of polycystin-1 consists of 16 copies of a novel protein module called the PKD domain. This domain has a β-sandwich fold, also called an immunoglobulin (Ig)-

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like fold. Nevertheless, the polycystins do not belong to the Ig family. Distinct domains of polycystin are encoded by separate exons. This opens the possibility of explaining differential functions in various tissues. Polycystin expression is confined mainly to epithelial cells, supporting a role in epithelial cell differentiation. Part IX covers pulmonary physiology and disease. Chapter 23 reports on different membrane-stabilizing/destabilizing properties of some β-agonists compared with cromoglycate. To this end, mononuclear cells were isolated from buffy coats and EPR spin labeling using 5- and 16-doxyl stearic acid was carried out. Cellular viability was probed with trypan blue. It was found that fenoterol and salbutamol destabilized the mononuclear cell membrane, whereas cromoglycate and reproterol stabilized it. Protection of mast cells from degranulation by some β-agonists should be differentiated from cell membrane stabilization of mononuclear cells. Cystic fibrosis (CF) and the cystic fibrosis transmembrane conductance regulator (CFTR) is reviewed in Chapter 24. Traditionally, in lungs the CFTR functions as an apical membrane localized Cl⫺ channel that regulates airway transepithelial fluid balance. Proper fluid balance is critical for mucociliary clearance of inflammatory particles, including bacteria. Activity of CFTR is regulated by phosphorylation of the R domain and ATP hydrolysis at two nucleotide binding domains (NBDs). Recently, however, it has been shown that ATP and Cl⫺ conduction appear to be separated. The normal function of CFTR now appears to be (inter alia) inhibiting Na⫹ channel activation (epithelial Na⫹ absorption) and possibly stimulating Cl⫺ secretion. It has been known for a long time that CF goes along with increased Na⫹ absorption (viscous mucus). CFTR may regulate the channel by means of a mediator or by direct protein–protein interactions. In addition, CFTR, abundantly expressed in the kidney along nephrons, influences K⫹ transport. There is also an astonishing diversity of ion channels, which are known to be regulated or influenced by CFTR. In CF, therefore, mutations may lead to additional disturbance of ion homeostasis apart from that of Cl⫺. Lung surfactant, in addition to its role in keeping the alveolae and small airways open, provides a barrier against external toxic materials. In degradation of this lipid layer, a Ca2⫹-independent lysosomal phospholipase is active (Chapter 25). A specific inhibitor active at the intralysosomal pH 4 has been studied. Most interesting is the finding that this enzyme also functions as a GSH peroxidase, which supports the idea that surfactant provides an antioxidant barrier. The different isoforms of α- and β-subunits of Na⫹-K⫹-ATPase that exist in different tissues are described in detail in Chapter 26. Upon completion of synthesis, the α-subunit associates with the β-subunit within the lumen of the endoplasmic reticulum. A sorting signal located within the NH2 terminal region of the α-subunit brings about polarized distribution of Na⫹-K⫹-ATPase on the cell surface. Reversed polarity was observed in polycystic kidney disease (see

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Chapter 22) as well as during ischemia (probably redistribution of plasma membrane protein, too!). Interaction of Na⫹-K⫹-ATPase with the cytoskeleton is described. Binding sites for cardiac glycosides are characterized on hairpin loops TM 1-TM 2, TM 5-TM 6, and TM 7. There are multifold long- as well as short-term influences, possibly from hormones or druglike substances, and from ensuing membrane structure changes and redistribution of subunits. All this results in a high probability of activity changes of the Na⫹-K⫹-ATPase in many disease states. Its relevance for etiology or duration and outcome in each case, however, is difficult to prove.

1 Antibacterial and Hemolytic Activity of Amphipathic Helical Peptides Margitta Dathe Institute of Molecular Pharmacology, Berlin, Germany

I. INTRODUCTION Over the past decade a multitude of peptides with cell lytic activity has been discovered in almost all kinds of species (1–4). The diversity of physiological systems in which the peptides are expressed and their broad-spectrum activities have confirmed their role as offensive and defensive weapons of creatures. Many of these peptides exhibit antimicrobial specificity and have been suggested to constitute a system of innate immunity (2,5–8). These properties have made several compounds promising candidates for the development of a new class of antibiotics (6,9,10). Stimulated by the challenge of bacterial resistance to conventional antibiotics (11–13) and by evidence for anticancer activity (9), considerable effort is being made to understand the structural basis of membrane selectivity and to elucidate the molecular mechanism of action (14,15). The peptides exert their effect by permeabilization of the lipid matrix of the target cell. It is generally accepted that the peptide–membrane interaction is the consequence of several common structural features of the peptides, such as the tendency to adopt an amphipathic conformation, as well as of properties of the anisotropic cell membrane (1,16,17). However, the molecular mechanism of membrane disturbance by the diverse membrane-active peptides is still controversial. This review covers membrane-active peptides with an amphipathic helical structure. We address the question of what distinguishes those peptides that are capable of lysing bacteria or normal eukaryotic cells and focus on advances in 1

Dathe

2

understanding the mode of membrane permeabilization. Finally, the potential of peptides as therapeutic agents and as tools for pharmaceutical research is outlined.

II.

MEMBRANE-PERMEABILIZING PEPTIDES

A.

Classification and Biological Function

On the basis of their origin, activity, or structure, peptides with cell lytic activity can be classified into several groups. The peptides have been discovered in all organisms in which they have been sought: bacteria, fungi, insects, and vertebrates such as fish, amphibians, birds, and mammals including humans (4). With respect to their activity against eukaryotic or prokaryotic cells they can be classified as either hemolytic or antimicrobial compounds. Apart from sequence homology within peptide families, the sizes and amino acid compositions vary widely. With respect to secondary structure, the peptides can be categorized as (1) linear, mostly assuming an α-helical conformation under appropriate conditions, (2) peptides with one to three disulfide bonds leading to the formation loops or of stabilized antiparallel β-sheets, and (3) linear peptides rich in specific amino acid residues with a preferred structure distinct from the α-helix or βconformation. Table 1 summarizes some properties of representative peptides from diverse sources. δ-Hemolysin is a peptidic toxin from Staphylococcus aureus with high hemolytic activity (18). The alamethicin-producing fungus Trichoderma virides (19) and insect venoms are other sources of hemolytic peptides. Peptidic toxins produced by microorganisms or as components of bee and wasp venoms such as melittin (20) and mastoparan (21) serve preferentially as offensive weapons. However, in addition to their hemolytic activity, the latter are potent antibiotics. Other peptides from insect species such as the cecropins from the pupae of silk moths and flies (22,23) are produced as a response to microbial infections. The peptides possess pronounced antibacterial selectivity but are only weakly hemolytic. Insects, which have no highly specific immune response (24,25), use antimicrobial peptides to combat microbial invaders. Lytic peptides from vertebrates are often less potent against eukaryotic cells but act on a rather broad spectrum of microorganisms (Table 1). The antimicrobial peptides from amphibians comprise a major group (26). Two representatives are the magainins from a family of antibacterial peptides that are expressed in the skin and intestine of the African clawed frog (27,28) and buforin (29), which was recently purified from the stomach of an Asian toad. Magainin was discovered when Zasloff noticed that incisions in the frog skin healed without infection, inflammation, or notable scarring despite being exposed to a bacteria-filled envi-

a

⫹⫹

⫺ ⫺ ⫹

α-Helical β-Structure

31, linear; ⫹5 41, three SES bonds; ⫹7 13, linear; ⫹4

Cecropin P1 β-Defensin hBD-2 Indolicidin

Extended helix

⫹ ⫹

⫺ ⫺

α-Helical α-Helical

23, linear; ⫹4 21, linear; ⫹6

Magainin II Buforin II

⫹⫹ ⫹⫹

⫹⫹ ⫺

⫹⫹ ⫹⫹

⫺ ⫹

Grampos

α-Helical α-Helical

⫹⫹ ⫹

RBC

26, linear; ⫹6 37, linear; ⫹7

α-Helical α-Helical/310helix

Secondary structure

⫹⫹

⫹⫹ ⫹⫹

⫹⫹ ⫹⫹

⫹⫹ ⫹⫹

⫺ ⫺

Gramneg

Bacteria

Cytolytic activity

Melittin Cecropin A

27, linear; 0 20, linear; ⫺1

Number of residues; total charge

⫹⫹, ⫹, and ⫺ denote high, low, and practically no biological activity, respectively; RBC ⫽ red blood cells.

Bovine neutrophils

Insects Bee venom Hemolymph (Hyalophora cecropia) Amphibians Frog skin (Xenopus laevis) Toad stomach (Bufo bufo gagarizan) Mammals Porcine small intestine Human skin

δ-Hemolysin Alamethicin

Peptide

Origin and Properties of Representative Cytolytic Peptidesa

Microorganisms and fungi Bacterium: (Staphylococcus aureus) Fungus: (Trichoderma virides)

Origin

Table 1

34

30 33

27 29

20 22

18 19

Ref.

Amphipathic Helical Peptides 3

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ronment. Most of the amphibian peptides are linear and assume an α-helical conformation on interaction with the target membrane. Peptide-based antimicrobial activity is also well established in mammals. The first mammalian cecropin was isolated from the porcine small intestine (30). The potentially helical peptide exhibits high activity against gram-negative bacteria but is less active against gram-positive bacteria and practically inactive against erythrocytes. The largest group of mammalian antimicrobial peptides comprise the β-structured defensins (see reviews 31,32). The peptides are found in rabbit, rat, mouse, guinea pig, cattle, and human. While the α-defensins are stored in circulating macrophages and neutrophils, β-defensins, among them the recently found β-defensin hBD-2 (33), are produced in skin, salivary glands, and airway tissues and by epithelia of the urogenital tract in response to bacterial infections. Other mammalian antibacterial peptides with remarkable structural variability are derived from cathelicidin precursor peptides. All those peptides that incorporate a high proportion of specific amino acid residues such as arginine, proline, or tryptophan belong to this group. Among them is indolicidin (34), the smallest of the known naturally occurring linear antimicrobial peptides that may assume an extended helix (35). Antimicrobial peptides in vertebrates are expressed in the epithelia and are used to protect the epithelial surface from microbial infections or for local control of functionally important microorganisms. Furthermore, vertebrate antibiotic peptides play a decisive role in the microbicidal mechanism after delivery to phagocytic vacuoles containing ingested microorganisms. Actually, the peptides are poorly efficient compared to the more sophisticated toxins and generally function without high specificity and memory, but they allow the animal an immediate response to microbial invasion prior to mobilization of the adaptive immune system (3,8,25). They thus constitute a secondary, chemical immune system for host defense.

B. Mechanism of Action and Properties of the Target Membranes All available evidence suggests that peptides exert their effect by disrupting the barrier function of the target cell membrane (2,14,15,17). This activity is not mediated by membrane receptors. The facts that much higher peptide concentrations (in the micromolar range) are required than are needed for specific peptide receptor binding and that l-amino acid peptides and their all-d enantiomers are equipotent show that protein chirality does not play a role. Additionally, activity against a broad spectrum of biological membranes exhibiting varying protein and enzyme patterns as well as peptide-induced permeabilization of different lipid

Amphipathic Helical Peptides

5

model membranes suggest a rather nonspecific effect of the peptides on the membrane lipid matrix. According to our current knowledge, membrane perturbation appears to be determined by a balance of electrostatic and hydrophobic interactions between the peptides and target membranes. The envelope of gram-negative bacteria consists of the outer wall and the cytoplasmic target membrane, interspaced by a peptidoglycan layer (36) (Fig. 1a). Highly negatively charged lipopolysaccharides (LPSs) are the main constituents of the outer membrane, and metal cations help stabilize the membrane by decreasing electrostatic repulsion between the LPS molecules. This wall allows the direct passage of only small hydrophobic residues by direct crossing through the bilayer or hydrophilic permeation of small molecules through the water-filled porin channels. Antimicrobial peptides have been postulated to overcome this barrier by displacement of the native cations from their binding sites. Binding of the much larger peptides disrupts the LPS arrangement. The resulting transient lesions in the outer membrane are large enough to permit the passage of peptides (self-promoted uptake) (10,35). The pronounced negative charge of the outer wall favors the accumulation of basic host defense peptides while remaining a barrier for electrically neutral and hydrophobic sequences. The inner target membrane of gram-negative bacteria is rich in phosphatidylethanolamine (PE), a zwitterionic lipid with the tendency to destabilize membranes at low bilayer–hexagonal transition temperatures. Negatively charged phosphatidyglycerol (PG) and a markedly negative inside transmembrane potential of approximately ⫺170 mV (37) support binding and disturbance of the cytoplasmic membrane by cationic peptides. Gram-positive bacteria have no outer membrane barrier, thus leaving the cytoplasmic membrane directly exposed to the lytic peptides. Here the presence of negatively charged teichoic and teichuroic acids (38) may facilitate the peptide–membrane interaction. Taken together, electrostatic interactions should play a decisive role for the peptideinduced permeabilization of bacterial membranes. The lipid composition of the eukaryotic membrane is highly complex (Fig. 1b) (36). The presence of sphingomyelin (SM), phosphatidylcholine (PC), and phosphatidylethanolamine (PE) in the outer leaflet renders the membrane surface of red blood cells electrically neutral at physiological pH. A high content of cholesterol stabilizes the lipid bilayer. The transmembrane potential (⫺9 mV) (39) is rather low, and the anionic charge of sialic acid molecules is located too far from the bilayer surface (40) to be significant. Consequently, electrostatic forces are of minor importance and hydrophobic interactions should dominate the peptide-induced damage of eukaryotic membranes. The precise events that cause inhibition of bacterial growth and lysis of red blood cells are not clear. However, the distinct differences in the membrane properties of prokaryotic and eukaryotic cells favoring electrostatic or hydropho-

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Figure 1 (a) Architecture of the membrane of gram-negative bacteria. The outer membrane is rich in negatively charged lipopolysaccharides and lipoproteins; the inner membrane contains phosphatidylethanolamine (PE), cardiolipin (CL), and negatively charged phosphatidylglycerol (PG). Occupation of ion-binding sites on the outer membrane by cationic peptides initiates self-promoted peptide uptake. (b) The structure of the membrane of erythrocytes as a model of a eukaryotic cell membrane is characterized by a high amount of sphingomyelin (SM), phosphatidylethanolamine (PE), phosphatidylcholine (PC), and cholesterol (Ch). Negatively charged phosphatidylserine (PS) is preferentially located in the inner leaflet.

Amphipathic Helical Peptides

7

bic interactions appear to regulate the specificity of the many differently structured membrane-active peptides. To gain more insight into the mechanism of action, the role of structural properties of the peptides for membrane binding and membrane permeabilization has to be elucidated in detail.

III. STRUCTURAL AND FUNCTIONAL STUDIES OF AMPHIPATHIC HELICAL PEPTIDES A.

Model Membranes and Methods of Structural and Functional Investigation

Since the structures of the various target membranes are very complex and our knowledge of their exact properties is rather limited, various lipid model systems have been used for physicochemical studies of peptide–membrane interaction. These models include (1) vesicles that represent spherical lipid bilayers enclosing an aqueous compartment, (2) planar bilayers covering a small hole in a partition wall separating two compartments that contain aqueous solutions, and (3) oriented bilayers prepared by covering a solid support with lipid films. Lipid monolayers and micelles complete the spectrum of systems mimicking membrane properties. Nuclear magnetic resonance (NMR), circular dichroism (CD), Fourier transform infrared (FTIR), and fluorescence spectroscopy using small unilamellar vesicles or oriented bilayers are the preferred techniques for structural studies and the investigation of the location and orientation of peptides in the membranebound state. NMR, differential scanning calorimetry, and neutron in-plane scattering are valuable methods that provide information regarding peptide-induced changes in lipid bilayer properties. Peptide-induced membrane permeabilization is mostly studied by the release of water-soluble compounds such as fluorescent dyes encapsulated in large unilamellar vesicles and by measuring voltage-induced conductivity by formation of ion channels in planar bilayers (for review, see Ref. 15). By adopting various preparative techniques, along with a diversity of lipid composition, which influences charge, thickness, and fluidity of the model membrane, one obtains a rich arsenal for modeling the diversity of natural cell membranes. B. Structural Motifs: Helicity, Charge, Hydrophobicity, Hydrophobic Moment, and Size of the Polar/Hydrophobic Domain Peptides that assume an amphipathic α-helical structure in a membrane environment have the broadest spectrum of occurrence and activity (Table 1). Amphipathic helical peptides constitute structures that twist into two-sided, spiral mole-

8

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cules (Fig. 2). One face consists of hydrophobic residues soluble in the lipid acyl chains that comprise the membrane interior, and the other face is polar and watersoluble. Since the peptides are short (from 13 to about 40 amino acid residues) and easily synthesized, analogs of naturally occurring peptides as well as ‘‘designer peptides’’ engineered on the basis of current knowledge of structural principles have been subjected to extensive structure–activity studies. Figure 2 illustrates the properties of three well-investigated membraneactive peptides. The peptides differ in their amino acid composition, which results in differences in size, charge, and pattern of the hydrophobic/hydrophilic helix envelope. Alamethicin consists of 20 amino acid residues (Fig. 2) and is rather hydrophobic. Only a few polar residues and one negatively charged glutamic acid unit form a small polar domain. The amino acid composition of magainin 2 results in moderate hydrophobicity (Fig. 2). The N-terminal α-amino group is not protected, and the cationic side chains are well distributed over the whole sequence, resulting in a charged domain covering an angle (φ) of 120° on the helical coat. Porcine cecropin P1 is 31 residues in length (Fig. 2). It is charged at the N and C termini at physiological pH and contains a large number of charged side chains, rendering the peptide very hydrophilic and resulting in a large cationic domain (φ) of 200°. The amphipathic helix of the peptides is complementary to the anisotropic structure of the cell membrane, with a polar, water-exposed surface and a hydrophobic interior; this complementarity is thought to provide the basis for peptide incorporation into the lipid matrix and subsequent perturbation of the membrane structure. Indeed, increasing amphipathic helical structure has been found to be well correlated with enhanced hemolytic and antimicrobial activity of melittin (41,42) or the antibacterial effect of both magainin (43) and cecropin P1 (44). Inversely, substitutions in melittin that prevent its folding into a helical conformation resulted in a loss of both hemolytic and antibacterial activity (45) [see also Chap. 3 and compare results of Shai (Chap. 2) employing diasteriomers]. Like-

Figure 2 Helical wheel projections and schematic drawings of the amphipathic helices of alamethicin, magainin 2, and cecropin P1. The one-letter code for amino acids is used, U indicating α-aminoisobutyric acid. Hydrophobic residues are shown in white, polar residues in gray, and cationic residues in black circles. N is the number of residues. The total charge Q was calculated under the assumption that K, R, and the N-terminal NH2 are positively charged and E and the C-terminal COOH bear a negative charge. H is the total hydrophobicity per residue, calculated as the sum of the hydrophobicities of the individual amino acid residues on the basis of the Eisenberg (132) consensus scale of hydrophobicity; µ is the hydrophobic moment determined as the vector sum of all residues, with the figure |µ| being a measure of the separation of polar and hydrophobic residues on the helix coat. The angle subtended by the charged residues on the helix surface is denoted by Φ.

Amphipathic Helical Peptides

9

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wise, helix disturbance by the introduction of proline in cecropin A (46) and the incorporation of helix-breaking d-amino acid residues in magainin (47,48) reduce its antibacterial activity. In many cases enhanced antibacterial activity associated with increased peptide helicity has been shown to be connected with reduced specificity. This effect is related to the much more pronounced connection between peptide helicity and hemolytic activity. Using model peptides of different helicities we have shown different peptide–lipid interactions to be responsible for the induction of permeability of membranes bearing high and low negative surface charge (49). Disturbance of the negatively charged membrane is determined by high peptide accumulation via electrostatic interactions. Peptide secondary structure is of less importance. In contrast, peptide interaction with the neutral lipid membrane is dominated by hydrophobic interactions, and the formation of a hydrophobic helix domain is essential for activity. The results demonstrate that amphipathicity is much more important for the permeabilization of the neutral membrane of erythrocytes than for activity against negatively charged bacterial envelopes. More recent studies confirm that a helical conformation is necessary for hemolytic activity but is not a prerequisite for antibacterial activity (50–52). Inspection of the amino acid sequences of the membrane-permeabilizing peptides (Table 1, Fig. 1) reveals that compounds with high antimicrobial activity are highly cationic in nature. This positive overall charge was found to distinctly modulate the effect. Thus, substitution of two negatively charged amino acid residues in the electrically neutral δ-hemolysin by cationic lysines induced potent antibacterial activity (53) while conserving the hemolytic activity. Also, an increased net positive charge of melittin (54) and the addition of 10 lysine residues to the N-terminus of magainin 2 (55) substantially enhanced their antibacterial activity. A decrease and final loss of antibacterial activity of magainin by reduction of charge from ⫹6 to zero was recently described (56). Magainin analogs having an increased charge enhanced the outer membrane permeabilization and effectively permeabilized the inner membrane, thus demonstrating the importance of electrostatic interactions for the antimicrobial effect. Studies with model peptides confirmed that a positive overall charge is much more important for antibacterial activity than for lysis of the neutral erythrocyte membrane (57). Surprising in this connection is the report that with an increase in the number of lysine residues in model peptides the hemolytic activity disappeared, but independent of charge all analogs studied were completely inactive against various bacterial strains (58). Such results suggest that in addition to peptide helicity and overall charge other structural properties are important to activity. Studies of natural (59,60) and model (61–63) peptides showing a varied balance of hydrophobic and hydrophilic residues implicate activity-modifying effects of peptide hydrophobicity (H ), the hydrophobic moment (µ), and the size of the polar domain (Φ) (for explanation, see Fig. 2). Based on the observation that the amphipathic helix of cationic peptides is less important for the permeabil-

Amphipathic Helical Peptides

11

ization of highly negatively charged membranes but is essential for the disturbance of neutral membranes (48,49), our recent work was aimed at uncovering the structural basis of activity and selectivity by studying the influence of hydrophobicity, hydrophobic moment, and size of the polar domain on peptide binding and membrane permeabilization. Our studies were based on structural modifications of the antimicrobial magainin 2 amide (Fig. 1) (64–66) and of a nonselective model peptide (49,67,67a). The results provided the following picture: At negatively charged membranes, high affinity caused by electrostatic interactions with the cationic peptides determines the permeabilizing effect. H, µ, and Φ play minor roles. With reduced membrane charge, the potential of the structural parameters becomes more pronounced, thereby increasing the role of hydrophobic peptide– lipid interactions. Although membrane affinity decreases, the permeabilizing activity remains high owing to an increased efficiency of the bound molecule to perturb the lipid arrangement. The perturbing effect on neutral membranes further increases with enhanced H, µ, and Φ. The results obtained are in accordance with a model of peptide–lipid interactions that explains the different activities by changes in the amount and location of bilayer-bound peptides (49). On negatively charged membranes electrostatic interactions cause high peptide accumulation. The sequences are fixed at the polar membrane interface. But this surface state is less effective for permeabilization. On neutral membranes, hydrophobic interactions become significant, and despite low binding affinities the peptides may penetrate deeply into the hydrophobic membrane region and effectively disarrange the bilayer structure. The peptide-induced permeabilization of neutral model membranes largely correlates with the lytic effect on red blood cells (67). The parameters H, µ, and Φ modulate the activity on the neutral membrane of erythrocytes most strongly. Thus, their enhancement results in a loss of peptide selectivity. In a comparable way the parameters influence peptide activity on slightly negatively charged bacterial plasma membranes, where the hydrophobic effect is superimposed onto electrostatic interactions (67a). The pronounced affinity for the acidic outer membrane is responsible for the antibacterial specificity of cationic peptides for gramnegative bacteria. This mechanistic picture of changes in the dominance of electrostatic and hydrophobic peptide–membrane interactions provides an explanation for the different activity spectra (Table 1) of such peptides as δ-hemolysin and alamethicin, the hemolytic and antimicrobial melittin, and the antibacterial magainin and cecropin P1. δ-Hemolysin and alamethicin are inactive against gram-negative bacteria, since low net charge prevents peptide accumulation and self-promoted uptake through the outer wall. The extended hydrophobic N-terminus of alamethicin, however, allows binding to, and penetration into, the neutral membranes of erythrocytes (68) and the slightly negatively charged envelope of grampositive bacteria. Melittin activity against eukaryotic cell membranes is mediated by strong hydrophobic interactions with the extensive hydrophobic N-terminal

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12

peptide domain, and the effect on prokaryotic membranes is favored by ionic interactions with the charged C-terminus. In magainin, structural motifs favoring hydrophobic interactions to cause hemolysis are poorly developed; the charge is high, and the hydrophobicity and hydrophobic moment are moderate (Fig. 2). However, electrostatic contributions to membrane interaction result in antimicrobial activity against gram-positive bacteria, which is further increased by the strong electrostatic effect on the outer wall of gram-negative bacteria. Electrostatic forces mediated by the high peptide charge are also responsible for the high activity of cecropin P1 against gram-negative bacteria, but the low hydrophobicity, hydrophobic moment, and small hydrophobic domain seem to reduce its effect on the membrane of gram-positive bacteria and to prevent lysis of red blood cells. C. Modes of Membrane Permeabilization Membrane lysis is thought to be a two-step process determined by (1) peptide binding and (2) permeabilization of the lipid bilayer (Fig. 3). Accumulation of amphipathic helices may distinctly influence the physical properties of lipid layers. Thickness-matching to the inserted peptide, area expansion, changes in the lipid phase, and lipid motion causing the formation of domains impair membrane stability (69–71). However, binding is not sufficient for membrane permeabilization. Brasseur (72) and Brasseur et al. (73) suggested that helices of membrane-bound peptides may adopt various orientations relative to the bilayer and associate to differently structured units depending on how the hydrophobic and polar domains on the helix are separated. Both peptide location and association distinctly influence the mode of membrane permeabilization. At present three main mechanisms are favored (Fig. 3): (1) association of helical peptide monomers in transmembrane orientation with the polar helix domain to form the lumen of a channel that allows the passage of small ions (16); (2) formation of pores as complex peptide–lipid structures with lipid headgroups and the polar face of peptide helices lining the aqueous pore center (74,75); and (3) formation of peptidic monolayers or bundles creating transient defects in the bilayer packing (44,76). Finally, at high concentrations amphipathic peptides may exhibit detergent-like properties. The detergent effect results in the release of membrane fragments as mixed micelles, vesicles, or nonspecific clusters outside the cell (77). 1.

Alamethicin Channels

Alamethicin is a typical ion-channel-forming peptide (1, Fig. 3). The peptide bears a low charge and is quite hydrophobic with [H] ⫽ 0.55, and the low hydrophobic moment ([µ] ⫽ 0.23) corresponds to a significant but low degree of amphipathicity (Fig. 2). Although very flexible as a monomer, in solution the

Amphipathic Helical Peptides

13

Figure 3 Modes of peptide membrane interaction. Peptides bind to the membrane by assuming an amphipathic helix with the polar domain exposed to the membrane surface and the hydrophobic helix face buried in the lipid acyl chain region. Changes in the lipid bilayer structure and reorientation of the bound peptides lead to membrane permeabilization by (1) formation of ion channels, (2) complex peptide-lipid pores, or (3) a carpetlike mechanism.

peptide may form a largely helical rod in the N-terminal sequence with a less well defined C-terminus in the associated state or in a lipid environment as indicated by NMR studies (78,79). When arranged in an ideal α-helix, the 20-residue ˚ linear displacement per turn) is long enough to span sequence (5.5 turns ⫻ 5.4 A ˚ thick lipid membrane. Thus the peptide appears to be ideally suited for a 30 A N-terminal insertion into the membrane interior. The affinity of the monomeric alamethicin for neutral bilayers, reflected by a partition coefficient of 1000 M⫺1 (80) and an apparent binding constant of 4000 M⫺1 (68), is rather high (Table 2). The localization and orientation of bilayer-bound peptide is dependent on a critical peptide/lipid ratio, as shown by oriented circular dichroism studies (81). The effects observed are determined by the physical state of the bilayer and its

b

a

PC ⫽ intrinsic (hydrophobic) partition coefficient. Data refer to magainin 2 amide. c NBD-labeled.

Cecropin P1 c

Magainin 2

103 (PC)a (131) (68) 4 ⫻ 103 (PC) 1.9 ⫻ 105 (PG)b (65) 2 ⫻ 104 (PC/PG) 4 ⫻ 102 (PC) 3.1 ⫻ 104 (PC) (44) 1.2 ⫻ 105 (PC/PS)

Affinity Kapp (M⫺1) (88) (48)

(112)

44 75b

80

Helicity α (%) Cooperative binding, association at low peptide/lipid ratio (131) Noncooperative binding, association at high peptide/lipid ratio (103) No association (101) Noncooperative binding, no association (44)

Mode of interaction, association behavior

Structure and Properties of Lipid-Bound Peptides Derived from Studies with Model Membranesa

Alamethicin

Peptide

Table 2

Parallel (112)

Parallel or perpendicular (81,82,95) Parallel or perpendicular (102,105,106)

Orientation to the membrane surface

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Amphipathic Helical Peptides

15

hydration level (82) (Table 2). At low peptide concentrations the majority of alamethicin molecules are in the surface state, whereas at higher concentrations or under high hydration they are in the membrane-inserted state. More recent structural investigations using solid-state 15N NMR studies (83,84), electron paramagnetic resonance spectroscopy (85), and molecular dynamic simulations (86) confirm the high flexibility of the central glycine/proline-containing sequence. The two residues allow bending of the more polar C-terminus away from the helical axis and its alignment, in the absence of a membrane potential, with the N-terminal helical segment inserted perpendicular to the bilayer surface. In this configuration alamethicin does not completely cross the bilayer, and the C-ter˚ from the membrane minus remains exposed to the aqueous solution, 3–4 A interface. It is under conditions of low concentration with largely monomeric peptide (85) that alamethicin induces channel activities in lipid bilayers. Well-defined conduction states are a convincing argument for the ‘‘barrel stave’’ model (87). Association of up to 12 monomers driven by dipole orientation of the alamethicin helix in the transmembrane potential field has been suggested to determine the different conductance levels (88). The channel diameters derived from geometrical considerations and conductivity experiments range between 0.2 and 2 nm (89,90). In addition to the passage of monovalent ions, Ca2⫹ exchange in chromaffin cells appears to be highly probable (68), and Mn2⫹ and Ni2⫹ influx has been suggested to be evoked by alamethicin channels (91). However, even the largest channels were found to be impermeable to water-soluble polymers (92,93). Thus, voltage-induced channel formation may disturb the ion balance of cells with a high transmembrane potential such as gram-positive bacteria (⫺190 mV) and chromaffin cells (⫺60 to ⫺80 mV) (68) and interrupt oxidative phosphorylation in mitochondria (94) but does not directly induce the release of membrane components. In addition, alamethicin channel activity on cell membranes lacking a transmembrane potential, such as red blood cells, remains unproven. Lysis occurs if the peptide concentration exceeds a critical membrane concentration. High alamethicin accumulation has been described as changing the membrane properties. Lipid chains become disordered, and the bilayer thickness decreases proportionally with increasing peptide concentration over a large area around the binding site (95). Overlapping of zones of metastability at high peptide accumulation may be the reason for membrane disruption. 2. Magainin Pores Voltage-induced channel activity in lipid bilayers has also been reported for the antimicrobial peptide magainin. However, the events are rare, short-lived, and poorly reproducible (96,97). From studies on model membranes no evidence exists that magainin forms distinct ion-sized transmembrane channels. On the other

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hand, bacterial cells are depolarized immediately on exposure to magainin (98,99), and decoupling of the respiratory free energy transduction of mitochondria and spermatozoa (100) has been reported, although the cells are not physically lysed. The peptide thus exhibits cytotoxic activity without disrupting the membrane. Channel formation is generally connected with an association of peptide monomers in a transmembrane orientation. However, magainin has been found to be monomeric in the lipid-bound state (101) and to be located mainly parallel to the bilayer surface (102,103) (Table 2). Several other results conflict with a classical channel model. The release of large dye molecules from potentialfree lipid vesicles, lipid flip-flop between the outer and inner layers of liposomes, and magainin translocation through the bilayer (74) cannot be explained by an ion channel model. It would appear that other driving forces besides a transmembrane potential are responsible for magainin-induced ‘‘holes,’’ the size of which ranges from those of ions to the size of small molecules. Transmembrane pores composed of phospholipids and magainin molecules (2, Fig. 3) were modeled (74) and detected by in-plane neutron scattering (75). Pore diameters reach 3 nm, about twice as large as the alamethicin channel. Because of the cationic charge carried by the peptide (Fig. 2), magainin binding to negatively charged membranes is high (Table 2). The adsorbed peptide pushes the lipid headgroups aside and thins the surface at the site of binding (104). The chain disorder caused by the bound peptide tends to spread over an ˚ in diameter. Enhanced surface expansion and curvature area as large as 100 A modulation lead to unfavorable strain between the outer and inner lipid layers. The outer membrane leaflet tends to exclude the surface-lying peptides and the magainin helix becomes aligned perpendicular to the membrane surface at high peptide concentration (105), and pore formation follows (106). Involvement of negatively charged lipids in the pore will stabilize the structure by reducing electrostatic repulsive forces between the peptides. In the pore state the outer and inner lipid layers form a continuum and peptides and lipids may be exchanged. Pore formation occurs at roughly the same concentration as that required for liposome leakage and cytolysis (105). This implies that membrane insertion and pore formation are also responsible for the cell lytic activity of magainin. The mechanism suggested is supported by the ability of magainin to induce positive curvature strain in planer bilayers (64). Pure lipid layers with a high negative curvature strain [PS, PE, cardiolipin (CL)] resist pore formation (56). Indeed, differences in the lipid composition of the inner membrane of gram-negative bacteria correlate well with the lytic effect. The membrane of Acinetobacter calcoacetisus containing a low proportion of PE and CL and being rich in negatively charged PG, which favors peptide accumulation, is most sensitive to magainin, whereas spheroplasts of Proteus vulgaris with a high PE and CL and low PG content show low magainin-induced lysis. Magainin-induced dye leakage from liposomes takes place at a peptide concentration of about 3 mol%. Based on weight values the critical magainin concen-

Amphipathic Helical Peptides

17

tration (81 g/mol lipid) is roughly equivalent to a detergent concentration that causes an increase in permeability (107). This fact points to a close biophysical relationship between the amphiphilic peptide and detergent. 3. Cecropin P1 Carpet A non-channel, non-pore mechanism has been suggested for the cecropins (44,108), dermaseptin (76), and pardaxin (109). These peptides disrupt the bilayer organization by a ‘‘carpetlike’’ mechanism (3, Fig. 3) (110). In this model strong binding of highly cationic peptide monomers, oriented parallel to the membrane surface with shallow penetration of the hydrophobic face into the nonpolar core, is followed by formation of peptide monolayers or bundles. High surface tension leads to disruption of bilayer packing and finally membrane collapse. Characteristic of this mechanism is a very high amount of bound peptide per lipid. Thus, cecropin P1 was described as exerting maximal membrane-disturbing activity at a peptide/lipid ratio of 1/7 in the outer leaflet (44). The 31-amino acid peptide cecropin P1 (Fig. 2) from the porcine small intestine (30) is not hemolytic. It is as potent as insect cecropins in inhibiting growth of gram-negative bacteria but is rather inactive against gram-positive bacteria (Table 1). The N-terminal peptide sequence is characterized by a high cationic charge density, while the C-terminus is more hydrophobic. The peptide forms an amphipathic helix over almost the entire length of the molecule (111). Cecropin P1 binds to phospholipid bilayers in a noncooperative manner, like insect cecropins. However, in contrast to the latter, cecropin P1 does not aggregate even at very high peptide concentration in the membrane, as confirmed by fluorescence energy transfer experiments (44), but remains oriented parallel to the membrane surface as shown by FTIR spectroscopy (112) (Table 2). Pore formation resulting in destruction of the energy gradient across the membrane has been suggested as the initial step in cell lysis induced by insect cecropins (113). The question of why porcine cecropin does not insert into the membrane may be answered in part by its particular distribution of charged and hydrophobic residues and rigid rod shape (Fig. 2). For the C-terminal hydrophobic moiety to insert into the membrane, a flexibility is required in the chain center, which, however, is provided only in the insect cecropins by proline and glycine. Furthermore, dipole orientation in the peptide helix makes such an insertion into a membrane with a negative potential inside the cell very unfavorable.

IV. SECONDARY EFFECTS AND PROSPECTIVE APPLICATIONS Besides the suggested main effects of membrane-permeabilizing peptides, other properties continue to emerge. Over and above the involvement of antimicrobial peptides in the prevention of infection, such peptides have been found to promote

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wound healing (9,114). In addition to evidence of antimicrobial synergy between different members of the magainin (100) family, magainin (115) and related model peptides (52,116) were observed to render bacteria more susceptible to classical lipophilic and amphiphilic antibiotics. The latter effect is related to the ability of cationic peptides to permeabilize the outer wall of gram-negative bacteria, thus improving uptake of antibiotics into the cell (6). Another property of potential therapeutic value is the ability of several of the cationic peptides to bind to bacterial lipopolysaccharides and to depress their toxic effects (6,117). Furthermore, a number of tumor-derived cell lines have been described to be very sensitive to magainin peptides (118–121) and cecropin analogs (122,123). Thus, the compounds bear the potential for the development of peptidic anticancer agents. On the other hand, the broad-spectrum histamine-releasing activity of antimicrobial peptides such as mastoparan (124) and magainin analogs (125) is not advantageous for purposes of drug development. Nevertheless, the activities of membrane-active peptides offer the hope that some compounds might be developed into agents of therapeutic relevance, in the treatment of skin and eye infections or skin carcinoma, for example. In 1998, Magainin Pharmaceuticals Inc. (USA) submitted an application with the U.S. Food and Drug Administration for a magainin analog, a topical antibiotic cream being evaluated for the treatment of infections in diabetic foot ulcers (Magainin Pharmaceuticals Inc. web site: http//www.magainin.com/home.htm). More recent reports on the uptake of membrane-active model peptides by endothelial cells at nonlytic concentration by a largely nonendocytotic mechanism (126) and the intracellular accumulation of the antibacterial buforin II without significant membrane damage but binding to DNA connected with rapid cell death (127) as well as binding of other membrane-penetrating amphipathic peptides to DNA (128) mark these compounds as potential tools for the transport of genetic material. Thus, the introduction of the respective DNAs or the expression of antibacterial peptides at the site of infection or of toxic peptides in carcinoma cells might open new approaches to therapy (129). Furthermore, the translocation phenomena of cationic peptides can also give a molecular basis for their putative direct interaction with G-proteins inside the cell (130), rendering such peptides valuable tools for studying and influencing G-protein-mediated signal transduction. A major challenge for development in all these areas is to translate the interesting properties of the peptides described here into clinically useful drugs.

ACKNOWLEDGMENTS I thank all members of our research group for their engaged work. The continuous financial support of the Deutsche Akadamie der Naturforscher Leopoldina is

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gratefully acknowledged. Heike Nikolenko is thanked for excellent technical assistance over many years and for her help in preparing the final version of this manuscript. I am grateful to John Dickson for critical reading of the manuscript.

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2 Bacterial Membrane as a Target for a Novel Class of Diastereomers of Cytolytic Peptides Yechiel Shai The Weizmann Institute of Science, Rehovot, Israel

I. CYTOLYTIC PEPTIDES Permeation of the cell wall membrane leading to cell death is a mechanism used by a large number of cytolytic polypeptides. The majority of these peptides can be classified into two main groups: (1) linear peptides, mostly helical, which do not contain cysteines, and (2) peptides with one or more disulfide bonds forming β-sheet or both β-sheet and α-helix structures (1). The most studied group contains host-defense short linear polypeptides (ⱕ40 amino acids). They vary considerably in sequence, chain length, hydrophobicity, and overall distribution of charges. The list includes (1) cytolytic peptides that are toxic to bacteria only, such as cecropins, isolated from the cecropia moth (2), and magainins (3) and dermaseptins (4), both isolated from the skin of frogs; (2) cytolytic peptides that are selectively cytotoxic to mammalian cells but not to bacteria, such as δ-hemolysin isolated from Staphylococcus aureus (5); and (3) cytolytic peptides that are not cell-selective, such as the bee venom melittin (6) and the neurotoxin pardaxin (7–9), which lyse bacteria and mammalian cells. Table 1 shows as an example the sequences and sources of several cytolytic peptides. Despite their broad spectrum of activity, most of the linear cytolytic peptides share a common structure in hydrophobic environments, namely, an amphipathic α-helix structure (10). In this structure, polar amino acids are arranged along one side of the helix as a consequence of 1,3 and 1,4 periodicities, and the 27

Hyalophora cecropia Hyalophora cecropia Porcine Xenopus laevis Xenopus laevis Xenopus laevis Xenopus laevis Phyllomedusa sauvagi Phyllomedusa sauvagi Phyllomedusa bicolor Phyllomedusa bicolor

Cecropin B Cecropin D

Cecropin P1 Magainin 1 Magainin 2 PGLa XPF Dermaseptin S1

Dermaseptin S2

Dermaseptin B1 Dermaseptin B2 Non-cell-selective Pardaxin Melittin LL-37 Moses sole fish Bee venom Human

Hyalophora cecropia

Source

G F F A L I P K I I S S P L F K T L L S A V G S A L S S S G G Q E-cooh G I G A V L K V L T T G L P A L I S W I K R K R Q Q-nh2 L L G D F F R K S K E K I G K E F K R I V Q R I K D F L R N L V P R T-cooh

KWKLFKKIEKVGQNIRDGIIKAGPAVAVVGQATQI A K-nh2 K W K V F K K I E K M G R N I R N G I V K A G P A I A V L G E A K A L-nh2 WNPFKELEKVGQRVRDAVISAGPAVATVAQATAL A K-nh2 S W L S K T A K K L E N S A K K R I S E G I A I A I Q G G P R-cooh G I G K F L H S A G K F G K A F V G E I M K S-cooh G I G K F L H S A K K F G K A F V G E I M N S-cooh G M A S K A G A I A G K I A K V A L K A L-nh2 G W A S K I G Q T L G K I A K V G L K E L I Q P K-cooh ALWKTMLKKLGTMALHAGKAALGAAADTISQG T Q-cooh ALWFTMLKKLGTMALHAGKAALGAAANTISQG T Q-cooh A M W K D V L K K I G T V A L H A G K A A L G A V A D T I S Q-nh2 G L W S K I K E V G K E A A K A A A K A A G K A A L G A V S E A V-nh2

Sequence

Sequences of Representative Bacteria-Selective and Non-Cell-Selective Cytolytic Peptides

Cell-selective Cecropin A

Peptide

Table 1

28

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Cytolytic Peptides

29

hydrophobic amino acids are arranged along the other side. This structure can be visualized schematically by using the Shiffer and Edmondson wheel projection (11). In this wheel, consecutive amino acids are 100° from each other such that each cycle contains 3.6 amino acids. Figure 1 shows the wheel projection of three peptides that represent the bacteria-specific cytolytic peptides dermaseptin and cecropin and a non-cell-selective peptide melittin. It is evident from the figure that there is no sequence homology between either the hydrophilic or the hydrophobic surfaces of these peptides, which might explain the difference in the spectra of their target cells. However, a major common feature for the bacteria-specific cytolytic peptides is the distribution of a relatively large number of positive charges along the hydrophilic face of their amphipathic helix. This net positive charge has been shown to be a prerequisite for selective binding to bacteria (discussed in the following paragraphs). Nevertheless, such charge distribution occurs also in non-cell-selective cytolytic peptides such as the human lytic peptide LL-37 (12). These observations suggest that other properties, rather than merely the amphipathic structure, contribute to the selective action of antibacterial peptides toward bacteria and not normal mammalian cells. Major questions concern what these properties are and whether a particular mechanism can be assigned to a particular biological function (i.e., antibacterial activity versus cytotoxicity to mammalian cells). This chapter focuses mainly on the structure and function of bacteria-selective cytolytic peptides.

Figure 1 Schiffer–Edmundson wheel projection of the N-terminal 22 amino acids of dermaseptin-S, cecropin-P, and melittin. Number 1 represents residue 1 of the peptides. Nonshaded areas indicate hydrophobic amino acids and dark areas indicate hydrophilic amino acids.

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30

II.

THE NEGATIVELY CHARGED OUTER SURFACE OF BACTERIA AS A TARGET FOR ANTIMICROBIAL PEPTIDES

Despite a significant number of studies aimed at understanding the function and structural requirements for the antibacterial activity of cytolytic peptides, their precise mode of action is yet not fully understood. Nevertheless, it has been suggested that peptide–lipid interactions, rather than receptor-mediated recognition processes, play a major role in their ability to lyse bacteria. This was demonstrated by the finding that analogs of cecropin and magainin, composed entirely of d-amino acids (enantiomers), possess antibacterial activity indistinguishable from that of the parent molecules (13–15). These enantiomers preserved the amphipathic α-helical structure of the wild-type peptides, a structure proposed to be requisite for their function. Another common feature found in native antimicrobial peptides is their net positive charge contributed by a large number of basic amino acids that are distributed along the hydrophilic face of the amphipathic α-helix (Fig. 1). This feature has been proposed to account for their preferential binding to bacteria and not to normal mammalian cells. One major difference between the outer surface of bacteria and normal mammalian cells is in their net charge. The outer surface of gram-negative bacteria contains lipopolysaccharides (LPSs), and that of gram-positive bacteria, acidic polysaccharides (teichoic acids), giving the surface of both gram-positive and gram-negative bacteria a negative charge (16). Therefore, the net positive charge of the antibacterial peptides facilitates their initial binding to the bacterial surface. In contrast, the outer leaflet of human erythrocytes (representative of normal mammalian cells) is composed predominantly of zwitterionic phosphatidylcholine (PC) and sphingomyelin phospholipids (17). Studies on the interaction of antimicrobial peptides with model phospholipid membranes revealed low affinity to zwitterionic phospholipids compared to acidic phospholipids. This has been demonstrated with cecropins (18,19), magainins (20–24), dermaseptins (25,26), and others (9,27). The low affinity of antimicrobial peptides to zwitterionic membranes might explain their inability to lyse erythrocytes.

III. MODE OF ACTION OF CYTOLYTIC PEPTIDES The amphipathic α-helical structure of cytolytic peptides permits simultaneous interaction with both the lipidic milieu of the membrane and the water-exposed surrounding, leading to permeation of the target cell. Two alternative models have been proposed to describe these interactions: (1) transmembrane pore formation via a ‘‘barrel stave’’ mechanism, and (2) membrane destruction/solubilization via a ‘‘carpet-like’’ mechanism.

Cytolytic Peptides

A.

31

Transmembrane Pore Formation via a ‘‘Barrel Stave’’ Mechanism

A ‘‘barrel stave’’ mechanism (28) describes a situation in which amphipathic α-helices insert into the membrane and form bundles. In these bundles the hydrophobic surfaces interact with the lipid core of the membrane and the hydrophilic surfaces point inward, producing a pore (right-hand panel of Fig. 2). Since these peptides can insert into the hydrophobic core of the membrane, their interaction with the target membrane is predominantly driven by hydrophobic forces. As a consequence, they can bind to both zwitterionic and charged phospholipid membranes. It was suggested that the ‘‘barrel stave’’ mechanism involves four major steps: (1) binding of the monomers to the membrane in an α-helical struc-

Figure 2 A cartoon illustrating the ‘‘barrel stave’’ (to the right) and ‘‘carpet-like’’ (to the left) models suggested for membrane permeation. In the carpet-like model the peptides are bound to the surface of the membrane with their hydrophobic surfaces (shaded area) facing the membrane and their hydrophilic surfaces (dark area) facing the solvent (step A). When a threshold concentration of peptide monomers is reached, the membrane breaks up into pieces (steps B and C). At this stage a transient pore is formed. (From Ref. 68.)

32

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ture; (2) molecular recognition between membrane-bound monomers, which leads to their assembly even at low surface density of the bound peptide; (3) insertion of at least two assembled monomers into the membrane to initiate the formation of a pore; and (4) progressive recruitment of additional monomers to increase the pore size. A prerequisite condition for this mechanism is that initial assembly of monomers on the surface of the membrane must occur before the peptide is inserted, since it is energetically unfavorable for a single amphipathic α-helix to traverse the membrane as a monomer. In the latter case the low dielectric constant and inability to establish hydrogen bonds will not allow the fatty acyl region of a lipid bilayer to be in direct contact with a polar surface of a single amphipathic α-helix. Two examples of cytolytic peptides that have been shown to insert into membranes via the barrel stave mechanism are alamethicin and pardaxin. Alamethicin is an amphipathic α-helical peptide containing 20 amino acids and produced by the fungus Trichoderma viride (29,30). The middle of the peptide and the C-terminus possess H-bonding patterns characteristic of a 310 helix (31) with a bend around Pro14. Alamethicin has been studied for over 20 years as a model for voltage-gated ion channels. The high concentration dependence of conduction and the multistep conductances seen in single-channel recordings were interpreted in terms of a barrel stave model for the channel pore (for reviews, see Refs. 32–34). Biophysical studies on the interaction of alamethicin with model membranes also support this mechanism (35). Pardaxin is an excitatory neurotoxin that has been purified from the Red Sea Moses sole, Pardachirus marmoratus (7,8), and from the peacock sole of the western Pacific, Pardachirus pavoninus (36) (reviewed in Ref. 37). Pardaxin is composed of 33 amino acids and adopts an amphipathic α-helical structure in hydrophobic environments. The peptide is cytolytic to both bacteria and mammalian cells (8,9,38). Similarly to alamethicin, pardaxin also has a helix–hinge– helix structure; the N-helix includes residues 7–11 and the C-helix includes residues 14–26. The helices are separated by a proline residue situated at position 13 similarly to alamethicin (39). Biophysical and functional studies with pardaxin and its analogs revealed that the peptide inserts into the membrane and specifically self-associates to form oligomers of different sizes, which support a barrel stave mechanism for its insertion and organization in the membranes (40,41) (reviewed in Ref. 37). B. Detergent-Like Membrane Solubilization via a ‘‘Carpet-Like’’ Mechanism The ‘‘carpet-like’’ mechanism (19,25) describes a situation in which amphipathic α-helical peptides initially bind onto the surface of the target membrane and cover it in a carpet-like manner (left-hand panel of Fig. 2). The target membrane can be permeated only after a threshold concentration has been reached. In con-

Cytolytic Peptides

33

trast to the barrel stave mechanism, the peptide does not insert into the hydrophobic core of the membrane but rather binds to the phospholipid headgroups (lefthand panel in Fig. 2). Initial interaction with the negatively charged target membrane is electrostatically driven, and therefore the active peptides are positively charged. The four steps possibly involved in this model are (1) preferential binding of positively charged peptide monomers to the negatively charged phospholipids, (2) laying of amphipathic α-helical monomers on the surface of the membrane so that the positive charges of the basic amino acids interact with the negatively charged phospholipid headgroups or water molecules, (3) rotation of the molecules leading to reorientation of the hydrophobic residues toward the hydrophobic core of the membrane, and (4) disintegration of the membrane by disruption of the bilayer curvature, leading to micellization. An initial step before the collapse of the membrane packing may include the appearance of transient holes in the membrane. Holes like these may enable the passage of low molecular weight molecules prior to complete membranal lysis. Such holes were described as a toroidal (or wormhole) model (23,42,43), which differs from the barrel stave model in that the lipid bends back on itself like the inside of a torus (step B in left-hand panel of Fig. 2). As seen in Figure 2, these holes may allow the passage of peptide molecules from the outer membrane into the inner membrane of, for example, gram-negative bacteria, in a process that may be referred to as selfpromoting uptake (44–47).

IV. ANTIBACTERIAL PEPTIDES KILL BACTERIA VIA A CARPET-LIKE MECHANISM Functional and structural studies with several antibacterial α-helical peptides support a ‘‘carpet-like’’ mechanism rather than a ‘‘barrel stave’’ mechanism for their mode of action. This conclusion is based on the following observations: 1. Various biophysical studies suggest that antibacterial peptides prefer to lie tangent to the surface of the membrane rather than take a transmembrane orientation. For example, one- and two-dimensional solid-state 15N NMR spectra of specifically 15N-labeled magainin 2 in oriented bilayer samples showed that the secondary structure of essentially the entire peptide is α-helix, immobilized by its interactions with the phospholipids and oriented parallel to the membrane surface (48). Another approach utilized peptides containing fluorescent probes that are sensitive to their environment. Using this approach, dermaseptin, cecropins (18,19,25,26,49), and the human LL-37 (69) antimicrobial peptides were found to lie on the surface of the membrane. In other studies, tryptophan-containing analogs of magainin were used, and the

Shai

34

results suggested that the orientation of the magainin 2 α-helix is parallel to the membrane surface (50). 2. In contrast to channel-forming peptides, most antimicrobial peptides do not self-associate in their membrane-bound state unless a threshold concentration has been reached (18,19,25,26,49). Furthermore, since antimicrobial peptides are highly positively charged amphipathic α-helices, it is energetically unlikely that they will remain monomeric within the hydrophobic core of the lipid membrane. The observed limited degree of aggregation may explain the formation of ion channels by insect cecropins (51) and magainin (52) in planar lipid bilayers. 3. Transient holes in the membrane are formed and are described by a toroidal (or wormhole) model for pore formation, which differs from the barrel stave model in that the lipid bends back on itself like the inside of a torus (23,42,43). These holes can be described as shown in Figure 2, left-hand panel. The bending requires a lateral expansion in the headgroup region of the bilayer.

V.

DIASTEREOMERS OF NON-CELL-SELECTIVE CYTOLYTIC PEPTIDES: A NOVEL GROUP OF ANTIMICROBIAL PEPTIDES

Numerous studies led to the conclusion that an amphipathic α-helix is a prerequisite structure for the biological activity of most linear lytic peptides. For example, previous studies revealed that incorporation of d-amino acids into the non-cellselective lytic peptide pardaxin preserved significant affinity to negatively charged phospholipid membranes but reduced affinity to zwitterionic ones (53). Studies on the structure and organization of the resulting diastereomers in the membrane-bound state revealed that they had reduced α-helical structure and could not penetrate and assemble in the membrane via a barrel stave mechanism. However, whereas pardaxin was lytic to both bacteria and normal mammalian cells, the diastereomers were devoid of biological activity (54). Similarly, the introduction of a single d-amino acid into the antibacterial peptide magainin, which is not lytic to normal mammalian cells, resulted in complete loss of its antibacterial activity (55). Despite the fact that pardaxin diastereomers have significant affinity to negatively charged membranes, they have only ⫹1 net positive charge compared to the very high net positive charge characteristic of antimicrobial peptides. This might explain the findings that pardaxin diastereomers were devoid of antimicrobial activity. In line with this assumption, we have synthesized a pardaxin analog in which the C-terminal glutamic acid was transamidated and the resulting analog had a net charge of ⫹5 (termed TApar, Table 2) (56). TApar has an α-helical structure (Fig. 3A) and is endowed with high antibacterial activity on gram-negative and gram-positive bacteria (Table 3) and with high hemolytic activity on human erythrocytes (Table 3). However, d-amino acids

Cytolytic Peptides

35

Table 2 Sequences and Designations of Diastereomers of Pardaxin, Melittin, and Model Peptides Peptide designation TAparc [d ]P7L18L19-TApar Melittinc [d ]-V5,8I17K21-melittin [d ]-L3,4,8,10-K3L9 [d ]-L3,4,8,10-K4L8 [d ]-L3,4,8,10-K5L7 [d ]-L3,4,8,10-K7L5

Sequencea,b GFFALIPKIISSPLFKTLLSAVGSAL S S S G G Q E-(nh2)2 GFFALIPKIISSPLFKTLLSAVGSAL S S S G G Q E-(nh2)2 GIGAVLKVLTTGLPALISWIKRK RQQ GIGAVLKVLTTGLPALISWIKRKRQ Q-nh2 K L L L L L K L L L L K-nh2 K L L L K L L L K L L K-nh2 K L L L K L K L K L L K-nh2 K K L L K L K L K L K K-nh2

a

Underlined and bold amino acids were substituted with their d enantiomers. E-(nh2)2 stands for glutamic acid in which the two ECOOH carboxylate groups were modified to two ECOEnhEch2ch2Enh2 groups by transamination with diamino ethanol. c Underlined sequences designate the N- and C-helices, respectively. Source: Refs. 56, 57, 64. b

incorporated into TApar dramatically reduced its α-helical structure in 40% trifluoroethanol–water (Fig. 3A) (56). This in turn reduced the hemolytic activity of the diastereomeric analogs (Table 3), which indicates the importance of this structure in the cytotoxicity of the peptide to mammalian cells. In contrast, the amphipathic α-helical structure seems not to be crucial for antibacterial activity, since with most of the bacteria tested there was no significant decrease in the antibacterial activity of the peptides (Table 3). The lack of a significant α-helical structure should prevent the diastereomers from inserting and forming a transmembrane pore, and hence a barrel stave mechanism is not favored as their mode of action. The effect of the diastereomers is total lysis of the bacterial wall, as revealed by electron microscopy (Fig. 4). The properties of the diastereomers suggest that they act via a carpet-like mechanism. To test the generality of the diastereomer concept we used native melittin as a second example (57). Melittin is highly toxic to both bacteria and mammalian cells. d-Amino acids were incorporated into the N- and C-terminal helices of melittin in a manner that would cause the greatest disruption to these helices (Table 2). Indeed, the diastereomers had a dramatically reduced α-helical structure (Fig. 3B). The low residual α-helical content can be attributed to the limited effect of a single d-amino acid on the α-helical structure (58). As found with pardaxin, the disruption of the α-helical structure of melittin totally abolished

36

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Figure 3 (A) Circular dichroic spectra of pardaxin and its diastereomer. (B) CD spectra of melittin and its diastereomer. Spectra were taken at peptide concentrations of 0.8 ⫻ 10⫺5 –2.0 ⫻ 10⫺5 M in 40% TFE/water. (——) Native molecules; (- - -) diastereomers. (Modified from Refs. 56 and 57.)

Cytolytic Peptides

37

Table 3 Minimal Inhibitory Concentration (µM) and Hemolytic Activity of Melittin, Pardaxin, Their Diasteriomers, and Model Peptidesa Minimal inhibitory concentration (µM)

Peptide designation Melittin [d]-V5,8,I17,K21-melittin TApar [d ]P7L18L19-TApar [d ]-L3,4,8,10-K3L9 [d ]-L3,4,8,10-K4L8 [d ]-L3,4,8,10-K5L7 [d ]-L3,4,8,10-K7L5 Dermaseptin-S Tetracycline

E. coli (D21)

A. calcoaceticus (Ac11)

B. megaterium (Bm11)

B. subtilis (ATCC 6051)

Hemolysis of RBCs (at 20 µM)

5 12 3 6 9 3.5 7 80 6 1.5

20 12 3 6 20 4 20 200 3 1.5

0.3 0.8 0.8 0.9 0.7 0.4 0.2 1 0.5 1.2

0.4 3.5 0.5 3 1.1 0.5 1 100 4 6.5

100% 0% 100% 0% 10% 0% 0% 0% 0% 0%

a

Results are the mean of three independent experiments, each performed in duplicate, with a standard deviation of 20%. Source: Refs. 56, 57, 64.

the hemolytic activity of the diastereomers but preserved the antibacterial activity (Table 3). The interaction of pardaxin, melittin, and their diastereomers with model phospholipid membranes was examined in order to elucidate the basis of the selective lytic ability of the diastereomers against bacteria. Pardaxin (37,38) and melittin (59–63) bind strongly to both zwitterionic and negatively charged membranes. The α-helical regions of both peptides are predominantly hydrophobic and contain only a few charged amino acids. Therefore, the binding forces between these helices and phospholipid membranes consist mainly of hydrophobic interactions between the hydrophobic faces of the amphipathic helices and the lipid constituent of the membrane. In contrast to native pardaxin and melittin, which bind strongly to both negatively charged and zwitterionic phospholipids, their diastereomers (56,57) bind and permeate strongly only negatively charged phospholipids. Thus, electrostatic interactions appear to play an important role in the initial binding of the diastereomers to negatively charged membranes. The disruption of the α-helical structure of pardaxin and melittin, which prevents manifestation of hydrophobic forces, is probably responsible for the inability of the diastereomers to bind zwitterionic phospholipids. Interestingly, despite the disruption of its α-helical structure, the all-l form and the diastereomer of either pardaxin or melittin have similar activities on negatively charged membrane. In the case of melittin, the wild type and the diastereomer exhibited similar partition coefficients with negative cooperativity, had the same potency for dissipating

38

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Figure 4 Electron micrographs of negatively stained E. coli untreated and treated with the various peptides at 80% of their MIC. Panels A,F, control bacteria; panel B, treated with [d ]-L3,4,8,10-K3L9; panels C,G, treated with [d ]-L3,4,8,10-K4L8; panels D,H, treated with [d ]-L3,4,8,10-K5L7; panel E, treated with [d ]-L3,4,8,10-K7L5. (From Ref. 57.)

Cytolytic Peptides

39

diffusion potentials, and penetrated to the same depth into the hydrophobic core of a mixture of phosphatidylcholine (PC) and phosphatidylglycerol (PG) vesicles, as seen in the tryptophan-quenching experiments using brominated phospholipids (57). To further investigate whether a balance between hydrophobicity and a net positive charge may be a sufficient criterion for selective bacterial lysis, and to gain insight into the mechanism underlying this effect, four diastereomers of linear and short (12 amino acids long) model peptides composed of varying ratios of lysine to leucine were synthesized (Table 2) (64). The location of d-amino acids remained constant in all peptides which was constructed for maximum disruption of α-helical structure. A direct correlation was found between hydrophobicity (measured according to Ref. 65) and the retention time of the peptides using reversed-phase high-performance liquid chromatography (RP-HPLC). This suggests that structure does not significantly contribute to overall hydrophobic interactions with the stationary phase. This supports our hypothesis that the effect of the helical structure was eliminated and therefore permitted the study of only two parameters, namely, hydrophobicity and net positive charge, by varying the ratio of leucine and lysine. Similarly to what has been found with the diastereomers of pardaxin and melittin, circular dichroism (CD) spectroscopy revealed that the model diasteriomers were devoid of α-helical structure (64). Nevertheless, they exhibited potent antibacterial activity similar to or greater than that of native antibacterial peptides such as dermaseptin S or the antibiotic drug tetracycline (Table 3). Moreover, their hemolytic activity was reduced or abolished (64). Interestingly, [d ]L3,4,8,10-K3L9, which is devoid of α-helical structure, had some hemolytic activity. This could indicate that the balance between hydrophobicity and positive charge compensates for the amphipathic α-helical structure. However, increasing the positive charge drastically reduced the hemolytic activity while antibacterial activity was preserved, demonstrating that the amphipathic α-helical structure is not required for antibacterial activity. The interaction of the model diastereomers with both negatively charged and zwitterionic phospholipid membranes correlated with their cytolytic activity on erythrocytes and bacteria. Similarly to what has been found with native antimicrobial peptides and diastereomers of melittin and pardaxin, the nonhemolytic diastereomers could bind strongly only to negatively charged membranes. Therefore, electrostatic interactions between the positively charged diastereomers and the negatively charged phospholipid membranes seem to have an important role in initial interactions and selectivity, but biological activity appears to be driven by the hydrophobic interactions between the nonpolar amino acids and the hydrophobic core of the lipid bilayer. High hydrophobicity may force the immersion of the peptide into the hydrophobic core of the lipid bilayer, regardless of the phospholipid headgroup, thereby permeabilizing membranes of eukaryotics and prokaryotics, as in the case of [d ]-L3,4,8,10-K3L9. On the other hand, high positive

40

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charge may prevent an efficient immersion step, so that the peptide interacts predominantly with the negatively charged lipid headgroup as in the case of [d ]-L3,4,8,10-K7L5 (Table 2). In contrast to these results, when the α-helical structure of magainin was disrupted by the introduction of three d-amino acids, the resulting diastereomer had no antibacterial activity (55), even though the net positive charge was not altered. Thus, an optimal balance that already exists between the α-helical structure, hydrophobicity, and net positive charge of native magainin allows selective antibacterial activity, and any change in one of these properties could cause a loss in magainin’s antibacterial activity. In contrast, hydrophobicity appears to play a major role in compensating for the loss of α-helical structure in melittin, pardaxin, and the diastereomers studied by Chen et al. (55). Diastereomeric peptides should have several advantages over known alll- or all-d-amino acid antibacterial peptides: 1. The peptides should lack the diverse pathological and pharmacological effects induced by α-helical cytolytic peptides. For example, Staphylococcus δ-toxin, the antibacterial peptide alamethicin, cobra direct lytic factor, and pardaxin exert several histopathological effects on various cells due to pore formation and activation of the arachidonic acid cascade. However, pardaxin diastereomers do not exert these activities (54). In addition, many amphipathic α-helical peptides bind to calmodulin and elicit several cell responses, and even all-d-amino acid α-helices, including melittin, have similar activity (66). Diasteriomers with disrupted α-helical structure are not expected to bind to calmodulin. 2. Local d-amino acid substitution would result in controlled clearance of the antibacterial peptides by proteolytic enzymes, as opposed to the total protection acquired by complete d-amino acid substitution (13). Total resistance of a lytic peptide to degradation is disadvantageous for therapeutic use. Furthermore, the antigenicity of short fragments containing d,l-amino acids is dramatically altered compared to their wholly l- or d-amino acid parent molecules (67). 3. Total inhibition of bacterial growth induced by the diastereomers is associated with total lysis of the bacterial wall, as shown by electron microscopy (Fig. 4). Therefore, bacteria might not easily develop resistance to drugs that trigger such a destructive mechanism. In summary, the results obtained with pardaxin, melittin, and the model peptide diastereomers indicate that neither a specific sequence, length, or position of d-amino acids is required for a polypeptide to have antibacterial activity. However, these factors seem to be more crucial for cytotoxicity toward mammalian cells. This novel family of antibacterial peptides can not act on the bacterial

Cytolytic Peptides

41

membrane via a barrel stave mechanism but via a carpet-like mechanism. Our results indicate that only modulating the hydrophobicity and net positive charge of linear cytotoxic polypeptides is sufficient in the design of a repertoire of potent antibacterial diastereomeric polypeptides for the treatment of infectious diseases.

REFERENCES 1. Boman HG. Peptide antibiotics and their role in innate immunity. Annu Rev Immunol 1995; 13:61–92. 2. Steiner H, Hultmark D, Engstrom A, Bennich H, Boman HG. Sequence and specificity of two antibacterial proteins involved in insect immunity. Nature 1981; 292: 246–248. 3. Zasloff M. Magainins, a class of antimicrobial peptides from Xenopus skin: isolation, characterization of two active forms, and partial cDNA sequence of a precursor. Proc Natl Acad Sci USA 1987; 84:5449–5453. 4. Mor A, Nguyen VH, Delfour A, Migliore SD, Nicolas P. Isolation, amino acid sequence, and synthesis of dermaseptin, a novel antimicrobial peptide of amphibian skin. Biochemistry 1991; 30:8824–8830. 5. Dhople VM, Nagaraj R. δ-Toxin, unlike melittin, has only hemolytic activity and no antimicrobial activity: rationalization of this specific biological activity. Biosci Rep 1993; 13:245–250. 6. Habermann E, Jentsch J. Hoppe Seyler’s Z Physiol Chem 1967; 348:37–50. 7. Lazarovici P, Primor N, Loew LM. Purification and pore-forming activity of two hydrophobic polypeptides from the secretion of the Red Sea Moses sole (Pardachirus marmoratus). J Biol Chem 1986; 261:16704–16713. 8. Shai Y, Fox J, Caratsch C, Shih YL, Edwards C, Lazarovici P. Sequencing and synthesis of pardaxin, a polypeptide from the Red Sea Moses sole with ionophore activity. FEBS Lett 1988; 242:161–166. 9. Oren Z, Shai Y. A class of highly potent antibacterial peptides derived from pardaxin, a pore-forming peptide isolated from Moses sole fish Pardachirus marmoratus. Eur J Biochem 1996; 237:303–310. 10. Segrest JP, De LH, Dohlman JG, Brouillette CG, Anantharamaiah GM. Amphipathic helix motif: classes and properties. Proteins 1990; 8:103–117. 11. Schiffer M, Edmundson AB. Use of helical wheels to represent the structures of protein and to identify segments with helical potential. Biophys J 1967; 7:121–135. 12. Agerberth B, Gunne H, Odeberg J, Kogner P, Boman HG, Gudmundsson GH. FALL-39, a putative human peptide antibiotic, is cysteine-free and expressed in bone marrow and testis. Proc Natl Acad Sci USA 1995; 92:195–199. 13. Wade D, Boman A, Wahlin B, Drain CM, Andreu D, Boman HG, Merrifield RB. All-d amino acid-containing channel-forming antibiotic peptides. Proc Natl Acad Sci USA 1990; 87:4761–4765. 14. Bessalle R, Kapitkovsky A, Gorea A, Shalit I, Fridkin M. All-d-magainin: chirality, antimicrobial activity and proteolytic resistance. FEBS Lett 1990; 274:151– 155.

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32. Woolley GA, Wallace BA. Model ion channels: gramicidin and alamethicin. J Membr Biol 1992; 129:109–136. 33. Sansom MS. Alamethicin and related peptaibols: model ion channels. Eur Biophys J 1993; 22:105–124. 34. Cafiso DS. Alamethicin: a peptide model for voltage gating and protein-membrane interactions. Annu Rev Biophys Biomol Struct 1994; 23:141–165. 35. Rizzo V, Stankowski S, Schwarz G. Alamethicin incorporation in lipid bilayers: a thermodynamic study. Biochemistry 1987; 26:2751–2759. 36. Thompson SA, Tachibana K, Nakanishi K, Kubota I. Melittin-like peptides from the shark-repelling defense secretion of the sole Pardachirus pavoninus. Science 1986; 233:341–343. 37. Shai Y. Pardaxin: channel formation by a shark repellant peptide from fish. Toxicology 1994; 87:109–129. 38. Shai Y, Bach D, Yanovsky A. Channel formation properties of synthetic pardaxin and analogues. J Biol Chem 1990; 265:20202–20209. 39. Zagorski MG, Norman DG, Barrow CJ, Iwashita T, Tachibana K, Patel DJ. Solution structure of pardaxin P-2. Biochemistry 1991; 30:8009–8017. 40. Rapaport D, Shai Y. Interaction of fluorescently labeled pardaxin and its analogues with lipid bilayers. J Biol Chem 1991; 266:23769–23775. 41. Rapaport D, Danin M, Gazit E, Shai Y. Membrane interactions of the sodium channel S4 segment and its fluorescently-labeled analogues. Biochemistry 1992; 31: 8868–8875. 42. Ludtke SJ, He K, Heller WT, Harroun TA, Yang L, Huang HW. Membrane pores induced by magainin. Biochemistry 1996; 35:13723–13728. 43. Matsuzaki K, Murase O, Fujii N, Miyajima K. An antimicrobial peptide, magainin 2, induced rapid flip-flop of phospholipids coupled with pore formation and peptide translocation. Biochemistry 1996; 35:11361–11368. 44. Sawyer JG, Martin NL, Hancock RE. Interaction of macrophage cationic proteins with the outer membrane of Pseudomonas aeruginosa. Infect Immun 1988; 56:693–698. 45. Piers KL, Brown MH, Hancock RE. Improvement of outer membrane-permeabilizing and lipopolysaccharide-binding activities of an antimicrobial cationic peptide by C-terminal modification. Antimicrob Agents Chemother 1994; 38:2311–2316. 46. Piers KL, Hancock RE. The interaction of a recombinant cecropin/melittin hybrid peptide with the outer membrane of Pseudomonas aeruginosa. Mol Microbiol 1994; 12:951–958. 47. Falla TJ, Karunaratne DN, Hancock REW. Mode of action of the antimicrobial peptide indolicidin. J Biol Chem 1996; 271:19298–19303. 48. Bechinger B, Zasloff M, Opella SJ. Structure and orientation of the antibiotic peptide magainin in membranes by solid-state nuclear magnetic resonance spectroscopy. Protein Sci 1993; 2:2077–2084. 49. Gazit E, Shai Y. Structure and orientation of the mammalian antibacterial peptide cecropin P1 within phospholipid membranes: ATR-FTIR study. Presented at 40th Annual Meeting of the Biophysical Society, Baltimore, 1996. 50. Matsuzaki K, Murase O, Tokuda H, Funakoshi S, Fujii N, Miyajima K. Orientational and aggregational states of magainin 2 in phospholipid bilayers. Biochemistry 1994; 33:3342–3349. 51. Christensen B, Fink J, Merrifield RB, Mauzerall D. Channel-forming properties of

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Shai cecropins and related model compounds incorporated into planar lipid membranes. Proc Natl Acad Sci USA 1988; 85:5072–5076. Cruciani RA, Barker JL, Durell SR, Raghunathan G, Guy HR, Zasloff M, Stanley EF. Magainin 2, a natural antibiotic from frog skin, forms ion channels in lipid bilayer membranes. Eur J Pharmacol 1992; 226:287–296. Pouny Y, Shai Y. Interaction of d-amino acid incorporated analogues of pardaxin with membranes. Biochemistry 1992; 31:9482–9490. Abu RS, Bloch SE, Shohami E, Trembovler V, Shai Y, Weidenfeld J, Yedgar S, Gutman Y, Lazarovici P. Pardaxin, a new pharmacological tool to stimulate the arachidonic acid cascade in PC12 cells. J Pharmacol Exp Ther 1998; 287:889–896. Chen HC, Brown JH, Morell JL, Huang CM. Synthetic magainin analogues with improved antimicrobial activity. FEBS Lett 1988; 236:462–466. Shai Y, Oren Z. Diastereoisomers of cytolysins, a novel class of potent antibacterial peptides. J Biol Chem 1996; 271:7305–7308. Oren Z, Shai Y. Selective lysis of bacteria but not mammalian cells by diastereomers of melittin: structure-function study. Biochemistry 1997; 36:1826–1835. Rothemund S, Beyermann M, Krause E, Krause G, Bienert M, Hodges RS, Sykes BD, Sonnichsen FD. Structure effects of double d-amino acid replacements: a nuclear magnetic resonance and circular dichroism study using amphipathic model helices. Biochemistry 1995; 34:12954–12962. Batenburg AM, Hibbeln JC, Verkleij AJ, De Kruijff B. Melittin induces HII phase formation in cardiolipin model membranes. Biochim Biophys Acta 1987; 903:142–154. Batenburg AM, Hibbeln JC, De Kruijff B. Lipid specific penetration of melittin into phospholipid model membranes. Biochim Biophys Acta 1987; 903:155–165. Batenburg AM, Van Esch JH, De Kruijff B. Melittin-induced changes of the macroscopic structure of phosphatidylethanolamines. Biochemistry 1988; 27:2324–2331. Beschiaschvili G, Seelig J. Melittin binding to mixed phosphatidylglycerol/phosphatidylcholine membranes. Biochemistry 1990; 29:52–58. Dufourcq J, Faucon JF. Intrinsic fluorescence study of lipid-protein interactions in membrane models. Biochim Biophys Acta 1977; 467:1–11. Oren Z, Hong J, Shai Y. A repertoire of novel antibacterial diastereomeric peptides with selective cytolytic activity. J Biol Chem 1997; 272:14643–14649. Eisenberg D. Three-dimensional structure of membrane and surface proteins. Annu Rev Biochem 1984; 53:595–623. Fisher PJ, Prendergast FG, Ehrhardt MR, Urbauer JL, Wand AJ, Sedarous SS, McCormick DJ, Buckley PJ. Calmodulin interacts with amphiphilic peptides composed of all d-amino acids. Nature 1994; 368:651–653. Benkirane N, Friede M, Guichard G, Briand JP, Van RM, Muller S. Antigenicity and immunogenicity of modified synthetic peptides containing d-amino acid residues. Antibodies to a d-enantiomer do recognize the parent l-hexapeptide and reciprocally. J Biol Chem 1993; 268:26279–26285. Oren Z, Shai Y. Mode of action of linear amphipathic α-helical antimicrobial peptides. Biopolymers 1999; 47:451–463. Oren Z, Lerman JC, Gudmundsson GH, Agerberth B, Shai Y. Stucture and organization of the human antimicrobial peptide LL-37 in phospholipid membranes: relevance to the molecular basis for its non-cell-selective activity. Biochem J 1999; 341: 501–513.

3 Bee Venom Toxicity Synergistic Action of Melittin on Interfacial Hydrolysis by Phospholipase A2 Yolanda Cajal University of Barcelona, Barcelona, Spain

Mahendra Kumar Jain University of Delaware, Newark, Delaware

I. INTRODUCTION Self-defense strategies, such as venomous sting or bite, are used by a variety of organisms. Insects like bees, yellow jackets, and other wasps inject about 100– 150 µg (dry weight) of venom in a single sting (1). Yellow jackets and other wasps can inflict multiple stings, but during its lifetime a bee inflicts only one sting, during which it leaves its venom sac, from which 95% of the venom is delivered within 20 s, at the site. Although a European honeybee (Apis mellifera) has more venom of higher potency, the ferocity of Africanized honeybees is in the multiple stings delivered by a swarm. Severity of the toxic effect of bee venom varies widely within the human population. Although rarely lethal, the inflammatory effects of the venom spread well beyond the site of the sting. Single stings often cause edema and local inflammation that can last for several days. In a human subpopulation, multiple stings can cause anaphylactic shock. Venoms of bees and wasps contain numerous biologically active substances that elicit inflammation and immune response. Two of the major components (2– 4), PLA2 (12% dry weight in bee venom) and up to 50% dry weight melittin (also spelled mellitin and mellittin) account for most of the cellular events leading to toxicity (Table 1). Hyaluronidase, apamin, mast cell degranulating peptide, histamine, and other amines present in the venom probably modulate the cellular responses initiated by PLA2 and melittin. The toxic effect of bee venom cannot 45

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46 Table 1 Toxicity-Related Cellular Responses of Bee Venom Components Function Hydrolysis of phospholipids Phospholipid exchange Cytolysis Interleukin production Degranulation in mast cells Antigenicity Allergenicity Affinity for neurotoxin receptor

PLA2

Melittin

Yes No No Yes Yes Yes Yes Yes

No Yes Weak Weak Yes Weak Weak Weak

be attributed to melittin-induced hemolysis, since lethal intravenous doses do not cause harm to circulating erythrocytes (5). Similarly, the inhibitory effect of apamin on the calcium-activated potassium channel does not account for the venom toxicity. In this chapter we examine the significance of cosecretion of PLA2 and melittin, which together account for most of the cellular responses to bee venom (Table 1). A functional synergism between the effects of these two components in the target tissue is implicated by enhanced PLA2 activity in the presence of melittin (6–8). As its basis we have shown that melittin promotes presentation of phospholipid substrate on excess vesicles for the interfacial catalysis by PLA2 bound essentially irreversibly to the product-containing vesicles (9). Thus a rapid substrate replenishment, through melittin contact with unhydrolyzed vesicles, leads to an apparent increase in the observed rate and extent of hydrolysis. We estimate that four melittin molecules form a contact. The amounts of PLA2 and melittin in venom from a single bee sting are about 0.2 and 10 nmol, respectively. Thus, a 50-fold molar excess of melittin over PLA2 could spread the initial hydrolytic damage to several neighboring cells by promoting substrate replenishment. This could be a critical step for spreading the initial rapid hydrolytic damage at the site of the sting to the blood capillaries and as a prelude to the downstream responses of the venom.

II.

THE ONSET OF PATHOPHYSIOLOGY BY PLA2

A.

Inflammatory Effects

Inflammation is the cumulative effect of a variety of responses of the inflammatory and immune cells attracted to the site of a sting by a range of chemical mediators. Although such responses are also seen under conditions related to

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infection and allergic reactions, the underlying interactions are only partially delineated in all such cases. As a general consensus view, such disorders are orchestrated by neutrophils and lymphocytes by releasing mediators in response to the initial assault, which may involve binding of a certain causative agent or damage by hydrolytic enzymes such as proteases and phospholipases (10). At this level of detail, the pathophysiology of bee venom toxicity is probably analogous to that of other inflammatory disorders. As attested by the presence of PLA2 in venoms of scorpions, lizards, and snakes, its hydrolytic action could be an important first step in venom toxicity because the products are both metabolic precursors for signals and modulators of cellular responses. Lysophospholipid is a potent chemotactic factor for human monocytes (11), and it is also a precursor for platelet activating factor. The second product, the unsaturated fatty acid from the sn-2 position of phospholipids, are weak chemoattractants for inflammatory cells. However, rapid enzyme-catalyzed oxidative metabolism of arachidonate, and possibly other polyunsaturated fatty acids, generate highly potent signal molecules collectively known as eicosanoids. Since arachidonate is not found free under most conditions, its release from phospholipids is a rate-limiting obligatory step for initiating cellular responses through eicosanoids. The enzymes of the eicosanoid pathway are normally present on the cytoplasmic side of certain differentiated cells. However, eicosanoids also appear to be produced in response to exogenous PLA2, presumably because arachidonate is taken up by these cells. Cytoplasmic eicosanoid production is also triggered by inflammatory cells activated by histamine and other chemoattractants. For example, both PLA2 and melittin promote mast cell degranulation (7,12) and the release of other mediators (13–15). In ras-transformed cells melittin induces Ca2⫹ influx (16). In turn, as a trigger for a second-phase response on the time scale of hours, eicosanoids and other messengers promote expression of protein modulators such as interleukins and acute-phase proteins (such as 14 kDa group II PLA2) whose secretion potentiates inflammation by inducing an IgE-independent degranulation in mast cells (17). In short, the vasoactive eicosanoids cause edema at the site of a bee sting by increasing the influx of fluid and inflammatory cells from the blood. The chemoattractant eicosanoids and the signaling molecules produced by inflammatory cells initiate the phagocytic and immune responses. They promote production of active oxygen species by neutrophils and mediate antibody-independent mast cell degranulation and histamine release. B. Antigenicity and Allergenicity Both PLA2 and melittin are antigenic as well as allergenic. Lymphocytes and other immune cells attracted to the inflammation site recognize the venom anti-

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gens. Production of weak IgG against PLA2 is a common response, although a human subpopulation also produces IgE (18). PLA2 has several epitopes (19); however, catalytically inactive PLA2, in which the catalytic histidine has been mutated, does not produce antibodies (20). Allergic reactions to bee venom are not rare. In about 3% of the human population a second sting can lead to anaphylactic shock. Even though the pathogenic mechanisms for allergy are not fully understood, severe systemic reactions in some patients have been linked to increased levels of IgE antibodies to components of bee venom, to a high level of release of mediators (13), and to a different profile of cytokine production from T-cells compared to nonallergic individuals (21). Tissue mast cells are attracted to the inflammation site by the released chemoattractants after PLA2 action on membranes; these cells play an important role in anaphylactic shock, producing not only histamine but also other preformed inflammatory mediators. Mast cell degranulation is promoted not only by PLA2 (14) and melittin (4,12), but also by apamine, mast cell degranulating peptide, and other amines found in bee venom. In addition, IgE-dependent degranulation is also induced when antigens bound to receptors in the surface of mast cells are recognized by IgE. It is not clear whether melittin and PLA2 bind directly to such receptors on immune cells. Melittin induces an IgE response in about one-third of bee venom allergic patients (22). Melittin structure has both a T-cell epitope, located in residues 7– 19, and a B-cell epitope in the hydrophilic C-terminus, residues 20–26 (23). By using melittin and several analogs it has been established that two structural elements of the peptides correlate with their immunogenicity: the peptide length and the presence of cationic residues in the C-terminus. A good correlation of the immunogenicity of the peptides is found with their propensity to self-associate and to bind to cell membranes, suggesting that the immunogenicity of melittin has the same physical basis as its propensity to bind cell membranes (24). To recapitulate, the pathophysiology of bee venom is most probably related to the initial hydrolytic event at the site of sting. Melittin amplifies the effect by presenting excess substrate from the membranes of neighboring cells to the membrane of a cell with bound PLA2. The products of hydrolysis produce powerful chemoattractants for inflammatory cells that unleash a cascade of cellular signaling events with damaging consequences for the whole organism. Note that a comparable sequence of cellular events is elicited not only by other allergens but also by invading viruses, bacteria, and parasites. Selectivity of the underlying pathways remains to be established; however, it appears that at least some of these are controlled at the expression level in a differentiation-dependent way. C. Phospholipase-Activating Peptides Melittin enhances cellular PLA2 activity (25,26) as well as the activity of other lipolytic enzymes (27,28). Polymyxins (29) and mastoparans (12) also activate

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PLA2, mediate phospholipid exchange (30,31) (see also Chap. 4), and promote release reactions in inflammatory cells. Several other phospholipase-activating peptides and proteins have also been described (32–34). However, in most cases alternative mechanisms are not ruled out; for example, cellular PLA2 is activated by leakage of Ca2⫹ into the cytoplasmic compartment (16,35). Activation of PLA2 and eicosanoid generation in mammalian cells have been correlated to an increased level of PLAP, a 28 kDa PLA2-activating protein (34,36–38) that was originally identified on the basis of its immunological cross-reactivity with antibodies to melittin (39). PLAP also activated PLA2 in vesicles (39). A synthetic PLAP peptide that spans a region of homology between natural PLAP and melittin showed the same activity as the whole protein in activating PLA2 from human monocytes, and it was more effective than melittin (37).

III. MELITTIN MEDIATES DIRECT PHOSPHOLIPID EXCHANGE BETWEEN VESICLES A.

Melittin at the Interface

Melittin is a cationic amphiphilic peptide (Fig. 1). It exists as an almost unstructured monomer in dilute aqueous solution and in media of low ionic strength (40–42). At higher concentrations, in solutions of high ionic strength or high pH, it acquires a bent α-helical structure. It also assumes a variety of higher order structures and aggregated forms depending on the pH, salt concentration, polarity, lipid composition, and peptide/lipid ratio (43–46). The structural basis for melittin is not established; however, the emerging consensus is that the conformation, orientation, and aggregation of melittin at the interface and in the bilayer play an important role in eliciting the functional forms. Given the cationic amphipathic character of melittin, electrostatic and hydrophobic effects play a role in determining the equilibrium between the various conformers of melittin. The plasticity of the melittin structure is attested by an ensemble of local minimum energy conformers for the truncated sequence of (1-20) melittin (47,48), which is mostly hydrophobic (Fig. 1). If only the hydrophobic effects are considered for the folding energetics, over 300 (⫽k) conformers are generated within the 5 kcal range of ⫺90 to ⫺86 kcal/mol. Within this ensemble of con-

Figure 1 The primary sequence of melittin. The first 20 residues at the N-terminus appear to dominate the folding in the hydrophobic environment. The cationic C-terminus appears to remain unstructured in the aqueous phase.

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formers, the φ and Ψ values of only a few residues differ, yet they remain within the permitted torsion range. For understanding the behavior of melittin, as a heuristic guide we choose three representative structures from this ensemble to speculate on the functional states of melittin. As shown in Figure 2a, the structure k99 is a helix disrupted by Pro-14. On the other hand, a right-angled helix (k25) is generated with a change in the torsion angles for Leu-13: from (⫺55, ⫺63) in k99 to (⫺140,⫹69) in k25 (Fig. 2b). A set of more compact structures is formed by an additional change at Leu-6: from (⫺64, ⫺41) for k99 to (⫺87, ⫹82) for k246. The only other significant departure in the calculated structures is seen in Gly-12. The NMR evidence also suggests significant departures from the extended helical form of melittin, and the six polar residues at the C-terminus seem to remain disordered (49–52). In short, considering the range of near isoenergetic structures generated for the truncated melittin, a preference for any of the forms will ultimately depend on the energetic contributions from the local environment and the steric factors. We believe that the distribution of the polar residues (Lys-7, Thr-10, Thr-11, Ser-18, and Lys-Arg-Lys-Arg-Asn-Asn at the C-terminus) on the hydrophobic sequence (Fig. 1) determines the functional forms of melittin (9). Operationally, steps leading to the two functional states of melittin at the interface are outlined in Scheme 1. At low mole fractions, melittin bound to the anionic interface forms vesicle–vesicle contacts that promote a direct and rapid exchange of phospholipids between the outer layers of the vesicles in contact, presumably through the contact-forming Mc form. For the discussion below, we suggest that an aggregated form of k246 type of structure(s) of melittin at the anionic phospholipid bilayer interface mediates vesicle–vesicle contacts capable of phospholipid exchange (Fig. 3). On the other hand, the pore-forming MP form, formed at higher mole percent melittin in zwitterionic vesicles, promotes leakage and possibly fusion and disruption of the bilayer. The amphipathic bent helix forms of melittin, k99 and k25, are well suited for the formation of transmembrane pores at a zwitterionic interface (Fig. 2). B. Melittin Binds to Anionic and Zwitterionic Vesicles The apparent dissociation constant for melittin at the anionic interface is at least 10 times lower than the value of about 30 µM at zwitterionic interfaces (9,53,54). Under conditions where the total lipid concentration is more than 200 µM, most of the melittin will be at the interface (55). It is not clear whether the stoichiometry of phospholipid to melittin at the interface is constant or depends on the presence of the negative charge at the interface. Depending on the conditions, about 3–30 phospholipid molecules are required for the binding of a melittin molecule. Spectroscopic properties suggest the existence of two or more forms of bound melittin (9,43,44,54,55); however, their relationship to the functional species remains to be resolved.

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Figure 2 Two of the predicted global minimum free energy conformers of the hydrophobic 1-20 segment of melittin: (a) k99 and (b) k25. (c) A side view of four k99 amphiphilic helix bundles as a conceptualization for the cross-section of a transmembrane pore. In this tetrameric arrangement Lys-7 (ball) is oriented in the aqueous pore. The hydrophobic residues of the helix form the outer surface, presumably in contact with the acyl chains of the bilayer. (From Refs. 47 and 48.)

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Scheme 1 An operational scheme for the interaction of melittin with phospholipid vesicles. At low mole fractions, the bound melittin, especially at anionic interfaces, forms stable contacts between vesicles. At higher mole fractions, aggregated melittin forms pores across zwitterionic bilayers.

C. Melittin Enhances Substrate Replenishment for Interfacial Catalysis in the Scooting Mode The structure (56) and kinetic properties (57) of bee venom phospholipase A2 (PLA2) are established. It requires calcium for the catalytic turnover but not for the binding to the interface. It shows little substrate specificity for the headgroup, and the chemical step is rate-limiting with a turnover number of about 100 s⫺1 for the natural phospholipids. An important feature of interfacial catalytic turnover by PLA2 is that only the substrate in the outer monolayer of vesicles is hydrolyzed and the vesicle integrity is retained, i.e., small solutes and PLA2 do not pass through the hydrolyzed bilayer (57). This is expected to be the case with natural membranes unless, of course, the products of hydrolysis are extracted from the hydrolyzed membrane. The exchange of substrate with the products of hydrolysis through melittinmediated vesicle–vesicle contacts (Fig. 4) is best demonstrated by monitoring the time course and the extent of hydrolysis of phospholipid vesicles by PLA2 (29). Such effects are quantitatively accounted for in terms of the primary kinetic steps in Scheme 2 (58,59). The fraction of the bound enzyme (E*) depends on the bulk concentration of the interface. In the scooting mode, all the enzyme is in the E* form at the interface. Since the catalytic turnover occurs only at the interface, the rate and extent of hydrolysis of vesicles are determined by the E*. In addition, the mole fraction of the substrate on the enzyme-containing vesicle determines the binding of the substrate to the active site and therefore the interfacial turnover rate. If the substrate and the bound enzyme do not exchange between the vesicle populations as is the case in the scooting mode, only the outer monolayer of the enzyme-containing vesicles is hydrolyzed and the excess vesicles remain unhydrolyzed. Melittin (9), as well as several other contact-forming peptides such as polymyxins (30,31) and mastoparans (Cajal and Jain, unpublished), promotes substrate replenishment for the catalytic turnover by PLA2 in the scooting mode.

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Figure 3 Conceptualization of the melittin-mediated vesicle–vesicle contact through which phospholipids selectively exchange. (a) A stable contact between two vesicles permits exchange of the products of hydrolysis from the enzyme-containing vesicle with the substrate on the unhydrolyzed vesicle. (b) The melittin-mediated contact (shown as a ring) between the outer monolayer of vesicles permits selective exchange of phospholipids. (c) A compact global minimum conformer, k246, of melittin (49) in which the hydrophobic residues form the back surface of the fold. (d) Juxtaposition of the hydrophobic surface of the k246 conformer to form a dimer. It is conceivable that such a dimer, formed with melittin monomers bound to anionic vesicles through the charged groups, could form stable contacts that permit exchange of phospholipids from one interface to the other.

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Figure 4 Effect of melittin (4.5 nmol) on the reaction. Bee venom PLA2 (18 nmol) was added to small sonicated vesicles of POPC (520 nmol in 4 mL of 0.5 mM CaCl2 and 1 mM NaCl at pH 8.0 and 23°C). The order of addition of melittin and enzyme has no discernible effect on the progress of the reaction; only the extent of hydrolysis changes with the amount of melittin (see Fig. 5). For details see Ref. 9.

Scheme 2 A kinetic scheme for interfacial catalysis by PLA2. The catalytic turnover occurs through E* and E*S, without the enzyme leaving the interface (represented by a box). Thus, in the scooting mode, the enzyme bound to the interface with high affinity hydrolyzes all the substrate molecules on the enzyme-containing vesicle with virtually infinite processivity. Under these conditions, hydrolysis of excess substrate, present as vesicles without bound enzyme, occurs only if the substrate is transferred to the enzymecontaining vesicles. An example is the melittin-mediated direct vesicle–vesicle exchange of substrate with products of hydrolysis.

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Since the rate of unmediated spontaneous exchange of natural phospholipids between vesicles is exceedingly slow, the reaction progress ceases shortly after the addition of PLA2 to 1-palmitoyl-2-oleoylglycero-sn-3-phosphocholine (POPC) vesicles (Fig. 4). Cessation of the reaction after an initial burst of hydrolysis is due to a higher affinity of PLA2 for the product-containing anionic vesicles (29,58,60). As summarized in Figure 5, excess substrate vesicles become accessible for the hydrolysis after the addition of melittin. The extent of hydrolysis increases with the melittin concentration until a plateau is reached, where about 65% of the total substrate present in the reaction mixture is hydrolyzed, i.e., all the substrate is in the outer monolayer of the vesicles. These results show that at ⬃1.5 mol% melittin, virtually all the substrate molecules present in the outer monolayer of all the vesicles are hydrolyzed by the same amount of enzyme, which in the absence of melittin hydrolyzed only ⬃5% of the total substrate. Higher melittin concentrations do not result in the formation of additional product, as if the inner monolayer were not accessible. Comparable results are obtained with anionic vesicles, where the affinity of PLA2 and melittin is high even before the formation of products. Independent results summarized in the next subsection confirm that melittin promotes presentation of excess substrate to the enzyme sequestered on the product-containing vesicles.

Figure 5 The extent of hydrolysis of POPC (520 nmol) vesicles as a function of melittin added to the reaction mixture containing excess vesicles. The reaction was initiated by bee PLA2 (21.4 pmol) as in Figure 4.

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A direct effect of melittin on the primary catalytic parameters of PLA2 is ruled out (9). For example, melittin does not have a discernible effect on the rate of hydrolysis of 1,2-diheptanoylglycero-sn-3-phosphocholine (DC7PC). Resonance energy transfer and gel filtration studies show that melittin does not enhance the binding of PLA2 to the interface. Independent controls also rule out a direct interaction of melittin with bee venom PLA2 as the basis for the observed kinetic effects. A similar increase in the extent of hydrolysis by melittin is observed with evolutionarily divergent PLA2 from pig pancreas or Crotalus adamentus and Naja nigricola venom, which suggests that the effect is not on the intrinsic kinetic properties of the enzyme. D.

Evidence for Phospholipid Exchange Through Melittin Contacts

Light-scattering studies show that melittin induces aggregation of vesicles, a step necessary for the vesicle–vesicle contact formation. The melittin-induced exchange of phospholipids between aggregated vesicles without (hemi)fusion is shown by monitoring the kinetics of exchange of fluorescently labeled phospholipid probes through melittin contacts between vesicles (9). Typically, the rate of exchange is far in excess of the catalytic turnover rate of about 100 s⫺1. Also, as suggested by results in Figure 5 and shown in Figure 6, only the extent of exchange increases significantly with the amount of melittin. This would be expected if melittin contacts are stable, i.e., the vesicles in contact do not exchange with excess vesicles. The presence of PLA2 does not have a significant effect on the time course of the exchange, nor does it exchange through melittin contacts. These contacts show modest selectivity for the exchange of certain phospholipids. For example, anionic and zwitterionic phospholipids and phospholipids labeled with pyrene, N-(7-nitro-2,1,3-benzoxadiazol-4-yl)dioleyl (NBD), and rhodamine are readily exchanged, but cationic octadecylrhodamine present in the same vesicles does not. This selectivity suggests that the exchange occurs through intervesicle molecular contacts (Fig. 3), where melittin acts as a ‘‘filter’’ or ‘‘sorter’’ for certain probes, and in effect it rules out nonspecific lipid mixing due to (hemi)fusion. Independent results with asymmetrically labeled vesicles showed that only lipids in the outer monolayer exchange; those in the inner layer do not (Fig. 6). Also, the trapped aqueous compartment remains intact. Collectively, these results show that even after hydrolysis by PLA2, ⬍1 mol% melittin does not induce flip-flop or solubilize phospholipids or make anionic vesicles leaky. E.

Melittin-Induced Lysis

Lethal intravenous doses of melittin do not cause harm to circulating erythrocytes (5). The in vitro hemolytic activity of melittin is attributed to its ability to perturb

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Figure 6 Melittin-mediated lipid exchange between anionic vesicles. (䊊) Fluorescence intensity of pyrene monomer as a function of mole percent melittin in a mixture of vesicles of POPC/pyPM (3:7) with POPC/DMPM (3:7). Bulk lipid concentration 1.7 µM for pyPM vesicles and 211 µM for unlabeled acceptor vesicles. (■) A lack of the inner monolayer mixing. RET fluorescence change of a 1:1 mixture of vesicles with NBD-PE labeled only in the inner monolayer and vesicles with Rh-PE (110 µM total lipid) as a function of melittin. (䊉) Vesicles do not become leaky. Reaction of anionic POPC/DMPM vesicles containing 0.6 mol% NBD-PE (110 µM) with dithionite (10 mM): percent of total lipid reduced by dithionite as a function of mole percent of melittin. PLA2 (2.14 pmol) treated vesicles showed the same behavior. (From Ref. 9.)

the barrier function of membranes. In red blood cells, anionic lipids are in the cytoplasmic leaflet of the membrane, and the outer zwitterionic surface could promote the hemolytic activity. A higher affinity of melittin for anionic vesicles is favored by electrostatic interactions; however, anionic charges in the interface decrease the lytic activity of melittin (61,62). The structure and interactions of melittin with aqueous dispersions of zwitterionic phospholipids have been extensively studied (44,45,63,64). Typically, such changes in the polymorphism of zwitterionic phospholipids occur at ⬎1 mol% melittin (53) and ultimately lead to leakage of trapped solutes (61,65), fusion (66,67), and a change in the bilayer organization (53,69–71). Since aggregation of melittin in the bilayer appears to be the driving force for such changes, the ratio of membrane-associated peptide per lipid could be the controlling factor for lysis and other disruptive changes in the bilayer.

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To recapitulate, as discussed by Jain and Bruch in Chapter 4, the polymyxin-mediated direct and rapid exchange of phospholipids between bilayers has provided significant insight into the mechanism of action of the cationic peptide antibacterials. The protein-mediated phospholipid exchange is also useful for understanding the function of myelin basic protein (71) and the lung surfactant protein A (72). As developed in this chapter, the melittin-mediated substrate replenishment could be the first step that ultimately leads to inflammation and the immune response. Melittin amplifies the extent of hydrolytic action of PLA2. If a melittin contact is made up of four molecules, the melittin/PLA2 molar ratio of ⬎50 in the venom would suggest that the hydrolytic effect of one PLA2 molecule could spread over 10 other cells. The products of PLA2 action and mast cell degranulation initiate responses that ultimately lead to inflammation and anaphylaxis. In this model, the synergistic effect of melittin and PLA2 is limited to the initial events whose integrated effect leads to an amplified effect of a small amount of locally injected venom. Also, it is not surprising that the synergistic effect of melittin and PLA2 is not seen in the end effects (73), because such metabolic responses are far removed from the initial limiting events.

ACKNOWLEDGMENTS We thank Drs. Scheraga and Lee for providing us the computed coordinates of truncated melittin shown in Figures 2 and 3. This work was supported by the Ministerio de Educacio´n y Cultura of Spain (contrato Doctores y Tecno´logos Y.C.) and PHS-NIH (M.K.J.). Preparation of the manuscript was facilitated by a DuPont Educational Aid Grant.

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7. Argiolas A, Pisano JJ. Bombolitins, a new class of mast cell degranulating peptides isolated from the venom of bumble bee Megabombus pennsylvanicus. J Biol Chem 1985; 260:1437–1441. 8. Mollay C, Kreil G, Berger H. Action of phospholipases on the cytoplasmic membrane of Escherichia coli. Biochem Biophys Acta 1976; 426:317–324. 9. Cajal Y, Jain MK. Synergism between mellitin and phospholipase A2 from bee venom: apparent activation by intervesicle exchange of phospholipids. Biochemistry 1997; 36:3882–3893. 10. Jansen-Timmen U, Tomic I, Specht E, Beilecke U, Habenicht AJR. The arachidonic acid cascade, eicosanoids, and signal transduction. Annu Rev NY Acad Sci 1994; 733:325–334. 11. Quinn MT, Pathasarathy S, Steinberg D. Lysophosphatidylcholine: a chemotactic factor for human monocytes and its potential role in atherogenesis. Proc Natl Acad Sci USA 1988; 85:2805–2809. 12. Argiolas A, Pisano JJ. Facilitation of phospholipase A2 activity by mastoparans, a new class of mast cell degranulating peptides from wasp venom. J Biol Chem 1983; 258:13697–13702. 13. Reisman RE, Arbesman CE, Lazell M. Observation of the aetiology and natural history of stinging insect sensitivity: application of measurements of venom-specific IgE. Clin Allergy 1979; 9:303–311. 14. King TP, Sobotka AK, Konchoumion L, Lichtenstein LM. Allergens of honey bee venom. Arch Biochem Biophys 1976; 172:661–671. 15. Hirata F, Axelrod J, Crews FT. Concanavalin A stimulates phospholipid methylation and phosphatidylserine decarboxylation in rat mast cells. Proc Natl Acad Sci USA 1979; 76:4813–4816. 16. Sharma SV. Melittin-induced hyperactivation of phospholipase A2 activity and calcium influx in ras-transformed cells. Oncogene 1993; 8:939–947. 17. Murakami M, Hara N, Kudo I, Inoue K. Triggering of degranulation in rat mast cells by exogenous type II phospholipase A2. J Immunol 1993; 151:5675– 5680. 18. Kemeny DM, Harries MG, Youlten LJF, Mackenzie-Mills M, Lessof MH. Antibodies to purified bee venom proteins and peptides. J Allergy Clin Immunol 1983; 71: 505–514. 19. Schneider T, Dudler T, Annand RR, Gelb MH, King TP, Suter M. Comparison of the antibody response to bee venom phospholipase A2 induced by natural exposure in humans or by immunization in mice. J Mol Recognit 1997; 10:93–100. 20. Dudler T, Machado DC, Kolbe L, Annand RR, Rhodes N, Gelb MH, Koelsch E, Suter M, Helm BA. A link between catalytic activity, IgE-independent mast cell activation, and allergenicity of bee venom phospholipase A2. J Immunol 1995; 155: 2605–2613. 21. Bonay M, Echchakir H, Lecossier D, Laine´ P, Herman D, Hance AJ, Soler P, Tazi A. Characterization of proliferative responses and cytokine mRNA profiles induced by Vespula venom in patients with severe reactions to wasp stings. Clin Exp Immunol 1997; 109:342–350. 22. King TP, Kochoumian L, Joslyn A. Melittin specific monoclonal and polyclonal IgG and IgG1 antibodies from mice. J Immunol 1984; 133:2668.

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23. Fehlner PF, Berg RH, Tam JP, King TP. Murine T-cell responses to melittin and its analogs. J Immunol 1991; 146:799. 24. Fehlner PF, Kochoumian L, King TP. Murine IgE and IgG responses to melittin and its analogs. J Immunol 1991; 146:2664–2670. 25. Feinmark SJ, Bailey JM. Lipid metabolism in cultured cells: activators of endogenous thromboxane A2 synthesis in cultured lung fibroblasts. J Biol Chem 1982; 257: 2816–2821. 26. Emmerling MR, Moore CJ, Doyle PD, Carroll RT, Davis RE. Phospholipase A2 activation influences the processing and secretion of the amyloid precursor protein. Biochem Biophys Res Commun 1993; 197:292–297. 27. Zeitler P, Wu YQ, Handwerger SJ. Melittin stimulates phosphoinositide hydrolysis and placental lactogen release: arachidonic acid as a link between phospholipase A2 and phospholipase C signal-transduction pathways. Life Sci 1991; 48:2089– 2095. 28. Rao NM. Differential susceptibility of phosphatidylcholine small unilamellar vesicles to phospholipase A2, C, and D in the presence of membrane active peptides. Biochem Biophys Res Commun 1992; 182:682–688. 29. Jain MK, Rogers J, Berg OG, Gelb MH. Interfacial catalysis by phospholipase A2: activation by substrate replenishment. Biochemistry 1991; 30:7340–7348. 30. Cajal Y, Rogers J, Berg OG, Jain MK. Intermembrane molecular contacts by polymyxin B mediate exchange of phospholipids. Biochemistry 1996; 35:299–308. 31. Cajal Y, Ghanta J, Easwaran K, Surolia A, Jain MK. Specificity for the exchange of phospholipids through polymyxin B mediated intermembrane molecular contacts. Biochemistry 1996; 35:5684–5695. 32. Tsunoda Y, Owyang Ch. A newly cloned phospholipase A2-activating protein elicits Ca2⫹ oscillations and pancreatic amylase secretion via mediation of G protein beta/ phospholipase A2/arachidonic acid cascades. Biochim Biophys Res Commun 1994; 203:1716–1724. 33. Koumamov K, Wolf C, Bereziat G. Modulation of human type II secretory phospholipase A2 by sphingomyelin and annexin VI. Biochem J 1997; 326:227–233. 34. Peterson JW, Dickey WD, Saini SS, Gourley W, Klimpel GR, Chopra AK. Phospholipase A2 activating protein and idiopathic inflammatory bowel disease. Gut 1996; 39:698–704. 35. Okamoto T, Isoda H, Kubota N, Takahata K, Takahashi T, Kishi T, Nakamura TY, Muromachi Y, Matsui Y, Goshima K. Melittin cardiotoxicity in cultured mouse cardiac myocytes and its correlation with calcium overload. Toxicol Appl Pharmacol 1995; 133:150–163. 36. Bomalaski JS, Steiner MR, Simon PL, Clark MA. IL-1 increases phospholipase A2 activity, expression of phospholipase A2-activating protein, and release of linoleic acid from the murine T-helper cell line EL-4. J Immunol 1992; 148:155–160. 37. Bomalaski JS, Ford T, Hudson AP, Clark M. Phospholipase A2-activating protein induces the synthesis of IL-1 and TNF in human monocytes. J Immunol 1995; 154: 4027–4031. 38. Clark MA, Ozgur LE, Conway TM, Dispoto J, Crooke ST, Bomalaski JS. Cloning of a phospholipase A2-activating protein. Proc Natl Acad Sci USA 1991; 88: 5418.

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39. Clark MA, Conway TM, Shorr RGL, Crooke ST. Identification and isolation of a mammalian protein which is antigenically and functionally related to the phospholipase A2 stimulatory peptide melittin. J Biol Chem 1987; 262:4402–4406. 40. Gauldie J, Hanson JM, Ramjanek FD, Shipolini RA, Vernon CA. The peptide components of bee venom. Eur J Biochem 1976; 61:369–376. 41. Faucon JF, Dufourq J, Lussan C. The self-association of melittin and its binding to lipids. FEBS Lett 1979; 102:187–190. 42. Talbot JC, Dufourcq J, de Bony J, Faucon JF, Lussan C. Conformational change and self-association of monomeric melittin. FEBS Lett 1979; 102:191. 43. Podo F, Strom R, Crifo C, Zulauf M. Dependence of melittin structure on its interaction with multivalent ions and lipid membranes. Int J Peptide Protein Res 1982; 19: 14–27. 44. Cornut I, Thiaudiera E, Dufourcq J. In: Epand R, ed. The Amphipathic Helix. New York: Academic Press, 1993:173–219. 45. Saberwal G, Nagaraj R. Cell-lytic and antibacterial peptides that act perturbing the barrier function of membranes: facets of their conformational features, structurefunction correlations and membrane-perturbing abilities. Biochim Biophys Acta 1994; 1197:109–131. 46. Dempsey CE. The actions of melittin on membranes. Biochim Biophys Acta 1990; 1031:143–161. 47. Ripoli DR, Liwo A, Scheraga HA. New developments of the electrostatically driven Monte Carlo method: test on the membrane-bound portion of melittin. Biopolymers 1998; 46:117–126. 48. Lee J, Scheraga HA, Rackovsky S. Conformational analysis of the 20-residue membrane-bound portion of melittin by conformational space annealing. Biopolymers 1998; 46:103–115. 49. Smith R, Separovic F, Milne TJ, Whittaker A, Bennett FM, Cornell BA, Makriyannis A. Structure and orientation of the pore-forming peptide, melittin, in lipid bilayers. J Mol Biol 1994; 241:456–466. 50. Okada A, Wakamatsu K, Miyazawa T, Higashijima T. Vesicle-bound conformation of melittin: transferred nuclear Overhauser enhancement analysis in the presence of perdeuterated phosphatidylcholine vesicles. Biochemistry 1994; 33:9438–9446. 51. Dempsey EE, Butler GS. Helical structure and orientation of melittin in dispersed phospholipid membranes from amide exchange analysis in situ. Biochemistry 1992; 31:11973–11977. 52. Yuan P, Fisher PJ, Pendergast FG, Kemple MD. Structure and dynamics of melittin in lysomyristoyl phosphatidylcholine micelles determined by nuclear magnetic resonance. Biophys J 1996; 70:2223–2238. 53. Batenburg AM, Van Esch JH, De Kruijff B. Melittin-induced changes of the macroscopic structure of phosphatidylethanolamines. Biosci Rep 1998; 8:299–307. 54. Pe´rez-Paya´ E, Dufourq J, Braco L, Abad C. Structural characterization of the natural membrane-bound state of melittin: a fluorescence study of a dansylated analogue. Biochim Biophys Acta 1997; 1329:223–236. 55. Rex S, Schwarz G. Quantitative studies of the melittin-induced leakage mechanism of lipid vesicles. Biochemistry 1998; 37:2336–2345. 56. Scott DL, Otwinowski Z, Gelb MH, Sigler PB. Crystal structure of bee venom phos-

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4 Polymyxins Prototype for a New Class of Antibiotics Mahendra Kumar Jain and Martha Bruch University of Delaware, Newark, Delaware

I. INTRODUCTION Antimicrobial peptides and proteins are produced by virtually all organisms (1– 3) as a part of their innate immune defense system (4). Several hundred antimicrobial peptides have been characterized from organisms ranging from bacteria and fungi to plants and animals. Examples include defensins from human plasma, magainins from frog skin, cecropins from insect larvae, and thionins from plants. The evolutionary success of such antimicrobials implies strategies for selectivity and for overcoming drug resistance. As a class they do not have a deleterious effect on the organisms that produce them. This fact alone rules out nonspecific mechanisms of action, such as leakage of the cytoplasmic content. Not all cationic peptides show antibacterial activity, and the membrane-permeabilizing effect of amphiphilic cationic peptides (1,5) does not account for the selectivity of polymyxins against gram-negative organisms. In fact, the membrane disruption by polymyxins is observed at a mole fraction that is more than 100 times the minimum inhibitory concentrations (6–8). Results reviewed in this chapter show that the antibacterial effect of polymyxin B (PxB) is due to a novel mechanism with far-reaching implications. II.

POLYMYXIN B: A PROTOTYPE FOR ANTIBACTERIAL CATIONIC PEPTIDES

The focus of our work is on a subgroup of cationic peptides, of which polymyxin B (Fig. 1) is the best studied. Polymyxins are produced by gram-positive Paeni63

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Figure 1 The structure of polymyxin B minimized to satisfy the NMR constraints. Dab ⫽ 1,3-diaminobutyric acid. (From Ref. 30.) A side view of this structure is shown in Figure 5.

bacillus species and are highly selective against a broad spectrum of gram-negative organisms (9). The ability of polymyxin to bind lipopolysaccharide (LPS) and disrupt the outer membrane of gram-negative organisms is a necessary, but not sufficient, condition for the antibacterial action. For example, the polymyxin nonapeptide (NP) is not antibacterial, yet it disrupts the LPS layer and promotes entry of other solutes into the periplasmic space (9–11). Agarose-bound polymyxin shows antibacterial activity, which rules out entry of the peptide into the cytoplasm as a condition for the antimicrobial action (9). Similarly, binding of PxB to anionic phospholipid is not a sufficient basis for its antibacterial action. A.

PxB Induces a Highly Selective Expression of the Promoter of the osmY Gene

We have adopted a novel strategy to identify the locus of metabolic stress induced by antimicrobial agents in growing organisms (12–14). The basis for the assay lies in the fact that organisms often respond and adapt to sublethal environmental adversities by increased expression of stress proteins to restore homeostasis (15,16). By obligatorily coupling the transcription of a specific promoter to a bacterial luminescence reporter luxCDABE operon on a plasmid introduced in

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E. coli, such fusion strains produce luminescence in response to a specific stress at sublethal concentrations (17,18). Thus, the stress response is measured with viable cells below the lethal concentration of an antibacterial. Since most stress genes are not normally highly expressed under normal growth conditions, the transcriptional luminescence response to stress in real time typically yields an excellent ratio of signal to background noise. Extension of the stress response results to establish the antimicrobial mechanism is based on the assumption that continued excessive stress leads to bacteriostasis and ultimately to cell death. While positive results from the stress promoter/luminescence technique permit identification of a specific stress, negative results with carefully selected stress promoters also rule out possible roles for the corresponding stresses. Results summarized in Table 1 show that osmY expression, also induced by the hyperosmotic stress (19,20), is the primary stress response of PxB in Escherichia coli, without a significant expression of other stress promoters. Each strain contains a plasmid with a specific stress promoter coupled to the expression of luxCDABE, which elicits response only to a specific stress. For example, the effect of the stress induced by PxB on the lux luminescence response, from the hyperosmolarity-sensitive E. coli fusion with osmY-lux reporter, is shown in Figure 2. As shown in the upper panel for the optical density (OD) change, PxB lowers the growth rate above 0.25 µM. The selectivity of the transcription of the osmY gene response is shown by the biphasic change near the minimum inhibitory concentration (MIC) of PxB. The luminescence responses to carbonylcyanide m-chlorophenylhydrazone (CCCP), a proton translocator or uncoupler that depletes the proton gradient, on the uncoupler-sensitive and the hyperosmolaritysensitive strains are compared in Figure 3. Here, luminescence increases with CCCP up to its MIC and then decreases as cells become metabolically nonviable. Only a monotonic decrease above the MIC is seen with the hyperosmolarity-

Table 1 Response of the Stress-Sensitive Strains of E. coli to PxB and Other Additives Fusion (gene : : lux #) strain grpE′ recA′ katG′ inaA′ lac micF/rob⫺ osmY′

TV1061 DPD2794 DPD2511 DPD2146 TV1048 DPD2194 DPD2170

Response to Stress

PxB

NaCl

Sucrose

CCCP

NP

Protein folding DNA damage Peroxide damage Proton leakage Limited carbon source Superoxide damage Hyperosmotic shock

⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹

⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹

⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫹

⫺ ⫺ ⫺ ⫹ ⫺ ⫺ ⫺

⫺ ⫺ ⫺ ⫺ ⫺ ⫺ ⫺

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Figure 2 Effect of PxB concentration on the change in the optical density (top panel) or luminescence (bottom panel) at 60 min after the addition of PxB to a hyperosmotic stress-sensitive fusion strain of E. coli in the early log phase of growth. Luminescence increase was calculated as 100 ⫻ (L-Lc)/Lc where Lc is luminescence of control at 60 min.

sensitive strain. The biphasic stress response is typical of that seen for lux fusions; the increase reflects the specific response that promotes expression of osmY by PxB, whereas the decrease above the MIC is a nonspecific response associated with the loss of viability due to reduced ATP levels, i.e., the concentration window at or below the MIC is useful for monitoring the stress response. Results in Table 1 clearly show that the stress profile for PxB is similar to that for hyperosmolar NaCl and sucrose, and the exceptions are instructive. The 2-10 peptide of PxB, NP, disrupts the outer membrane at 10 µM, yet it does not induce osmY expression. CCCP does not induce expression of osmY, although it elicits luminescence response from the strain that responds to changes in the proton gradient. These results show that the PxB-induced expression of the osmY gene is due neither to the disruption of the outer membrane nor to the stress of proton leakage through the cytoplasmic membrane. Results with sucrose also rule out the possibility that the effect of PxB and NaCl is on an ion channel or some

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Figure 3 Effect of carbonylcyanide m-chlorophenylhydrazone (CCCP) concentration on the change in the optical density (䊊) or the luminescence 60 min after the addition of CCCP to (䉱) the uncoupler-sensitive or (䊐) the hyperosmotic stress-sensitive fusion strains of E. coli. Other conditions as in Figure 3.

such osmoregulatory mechanism. The possibility that PxB directly interacts with a cytoplasmic regulator of osmY can also be discounted by the observation that PxB immobilized on agarose beads is antibacterial and also induces the expression of osmY promoter. B. Rapid Responses of PxB The osmY promoter is expressed in response to hyperosmolar stress on the time scale of cell division. Since the PxB and hyperosmotic stress induce the same transcriptional end effect, we explored the effect of PxB on the early time events associated with hyperosmotic shock. The rapid response of bacteria to hyperosmotic shock is a rapid wrinkling of the cytoplasmic membrane due to efflux

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of water from the cytoplasmic compartment (21). The initial shrinkage of the cytoplasmic compartment due to water efflux is essentially complete in less than 5 s, and it is followed by the onset of osmoregulatory mechanisms that restore the cytoplasmic volume in about 5 min. These plasmolytic events in response to hyperosmolar NaCl are readily monitored as a change in turbidity or scattering; however, the optical signal is not readily discernible in sucrose solutions due to a change in the refractive index of the medium. As shown in Figure 4, the extent of the initial rapid cytoplasmic shrinkage is significantly smaller for the cells pretreated with polymyxin. The magnitude of the initial rapid change depends on the magnitude of the osmotic stress, and PxB lowers the magnitude of the change with PxB concentration. In contrast, PxB added before or after the osmotic shock does not have a significant effect on the time course of the recovery phase. Collectively, results of the transcriptional response induced by PxB below its MIC clearly show that PxB induces the osmY promoter without entering the cytoplasm. A similar response is also induced by other peptide antimicrobials (13,14) as well as by hyperosmolar NaCl or sucrose (12,13). The promoter-coupled lux response is biphasic, which rules out the possibility that PxB-mediated stress may have its origin in the loss of the energy-requiring mechanisms that

Figure 4 The time course of hyperosmolarity-induced increase in the 90° scattering at 600 nm of E. coli without (top) or treated with 0.25 µM PxB. Note that the major effect of PxB is on the initial rapid increase in the scattering, and the effect on the first-order recovery process is virtually negligible.

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maintain the osmotic balance. A possible role for an effect of the binding to or disruption of the LPS layer is also ruled out by the fact that PxB and PxB nonapeptide bind to and disrupt the LPS layer of the outer membrane with equal efficacy. However, the nonapeptide is not antimicrobial, nor does it induce osmY promoter, nor does it have any effect on the plasmolysis time course. Thus the antimicrobial effect must be due to a step after the disruption of the LPS layer. Osmotic changes occur during growth, and their control in eukaryotes (22) and prokaryotes (23) is finely tuned by a host of genes with roles in feedback metabolic loops that determine the recovery phase of the plasmolysis time course (24). The osmY gene product is a periplasmic protein with an unknown function. Transcription of osmY is regulated and induced by hyperosmolarity, nutrient starvation, and possibly other regulatory circuits that ultimately lead to the stationary phase (19,20). It is intriguing that not only are the transcriptional end results of antimicrobial action and hyperosmotic stress the same (Table 1), but the rapid scattering change induced by hyperosmotic stress is blocked in the presence of PxB. So the key issue is the identity of the common initial locus of action of these two apparently diverse stresses. Based on the experiments described below, our working hypothesis is that the interaction of PxB with the cell must somehow rapidly prevent the hyperosmolarity-induced wrinkling of the cytoplasmic membrane by the formation of stable PxB contacts between the cytoplasm and the outer membrane. In such a model, the regulatory function is ascribed to membrane-localized stretch receptors that signal the stress on the membrane to the cytoplasm. It is intriguing to consider whether the osmY gene product in the periplasmic space is such a receptor. C. PxB-Induced Contacts in the Periplasmic Space Polymyxin B-mediated contacts between the interfaces surrounding the periplasmic space could account for the effects described in the preceding section (6,26). Phosphatidylglycerol is a significant component of the two interfaces that surround the periplasmic space. PxB could form a stable contact between the interfaces that come close enough to permit such contacts. Many of the lipid components of the outer membrane are synthesized in the cytoplasm and on the inner membrane. Thus contact between the two membranes appears to be an obligatory step during the cell growth. If so, PxB could stabilize such contacts and cause stasis. For the prevention of shrinkage the minimum condition is that the cytoplasmic membrane remain in stable contact with a stable interface. We have found a significant correlation between the antimicrobial efficacy of the various peptides and their ability to induce osmY expression or to promote exchange of phospholipid (14). Since contact formation is a prelude to the exchange, it is not obvious whether the ability to exchange phospholipid is a critical requirement for the rapid effect on the hyperosmotic shrinkage or even for the

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antimicrobial effect. So far we have not found an antimicrobial peptide that binds to LPS, enters the periplasmic space, and forms vesicle–vesicle contacts without mediating a phospholipid exchange. PxB-mediated exchange of phospholipid may have significant consequences for cellular homeostasis, because most organisms maintain a remarkably specific phospholipid composition in their membranes (27), and microorganisms adapt to a changing environment by changing their phospholipid composition. A role for the phospholipid degradation by the phospholipase A2 present in the outer membrane (OMPLA) is ruled out as the basis for the antimicrobial effect, because the OMPLA⫺ mutant remains susceptible to PxB. In short, PxB contacts between the two anionic phospholipid monolayers that enclose the periplasmic space could form a basis for bacteriostasis and ultimate loss of viability of gram-negative organisms.

D.

Resistance to PxB Could Not Be Induced by Point Mutagenesis

A phospholipid target as the primary basis for the antimicrobial action implies that organisms cannot become genetically resistant by mutation of the target gene product. Polymyxins have been in use for decades against topical infection; however, we have not found any reported instance for a target-based genetically stable resistance against PxB (5). Our repeated attempts to isolate resistant strains from E. coli or Pseudomonas aeruginosa mutagenized by ethylmethane sulfonate or UV irradiation have been unsuccessful (14). Moreover, the fact that PxB does not enter the cytoplasm suggests that PxB may not succumb to resistance by drug efflux mechanisms.

E.

Nonmutational Adaptation to PxB

Several polymyxin-resistant gram-negative organisms are known to have PxBsensitive strains. PxB-adapted strains of the sensitive gram-negative species have also been characterized. Some of these inducible changes appear to modify lipid A: The 4′ phosphate is free in the sensitive strains, and it is esterified with 4amino-4-deoxy-l-arabinose (AraN) in the resistant strains. Apparently, the modified LPS has a lower affinity for PxB. Such a modification of lipid A is due to constitutive enzymes (28). Expression of certain outer membrane proteins has also been correlated to PxB resistance (3), and several genetic loci that confer resistance against peptide antimicrobials have been reported (29). Our attempts to isolate PxB-adapted strains by successive transfer to higher PxB concentrations were successful only with the type strain of P. aeruginosa, but the adopted culture rapidly reverted to the PxB-susceptible form in the absence of PxB (14). Several other strains could not be adapted by such a protocol. Also, long periods of incu-

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bation with E. coli do not degrade PxB, but cecropin is apparently degraded, suggesting a possible role for the surface protease. F. Untoward Effects of Polymyxins High affinity of PxB for LPS makes it an attractive agent for the treatment of septic shock; however, systemic use of polymyxins is limited by its nephro- and neurotoxicity. Cellular effects inhibited by PxB include the release reaction of monocytes and macrophages, mitogenicity, Shwartman reaction, and anticomplement reaction. The ability of polymyxins to mediate phospholipid exchange could account for some of the effects that require membrane–membrane interactions. The structure–activity correlation for the various effects of polymyxins may ultimately provide a basis for minimizing unfavorable effects of such antimicrobials.

III. PxB-INDUCED CONTACTS BETWEEN PHOSPHOLIPID VESICLES Our interest in the effect of PxB on microbial physiology was elicited by a chance observation that PxB promotes a direct and rapid exchange of phospholipid between anionic vesicles without solubilization, fusion, or leakage of vesicles (6– 8,25,26). As a prelude for the phospholipid exchange, PxB forms stable contacts between vesicles of monoanionic phospholipid, including phosphatidylglycerol. PxB also binds to vesicles of dianionic phosphatidic acid, but the contacts are not stable and its exchange is not observed. None of these three functions is seen with phosphatidylcholine vesicles. Exchange of the phospholipid between the covesicles in contact also shows selectivity, i.e., exchange of only monoanionic phospholipid exchange, not the zwitterionic or dianionic phospholipid. Operationally, for the phospholipid exchange the initial binding of the peptide to vesicles must lead to contact formation between vesicles, followed by the phospholipid exchange through vesicles in contact through the peptide. Control of the phospholipid exchange could occur at any of the three stages. A.

Binding of Phosphoesters to PxB as the Primary Molecular Interaction

The higher order structure of a peptide is critical for understanding the molecular basis of its action. PxB does not assume a stable higher order structure in the aqueous phase. A family of higher order structures of PxB is formed in aqueous trifluoroethanol, where its circular dichroism (CD) signatures are similar to those from PxB bound to anionic phospholipid vesicles (30). A low energy structure obtained from the family of structures satisfying the nuclear magnetic resonance

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(NMR) constraints is shown in Figure 1. Fixed positions for the backbone of Phe-6, Leu-7, and α,γ-diaminobutyric acid residue 10 (Dab-10), coupled with a flexibility for the other ring residues, gives rise to ring pucker and a shift in the position of the side chains. Only one NOE is observed for the acyl chain on the N-terminus, suggesting that it is extremely floppy. The range of structures assumed by PxB in solution as well as at membrane interfaces is relevant for the multiplicity of interactions. Specific interactions of PxB with anionic phosphoester groups appear to be a common basis for the interaction with cationic phospholipids and with LPS. Cationic PxB is expected to interact with anionic species at the interface and possibly compete for the binding of divalent cations at the cell surface. Additional factors must also be at work, because cationic peptides, such as NP, are not antimicrobial. On the other hand, a solitary role for the hydrophobic interactions dominated by the alkyl chain of PxB alone can be ruled out. The CD spectrum of PxB does not change in the presence of zwitterionic vesicles. Also, PxB does not induce contacts between zwitterionic vesicles, and phosphatidylcholine present in anionic vesicles does not exchange through functional contacts formed between covesicles of anionic and zwitterionic phospholipids. PxB disrupts the outer membrane by binding to LPS, presumably through strong interactions with the phosphate groups in the lipid A moiety of LPS. Similarly, interaction with phosphatidylglycerol through the glycero-3-phosphate group appears to be important for the formation of PxB contacts and for the exchange. Both of these interactions of PxB to form functional intermembrane contacts are critical for its antimicrobial action. As summarized in Figure 5, molecular docking studies of phosphoesters on PxB within the NMR constraints suggest that there are two possible binding sites (30): the first with Dab-3 and -5 with a possible contribution from Dab-1; and the second with Dab-8 and -9 with an H-bonding contribution from NH and OH of Thr-10. Both mono- and dianionic phosphoesters bind to these sites, although the affinity for the monoanionic forms is considerably lower. The most intriguing functional feature of this geometry is that although the two sites could be occupied by two separate phosphoester molecules, they can also be simultaneously occupied by the two phosphates of lipid A. For example, in phosphatidylglycerol bound to PxB (Fig. 5a), the phosphate is oriented toward the peptide, whereas both the acyl chains point away. This arrangement is well suited for the localization of PxB at the phospholipid interface, where not only could it bind to two different phospholipid molecules, but also the two sites could be temporally differentiated by the migration of one ligand from one site to another during the phospholipid exchange. A significant difference in the calculated affinities for the mono- and dianionic species for these sites also offers certain functional insights. The fact that PxB does not mediate the exchange of phosphatidic acid would imply that in

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Figure 5 A side view of the energy-minimized structure of PxB docked with (a) diacetylphosphatidylglycerol and (b) the glucosamine disaccharidephosphate of lipid A.

this case the temporal migration is not favored. Also, the fact that PxB does not form stable contacts between phosphatidic vesicles implies that both sites are occupied by ligands from the same surface. Similar factors could be at work during the binding of PxB to lipid A (Fig. 5b) with two phosphate groups: one at the 1-position and the other at the 4′-position of the glucosamine disaccharide backbone. Both of these groups snugly fit simultaneously on the same PxB, which may be an early step during the self-promoted uptake in the periplasm and disruption of the bacterial outer membrane. In a PxB-resistant organism the 1- or 4′phosphates are substituted by amino-arabinose (28), which would interfere with the binding to PxB and thus hinder its uptake. IV. IS PxB A PROTOTYPE FOR A NEW CLASS OF ANTIBIOTICS? It has been noted that no new class of antibiotic has been discovered in the last 25 years (5). If so, the hyperosmotic stress mechanism proposed for cationic

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peptides may represent a major breakthrough in identifying antibacterial agents with highly desirable properties. Their attractive features include (1) they have a rapid mode of action, virtually on contact; (2) their action though the periplasmic space makes them an unlikely target for the drug resistance through efflux mechanisms; and (3) with phospholipid as the primary target, such agents are unlikely to develop resistance by point mutation. Furthermore, growth inhibition and bacteriostasis induced by such agents, without a direct bactericidal effect, offer novel possibilities for managing infection through the normal immune system. Not only can a modest cell population from the initial infection, induced to a stasis mode, be readily neutralized by the innate immune system of the host, but also a lower endotoxin level will prevent secondary complications. To recapitulate, the very existence of natural antimicrobial peptides as part of an innate defense mechanism adopted by virtually all organisms suggests strategies toward target selectivity. Our suggested mechanism for their action, through the contacts between phospholipid bilayers mediated by antimicrobial agents, offers a possible solution to the problems of antibiotic resistance. Their very existence and the underlying evolutionary experience that gave rise to such antimicrobials may also offer some very fundamental lessons for managing infection. The osmY-lux response provides a basis for a unique assay for such novel antimicrobial agents with a highly desirable pharmacological profile. The action of polymyxins, by inducing stasis of growth through the osmoregulatory mechanism, bears on a very fundamental aspect of cell cycle, which remains to be elucidated. Another challenge is that of eliminating untoward and toxic effects of these agents, which have limited their clinical use.

ACKNOWLEDGMENTS The work described from our laboratory was supported by NIH, and manuscript preparation was supported by a DuPont Educational Aid Grant.

REFERENCES 1. Hancock REW, Falla T, Brown M. Cationic bactericidal peptides. Adv Microb Physiol 1995; 37:135–175. 2. Hoffman JA. Innate immunity of insects. Curr Opin Immunol 1995; 7:4–10. 3. Young ML, Bains M, Bell A, Hancock REW. Role of P. aeruginosa outer membrane protein OprH in polymyxin and gentamicin resistance: isolation of an OprH-deficient mutant by gene replacement techniques. Antimicrob Agents Chemother 1992; 36: 2566–2568.

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4. Zasloff M. Antibiotic peptides as mediators of innate immunity. Curr Opin Immunol 1992; 4:3–7. 5. Hancock REW, Piers K, Brown M, Falla T, Gough M, Wu M, Fidai S. Cationic peptides: a class of antibiotics able to access the self-promoted uptake pathway across the Pseudomonas aeruginosa outer membrane. In: Nakazawa T, Furukawa K, Haas D, Silver S, eds. Molecular Biology of Pseudomonas. Washington DC: ASM Press, 1996:441–450. 6. Cajal Y, Berg OG, Jain MK. Direct vesicle-vesicle exchange of phospholipid mediated by polymyxin B. Biochem Biophys Res Commun 1995; 210:746–752. 7. Cajal Y, Rogers J, Berg OG, Jain MK. Intermembrane molecular contacts by polymyxin B mediate exchange of phospholipid. Biochemistry 1996; 35:299–308. 8. Cajal Y, Ghanta J, Easwaran K, Surolia A, Jain MK. Specificity for the exchange of phospholipid through polymyxin B mediated intermembrane molecular contacts. Biochemistry 1996; 35:5684–5695. 9. Storm DR, Rosenthal KS, Swanson PE. Polymyxin and related peptide antibiotics. Ann Rev Biochem 1977; 46:723–763. 10. Kubesch P, Boggs J, Luciano L, Maass G, Tummler B. Interaction of polymyxin B nonapeptide with anionic phospholipid. Biochemistry 1997; 26:2139–2149. 11. Vaara M. Agents that increase the permeability of the outer membrane. Microbiol Rev 1992; 56:395–411. 12. Oh J-T, Van Dyk TK, Cajal Y, Dhurjati PS, Sasser M, Jain MK. Osmotic stress in viable Escherichia coli as the basis for the antibiotic response by polymyxin B. Biochem Biophys Res Commun 1998; 246:619–623. 13. Oh J-T, Cajal Y, Dhurjati PS, VanDyk TK, Jain MK. Cecropin induces hyperosmotic stress response in viable Escherichia coli through membrane contacts in the periplasmic space. Biochim Biophys Acta, in press. 14. Oh J-T, Cajal Y, Skowronska EM, Belkin S, Van Dyk TK, Sasser M, Jain MK. The cationic peptide antimicrobials interfere with the hyperosmotic stress response in Escherichia coli. Biochim Biophys Acta, in press. 15. Gottesman S. Bacterial regulation: global regulatory networks. Annu Rev Genet 1984; 18:415–441. 16. Welch WJ. How cells respond to stress. Sci Am May 1993:56–64. 17. Belkin S, Van Dyk TK, Vollmer AC, Smulski DR, LaRossa RA. Monitoring subtoxic environmental hazards by stress-responsive luminous bacteria. Environ Toxicol Water Qual 1996; 11:179–185. 18. Van Dyk TK, Ayers BL, Morgan RW, LaRossa RA. Constricted flux through the branched-chain amino acid biosynthetic enzyme acetolactate synthase triggers elevated expression of genes regulated by rpoS and internal acidification. J Bacteriol 1998; 180:785–792. 19. Yim HH, Brems RL, Villarejo M. Molecular characterization of the promoter of osmY, an rpoS-dependent gene. J Bacteriol 1994; 176:100–107. 20. Lange R, Barth M, Hengge-Aronis R. Complex transcriptional control of the σs-dependent stationary-phase-induced and osmotically regulated osmY (csi-5) gene suggests novel role for Lrp, cyclic AMP (cAMPM) receptor protein-cAMP complex, and integration host factor in the stationary-phase response of Escherichia coli. J Bacteriol 1993; 175:7910–7917.

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21. Koch AL. The biophysics of the gram-negative periplasmic space. Crit Rev Microbiol 1998; 24:23–59. 22. Waldegger S, Lang F. Cell volume and gene expression. J Membrane Biol 1998; 162:95–100. 23. Csonka L. Physiological and genetic responses of bacteria to osmotic stress. Microbiol Rev 1989; 53:121–147. 24. Hoffman EK, Dunham PB. Membrane mechanisms and intracellular signalling in cell volume regulation. Int Rev Cytol 1995; 161:173–262. 25. Jain MK, Rogers J, Berg OG, Gelb MH. Interfacial catalysis by phospholipase A-2: activation by substrate replenishment. Biochemistry 1991; 30:7340–7348. 26. Cajal Y, Jain MK. Membrane-membrane contacts: a ‘‘sorting’’ role for proteins? J NIH Res 1996; 8:35–39. 27. Raetz CRH. Biochemistry of endotoxins. Annu Rev Biochem 1990; 59:129–170. 28. Groisman EA. How bacteria resist killing by host-defence peptides. Trends Microbiol 1994; 2:444–449. 29. Bruch M, Cajal Y, Koh JT, Jain MK. Higher order structure of polymyxin B and its functional significance. J Am Chem Soc, in press.

5 Environmental Effects on Skin Lipids and Impairment of Barrier Function Ju¨rgen Fuchs Johann Wolfgang Goethe University, Frankfurt, Germany

I. INTRODUCTION Skin is a biological interface with the environment, functions as the first line of defense against noxious stimuli, and protects the body’s interior from external insults. This protective mechanism can break down under stresses caused by environmental extremes. The skin is one of the largest body organs and serves as a major portal of entry for many environmental pollutants, some of which are free radical generaters. Lipid peroxidation is one of the molecular consequences of free radical reactions. Skin is a potential target organ of oxidative injury because it is continuously exposed to environmental pollutants, visible and ultraviolet radiation, and high oxygen concentrations and contains a variety of oxidizable lipids critical for maintenance of epidermal barrier function. Peroxidation of skin lipids may cause disturbances of barrier function, induction of inflammatory reactions, impairment of immune surveillance, and predisposition to cancer. Intact skin barrier has important health implications, and barrier function can be measured easily by noninvasive techniques (1). It is established that homeostasis of the physiological skin lipid composition is essential to an intact barrier function, and this field has been thoroughly investigated in past decades. The concentration of lipid peroxidation products in skin is significantly influenced by nutrition (2), cell turnover (3), and environmental factors, solar radiation being the most important determinant. A potential role of environmental oxidants as modulators of skin lipid integrity is emerging, and this chapter is designed to give a brief introduction into this field. 77

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II.

SKIN BARRIER FUNCTION

The skin’s most important function is to act as a barrier against fluid loss, microorganism infections, and percutaneous absorption. To fulfill this role, keratinocytes proliferate and differentiate to produce a protective layer, the stratum corneum, which is part of the epidermis. The morphological structure of skin is depicted in Figure 1, which shows the different skin layers of the epidermis, the outermost part of the skin, the dermis, and the subcutis, the innermost part of the skin. The stratum corneum controls the diffusion and penetration of chemical substances into and through the skin. The barrier function of the stratum corneum seems primarily to be regulated by the lamellar lipid bilayers between the corneocytes, which originate largely from polar lipid precursors provided by the cells of stratum granulosum via exocytosis of the lamellar body content. In particular, the structural organization of these intercellular lipid lamellae seems to be responsible for the very low water permeability of the intact skin. Disruption of the epidermal barrier triggers a homeostatic response that is designed to rapidly restore functional integrity to the disturbed site. The major lipid components isolated from the cornified epidermal layers are ceramides, which belong to the class of sphingolipids, cholesterol, and free fatty acids. The stratum corneum lipids consist of ceramides 1–6 (⬃55%), cholesterol (⬃25%), and free fatty acids (⬃10%). All three stratum corneum lipids (ceramides, cholesterol, and free fatty acids) are required for permeability barrier homeostasis. Recent studies have shown that application of one or two of these lipids to perturbed skin delays

Figure 1 Morphological structure of human skin showing the epidermis, dermis, and subcutis. The outermost layer, the epidermis, consists of the stratum corneum, stratum spinosum, stratum granulosum, and stratum basale.

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barrier recovery; only equimolar mixtures allow normal recovery (4). Their biosynthesis is in tight relationship with the cutaneous barrier function. In studies in which the barrier is artificially disturbed, lipid biosynthesis is found to be directly regulated by barrier permeability. The ceramides involved in this process are located in the extracellular spaces of the upper epidermal layers, whereas sphingomyelin, the most common sphingolipid, is an integral part of the bilayer plasma membrane of the keratinocytes (5). Stratum corneum lipid composition is essential for barrier integrity, and it was suggested that oxidation and degradation of stratum corneum lipids could affect the barrier function of the stratum corneum (6). Disturbed barrier function then can cause upregulation of the cytokine cascade in deeper skin layers, thereby triggering an inflammatory response (7). Calcium, a regulator of a variety of physiological and biochemical functions, plays an important role in the regulation of cellular differentiation and desquamation of epidermal keratinocytes and is essentially involved in modulating stratum corneum barrier homeostasis (8–11). Lipid peroxidation and Ca2⫹ were suggested to be linked as mediators of cell homeostasis as well as of cell damage. The Ca2⫹ ion was shown to enhance as well as inhibit lipid peroxidation, and the interactions between Ca2⫹ and lipid peroxidation seem to be complex (12).

III. SKIN LIPIDS Skin lipids can artificially be divided into lipids of the epidermis, epidermal appendages (hair, nail), sebaceous glands, and subcutaneous tissue. Human skin surface lipids emanate from two sources, the keratinizing epidermis and the sebaceous gland (13). A.

Epidermal Lipids

Epidermal lipids account for about 8% of the cell’s dry weight (14). Regional variations of human epidermal lipids may underlie local variations in skin permeability to transepidermal water loss (15). The overall composition of lipids is quite similar in different species but markedly different in the successive layers of the epidermal cells (16). As epidermal cells progress from the basal cell layer and keratinize to form the stratum corneum, they undergo major changes in their lipid composition. Phospholipids and sterols are constituents of cellular membranes. The epidermal phospholipids represent a major part of the basal cell lipids (45%), but phospholipase-mediated degradation results in a low phospholipid content in the upper cell layers. In contrast, neutral lipids increase in successive upper layers. Squalene, synthesized in the lower epidermis, is converted into vitamin D precursors or cholesterol, followed by sulfation and esterification. Cho-

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lesterol sulfate plays a role in the organization of membrane lipids, and cholesterol sulfatase regulates this function (17). During the process of keratinization, elongation and desaturation of fatty acids occur, sterols are esterified and sulfated, and sphingolipids become acylated. Striking amounts of sphingolipids are present as the cells mature (15). These lipids consist predominantly of sphingomyelins, glucosylceramides, and ceramides. Glucosylceramide A accounts for about 50% of the total mammalian epidermal glycolipids. Of the glucosylceramides, 50% are acylglycosylceramides, and the most prominent fatty acid esterified to this carbohydrate is the essential fatty acid linoleic acid (C18:2) (18). This unsaturated fatty acid, essential to normal epidermal barrier function, is highly susceptible to oxidative injury. All other ceramides are highly saturated; therefore, these molecules seem to be well suited to resisting oxidative damage. Together with sterols the sphingolipids form stable bilayers in the intercellular space of the stratum corneum, thereby forming a barrier to transepidermal water loss and replacing the phospholipids as they are lost with differentiation (14). B. Sebaceous Gland Lipids Quantitatively, lipids produced by sebaceous glands (sebum) are much more significant than lipids derived from the keratinizing epidermis, although on a weight basis epidermal cells are more active in lipid biosynthesis than dermal cells. The amount of recovered surface lipids is directly proportional to the number of sebaceous glands in the skin, which vary widely in size and density of distribution on the surface of the body. On the scalp and face, sebaceous glands are abundant, while in other locations (limbs and trunk apart from the midline) they are less numerous. Surface lipids collected from areas rich in sebaceous glands represent the approximate composition of the sebum: triacylglycerols and free fatty acids 58%, wax esters 26%, squalene 12%, cholesterol esters 3%, and cholesterol 1.5%. Pure sebum has no free fatty acids. Free fatty acids result primarily from triacylglycerol hydrolysis by Propionibacteria in the sebaceous glands and on the skin surface. Sebaceous gland fatty acids consist of saturated and unsaturated, branched and unbranched structures with chain lengths ranging from six to 28 carbons (19). However, most of these fatty acids are composed of C12 or C14 acids. The concentration of triacylglycerols and free fatty acids varies, depending upon the activity of the bacteria. Free fatty acids may compose up to 20% of the sebum lipid mixture in humans, and about 50% of the sebum fatty acids are unsaturated, providing substantial substrate for lipid peroxidation processes. In humans, surface wax esters and sterols originate uniquely from the sebaceous glands (20). It should be noted that sebum composition is species-specific (21). In humans, sebum fatty acid composition may even differ greatly between individuals (22). In contrast to most mammalian skin, large amounts of squalene, a

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sterol compound with six unsaturated double bonds, accumulates in human skin. Squalene represents about 10–15% of total skin surface lipids. The physiological presence of a high sterol concentration in the outer skin layers may serve a protective function, e.g., that of an antioxidant. Recently it was demonstrated that human stratum corneum cholesterol effectively competes with the peroxidation of other human skin lipids such as unsaturated ceramides and free fatty acids (23) as well as unsaturated phospholipids (24). On the other hand, some sterol compounds may exert phototoxic reactions in skin by acting as photosensitizers. 5,7,9(11),22-Cholestartien-3-beta-ol, which is present in human skin surface lipids, was identified as a photosensitizer, generating singlet oxygen (25). Since 25% of the total skin surface lipids are unsaturated, the physiological presence of lipid peroxidation products at the epidermal surface should be suspected. Species differences do also exist in skin content of lipid peroxidation products; e.g., in contrast to the rat, only small amounts of lipid peroxidation products are found in mice, guinea pigs, and hamsters. The exceedingly high lipid peroxide content of the rat is explained by the fact that the rat is the only common rodent that excretes large amounts of linoleic acid in the skin surface (2,26). But in healthy human skin also, lipid peroxidation products are detected in considerable quantities (27–29).

IV. ENVIRONMENTAL OXIDANTS Free radical mediated reactions have been discussed as one of the basic mechanisms of xenobiotic toxicity (30–33). Induction of oxidative stress may be an epiphenomenon or just a consequence of tissue damage following exposure to xenobiotic substances; however, in some cases it may represent a primary mechanism of tissue injury. Exogenous sources of oxidizing chemicals comprise redox cycling xenobiotics, charge transfer complexes, oxidizing and alkylating chemicals, redox-active metal ions, free radical initiators, and photosensitizers as well as ultraviolet and ionizing irradiation. The activity of a chemical toxicant is sometimes a complex balance between metabolic activation by cytochrome P450 monooxygenase and detoxification by enzymes such as glutathione-S-transferase. Skin can be a target organ for toxic effects of xenobiotics, and cutaneous metabolism via cytochrome P450, peroxidases, and flavoprotein dehydrogenases can lead to bioactivation of xenobiotics to pro-oxidant substances and induce peroxidation of skin lipids and proteins. Macromolecular carbonyls, as markers for oxidatively damaged proteins in human stratum corneum, were suggested to be a sensitive indicator of skin exposure to environmental oxidants such as ozone, ultraviolet radiation, and hypochlorous acid (34). There is experimental evidence that reactive oxidants are involved as trigger factors as well as mediators of inflammatory reactions. For instance, free radicals can trigger cutaneous inflammation (35–

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37), and the cellular redox status modulates the expression of proinflammatory cytokines. In particular, the upregulation of interleukin 8 expression through an oxidative stress indicates that reactive oxidants participate in proinflammatory cytokine activation in human cells (38). The radical mechanism has gained interest in the discussion of the mechanism of hapten–protein binding (39). Studies indicating that radical reactions are important for haptens containing hydroperoxide groups have been published (40–43). A.

Quinones, Imminium Ions, and Alkylating Agents

The charge transfer mechanism for generation of reactive oxygen species was suggested as a unifying concept for the mechanism of action of quinone drugs, metal complexes, imminium ions, and alkylating agents (44–46). A charge transfer complex is formed when an electron donor interacts with an electron acceptor in which electronic charge passes from the donor to the acceptor. By this mechanism, the binding of a xenobiotic to a target biomolecule results in formation of a conducting salt that can catalytically generate reactive oxygen species via electron transfer (46). Particularly in the case of imminium ions, the positive charge enhances electron abstraction from donor molecules (47). Chemicals that may cause skin damage (e.g., contact dermatitis) via the generation of reactive oxidants, induction of lipid peroxidation, and/or inhibition of antioxidant systems are quinones, as demonstrated for doxorubicin and adriamycin (48–53), and imminium compounds such as the herbicides paraquat and diquat (54–56). Reactive oxygen radicals may be generated from quinones as a result of semiquinone radical generation via autoxidation of hydroquinones, metabolic activation, or comproportionation. Paraquat (1,1′-dimethyl-4,4′-bipyridyl) and diquat (1,1′ethylene-2,2′-bipyridyl) are nonselective contact herbicides. They are almost exclusively used as a dichloride or dibromide salt, respectively. Both the herbicidal and toxicological properties of paraquat are dependent on the ability of the parent cation to undergo a single-electron addition to form a free radical that reacts with molecular oxygen to re-form the cation and concomitantly produce a superoxide. Diquat undergoes a single-electron addition to form a radical that also reacts with molecular oxygen to re-form diquat and concomitantly produce a superoxide anion. A further example of a xenobiotic that causes skin damage via mechanisms involving oxidative stress is the chemical warfare agent and vesicant sulfur mustard [bis(2-chloroethyl)sulfide], which is a potential threat to both battlefield and civilian populations. This alkylating agent depletes cellular glutathione, forms stable DNA adducts, and may also stimulate formation of reactive oxidants (57– 59). B. Ultraviolet Radiation Biological effects induced by UVB and UVA radiation involve different chromophores leading to different photobiological responses. The bulk of UVB-mediated

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photochemistry of cellular components occurs by direct excitation and is almost an oxygen-independent process. However, there is also evidence of oxidative damage triggered by UVB. In contrast to UVB, UVA is very efficient in inducing oxidative stress in human skin cells and in mimicking the action of ionizing irradiation. The effects of UVA and visible radiation seem to be mediated by endogenous photosensitizers, which comprise flavins, tryptophan and tyrosine, ubiquinones, porphyrins, NADH and NADPH, nucleosides, and iron sulfur clusters. Exogenous photosensitizers deriving from environmental pollution, plants, drugs, and cosmetics may become biologically relevant as photosensitizers as well. Ultraviolet radiation increases thiobarbituric acid reactive substances (TBARS) in the skin of hairless mice (60–65), in isolated human skin cells (66– 69), in human skin (70,71), and in human skin surface lipids (72,73). However, the TBARS assay is notoriously fraught with artifacts and has a relatively high background (74). More substantial information on the occurrence of lipid peroxidation in skin cells was obtained by measurement of lipid hydroperoxides, malondialdehyde (MDA), and 4-hydroxynonenal by high performance liquid chromatography (HPLC) (75). Several different types of lipid hydroperoxides were increased in high concentration immediately after simulated solar radiation of mouse skin. Before irradiation no hydroperoxides were measurable in mouse skin (76). The action spectrum of UV radiation-induced lipid peroxidation in human skin fibroblasts shows a continuously decreasing response from 280 to 420 nm. The UVB/UVA effectiveness ratio for this effect ranges from 10 to 100 (69). Given the solar spectral distribution, the amount of UVA reaching the earth’s surface is about 20 times greater than that of UVB. Thus solar UVA is a significant factor in inducing skin lipid peroxidation (69). Low doses of UVA (suberythemal and minimally erythemal doses) generated MDA and 4-hydroxyalkenals in skin fibroblasts (75) and lipid hydroperoxides in keratinocytes (77) as determined by HPLC. Diminished barrier function (e.g., increase in transepidermal water loss) in response to ultraviolet irradiation has been reported in animal and human skin (77–84). C. Hydroperoxides, Peroxides, and Lipid Peroxidation Products Organic peroxides are widely used as initiators for free radical polymerization of monomers to thermoplastic polymers, for curing thermoset polyester resins, and for cross-linking elastomers and polyethylene. Organic peroxides are compounds containing carbon and one or more peroxy bonds in the molecule. Their usefulness is based on their thermal instability. They decompose readily into highly reactive free radicals. The major structural classes of organic peroxides are diacyl peroxides, peroxyacids, ketone peroxides, dialkyl peroxides, peroxy esters, hydroxy peroxides, and peroxydicarbonates. The main toxic effect is irritation of the skin, mucous membranes, and eyes (85–87), and some hydroperoxides

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and peroxides are tumor promoters. The cellular effects of tert-butyl-hydroperoxide, cumene hydroperoxide, dicumyl peroxide, and benzoyl peroxide were examined in human keratinocytes (88–91). These hydroperoxides and peroxides undergo metal ion dependent activation in keratinocytes to form alkoxyl, alkyl, and aryl radicals. But other non-radical products of peroxide metabolism, such as electrophilic quinone methides, are toxicologically important reactive intermediates of some hydroperoxides (92). The products of lipid peroxidation (lipid hydroperoxides and their degradation products, e.g., malondialdehyde, 2-alkenals, and 4-hydroxyalkenals) are also toxic; systemic toxicity of various lipid peroxidation products has been documented (93–95). Lipid peroxidation products also exert toxic effects in mammalian skin (96–100). Species differences in skin irritability to peroxidation products are noted; e.g., the skin of guinea pigs is usually more sensitive to irritation by lipid peroxidation products than human skin (96). In conclusion, several environmental organic peroxides and hydroperoxides as well as endogenous lipid peroxidation products are potent skin irritants and may cause disturbances of the epidermal barrier integrity. However, detailed clinical studies on the impairment of skin barrier function by these compounds are not yet available. D.

Oxidizing Gases and Dusts

Volatile chemicals and dust are common sources of environmental exposure, and some of these substances cause oxidative damage. Oxidants of photochemical smog generally include ozone, nitrogen dioxide, peroxyacyl nitrates, hydrogen peroxide, alkyl peroxides, nitrous and nitric acids, formaldehyde and formic acid, and traces of other compounds. Peroxyacetyl nitrate and peroxypropionyl nitrate are the two most abundant peroxyacyl nitrates, their concentration being highest in photochemical smog (101), and they are potential eye irritants. Typical indoor air pollutants are the nitrogen oxides, tobacco smoke, volatile organic peroxides, and mineral fibers such as asbestos. Of all the gaseous oxidants the toxicity of ozone and nitrogen dioxide has been studied in detail because of the abundance and earlier identification of these gases as constituents of photochemical smog and as having enormous potential effects on humans and ecosystems. Ozone and nitrogen dioxide are toxic gases, and their inhalation can cause respiratory irritation and injury; therefore, mainly the pulmonary effects of these gases have been investigated. The toxic effects of the gases are attributed to their ability to induce free radical reactions in the biological system. E.

Ozone

Ozone has long been recognized as a natural constituent of the atmosphere. Ozone in the upper atmosphere (stratosphere) occurs naturally and protects skin by fil-

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tering out solar ultraviolet radiation. The main sources for ozone in the troposphere, where about 10% of the atmospheric content is found, are influxes from the stratosphere and photochemical production involving nitrogen oxides and volatile organic substances from natural and anthropogenic sources. Under particular meteorological conditions ozone can be formed within the atmosphere as a result of human activities, e.g., automobile-dependent transportation and petroleumdependent energy production. Ozone at ground level (troposphere) is a noxious, highly reactive oxidant pollutant and is one of the major components of photochemical smog. Ozone concentrations of up to 0.1–0.8 ppm are found in polluted air, and long exposures to 10 ppm cause lethal damage in rodents due to failure of the pulmonary system. The lung and the skin are the organs most directly exposed to ozone. Ozone is probably the most reactive chemical to which these organs are routinely exposed in the environment. The biological effects of ozone are attributed to its ability to cause ozonization, oxidation, and peroxidation of biomolecules, both directly and via secondary toxic reactions (102–104). Numerous studies have documented the effects of ozone on the respiratory tract (102,104,105), but there are only fragmentary reports (106,107) on possible ozone-induced oxidative damage in skin. Recently, ozone-induced oxidative damage in murine epidermal skin was demonstrated. Short-term (2 h) exposure of hairless mice to high ozone concentrations (1–10 ppm) depleted ascorbic acid and α- and γ-tocopherol in the upper epidermal layers and induced lipid peroxidation in a dose-dependent manner. Repeated ozone exposures (1 ppm) for 2 h on six consecutive days decreased only stratum corneum α-tocopherol but did not cause lipid peroxidation (108–110). It remains to be seen whether more sensitive techniques will demonstrate oxidative damage in susceptible skin compartments at more relevant ozone concentrations. Ozone-induced lipid peroxidation in the stratum corneum may be harmful to the skin in two ways. Stratum corneum lipid peroxidation could affect the barrier function of the epidermis, because the stratum corneum lipid composition is essential for barrier integrity (111,112). Perturbation of the stratum corneum lipids may be an important trigger factor for a number of inflammatory skin diseases such as endogenous eczema, psoriasis, and irritant dermatitis (7). Increased lipid peroxidation products in the upper skin layers could trigger inflammatory responses in deeper skin layers. This finding may have implications for the pathophysiology of skin disorders that occur with increasing incidence in polluted air urban areas, such as endogenous eczema (113). F. Nitrogen Dioxide Nitrogen dioxide is one of the important indoor oxidizing gases, and the oxides of nitrogen, generally a mixture of nitrogen oxide and nitrogen dioxide, are produced by both natural and human processes. They are produced during combus-

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tion of fossil fuels such as oil, coal, and gasoline. Indoor concentrations of nitrogen dioxide can be found up to peak values of 0.1–0.5 ppm, e.g., in kitchens with gas stoves or kerosene gas heaters, but average values in homes are 0.025– 0.075 ppm. Short-term exposure (4 h) of healthy human skin to low concentrations of nitrogen dioxide (0.023–0.030 ppm) caused disturbances of epidermal barrier function with increased transepidermal water loss (114). Nitrogen dioxide is known to cause oxidative damage resulting in the generation of reactive oxidants that may oxidize amino acids in proteins and lipids.

V.

OUTLOOK

Environmental oxidants are potential risk factors for human health, and the skin is one of the main target organs, but the exact role of environmental oxidants in cutaneous pathophysiology has not yet been established. There are only a few reports showing that exposure to ozone or nitrogen dioxide causes skin damage such as lipid peroxidation and impairment of barrier function (e.g., increase of transepidermal water loss). Simultaneous or subsequent exposures to environmental oxidants may lead to an additive or synergistic effect. A clinically relevant reaction may occur in skin exposed to multiple oxidant factors (e.g., a combination of ozone, nitrous oxide, peroxyacyl nitrates, hydroperoxides, and ultraviolet light), although each factor alone would elicit only a minor reaction or none. On the other hand, the outcome of multiple subsequent or simultaneous exposures may cause a hardening effect, resulting in a decreased response. Furthermore, endogenous factors such as genetic predisposition and aging may significantly influence skin sensitivity to exogenous and environmental insults. It was shown that chronologically aged skin displays altered epidermal permeability, with increased susceptibility to environmental pollutants (115). In view of all this, detailed studies are required to investigate how chronic or intermittent parallel exposures to multiple environmental oxidants affects the skin’s antioxidant system, causes oxidative damage at which molecular level, which structural and functional deficiencies will result, and what role endogenous factors play in this scenario. These investigations may open up new ways in our understanding of environmentally triggered skin diseases and how to treat them most efficiently.

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104. Pryor WA, Church DF. Aldehydes, hydrogen peroxide and organic radicals as mediators of ozone toxicity. Free Radical Biol Med 1993; 122:483–486. 105. Lippmann M. Health effects of ozone. A critical review. JAPCA 1989; 39:672– 695. 106. Podda M, Koh B, Cross CE, Packer L. Ozone exposure depletes lipophilic antioxidants in murine skin. J Invest Dermatol 1995; 104:639A. 107. Fukase O, Hashimoto K. The effects of exposure to ozone on collagen in lungs and skin. Nippon Eiseigaku Zasshi 1982; 37:694–700. 108. Thiele JJ, Traber MG, Tsang K, Cross CE, Packer L. In vivo exposure to ozone depletes vitamins C and E and induces lipid peroxidation in epidermal layers of murine skin. Free Radical Biol Med 1997; 23:385–391. 109. Thiele JJ, Traber MG, Polefka TG, Cross CE, Packer L. Ozone exposure depletes vitamin E and induces lipid peroxidation in murine stratum corneum. J Invest Dermatol 1997; 108:753–757. 110. Thiele JJ, Traber MG, Packer L. Depletion of human stratum corneum vitamin E: an early and sensitive in vivo marker of UV induced photo-oxidation. J Invest Dermatol 1998; 110:756–761. 111. Bouwstra JA, Gooris GS, Cheng K, Weerheim A, Brass W, Poence M. Phase behavior of isolated skin lipids. J Lipid Res 1996; 37:999–1011. 112. Elias PM, Feingold KR. Epidermal lipids, barrier function and desquamation. J Invest Dermatol 1983; 80:44–49. 113. Schultz-Larsen F. Atopic dermatitis: a genetic-epidemiologic study in a population based twin sample. J Am Acad Dermatol 1993; 28:719–723. 114. Eberlein-Ko¨nig B, Przybilla B, Ku¨hnl P, Pechak J, Gebefu¨gi I, Kleinschmidt J, Ring J. Influence of airborne nitrogen dioxide or formaldehyde on parameters of skin function and cellular activation in patients with atopic eczema and control subjects. J Allergy Clin Immunol 1998; 101:141–143. 115. Ghadially R, Brown BE, Sequeira-Martin SM, Feingold KR, Elias P. The aged epidermal permeability barrier. J Clin Invest 1995; 95:2281–2290.

6 Oxidative Stress and Loose Coupling/Uncoupling Guido Zimmer Johann Wolfgang Goethe University, Frankfurt, Germany

I. INTRODUCTION In connection with the term ‘‘oxidative stress,’’ we consider ‘‘loose coupling/ uncoupling’’ in relation to mitochondria. In pathophysiology, ‘‘oxidative stress’’ basically means an imbalance of redox potential accompanied by a surplus of partially reduced oxygen atoms, for example, O2•⫺ , which to a great extent has used up the reduced forms of cellular defense, i.e., glutathione 2 RSH → RSSR. In other cases, the oxidation of reduced glutathione is not in the foreground. Further antioxidative enzymatic systems, i.e., glutatione peroxidase, superoxide dismutase, and catalase, and, apart from vitamins C and E and provitamin carotene, other thiol-containing smaller molecules, thioredoxin, and lipoic acid take part in the multifold scenery. Thioredoxin peroxidase has recently been added to this arsenal (1). But even this pool may become exhausted. Since the oxygen is no longer completely, but only partially, reduced, there is also diminished formation of water due to respiration in the respiratory chain. Instead, the protons are no longer taken up into the mitochondria and used to form ATP through the ATP synthase. However, protons are produced at a surplus rate due to increased ATP splitting. In addition there is an increase in the production of lactic acid from pyruvate or a step preceding the pyruvate dehydrogenase serving as the last reaction of the physiologically functioning glycolytic chain. Therefore, the intracellular pH is shifted downward from 7.2–7.4 to 7.0 and even lower. Thus, in the clinical situation the complexity of oxidative stress can in 95

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many cases become reduced to the situation of ‘‘lack of oxygen.’’ This is the pathophysiology we discuss within the framework of this chapter.

II.

‘‘LACK OF OXYGEN’’ EQUALS ISCHEMIA

For a long time, the lack of oxygen during hypoxia or ischemia has been considered to be the sole and exclusive cause of tissue damage (2–4). If only the blood supply could be restored within a certain time, then the tissue would recover. If the time was extended beyond the limit, reperfusion with oxygen-containing solution could no longer salvage tissue; the result then was irreversible damage after 4–6 h of ischemia. The time limit was set by different investigators from the beginning of ischemia; most of them agreed on a maximum of 4–6 h of ischemia resulting in irreversible damage. Diagnosis of ‘‘irreversibility’’ was mostly accomplished by electron microscopic work that relied completely on the available methods. The conventional embedment procedures, however, led to denaturation of the membrane proteins and thereby to distortion of membrane structure (5). In particular, when osmium tetroxide is used for fixation of the tissue, extensive structural rearrangements result, which already distort the ‘‘control’’ membranes even without any oxidative stress. In the absence of reliable control specimens, the conclusions drawn were necessarily ambiguous, especially with respect to judgments concerning the reversibility or irreversibility of damage. The only method known to avoid extensive structural change of membranes is the ‘‘low denaturation’’ technique developed by Sjo¨strand (5). Using this technique for the electron microscopic analysis of mitochondrial membranes, a generally intact structure even after 6 h of ischemia was revealed (6). This finding made it possible, for the first time, to nail down the ensuing damage occurring during conventional reperfusion as reperfusion damage (6).

III. ‘‘LOOSE COUPLING’’ OF MITOCHONDRIA In general, the overall intact structure of heart mitochondria after 6 h of ischemia does not preclude observation of slight, completely reversible changes. The usually planar cristae were transformed into cristae exhibiting a zigzag pattern in cross sections (6). The zigzag structures were deduced to result from a shift of material from one membrane region into a neighboring region. Thus there was a regular ‘‘accumulation’’ of membrane protein at one site resulting in ‘‘dilution’’ at a neighboring site. Such redistribution of membranous materials, especially proteins, was early recognized as resulting in permeability changes with increased leakiness to small

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ions (6). Such close apposition of membrane proteins could then also favor aggregation and, most significantly, the formation of covalent bonds between the aggregated proteins. Among these, disulfide bridges frequently appear and have been shown to occur in particular in one of the most abundant transport proteins, the AdN translocator (7). Moreover, there is evidence that in the hydrophobic interior of the ATP synthase molecule, a disulfide bridge between subunits and/or within subunit b is present under nearly all conditions of experimentation (8). More recent experimental results revealed a cross-link between Cys-197 of subunit b and Cys-91 of F1γ formed by addition of diamide. Both cysteines (9) are located in the ‘‘stalk’’ region of ATP synthase. It was also observed (9) that in the presence of ∆µH⫹ generated by respiration, neither cross-linking between the above-mentioned cysteines nor the associated effects on oligomycin-sensitive decay of ∆µH⫹ and the decoupling of ATP synthesis occurred. These observations were considered evidence of close contacts of the two protein regions on subunit b and F1γ in the resting state of the enzyme only. A structural change is imposed on these subunits by ∆µH⫹ , bringing those protein regions apart so that they can no longer become engaged in the formation of disulfide. ∆µH⫹ collapsing agents restored the cross-link formed by these two cysteines. This work coincides with the following observations. With the isolated, highly purified ATP synthase (10), our own experiments have revealed a significant decrease in the labeling of the regions of around 25 kDa as well as 10 kDa in the presence of oligomycin (11). This labeling was accomplished with radioactive [14C] α-lipoic acid under the conditions of Godinot et al. (8). The 10 kDa region includes the binding site for oligomycin. This means also that the purified enzyme behaves in a way that could be anticipated from the work with mitochondrial particles. Free sulfhydryl groups in the ‘‘b’’ subunit appear very important for the functioning of ATP synthase in the respiring mitochondrion. Conversely, nonrespiring mitochondria are prone to functional decay of ATP synthesis by cross-linking within subunit b and F1γ. Moreover, the region of low molecular weight, ‘‘c’’ subunits, appears to be involved (see above). Most recent results from yeast H⫹ ATP synthase, which holds two copies of subunit b per molecule, provided evidence for a spontaneous disulfide bridge between two subunits b (12). Obviously, the molecular surroundings (also primary structure) around the cysteines is decisive for their reactivity (13–15). The aforementioned results add to previous and recent findings: F1 F0 ATP synthase appears particularly vulnerable to, for instance, H2 O2 (16–18). Changes in mitochondrial permeability after ischemia of varying time (4– 12 h) have been observed in rat skeletal muscle mitochondria by our group (19,20). The only sign of some structural or functional disorganization of the kind described above could then be found in changes in respiration: respiratory

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control ratios (RCR) were decreased, which was generally due to an increase in state 4 of respiration. The state 3/state 4 ratio, accordingly, was decreased. Permeability changes of that kind, which are due to 1. Redistribution of membranous materials and aggregation (5) 2. Formation of disulfide bonds and, probably, increasingly hydrophobic interactions will first result in increased leakiness to small ions. In parallel, the membrane gains in polarity, particularly in the hydrophobic core. It was found that oxidative stress induced a large increase in hydrocarbon core molecular volume in the cardiac membrane bilayer (21). The interface and the adjacent region of highly ordered alkyl chains (22) become less ordered. The leakiness may then become symbolized by ‘‘longer existing holes,’’ due to an overall decrease in membrane fluidity in the hydrophobic core (Fig. 1). All this appears to be a gradual development, including first an increased permeability to K⫹ and H⫹ preceding the major changes alluded to above (23,24). Intracellular calcium in vivo, forming a gradient of about four orders of magnitude from the blood (above 1 mmol/L) to the intracellular compartment (0.1 µmol/L), is a main factor of oxidative/hypoxic change (25–27). Evidence has been obtained that during ischemia (4–6 h, heart muscle) the amount of intramitochondrial calcium can be doubled. It is basically around 20 nmol/mg protein, which can be increased in vivo to about 40 nmol/mg protein during ischemia (28) and experimentally to about 80 or 160 nmol/mg protein (29). In brain tissue, moreover, a cytosolic decrease of 0.2 µM of Ca2⫹ was found to become therapeutically relevant (30). Other observations in canine heart cells subjected to 3 h hypoxia in the absence of glucose followed by 5 min reoxygenation resulted in a rise of calcium from 3.3 to 6.6 nmol/mg in response to that stress (31). Similarly, Miyata et al. (32) reported that cardiac myocyte recovery after exposure to anoxia occurred only in cells in which cytosolic or mitochondrial Ca2⫹ remained below 250 nM before reoxygenation. Early during reoxygenation, mitochondrial Ca2⫹ remained higher in cells that revealed hypercontraction (305 nM) than in those that recovered (138 nM). Moreover, cytosolic free Ca2⫹ was measured in single rat heart cells during anoxia and reoxygenation. It was found that the ability of cells to restore Ca2⫹ homeostasis upon reoxygenation is exceeded if concentrations above 5 µM are attained. Sustained recovery was seen only in cells where Ca2⫹ was less than 1.5 µM (33). Even if we assume an opening of the ‘‘permeability transition pore’’ (PTP) under its low conductance state in vitro, the concentration of Ca2⫹ outside mitochondria does not decrease below about 2–5 µM before the opening (34). By contrast, the amounts of Ca2⫹ required to induce the membrane permeability transition (MPT) at the high conductance state are excessive; they appear far from saturated at 25 µM Ca2⫹ matrix free calcium (35). Considering the above

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Figure 1 The scheme exemplifies half of a bilayer membrane structure in (a) a fluid state and (b) a rigidified state. In the fluid system (a), protein passage is impaired by collisions and the tight lipid interface. In the rigid system (b), passage (first stage, H⫹ and K⫹ ions) is possible due to protein aggregation and the loosened lipid interface. Left center: Membrane-spanning protein is symbolized by open bottom. In (a) the protein’s lateral mobility is indicated by the shaded surroundings and the small double arrows. The lipids’ rotatory and wobbling motion is not shown. Instead, the tightness of the polar membrane region is symbolized by the straight and highly ordered part of the interfacial alkyl chains extending to about carbon 7, EPR and NMR spectroscopic results (69–71), and work of Xiang and Anderson (72,73) and Xiang et al. (74). In the rigid system (b), proteins tend to become aggregated (23,26) and the interfacial region becomes less ordered (75). The apolar part of the lipids is becoming less fluid (76). Thus, the existing differences in the borderline barrier (polar, highly ordered, apolar, highly disordered) as in (a) become diminished; instead, we find polar, intermediately ordered, apolar, intermediately disordered as in (b). This is further underlined by the increase in polarity of the apolar membrane core (77,78); these structural changes gradually diminish the barrier and render the membrane more and more leaky. In this scheme, observations on plasma membrane (77,78) confirmed by permeability studies (Zimmer et al., this volume) and on intracellular (mitochondrial) membrane (23,26,75,76) have been used.

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results as reported in Refs. 28,30–33 we realize that such a range of concentrations appears to be nearly two orders of magnitude (80-fold) lower than that determined for the k0.5 of 16 µM for Ca2⫹ at pH 7 (36) for opening the mitochondrial permeability transition pore or megachannel that allows molecules below 1.5 kDa to unspecifically permeate. For the above reasons, we suggested that the opening of the permeability transition pore (PTP) appears irrelevant for the reversible ischemia/reperfusion situation (24). Obviously, opening the PTP belongs to a pathophysiology leading to cellular apoptosis (37). Therefore, it was proposed that opening of the PTP may constitute an irreversible state of the effector phase of apoptosis (38). Accordingly, membrane permeability transition has been suggested to be decisive for reversibility or irreversibility of membrane damage (39). This view is similar to our own (24), since in the case of reversibility, the MPT pore (MPTP) has necessarily to be closed, at least for the majority of mitochondria. This can be attained up to a certain range of intracellular calcium (33): k0.5 for opening of the pore of approximately 16 µM, compared to the normal physiological solution, amounts to a surplus of intracellular cytosolic calcium of 150- to 160-fold. Unless such an intracellular range is reached, the MPTP may remain closed. At cytosolic Ca2⫹ above 16 µM, however, the majority of mitochondria will be overloaded by excess Ca2⫹ to such an extent that the pore opens. This may then result in the initiation of irreversibility where mitochondria are overloaded with calcium without being able to return to a cytosolic equilibrium suited for cellular survival and indeed become a ‘‘prelude to cell death,’’ or apoptosis (40). Investigations on sequestration of iontophoretically injected Ca2⫹ by living endothelial cells have revealed that even after long injection periods, mitochondria are not uncoupled, contrary to what is seen in isolated mitochondria (41). Conversely, measurements of free Ca2⫹ in single heart cells have confirmed the stability of Ca2⫹ of 2.5 ⫻ 10⫺7 M for 16 min even after metabolic interruption with cyanide and deoxyglucose. Before cell death, internal Ca2⫹ increased to 4 ⫻ 10⫺6 to 5 ⫻ 10⫺6 M. It was concluded that since severe injury to the cell occurs before the free Ca2⫹ concentration rises above 1 ⫻ 10⫺7 to 3 ⫻ 10⫺7 M, cell damage seems to be the cause, not a consequence, of a rise in free Ca2⫹ (42). Even during reoxygenation or reperfusion, such overloading of mitochondria due to excessive amounts of cytosolic Ca2⫹ appears improbable, provided the conditions for reoxygenation or reperfusion appear to be properly controlled (6,28). The question we have to deal with is that of time of ischemia or hypoxia versus damage. This specific problem of how long a certain tissue can stand a time of ‘‘no flow’’ or ‘‘low flow’’ is, however, not exactly known. For example, heart tissue can survive experimentally set ischemic/hypoxic conditions for more than 6 h (6). This contrasts very much with previous, though recently repeated (43), views.

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All measures that can be taken against the situation of oxidative stress, loose coupling, or the permeability increase that occurs during the reversible extent of Ca2⫹ accumulation in the cytosol have been described in detail during the last two decades (6,44–68). There is a chance to treat the reversible phase of structural change that is also becoming evident from several hours after induction of apoptosis in acute cancerous blood cells (lymphoblasts) until signs of plasma membrane derangement become apparent by spin labeling with 16-doxylstearate (see Chap. 20). In many cases the initiation of the apoptotic process, therefore, appears to originate in the intracellular region, i.e., in the milieu of organelles (mitochondria). The lack of oxygen, leading to a lack of energetic support of ATP-dependent Na⫹ /K⫹ exchange in the plasma membrane, which subsequently results in activation of H⫹ /Na⫹ exchange by increasing intracellular acidosis, is not directly visible as gross structural changes appearing in EPR spin labeling of the plasma membrane. Sodium ions then become consecutively exported by means of the Ca2⫹ /Na⫹ exchanger, which initiates the intracellular structural rearrangements.

ACKNOWLEDGMENTS I thank Dr. Klaus Zwicker for critically reading this chapter and for his valuable suggestions. Annette Hochberger patiently provided the many drafts of this chapter and carried out the expert artwork. Work in the author’s laboratory on mitochondrial ATP-synthase has been supported over many years by the Deutsche Forschungsgemeinschaft. EPR studies were supported by Paul and Ursula Klein Stiftung.

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65. Assadnazari H, Zimmer G, Freisleben H-J, Werk W, Leibfritz D. Cardioprotective efficiency of dihydrolipoic acid in working rat hearts during hypoxia and reoxygenation, 31P NMR investigations. Arzneim-Forsch/Drug Res 1993; 43:425–432. 66. Haramaki N, Packer L, Assadnazari H, Zimmer G. Cardiac recovery during postischemic reperfusion is improved by combination of vitamin E and dihydrolipoic acid. Biochem Biophys Res Commun 1993; 196:1101–1107. 67. Zimmer G, Beikler T-K, Schneider M, Ibel J, Tritschler H, Ulrich H. Dose/response curves of lipoic acid R- and S-forms in the working rat heart during reoxygenation: superiority of the R-enantiomer in enhancement of aortic flow. J Mol Cell Cardiol 1995; 27:1895–1903. 68. Bolli R, Zughaib M, Li X-Y, Tang X-L, Sun J-Z, Triana JF, McCay PB. Recurrent ischemia in the canine heart causes recurrent bursts of free radical production that have a cumulative effect on contractile function. A pathophysiological basis for chronic myocardial ‘‘stunning.’’ J Clin Invest 1995; 96:1066–1084. 69. Gaffney BJ, McConnell HM. The paramagnetic resonance spectra of spin labels in phospholipid membranes. J Mag Res 1974; 16:1–28. 70. Seelig A, Seelig J. The dynamic structure of fatty acyl chains in a phospholipid bilayer measured by deuterium magnetic resonance. Biochemistry 1974; 13:4839– 4845. 71. McConnell HM. Molecular motion in biological membranes. In: Berliner LJ, ed. Spin Labeling. Theory and Applications. New York: Academic Press, 1976:525– 560. 72. Xiang T-X, Anderson BD. The relationship between permeant size and permeability in lipid bilayer membranes. J Membrane Biol 1994; 140:111–122. 73. Xiang T-X, Anderson BD. Permeability of acetic acid across gel and liquid-crystalline lipid bilayers conforms to free-surface-area theory. Biophys J 1997; 72:223– 237. 74. Xiang T-X, Xu YH, Anderson BD. The barrier domain for solute permeation varies with lipid bilayer phase structure. J Membrane Biol 1998; 165:77–90. 75. Zwicker K, Dikalov S, Matuschka S, Mainka L, Hofmann M, Khramtsov V, Zimmer G. Oxygen radical generation and enzymatic properties of mitochondria in hypoxia/ reoxygenation. Arzneim-Forsch/Drug Res 1998; 48:629–636. 76. Scheer B, Zimmer G. Dihydrolipoic acid prevents hypoxia/reoxygenation and peroxidative damage in rat heart mitochondria. Arch Biochem Biophys 1993; 302:385– 390. 77. Biesert L, Adamski M, Zimmer G, Suhartono H, Fuchs J, Unkelbach U, Mehlhorn RJ, Hideg K, Milbradt R, Ru¨bsamen-Waigmann H. Anti-human immunodeficiency virus (HIV) drug Hoe/Bay 946 increases membrane hydophobicity of human lymphocytes and specifically suppresses HIV-protein synthesis. Med Microbiol Immunol 1990; 179:307–321. 78. Gu¨ldu¨tuna S, Zimmer G, Imhof M, Bhatti S, You T, Leuschner U. Molecular aspects of membrane stabilization by ursodeoxycholate. Gastroenterology 1993; 104:1736– 1744.

7 Uncouplers of Oxidative Phosphorylation Activities and Physiological Significance Yasuo Shinohara and Hiroshi Terada University of Tokushima, Tokushima, Japan

I. INTRODUCTION Bioenergy necessary for cell growth, cell division, and cell life is mainly provided as light energy from the sun or as food nutrients. However, most living organisms can use energy only in the form of chemical energy released by hydrolysis of ATP. Thus, their energy source must be converted into chemical energy in the form of ATP. The cellular organelles in higher organisms such as chloroplasts and mitochondria are responsible for this conversion. In these organelles, energy obtained in the form of light or nutrients is first converted into a proton electrochemical gradient (∆µH⫹) across their membranes by a redox enzyme complex consisting of the electron transport chain (in the case of mitochondria, also called the respiratory chain). Using the H⫹ gradient generated as a driving force, ATP can be synthesized from ADP and inorganic phosphate (Pi ) by ATP synthase. Energy conversions in chloroplasts and mitochondria are referred to as photophosphorylation and oxidative phosphorylation, respectively. Uncouplers are reagents that release the coupling between reactions of electron transport and phosphorylation of ADP by ATP synthase, thus inhibiting ATP synthesis. Inhibitors of the electron transfer chain, such as cyanide and antimycin A, block electron transport, while inhibitors of energy transfer, such as oligomycin, inhibit ATP synthase activity and its reverse reaction of ATPase activity. However, uncouplers do not inhibit these two machineries necessary for ATP synthesis. It is noteworthy that uncouplers inhibit ATP synthesis by dissipating ∆µH⫹ across the membranes (Fig. 1). 107

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Figure 1 Site of action of the uncouplers of oxidative phosphorylation. Unlike inhibitors of respiratory chain such as rotenone, antimycin A, and CN⫺ and those of ATP synthase such as oligomycin and aurovertin, uncouplers are thought to show their activities in the membrane rather than acting on particular proteins.

To achieve efficient energy conversion, energy-converting membranes are highly resistant to transport of ions, especially H⫹. The H⫹ in the cytosol enters the mitochondrial matrix space via the H⫹ channel in the H⫹-ATPase (ATP synthase) of the inner mitochondrial membrane for synthesis of ATP. Uncoupling occurs when the H⫹ electrochemical gradient across the inner mitochondrial membrane is collapsed by permeabilization to H⫹. Accordingly, uncoupling in mitochondria is accompanied by (1) acceleration of state 4 respiration, (2) inhibition of the ATP s Pi exchange reaction, (3) activation of the latent H⫹-ATPase activity, and (4) dissipation of transmembrane electrical potential difference, ∆ψ. These are the general characteristics of uncoupling. In a strict sense, uncoupling is triggered by an increase in the H⫹ permeability of the energy-transducing membrane without damage to the membrane structure. However, increase in the permeability of ions other than H⫹, such as permeability transition (PT) induction (see Sec. IV.B) and ionophore-mediated ion transport, cause dissipation of ∆ψ, leading to uncoupling. In addition, there is a unique ‘‘uncoupling’’ in which the coupling reaction of electron transport with phosphorylation is released, while ∆µH⫹ is not dissipated. The molecular mecha-

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nisms of uncoupling and the structural characteristics of uncouplers have been studied extensively (1–3), but the mechanisms are not yet fully understood. In this chapter we review recent developments in studies of various types of uncoupling.

II.

PROTONOPHORIC UNCOUPLERS

A.

Weakly Acidic Protonophores

Almost all well-known potent uncouplers are hydrophobic weak acids, and their activities are due to protonophoric actions in energy-transducing membranes. The chemical structures and properties of well-known weakly acidic uncouplers are summarized in Table 1. All these uncouplers have acidic dissociation groups such as phenolic OH and anilino NH groups, and electron-withdrawing groups such as ECN, ENO2 , ECF3 , and ECHCC(CN)2 . In addition, they have hydrophobic groups such as tert-butyl, Cl, and CF3 . Quantitative structure–activity relationship studies showed that high hydrophobicity and moderate electron-withdrawing abilities are important for their uncoupling activities (1,5–7). The most powerful uncoupler known to date is SF6847, in which the aciddissociable phenolic OH is surrounded by two bulky tert-butyl groups and the electron-withdrawing malononitrile group [ECH(CN)2] regulates the deprotona-

Table 1 Chemical Structures and Properties of Some Potent Uncouplers

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tion of the phenolic OH (5,6). Its concentration required for uncoupling is as low as 10 nM, showing that only about 0.06 mol of SF6847 is sufficient for one phosphorylation site (ATP synthase) or that less than 0.2 mol SF6847 per respiratory chain complex is sufficient for full induction of uncoupling (8). Therefore, the uncoupler molecule acts as a catalyst rather than attacking a certain protein site. These results support the idea that the uncoupler is a proton conductor across H⫹-impermeable energy-transducing membranes such as inner mitochondrial membranes, acting as a protonophore (1,2). The simplest protonophoric action of an uncoupler, known as the shuttle mechanism, is illustrated schematically in Figure 2, in which the anionic molecular form of the uncoupler, U⫺, binds H⫹ on one side of the membrane and then the neutral molecular UH crosses the membrane. On the other side of the membrane, UH releases H⫹ and the deprotonated U⫺ returns to the original side of the membrane. The cycle of the protonation and deprotonation of the uncoupler molecule results in the transport of H⫹ across the H⫹-impermeable membrane, thus dissipating the ∆µH⫹ . It is noteworthy that the turnover value of SF6847 of about 1000 cycles/s in the mitochondrial membrane is close to the theoretical maximum value according to Brownian motion of the uncoupler molecule (1,8).

Figure 2 Protonophoric action of weakly acidic uncouplers by shuttle mechanism. Both anionic and neutral forms of weakly acidic uncouplers, designated as U⫺ and UH, respectively, exist in the hydrophobic region of the phospholipid bilayer and show protonophoric activity.

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Apparently, the efficiency of the cycle depends on the stability of the polar U⫺ in the hydrophobic phospholipid bilayer membrane. In the case of SF6847, the bulky tert-butyl groups located at both ortho positions to the phenolic OH group hinder the polar anionic phenolic O⫺, and in addition the anionic electron is delocalized by the electron-withdrawing malononitrile group. The electron-withdrawing ability of the malononitrile group is very sophisticated (5,6,9). It is highest when the phenolic benzene ring and the malononitrile group are coplanar and lowest when they are oriented perpendicular to one another. The malononitrile group rotates around the axis of the benzene ring. Its rotational motion in the anionic form is much slower than in its neutral molecular form, resulting in a higher probability of its taking a coplanar configuration in the anionic form (Fig. 3). By this mechanism, the malononitrile group withdraws anionic electrons very efficiently, stabilizing the uncoupler anion in the hydrophobic phospholipid membrane (1,2). Recently, the phenylpyridylamine fungicide fluazinam (for chemical structure, see Fig. 4) was reported to be a more powerful uncoupler than SF6847 (10). However, it is biologically unstable, being rapidly metabolized by conjugation with glutathione during incubation with mitochondria. The action of fluazinam suggests that its uncoupling activity is not based on its binding to the hypothetical uncoupler binding protein, because otherwise it would be more stable in mitochondria (10).

Figure 3 Intramolecular rotation regulates the uncoupling activity of SF6847. SF6847 consists of two flat parts of the benzene ring and the malononitrile moiety connected by a single bond. From 1H NMR analysis and results of molecular orbital calculations, the freedom of intramolecular rotation is directly associated with protonation/deprotonation of the phenolic OH group.

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Figure 4 Chemical structure of fluazinam.

B. Cationic Protonophores As described above, the most powerful uncouplers are hydrophobic weak acids, and little is known about cationic protonophores. AU-1421 is a well-characterized cationic protonophoric uncoupler (11). It is maximally effective in rat liver mitochondria at about 20 µM, and its action is not dependent on Pi in the incubation medium, unlike the other cationic uncouplers described later (see Sec. III.A). These results together with the fact that AU-1421 shows protonophoric activity in mitochondria and liposomes indicate that it induces uncoupling by a protonophoric action like weakly acidic uncouplers. Therefore, its amine moiety seems to be responsible for its protonophoric action. The antitumor alkaloid ellipticine and its isomers, which contain an H ⫹dissociable amine moiety, are also reported to be protonophoric basic uncouplers, but their activities seem to be lower than that of AU-1421 (12). Some other amine compounds are reported to induce uncoupling by a protonophoric action. However, their actions are not well characterized. C. Recouplers Once uncoupling is induced in mitochondria, the uncoupling action continues as long as the uncoupler remains in the membrane in the presence of O2, and it is difficult to recover phosphorylation activity. The most common way to terminate uncoupling is to trap the membrane-bound uncoupler by addition of a chemical with high affinity for it but with no effect on ATP synthesis. Bovine serum albumin is usually used for extraction of weakly acidic uncouplers from the membrane. Starkov et al. (13) reported that the effects of potent uncouplers such as SF6847 and FCCP in mitochondria and chromatophores are reversed by the cholesterol derivative 6-ketocholestanol (kCh; for chemical structure, see Fig. 5). As the reversal action of kCh is not due to trapping of uncouplers, kCh is called a recoupler (13). However, kCh is not always effective; it reverses uncoupling caused by low concentrations of potent uncouplers, but it is ineffective with

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Figure 5 Chemical structure of 6-ketocholestanol.

(1) high concentrations of potent uncouplers, (2) weak uncouplers such as DNP, and (3) the pore opener gramicidin (Fig. 6). Furthermore, kCh was found not to restore the increased electrical conductance of black lipid membranes (BLMs) caused by weakly acidic protonophores but to restore that in proteoliposomes (14). Therefore, its restoration of uncoupling by arrest of proton conductance takes place only in membranes containing proteins, suggesting that uncoupling is mediated by one or more protein components in the membranes and that it

Figure 6 Effect of recoupler 6-ketocholestanol (kCh) on the uncoupling activities of SF6847 and gramicidin (gram). (From Ref. 8.) The membrane potential of mitochondria (mt) was optically monitored with safranin O. SF6847, kCh, and gramicidin (gram) were added at final concentrations of 30 nM, 80 µM, and 2 µg/mg of protein, respectively. (a) kCh completely restored the membrane potential dissipated by SF6847. (b) When kCh was added first, the membrane potential was not changed by addition of SF6847. In both cases, addition of gramicidine caused spontaneous dissipation of the membrane potential.

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modifies the interaction of the uncoupler with these proteins, possibly by changing membrane fluidity (3,14). The fact that uncoupling is mediated by some protein is inconsistent with the fact that the concentration necessary for induction of uncoupling in mitochondria is similar to that for an increase in proton conductivity in BLMs and liposomes (1). This could be explained by supposing that uncoupling is based on a simple shuttle mechanism when the uncoupler concentration is high whereas it is mediated by certain protein at low uncoupler concentration. The recoupler is thought to abolish uncoupling mediated by this protein but not uncoupling due to protonophoric action (3). From the difference between the effects of recouplers on biological membranes and phospholipid bilayer membranes, Starkov et al. (14) classified uncouplers into three types, as shown in Table 2. This classification seems to be available for every type of uncoupling action. Furthermore, the type of uncoupling is likely to depend, at least in part, on the membrane lipid composition having certain membrane fluidity. The above results suggest that there should be an enhancer of uncoupling. Although the bisbenzylisoquinoline alkaloid cepharanthine does not have any effect on oxidative phosphorylation in rat liver mitochondria, it greatly enhances the uncoupling action of SF6847 (Terada et al., unpublished observations). The action of cepharanthine could be due either to facilitation of uncoupler binding to the hypothetical protein or to its stabilization of the uncoupler anion in the membrane by their interaction. Detailed studies are necessary on the action mechanisms of the recoupler and uncoupling enhancers. In addition, it is not apparent which protein, if any, modulates uncoupling. It is interesting that a 30 kDa ‘‘uncoupler-binding protein,’’ which may trigger uncoupling by binding to an uncoupler, was proposed in the 1970s (15).

Table 2 Classification of Uncouplers Category of uncoupler Low concentrations of highly active uncouplers such as SF6847, FCCP, and CCCP Low concentration of FFAs and DNP

Gramicidin and high concentrations of uncouplers

Effective recouplers 6-Ketocholestanol

ADP, carboxyatractyloside and bongkrekic acid in muscle and liver: ADP, ATP, GDP, and GTP in BAT None

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III. ATYPICAL UNCOUPLERS In addition to the protonophoric uncouplers, compounds other than hydrophobic weak acids and bases also induce uncoupling. Although they release respiration and activate H⫹-ATPase, like protonophoric uncouplers, their features of uncoupling are different from those induced by protonophores. Thus, such uncouplers are referred to as atypical uncouplers. A.

Phosphate-Dependent Cationic Uncouplers

Cationic cyanine dyes such as Tri-S-C7 (5) (for chemical structure, see Fig. 7) induce uncoupling in rat liver mitochondria (16). The features of their uncoupling are similar to those of protonophores, such as release of state 4 respiration and oligomycin-inhibited respiration, activation of H⫹-ATPase, and dissipation of ∆ψ. However, Pi is necessary for this uncoupling, and the dose–response curve is sigmoidal, unlike the linear relation in protonophoric uncoupling. In addition, the uncoupling starts when binding of the dye to mitochondria attains a maximum followed by release of the bound dye into the incubation medium, and it is accompanied by swelling of the mitochondria, reflected by a decrease in the optical absorbance of the mitochondrial suspension (16–21). As described later (Sec. IV.B), these features are essentially the same as those induced by Ca2⫹. As cyanine dyes induce a large membrane current fluctuation of BLM containing phosphatidylserine only in the presence of Pi , and as they cause a leakagetype pathway in the membrane, as judged by noise analysis, their uncoupling activity is thought to be due to destabilization or turbulence of the phospholipid bilayer rather than dissipation of ∆ψ due to their direct transfer into the matrix space of mitochondria (17,21). Besides cyanine dyes, a copper–o-phenanthroline complex (22), crystal violet (23), and Cd2⫹ (24) also cause Pi-dependent uncoupling. These are thought to exert their uncoupling effects by a mechanism similar to that of cyanine dyes.

Figure 7 Chemical structure of the cyanine dye tri-S-C7 (5).

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Very recently, they were found to show the uncoupling activities by inducing PT (see Sec. IV.B) in the mitochondrial membrane like Ca2⫹ (25).

B. Decouplers (Slip Inducers) In 1983, Rottenberg (26) reported that the general anesthetics chloroform and halothane induce uncoupling of oxidative phosphorylation in mitochondria. However, these compounds do not collapse the ∆µH⫹. This uncoupling without accompanying dissipation of the ∆µH⫹ was named decoupling, and the compounds inducing decoupling were referred to as decouplers (27). Besides these general anesthetics, free fatty acids (FFAs) (27–29) and the local anesthetic bupivacaine (30–32) are reported to be decouplers that do not affect the ∆µH⫹ but do stimulate state 4 respiration. Much higher concentrations are required for the dissipation of ∆µH⫹ than for full release of mitochondrial respiration (Fig. 8). However, these decouplers are claimed to show uncoupling, not to induce decoupling. The local anesthetic bupivacaine was first reported to be an uncoupler (33), but later to be a decoupler (30,31). After that, it was reported to be just an

Figure 8 Decoupling effect of bupivacaine on mitochondrial oxidative phosphorylation. (From Ref. 39.) Mitochondrial Vox (the velocity of oxygen consumption) and ∆p (proton motive force) were measured in the absence and presence of various concentrations of bupivacaine. Unlike protonophoric uncouplers, bupivacaine at less than 2 mM accelerates mitochondrial respiration (䊉) without dissipation of ∆p (䊊).

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uncoupler (34,35), but subsequently was said to show decoupling activity under certain conditions (32). These discrepant reports could be largely due to differences in the experimental conditions including the integrity of mitochondria. Although the fatty acid palmitate was reported to show decoupling (27–29), it induces uncoupling in the incubation medium without Mg2⫹ (36). 1. Induction of ‘‘Slip’’ of the Primary Pumps It is most possible that the decoupling is due to the slip of the primary pumps, although its molecular mechanism is not well characterized. Decouplers accelerate substrate oxidation or ATP hydrolysis, but they do not stimulate the accompanying extrusion of H⫹ across the inner mitochondrial membrane. As a result, respiration is accelerated without affecting the ∆µH⫹. Molecular slip is discussed in Section III.C. 2. Release of Protons from the Intramembranal Proton Transfer Pathway Rottenberg (37) suggests that both the intramembranal proton pathway and the bulk proton pathway contribute to energy conversion. That is, parallel coupling by these two proton pathways establishes the ∆µH⫹ . Before this proposal, ‘‘localized hypotheses’’ were postulated in which pumped-out H⫹ was directly transferred between redox and ATPase H⫹ pumps (38) or localized at the surface of the membrane, which is not quickly equilibrated with bulk H⫹ (39). The parallel coupling hypothesis is very similar to these localized hypotheses, but the importance of ∆µH⫹ for energy conversion is more sophisticated. Decouplers either specifically collapse the localized H⫹ gradient (localized hypothesis) or release protons in the intramembranal proton pathway (parallel coupling mechanism). As a result, they induce uncoupling without affecting the apparent bulk ∆µH⫹. C. Molecular Slip of Hⴙ Pumps During substrate oxidation, H⫹ is extruded into the cytosolic space by components of the respiratory chain. The H⫹ extruded per electron (H⫹ /e⫺) has long been believed to have a fixed value. However, the results of H⫹ /e⫺ measurements under various conditions suggest that it is changeable (Fig. 9) (see Refs. 40 and 41). When this ratio is decreased by insufficient coupling between substrate oxidation and H⫹-extrusion reactions (i.e., slip), then a smaller fraction of substrate energy is converted to ∆p (⫽∆µH⫹ /F, where F is the Faraday constant). Bupivacaine is reported to cause a slip of cytochrome oxidase in mitochondria (31). Anilinonaphthalene sulfonate (ANS), which converts decoupling to uncoupling due to formation of a leakage-type ion pathway like cyanine dyes plus Pi , abolishes slip of the pump (31).

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Figure 9 Molecular slip in the primary pump. (a) Flows of electrons and protons in a tightly coupled primary pump. (b) When this pump slips, the electron flow/H⫹ efflux ratio increases. The molecular slip might lower the efficiency of energy conversion as well as uncoupling.

D.

Fatty Acids

The uncoupling mechanism of free fatty acids (FFAs) has long been a matter of debate. The fact that FFAs induce uncoupling was first reported in 1956 (42), but their mode of action seems not to have been established yet. The reported uncoupling actions of FFAs are diverse, suggesting that they have two or more modes of action (27–29,36,43–45). They are reported to cause (1) uncoupling based on the protonophoric shuttle mechanism, (2) decoupling, (3) both uncoupling and decoupling, (4) protonophoric uncoupling mediated by mitochondrial solute carriers such as the ADP/ATP carrier, and (5) induction of the permeability transition. As FFAs are important endogenous nutrients and they are substrates of oxidative phosphorylation in mitochondria, a full understanding of their ‘‘uncoupling’’ action is of importance. IV. INTRINSIC UNCOUPLING In mitochondria, substrate oxidation without formation of a ∆µH⫹ should result in energy expenditure as heat. Thus, uncoupling of oxidative phosphorylation in vivo is of importance for body weight control (antiobesity) and thermogenesis. The 30 kDa uncoupling protein acting as the proton channel is responsible for these energy expenditures. Uncoupling mediated by the uncoupling protein (UCP) is the major intrinsic uncoupling mechanism for heat production in animals. A.

Uncoupling Protein

In mammals, excess energy intake is exhausted as heat in brown adipose tissue (BAT). This unique function of BAT is mainly attributable to uncoupling by the

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type 1 uncoupling protein (UCP1) (for reviews, see Refs. 46 and 47). This 30 kDa protein increases the permeability of the inner mitochondrial membrane to H⫹ and thus uncouples oxidative phosphorylation, causing expenditure of excess energy as heat. The function of UCP1 is activated by FFAs and inhibited by guanine nucleotides such as guanosine di- and triphosphate (GDP and GTP). This protein is expressed in BAT especially when surplus heat production is necessary in such periods as (1) adaptation of newborns to outer circumstances, (2) arousal from hibernation, and (3) exposure to cold. In addition, there have been various trials to develop antiobesity drugs by stimulating expression of the UCP1 such as with stimulants of the β3-adrenergic receptor (48). As the amount of BAT, and hence the amount of UCP1, in human adults is low, possible involvements of other tissues in regulation of thermogenesis have been proposed. In 1997, two proteins having high homologies in amino acid sequences (59% and 57%) to that of human UCP1 were identified by a homology search of databases of expressed sequence tags, and they were named UCP2 and UCP3, respectively (49–53) (Fig. 10). UCP2 is expressed in various tissues, but UCP3 is expressed mainly in muscle. As UCP2 dissipates mitochondrial membrane potential in yeast cells, it is thought to act as a proton channel like UCP1. Quite recently, a fourth member of the UCP family of proteins was found to be expressed in the brain and was named BMCP1 (brain mitochondrial carrier pro-

Figure 10 Structural similarities of the human UCP family of proteins. The primary structures of members of the UCP family are cited from the EMBL/GenBank database. hBMCP1 indicates human brain mitochondrial carrier protein-1 (63). Note that the Nterminal extension of hBMCP1 with the amino acid sequence MGIFPGIILIFLRVKFATAAVIVSGHQ is not included in this figure. Amino acid sequences are aligned to show the highest similarity between four proteins by insertions of appropriate gaps (⫺). Three inner molecular repeated structures are also considered in the alignment. The symbols * and # indicate amino acids that are conserved in all four proteins and three UCP proteins (UCP1-3), respectively. Shaded characters indicate amino acids highly conserved in three repeated structures.

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tein-1) (54). Its N-terminal sequence is longer than those of other UCPs, but its structural features are similar to those of other UCPs. Although BMCP1 induces uncoupling in yeast cells, its physiological role in the brain is still unknown. Hence, energy expenditure is possible in other tissues besides BAT. However, it is unlikely that only UCP is responsible for energy expenditure in BAT and other tissues. For consumption of excess energy as heat, a certain molecular machinery could be working in BAT to achieve efficient uncoupling by UCP. To find proteins related to energy expenditure in BAT, a search for proteins specifically expressed in BAT but not in white adipose tissue (WAT) should be effective. Muscle-type carnitine palmitoyltransferase I (CPTI-M) (55) and muscle-type acetyl CoA carboxylase (mACC) (56) were found to be predominantly expressed in BAT. Furthermore, the transcript level of muscle-type fatty acid binding protein (H-FABP) is elevated about 100-fold by cold exposure of rats, but that of adipose-type FABP is independent of cold exposure (57). It is noteworthy that muscle-type protein isoforms and isozymes are preferentially used for fatty acid metabolism in BAT but not in WAT. Therefore, it is quite likely that an energy expenditure mechanism similar to that in BAT is effective in skeletal muscle (58–60). B. Permeability Transition* It has long been known that Ca2⫹ induces uncoupling in mitochondria. However, its uncoupling differs from that by protonophores; i.e., the uncoupling is supported by Pi , Ca2⫹ is transported into mitochondria before induction of uncoupling, the SHE reagents such as N-ethylmaleimide (NEM) cause protection against the uncoupling, arresting the uncoupling, and the uncoupling is accompanied by swelling of mitochondria. The uncoupling induced by Ca2⫹ is sometimes denoted as Ca2⫹ uncoupling to distinguish it from protonophoric uncoupling. Now, it is apparent that Ca2⫹ uncoupling takes place as a consequence of induction of permeability transition (PT) in the inner mitochondrial membrane. The PT pore enables the permeation of solutes with molecular weights of less than 1500 across the inner mitochondrial membrane and its formation is transient, closing again after extrusion of Ca2⫹ . The immunosuppressor cyclosporin A (CsA) is a very potent and effective inhibitor of PT pore formation. By PT pore formation, the components of the matrix such as small proteins and metabolites are effluxed and the configuration of the mitochondrial membrane is greatly altered, as shown in Figure 11. Although extensive studies have been carried out on the nature of PT pores and their induction (61–63), their physiological meanings are not yet fully understood. As mitochondria are thought to be ‘‘pools’’ of cellular Ca2⫹ , PT is likely

* The contents of Section IV.B are related to those of Section III.A.

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Figure 11 Drastic change in mitochondrial configuration induced by permeability transition. (From Ref. 77.) Configurations of rat liver mitochondria were analyzed by transmission electron microscopy. A and B indicate mitochondrial samples without addition of Ca2⫹ and that treated with 100 µM Ca2⫹, respectively. Bars indicate 1.0 µm. Addition of 100 µM Ca2⫹ caused PT with rupture of the mitochondrial inner membrane structure.

to act as a ‘‘reset’’ system to recover inactive mitochondria with accumulated Ca2⫹ (61). Furthermore, the PT pore is postulated to be associated with protein import into mitochondria (64). Recently, mitochondrial PT was found to be closely associated with the early stage of programmed cell death, apoptosis (for reviews, see Refs. 65–68). A wide variety of proteins in various locations such as hexokinase in the cytosol, porin located in the outer mitochondrial membrane, the ADP/ATP carrier located in the inner mitochondrial membrane, creatine kinase in the intermitochondrial

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space, and cyclophilin D in the matrix space are thought to be associated with PT induction (65). Quite recently, pro- and antiapoptotic members of the Bcl-2 family of proteins were found to interact with the ADP/ATP carrier and porin, and this interaction was found to modulate PT pore formation (69,70). As yeast cells are easily modified genetically, the use of yeast mitochondria could be very effective for understanding the molecular basis of the PT pore (71).

V.

CONCLUSION

As described above, various compounds induce uncoupling activity in energytransducing membranes such as mitochondria and chloroplasts. Potent uncouplers are hydrophobic weak acids acting essentially as protonophores, thus dissipating the ∆µH⫹ necessary for ATP synthesis. However, there are other types of uncoupling, such as that induced by fluctuation of membrane integrity or damage to membrane structure. In addition, there seem to be types of uncoupling mediated by certain proteins. Although there have been a number of studies on the mechanism by which uncoupling is induced, uncoupling is still a problem to be solved. The studies are helpful for elucidating the mechanism of ATP synthesis. Intrinsic uncoupling is very important in biochemical reactions under physiological and pathological conditions. Uncoupling mediated by UCP is indispensable for expenditure of surplus bioenergy, and that due to PT induction is likely to be closely associated with cellular activities such as programmed cell death. As these physiological and pathological events are complex, consisting of various biochemical reactions, more detailed studies are necessary for understanding the role of intrinsic uncoupling.

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8 Tetraether Lipid Liposomes Hans-Joachim Freisleben University of Indonesia, Jakarta, Indonesia

I. INTRODUCTION A.

Archaea

Gallia omnia divisa est in partes tres (all gallia are divided into three parts) was written almost 2000 years ago by the Roman Caesar about a part of the world that has since become better known as France. Likewise, in 1978, Woese and his coworkers divided the whole formerly dualistic world of prokaryotic and eukaryotic cells into three parts: eukaryotes, bacteria, and the new kingdom of archaea (singular: archaeon). I do not want to go into phylogenetic criteria of each of the three kingdoms; however, one characteristic issue that separates archaea from all other cells must be referred to in this context: phytanol ether lipids. In archaeal membranes, we find various types of diether and tetraether lipids, differently interdigitated or even covalently omega-linked. The latter case results in tetraether lipids with one or two membrane-spanning biphytanyl chains. Thermoplasma acidophilum is a thermoacidophilic archaeon that was first isolated by Darland et al. (1) from a steaming, self-heated coal refuse pile in Indiana. The temperature of these piles—at least in areas where Thermoplasma grows—is 56 °C, and oxidative degradation of pyrite-containing material generates sulfuric acid and an environmental milieu of pH 1–2. Since this archaeon does not have a cell wall, its cytoplasma membrane must be very resistant to acidic pH. Segerer et al. (2) found further species within the genus Thermoplasma in their natural environments, such as volcanoes, solfataric soils and mudholes, hot springs, etc., in Italy, Iceland, the United States (Yellowstone National Park), and Indonesia (Java: Dieng Plateau, Tangkuban Perahu, and Ciater). They named them Thermoplasma volcanium sp. nov., with three genotypes, T.v.1–T.v.3. All species grow anaerobically and aerobically and show pleomorphism and flag127

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ellation. In the laboratory, growth optimum of Th. acidophilum is at pH 2 and 59 °C in a sulfuric acid–containing Freund’s medium (3). B. Archaeal Lipids The lipids constituting archael membranes differ considerably from bilayer-forming phospholipids in all other cells. These structural differences have been used to characterize archaeal cells and to differentiate them from bacteria or eubacteria and eukaryotes (4). Archaeal lipids are glycerol (or another polyol, e.g., nonitol in Sulfolobus species) ethers in which only the phosphate, if present, is linked via ester bonds. Furthermore, archaeal lipids do not contain double bonds but do contain methyl side groups, and they form pentacycles in the hydrocarbon chains due to growth temperature. Ether bonds have advantages over esters in acidic environments because they are resistant to hydrolysis at low pH. The lack of double bonds increases the resistance of the lipids to oxidation; the methyl side groups exert a fluidizing effect similar to that known for double bonds in common lipids. Archaea use isoprenoid derivatives to synthesize the hydrocarbon moieties of their lipids, e.g., phytanol 3,7,11,15-tetramethylhexadecanol. Besides diether structures, tetraether lipids are found in several archaeal genera. In these latter lipids the phytanyl residues are covalently linked at their terminal (omega) carbons to form a dibiphytanyl tetraether macrocycle with 72 atoms participating in the cycle (5,6). The membrane lipids from Th. acidophilum consist of this tetraether macrocycle. According to Langworthy (7), in both biphytanyl chains symmetrical pentacyclation occurs, with up to two pentacycles per chain and in parallel with increasing growth temperature from 39 to 59 °C. We found a somewhat different pattern of pentacyclation that suggests different numbers of pentacycles in the two hydrocarbon chains (8,9). Due to the degree of pentacyclation and to the extent of distortion of the hydrocarbon chains, the distance between the two glycerol molecules is 3–4 nm, corresponding to the hydrophobic region. Thus, the dimensions of tetraether lipids (TELs) are sufficient to form monolayers of a thickness similar to that of common bilayer membranes. Recent investigations revealed a strong tendency of TELs to form coils or ‘‘horseshoes’’ with both polar headgroups coming closely together and thus a complex phase transition behavior and separation into clusters of different conformations (9,10). The chemical formula of the main tetraether phospholipid (TEL) from Th. acidophilum was given by Strobl et al. (11) as 2,3,2′,3′-tetra-Odibiphytanyldi-sn-glycerol-1′-glycosyl-1-phosphoryl-3′-sn-glycerol (Fig. 1). A space-filling model of TEL was published by Freisleben et al. (3). The molecular weight of TEL was calculated as 1638 Da. The 1′-glycosyl residue was meanwhile defined to be β-l-gulose (8,9,12,13).

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Figure 1 The main tetraether phospholipid (TEL) of Thermoplasma acidophilum.

Thermotropic main phase transition of TEL occurs below 0 °C (14); hence the lipid is sufficiently fluid at room temperature (RT) to form monomolecular films (10,11), black lipid membranes (15), and liposomes (16). A metastable phase transition or phase separation has been observed for TEL39 at 20 °C (17) and for TEL59 at 17 °C (8). TELs are able to form mixed phases with bilayer-forming lipids (e.g., DPPC) as detected by differential thermoanalysis (DTA) measurements (18). Hence, it should also be possible to prepare stable mixed liposomes with egg lecithin and tetraether lipid.

II.

METHODS AND MATERIALS

Thermoplasma acidophilum was grown at pH 2 and 59 °C in a 50 L Braun fermenter (3). The main phospholipid (TEL) was extracted and purified according to Blo¨cher et al. (14), modified by the application of medium pressure liquid chromatography (MPLC) and high performance liquid chromatography (HPLC) methods in order to obtain higher purity (8). Cell membrane stability was measured via the release of protein from the cells. Protein determination was accomplished according to Lowry et al. (19) and Bradford (20); phospholipid was measured according to Chen et al. (21). A.

Preparation of Liposomes

Irrespective of the preparation method, TEL was dissolved in a chloroform–methanol (2: 1 v/v) solution. The solvent was removed by a rotary evaporator (Rotavapor Bu¨chi, Flawil, Switzerland) at 40–50 °C. The dry lipid film was subjected to a high vacuum at room temperature for at least 12 h. For liposome preparation, TEL was suspended in one of the buffers indicated below; if possible, phosphatebuffered saline (PBS) was used. For comparison, liposomes of lecithin from frozen egg yolk (EYL; Sigma, Deisenhofen, Germany) were prepared with 2.5% (w/w) phosphatidic acid

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(Sigma, Deisenhofen, Germany) in order to stabilize the liposomes; furthermore, they become negatively charged, similar to the TEL liposomes. For preparation of mixed liposomes, various amounts of TEL were mixed with egg lecithin. After preparation, all liposomes were centrifuged in an Eppendorf 3200 centrifuge for 10 min to remove nonliposomal material, which may be present, especially in hand-shaken TEL preparations. The liposomes were contained in the supernatant. TEL liposomes were prepared by the following methods: 1. Hand-shaken liposomes. McIlvaine buffer or PBS was added to the lipid film and shaken by hand to disperse the lipid film until a homogeneous liposome suspension was formed (22). The lipid concentration was 20 mg/mL. 2. Sonication. Hand-shaken TEL liposomes were sonicated (Branson Sonifier B 15, Heusenstamm, Germany) at 20 kHz for 5 min (23,27). 3. Detergent solubilization and detergent dialysis. TEL was dissolved in PBS containing the detergent octyl-β-d-thioglucopyranoside (Calbiochem, Frankfurt, Germany). The TEL/detergent (L/D) molar ratio varied from 0.05 to 0.3. This micellar suspension was transferred to a Lipoprep dialysis cell (Diachema AG, Langnau, Switzerland) and dialyzed at RT for 24 h (24). The lipid concentration was 10–20 mg/ mL. Mixed liposomes made of TEL and egg lecithin (Sigma, Deisenhofen, Germany) were prepared in the same way as TEL liposomes. 4. Extrusion through polycarbonate filters. Hand-shaken liposomes from TEL were freeze-thawed three times and sonicated, or directly applied and extruded through 800 nm and finally through 200 nm polycarbon filters by means of a LiposoFast syringe (Avestin Inc., Ottawa, Canada) (25). 5. Extrusion through French pressure cell. Hand-shaken liposomes were transferred to a French pressure cell (max. vol. 3.6 mL; SLMAminco Inc., Urbana, IL). Four cycles at a pressure of 16,000 psi were applied (26,27). The lipid concentration was 10–20 mg/mL. B. Size Distribution and Zeta Potential of the Liposomes The size and size distribution of the liposomes were measured by means of a Malvern laser particle sizer, Autosizer II. The zeta potential was measured with a Zeta Sizer (Malvern Instruments Ltd., Malvern, UK). C. Electron Paramagnetic Resonance (EPR) Measurement Spin labels 3-doxylcholestane, 5-doxylstearic acid, 16-doxylstearic acid, 5-doxyldecane, and di-tert-butyl nitroxide were added to the primary lipid films at 1–5

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131

µM concentrations. The signal was recorded by means of a Bruker B-R 70 Digital EPR spectrometer with a B-E 25 magnet. Order parameters S were calculated according to Zimmer and Freisleben (28).

D.

Stability, Swelling, and Release from the Liposomes

Shelf life and stability of the liposomes were tested by measuring size and size distribution at various temperatures over 2 years. The leakiness was measured by the release of entrapped 6-carboxyfluorescein (CF; Sigma, Deisenhofen, Germany) (29). Swelling was determined by adding alcohols (polyols) and reading the optical density (OD) at 570 nm (16). Resistance against protons and detergents was determined by measuring the size distribution, order parameters, and the release of CF and [125I]insulin (Hoechst, Frankfurt, Germany) under the addition of bile salts and other detergents at varying pH values and temperatures (16,30–32). Proton permeability was measured as described in Ref. 33.

E.

Interactions of Tetraether Lipid Liposomes with Cell Membranes and Cells

Interactions of liposomes with cells and cell membranes were measured mainly by optical means using an Olympus fluorescence microscope with a camera/ photounit or a spectrofluorophotometer using fluorescence dyes such as carboxyfluorescein, Nile Red, and bromobimanes. The latter served as indicators of intermembrane exchange. Incorporation of thiol-reactive bromobimanes into liposomes should give low emission intensity; as soon as intermembrane exchange or release occurs from liposomes to cell membranes, bromobimanes can react with thiols present in the cellular membranes, resulting in a rapid increase in fluorescence (34,35).

F. Cytotoxicity, Mutagenicity, Toxicity, and Distribution in Mice Some toxicological methods can be found in Section III. Detailed methodology is described in Refs. 36 and 37.

G. Penetration into the Skin of Hairless Mice Methods used to determine the penetration of spin-labeled vitamin A acid into the skin of hairless mice (38) are described together with the results in the following section.

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III. RESULTS The size of TEL liposomes depends on the method of preparation. As shown in Table 1, the size decreases in the order hand-shaken ⬎ sonicated ⬎ detergentdialyzed ⬎ filter extruded ⬎ French cell extruded. The size of hand-shaken liposomes is in the range of several micrometers with a very broad size distribution. These liposomes are multilamellar and polymorphous, whereas smaller TEL liposomes tend to become unilamellar and round. Sonicated TEL liposomes are in the range of 450–600 nm, whereas detergent-dialyzed liposomes are 370–450 nm. Besides the method of preparation, the grade of purity of TEL is of considerable importance. Extended studies were carried out with different purification grades of TEL on size and size distribution of the liposomes. With the five methods investigated, the particle size decreased in parallel with higher purity of TEL, i.e., primarily, but not only, removal of chromophores. Hence, efforts were made in our laboratory to achieve a high level of TEL purity using an MPLC/HPLC purification procedure that yields more than 95% purity (8).

A.

French Pressure Cell Extruded vs. Sonicated Liposomes

Tetraether lipid liposomes were prepared by method 5, and the size distribution was determined by laser particle sizer. The average mean was 150 nm (SD ⫾ 22). In comparison, a liposomal diameter of 600 nm was obtained after sonication, and this difference was highly significant, demonstrating that the preparation method determines the size of TEL liposomes. TEL liposomes prepared by detergent solubilization and subsequent detergent dialysis (method 3) are around 400 nm. Extrusion through polycarbonate filters (method 4) yields liposomes similar in size to those obtained with the French pressure cell (100–200 nm) depending on the pore width of the final filter (100 or 200 nm, respectively). Using the various methods, the whole spectrum of TEL liposomes from a minimum size of 80 nm (Fig. 2) to 600 nm can be prepared with good reproduc-

Table 1 Sizes of TEL Liposomes Method of preparation 1. 2. 3. 4. 5.

Hand-shaken Sonication Detergent dialysis Filter extrusion French press

Size, average mean (SD) 7500 600 370 220 150

nm nm nm nm nm

(2500–⬎20,000) (⫾40) (⫾35) (⫾60) (⫾22)

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Figure 2 Laser light scattering of TEL liposomes (Malvern laser particle/Zeta Sizer PCS, Malvern Instruments, UK). Attempts to form the smallest possible TEL liposomes by several extrusion cycles through polycarbonate filters, 100 nm pore width (LiposoFast, Avestin Inc., Canada).

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ibility. Larger TEL liposomes (hand-shaken) vary in size over a wide range and are no longer uniform and reproducible in size, shape, and lamellarity (30). B. Zeta Potential The zeta potential of TEL liposomes was determined in a Malvern Zeta Sizer Version PCS 1.1 at 137.4 V to be ⫺58.2 (SD ⫾ 0.2) at pH 7.4 in phosphatebuffered saline that was diluted with distilled water 1:1 (v/v) for the measurement (Fig. 3).

Figure 3 Zeta potential profile of TEL liposomes (Malvern laser particle/Zeta Sizer PCS, Malvern Instruments, UK).

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C. Order Parameters of Liposomal Membranes as Determined by EPR Measurement The order parameters of the membranes of TEL liposomes as determined by means of various spin labels reporting from different (i.e., outer, polar to the inner, hydrophobic) regions of the membrane are given in Table 2.

D.

Mixed Liposomes from TEL and Egg Lecithin

Tetraether lipid is able to form mixed phases with bilayer-forming lipids as detected by DTA measurements (17). Hence, it should be possible to form mixed liposomes with TEL and egg lecithin. Mixed liposomes were prepared according to method 3 using n-octylglucoside as detergent (Table 3). Homogeneity of the preparations was evaluated from the uniform or nonhomogeneous size distribution (one or more peaks in the laser particle sizer). For egg lecithin liposomes, a molar lipid to detergent (L/D) ratio of 0.2–0.3 was the optimum, whereas with pure TEL a ratio of 0.05 was necessary, i.e., a higher detergent concentration. In the former case, liposomes with a distribution maximum of 153 nm (SD ⫾ 39) at L/D ⫽ 0.2 and 144 nm (SD ⫾ 41) at L/D ⫽ 0.3 were formed; in the latter (TEL/D ⫽ 0.05), the maximum was at 372 nm (SD ⫾ 35). At lower detergent concentrations TEL did not form homogeneous or stable liposomes. The size distribution of mixed liposomes showed homogeneity in the laser particle sizer at TEL/egg lecithin ratios of 75 :25 (L/D ⫽ 0.05), 50 :50 (L/D ⫽ 0.15), and 25 :75 (L/D ⫽ 0.15–0.2) (30).

Table 2 Order Parameters (S) of Liposomal Membranesa Spin label 3-Doxylcholestane 5-Doxylstearic acid 16-Doxylstearic acid 5-Doxyldecane Di-tert-butyl nitroxide a

S (TEL)

S (egg lecithin)

0.9 0.781 0.258 0.187 0.1

0.655 0.644 0.201 0.130 0.1

The reporter groups of the applied spin labels are located progressively from the polar region of the membrane (starting with 3-doxylcholestane) to the hydrophobic core, and/or the spin probes are decreasingly fixed to the polar region such as stearic acids vs. 5-doxyldecane and di-tert-butyl nitroxide.

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136 Table 3 Size (nm) of Mixed Liposomes from TEL and Egg Lecithina L/D molar ratio

TEL : egg lecithin ratio (w/w) 100: 0

75: 25

50: 50

25: 75

0:100

0.05 0.1 0.15 0.2 0.3

372 396–495 n.d.b 421–466 —

558 199–550 169–307 480–792 —

— 359–778 393–664 — —

— 422–645 453–489 325–510 —

— — — 145–162 137–150

L/D ⫽ lipid/detergent molar ratio. Preparations that were not stable in the size distribution determinations within the first 2 days were excluded from further investigations and are not mentioned in the Table (—). b Pure TEL liposomes were not prepared at a 0.15 molar ratio to detergent. a

E.

Long-Term Shelf Stability

Tetraether lipid liposomes were stored in PBS (10 mg lipid/mg) at 37 °C, at RT, and at 4–8 °C. The size distribution was followed in the laser particle sizer after 1, 2, 6, 9, 16, 29, 57, and 109 weeks. Furthermore, the stability was determined at high temperatures (60, 70, 80, 90, 100, and 120 °C) over 10 weeks (Table 4). For comparison, liposomes from egg lecithin were stable for only 1 to a few weeks in the refrigerator (4–8 °C) and for only days at higher temperatures. Mixed liposomes with TEL/egg lecithin ratios of 25 : 75 and 50: 50 (w/w) were stable in the refrigerator for up to 622 days (Table 5). However, samples with increasing amounts of TEL, especially the 75 :25 mixtures, separated during storage into two peaks corresponding to the size of pure egg lecithin and TEL liposomes. In general, mixing TEL with egg lecithin increased the shelf stability of the liposomes, especially at the 25 :75 (w/w) ratio, which correlates to 11–12

Table 4 Long-Term and High-Temperature Shelf Stability of TEL Liposomes At a temperature (°C) of 4–8 RT 37 60 70 size increased from 150 nm to 155 160 165 150 150 after . . . weeks 109 109 109 10 10 a

80

90

100

120

155

155

177

n.d.a

10

10

10

10

After storage of 10 weeks at 120 °C the size of TEL liposomes could no longer be determined.

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Table 5 Shelf Stability of Mixed Liposomes at 4–8 °C Days 25: 75 50: 50

1 387

2 398

6 396

9 407

16 411

20

27

49

408

389

387

79 405

86 446

622 488 498

25 :75 and 50 :50 are the TEL/lecithin ratios (w/w). At the days indicated, the size of the liposomes (nm) was measured in the laser particle sizer. Increase in size by 50% was set as the limit for ‘‘stability.’’

mol% TEL (30). Lipid concentration was determined via phosphate content according to Chen et al. (21). The molecular weight of TEL was calculated as 1638. F. Influence of Protons, Temperature, Alcohols, and Detergents Proton permeability of liposomal TEL membranes was measured by various methods as described in Ref. 33 and was shown to be about two magnitudes lower than in egg yolk lecithin (EYL) liposomes but could be increased by the uncoupler FCCP. In other experiments, the influence of temperature on proton permeability was investigated; even at 74 °C, the rate constant per second was 50% of that in EYL liposomes at RT. In these experiments, the TEL liposomal membrane turned out to be the most proton-impermeable reported so far (33,39). Swelling of liposomes after addition of polyols with three to six carbons (glycerol, erythritol, xylitol, and mannitol), if it occurred at all, was less than 5% with all TEL liposomes tested at RT. The swelling rates of sonicated liposomes from TEL, dipalmitoyl-phosphatidylcholine (DPPC), and mixed TEL/DPPC liposomes were previously shown by Ring et al. (16). TEL liposomes also resisted detergent and alcohol concentrations 8 times higher than those withstood by DPPC liposomes. Comparison of TEL with egg lecithin liposomes is shown in Table 6. G. Influence of High Concentrations of Bile Salts on the Size of Liposomes Mixtures of chenodeoxycholate/cholate/deoxycholate 2: 2: 1, as may occur in the small intestine, were incubated at total bile salt concentrations of 10 and 30 mM (32). Their influence on liposome size is demonstrated in Table 7. H. Sterilization of Liposomes Although there was no doubt that TEL liposomes are heat-stable and can be autoclaved or even heat sterilized (40), it was sufficient in our experiments to

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138 Table 6 Concentration of Alcohol (%) or Detergent (mM) at Half-Maximum (50%) CF Release Alcohol

% with TEL

% with egg lecithin

32 27 13 0.01

25 15 7 0.0025

mM with TEL

mM with egg lecithin

Methanol Ethanol n-Propanol Triton X-100 Detergent n-Octylglucoside Deoxycholate SDS a

12 0.7 n.d.a

8 0.2 0.5

Under the applied conditions and at all concentrations tested, sodium dodecylsulfate (SDS) did not stimulate TEL liposomes to release more than 10% CF. Hence, a half-maximum concentration could not be determined.

use sterile filtration, especially since this can be done in one run with the extrusion method for liposome preparation if polycarbonate filters with 100 or 200 nm pores are used. I. Interaction with Cells and Degradation of Tetraether Lipids Tetraether lipid liposomes containing carboxyfluorescein for detection are much more strongly attracted by cell surfaces than EYL liposomes. These experiments do not yet answer the question of whether TEL liposomes are taken up by living cells (and if so, by what mechanism) or whether they may just be attached to the cell surface without penetration.

Table 7 Influence of Bile Salt Concentrations on Liposome Sizea Bile salt conc. (mM) Liposome TEL Egg lecithin a

0

10

30

133.1 140.6

109.3 38.4

118.3 1302.6

The size (nm) is given as the mean of laser particle sizer measurements.

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The influence of TEL liposomes on electrolyte homeostasis, energy metabolism, DNA, and protein biosynthesis of Ehrlich mouse ascites tumor (EMAT) cells was investigated in detail. Significant alterations or deviations from control cells could not be detected (36). These experiments showed that there is no cytotoxic effect on EMAT cells, but this again could be due to a lack of cellular uptake. On the other hand, transport and release studies with lipophilic fluorescence dyes revealed that TEL liposomes exhibit intermembrane exchange and also fuse with cellular membranes. Successful gene expression in transfection experiments showed that TEL–DNA complexes are taken up into target cells. From other liposome-like lipid–DNA complexes it is known that uptake of these complexes mainly occurs via endocytosis; hence, it can be concluded that TEL liposomes have the same interactions with cell membranes as other liposomes, i.e., intermembrane exchange, fusion phenomena, and endocytosis. Degradation of TEL has not been clarified. Investigations with hepatocytes are still under way in cooperation with the University of Graz, Austria. Uptake in hepatocytes appears to occur; however, degradation products cannot easily be detected because of the minute amounts, and their identification is even more difficult because reference substances of TEL degradation products are not available. J. Cytotoxicity, Toxicity, and Distribution in Mice Mouse lymphoma cells L5178Y, EMAT cells, and permanent hamster fibroblasts V79 were used in our own and other laboratories, and a general Biotest cytotoxicity screening was accomplished by Dr. U. Bethge (Biotest GmbH, Dreieich, Germany). Mutagenicity and antimutagenic efficacy were tested with Salmonella typhimurium TA100 in the Ames plate-incorporation test. None of the testing showed cytotoxic or mutagenic properties of TEL or TEL liposomes, respectively (36). No toxicity was detected in mice. Two models were applied: central nervous system screening in NMRI mice and survival rate in immunosuppressed NMRI mice. The TEL-fed mice lived slightly longer than the controls (37). Organ distribution screening showed that TEL liposomes (100–200 nm), like other negatively charged liposomes of the same size, are rapidly cleared from the circulating blood within 15–30 min and appear mainly in the liver (80%) and in the spleen (8%) (37). K. TEL Liposomes as Vehicles or Drug Carriers Incorporation of compounds into liposomes that should be transported and delivered depends on the physicochemical properties of both the compound and the

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lipid. The easiest and most effective method is to incorporate sufficiently lipophilic compounds into the liposomal membrane. This can be achieved in one step with the preparation of the liposomes, provided that the compound to be incorporated does not disturb the formation and stability of the liposomes and the compound is not destroyed by the method of preparation. If the amount of compound that should be incorporated is not too high, it will be taken up virtually entirely into the liposomal membrane. Good results were achieved with incorporation of lipid-soluble dyes such as Nile Red, bromobimane (Thiolyte MB, Calbiochem), and spin-labeled retinoic acid. Incorporation of methylprednisolone has also been satisfactory. In these cases, delivery, i.e., release of the compounds to target cells, is also a clear mechanism, as was demonstrated by intermembrane exchange from TEL liposomes to erythrocyte ghosts and membranes of intact cells (34,35). Intermembrane exchange does not need any uptake of liposomes into cells, because lipophilic compounds migrate from the liposomal membrane and penetrate into the cell membrane if the liposomes touch the cell surface or are attached to it (Fig. 4). Most pharmaceutically relevant compounds are sufficiently lipophilic to follow this pattern of liposomal incorporation and release.

Figure 4 Intermembrane exchange of thiol-reactive fluorescence dye bromobimane from TEL liposomes to red blood cell ghosts. (⫹) Direct incubation of ghosts with bromobimane; (䊐) intermembrane exchange from TEL liposomes; (■) intermembrane exchange from EYL liposomes.

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The problem is quite different for the incorporation of water-soluble compounds. They are dissolved in the buffer and are enclosed in the liposomes as the latter are formed. Since the inner volume of the liposomal lumen is roughly 1% of the outer volume (10–20 mg lipid/mL), this will also be the amount of compound that will be found in the liposomes; about 99% will not be entrapped. Some amount of the compound may be attached to the outer surface of the liposomes. There are possibilities to increase the entrapped amount by osmosis or temporary pores (e.g., with bile salts), but none of these modifications can alter the result in general: The great majority will remain in the outer buffer. This has to be removed via a gel filtration column and should be regained for economic and other reasons. The liposomal suspension contains the entrapped amount of compound. Release of water-soluble compounds from liposomes may follow different patterns. (1) There is a certain amount of contact release as soon as liposomes touch cell membranes or get attached to them, or (2) the liposomal membrane fuses with the cell membrane with rapid release of the compound into the interior of the cell, or (3), which appears to occur more often, liposomes are actively taken up into the cell by endocytosis and the compound is released into the cytoplasm more slowly. In addition to intermembrane exchange, fusion phenomena and endocytosis have been observed with tetraether lipids (Balakireva and Balakirev, personal communication). The affinity to various cell lines—(1) tumor cell lines, i.e., hepatocarcinoma cells (HepG-2) and colon carcinoma cells (HT29); (2) immortalized cell lines, i.e., cystic fibrosis-pancreas carcinoma cells (CF-Pac), Chinese hamster ovary (CHO) cells, and baby hamster kidney (BHK) cells; and (3) a primary fibroblast (PFB) cell culture—were investigated at different lipid concentrations for 4.5 h. T84 colon carcinoma cells were used for time-dependent comparison between TEL and EYL liposomes. Two effects were detected. First, TELentrapped CF fluorescence was much faster and much more intensively associated with T84 colon carcinoma cells than EYL-entrapped fluorescence. Second, cellassociated fluorescence from TEL liposomes exerted varying concentrationdependent intensities. HT29 cells showed intensive fluorescence at 100 µg/mL, HepG-2 and CHO cells exhibited similar intensity at 250 µg/mL, whereas CFPac cells accumulated much less fluorescence and only at low TEL concentrations, below 200 µg/mL medium. HT29 did not exhibit concentration-dependent kinetics of cell-associated fluorescence above 100 µg/mL. Above a small peak at 100–150 µg/mL, fluorescence of CF-Pac cells was inverse to TEL concentration in the culture medium. HepG-2 cells showed a concentration-dependent increase in fluorescence intensity with some saturation effect above 250 µg/mL, whereas with CHO cells fluorescence increased more strongly above this concentration (Fig. 5). At 200 µg TEL/mL cell culture medium, two groups of different fluorescence intensities were seen, the first group at high intensity with HT29 ⬎

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Figure 5 Carboxyfluorescein fluorescence of various cell lines after incubation with TEL liposomes at different lipid concentrations. Cell cultures and cell lines: CF-Pac, cystic fibrosis-pancreas carcinoma; HT29, colon carcinoma; HEPG, liver cell carcinoma (HepG2); CHO, Chinese hamster ovary cells.

CHO ⫽ HepG-2 cells and the second at about half this intensity with PFB ⬎ CF-Pac ⫽ BHK cells (Fig. 6). L. Release of 6-Carboxyfluorescein (CF) and [125I]Insulin Data on the release of CF from sonicated TEL liposomes are given in Table 8; dependence on pH is shown in Table 9. Moreover, release of [125I]insulin was studied with hand-shaken liposomes of TEL and egg lecithin. Of the applied amount of insulin, 4–6% was associated with liposomes from egg lecithin after passage through a Sephadex G-50-300 column, and in the case of TEL liposomes, 4%. The degree of association to the liposomes depended on the pH of the medium in the case of egg lecithin (pH 3.5, 4%; pH 7, 6%) and was pH-independent

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Figure 6 Carboxyfluorescein fluorescence as in Figure 5, at a fixed liposome concentration of 200 mg TEL/mL. Cell cultures and cell lines: BHK, baby hamster kidney cells; CFPac, cystic fibrosis-pancreas carcinoma; PFB, primary fibroblast cell culture; HT29, colon carcinoma; HepG2, liver cell carcinoma; CHO, Chinese hamster ovary cells.

Table 8 Release of Carboxyfluorescein at pH 7.4 Within 24 ha Temperature (°C) 4 30 50 a

TEL (%)

Egg lecithin (%)

2.5 3.0 8.0

9 41 100

Experiments with release of carboxyfluorescein were successfully accomplished with sonicated liposomes because this method encapsulates sufficient amounts of CF. The data published by Ring et al. (16) for DPPC liposomes are consistant with those from egg lecithin.

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144 Table 9 Release of Carboxyfluorescein at Various pH Values Within 14 Mina pH

TEL (%)

Egg lecithin (%)

6.1 6.6 7.0 7.4

0 2.1 4.2 5.8

31.4 23.3 17.5 19.4

a

Experiments with release of carboxyfluorescein were accomplished as indicated in Table 8 at RT. After 14 min 100% release was achieved by addition of 2% Triton X-100.

in the case of TEL. Of the [125I]insulin associated with the liposomes, 90% was indeed entrapped intraliposomally with egg lecithin (10% was adhesive to the outer surface), and only 72% was entrapped intraliposomally by TEL. Release of [125I]insulin from the liposomes was investigated for 7 days with variation of temperature (RT, 8, 37, 47 °C), pH (3.3, 5.0, 7.4, 8.0–8.5), electrolytes (NaCl, MgCl2 , CaCl2), and osmolarity (PBS/NaPi and McIlvaine buffers, 150–350 mosmol). Release from liposomes of egg lecithin varied with these variations, whereas release from TEL liposomes was low and independent of the variation of external conditions. Figure 7 depicts release of [125I]insulin from TEL liposomes at neutral pH. In the beginning of the experiment, on the first day, there is some release as determined by a decrease in counts per minute in the pellet (liposome-associated) and a concomitant increase of [125I]insulin in the medium, which was supposedly detached from the outer liposomal surface. The greatest differences between release from liposomes of egg lecithin and those of TEL were observed at pH 3.3–5. Whereas the former released almost all the insulin during the first day, 53% of the insulin remained associated with TEL liposomes, and after 7 days, 46% at pH 5 and 35% at pH 3.3. At alkaline pH, the release from TEL liposomes was higher and similar to that from EYL liposomes (30). The results obtained with TEL liposomes were not influenced by variation of the electrolytes and osmolarity in the medium.

M.

In Vitro Intestinal Absorption Model

Two vitreous chambers were separated with excised porcine intestinal mucosa after removal of the serosa. The buffer in each chamber was gassed with carbogen [O 2 /CO 2 (95% :5%)] and pH varied from 6.5 to 8.2 at the intestinal lumen side. From preliminary experiments detecting liposomal test compounds in the oppo-

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145

Figure 7 Release of [125I]insulin from hand-shaken TEL liposomes in various buffer systems (see text) over a period of 7 days: cpm/mL ⫽ counts per minute per milliliter; (䊐). In the supernatant medium after centrifugation; (■) in the resuspended pellet containing the liposomes.

site, blood-side, chamber, it can be concluded that liposomal penetration and absorption through the mucosa may occur (41). N.

Special Applications

For delivery of special freight, such as DNA, special techniques have to be applied, including admixture of positively charged lipids with the negatively charged TEL or chemical modification of TEL itself by introduction of positive headgroups. Similar questions arise for special targeting with introduction of structures that can dock to specific receptors on cell membranes or in the circulating blood. In this respect, TEL can serve well as a membrane-spanning anchor if these structures are covalently or otherwise linked to the TEL molecule. O. Penetration into the Skin Penetration of spin-labeled retinoic acid into the skin of nude mice was investigated with liposomal and nonliposomal pharmaceutical preparations. Two conventional pharmaceutical preparations and three liposomal gels were compared: unguentum emulsificans aquosum (German Pharmacopoea, DABG); a modern

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hydrogel formulation; a soybean lecithin/cholesterol liposomal gel; a liposomelike soybean lecithin/oleoyl hydrolyzed animal protein (OHAP) gel; and a TEL liposomal gel. Retinoic acid spin label (RASL) was incorporated into the liposomes and into the gels to attain a final concentration of 2 mM. The preparations were then applied to the skin of nude mice, and the penetration of RASL was then followed by electron paramagnetic resonance (EPR) imaging using modulated simultaneous scan (MOSS) (42). In general, liposomal systems appear to have a higher penetration rate than conventional lipogels or hydrogels. The lowest penetration was exhibited from unguentum emulsificans aquosum and, among the liposomal preparations, from the soybean lecithin/cholesterol mixture. The highest penetration rate was seen between 10 and 20 min after application from TEL liposomes. At 45 and 60 min after application, differences emerged and penetration from the OHAP gel prevailed, TEL liposomes being between the two other liposomal systems (38).

IV. DISCUSSION The main tetraether lipid (TEL) from Th. acidophilum forms extremely stable liposomes down to a size of 100 nm. These are the smallest closed vesicles reported so far with a pure membrane-spanning lipid, considerably smaller than formerly expected (16). These results are in contrast to those obtained by Lelkes et al. (43) with tetraether lipids from Caldariella acidophila. They found that tetraether lipids form closed vesicles only with at least 25% other phospholipids (they used egg phosphatidylcholine). The size of TEL liposomes depends primarily on the method of preparation but also on the purity of the lipid. In the course of improvement of lipid purification, the size of liposomes was reduced by 25–40% using the same method of preparation. MPLC purification yielded more than 95% purity at sufficient amounts. The high purification grade achieved could be followed optically by removal of chromophores; i.e., highly purified TEL is totally white, whereas former purification had yielded a yellow or brownish lipid. Tetraether lipid mixes with EYL and soybean lecithin and stabilizes lecithin liposomes. Conventionally, phosphatidic acid is used in 2.5% addition to stabilize lecithin liposomes by providing negative charges. TEL adds negative charges to lecithin liposomes (as phosphatidic acid does) and, moreover, covalently connects both polar surfaces of the liposomal membrane bilayer. Thus, TEL is both an electrostatic and structural stabilizer of lecithin bilayers. Structural stabilization is generally provided by cholesterol and also by α-tocopherol; the latter, however, is used primarily as an antioxidant in liposomal membranes (44).

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147

However, TEL does not mix with egg lecithin at arbitrary ratios. Up to 25% TEL/75% lecithin (w/w) homogeneously mixed phases appear to form with egg lecithin, whereas mixtures with 50% or more TEL appear to undergo phase separation into domains or even differentiation into separated egg lecithin and TEL liposomes. DSC measurements with mixtures of TEL and DPPC had shown that ‘‘DPPC appears to be a somewhat better solvent for the tetraether lipid than tetraether lipid for dipalmitoyl phosphatidylcholine’’ (17). Apart from resistance to thermal, hydrolytic, and oxidative attack, tetraether lipid membranes are highly stable against mechanical influences. Furthermore, in freeze-fracture electron microscopy, these membranes do not break in the plane of the membrane (tangentially, as bilayers do) but perpendicular to the plane of the membrane (45). The zeta potential of liposomes depends on the pH and ionic strength of the bulk medium. Hence, the value of ⫺58 mV for TEL liposomes holds for 75 mM NaCl, 5 mM NaPi , pH 7.4. The net surface charge of the liposomes is negative, and the surface is also negatively charged toward the medium. Under these conditions one elementary charge is calculated per 14.15 nm2 , or about 5000 negative charges per liposome. At a calculation of 80,000 TEL molecules per liposome, one out of 15 molecules ought to be oriented with the phosphoglycerol ester to the outer surface, whereas the great majority of the molecules expose their uncharged sugar moiety to the external medium due to the higher demand of hydration. For the calculations, liposomes 150 nm in diameter, corresponding to a surface of 70,685 nm2, were applied. The order parameters of the liposomal membrane exhibit an unexpected phenomenon. Although TEL is membrane-spanning, the fluidity gradient from the polar region to the hydrophobic core of the membrane is steeper than in egg lecithin. The fluidity in the inner region is comparable in both liposomal membranes as demonstrated by spin labels 16-doxylstearic acid, 5-doxyldecane, and di-tert-butyl nitroxide. However, the polar region is much more rigid in TEL liposomes than in egg lecithin, as shown by 3-doxylcholestane and 5-doxylstearic acid. Results of the determination of order parameters in membranes depend very much on the methods used. Whereas fluidity measurements with a glass viscosimeter yielded results similar to those of EPR spectroscopic methods (28), polarization fluorometry and EPR spectroscopy may differ. In this review, only EPR data are presented, which remain to be compared with results obtained from experiments using different methods. The stability of TEL liposomes is higher at low pH than at neutral or alkaline pH, which was proven by release of the hydrophilic dye CF and insulin. This phenomenon does not depend on the method of preparation. It was observed in small unilamellar (French pressure cell and carbonate filter extruded), intermediate (sonicated), and large, multilamellar (hand-shaken) TEL liposomes and can

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also be compared to properties of the original cell membrane, which exhibits maximum stability at pH 3–4. From these data, it was concluded that TEL liposomes might be useful to transport drugs that are labile to acidity through the stomach. Hence, it was interesting to know if bile salts of composition and concentrations that occur in the small intestine would alter the TEL liposomes. Although about 2 mM deoxycholate caused a total release of CF, the size of TEL liposomes was only marginally altered by the mixture of bile salts applied up to 10 and 30 mM concentrations. Liposomes from egg lecithin changed tremendously under the same conditions in a range from 38 nm to 1.3 µm, indicating the formation of very small and very large particles, supposedly mixed (giant) micelles. TEL liposomes do not appear to be micellized by bile salts at high concentrations in the same way as egg lecithin liposomes. These experiments demonstrate that TEL liposomes are stable under all conditions tested in comparison to lecithin liposomes. However, under high concentrations of bile salts at alkaline pH the unique stability of TEL liposomes at acidic pH is diminished and the release of entrapped compounds is enhanced. These properties can well be used to transport acid-sensitive compounds through the stomach to release them at absorption sites in the intestine. Absorption of entire TEL liposomes has still to be tested. Results from interaction of TEL liposomes with cells and cell membranes indicate that lipophilic substances are rapidly transferred from TEL liposomes to cell membranes. Other experiments indicate that fusion and endocytosis also occur. The high fluorescence of TEL-entrapped water-soluble CF at cell membranes could speak for contact release at cell surfaces. However, fluorescence should then be more diffuse, which is not the case. Hence, the cell-associated high fluorescene speaks rather for a high affinity of TEL liposomes to cell surfaces and for direct uptake of CF into the cells. As previously indicated, cells and cell types differ in their kinetics to associate and accumulate fluorescent TEL liposomes. In other words, they obviously have varying affinity to TEL liposomes and/or a varying tendency for their uptake and/or varying tendency for uptake of fluorescence dyes from these liposomes. From the experiments conducted so far with less than 10 cell different types it cannot yet be concluded under what conditions preference to a certain cell type can be achieved for, say, tumor-targeted drug delivery. Since cell culture experiments of this kind are limited in their application to in vivo pharmacokinetics and pharmacodynamics, animal studies have to be conducted with tumor location facilitating evaluation of direct liposome–tumor cell interaction, e.g., colon or urinary bladder carcinomas. We are currently investigating another aspect: immunosuppression after organ transplantation in a monkey model with mixed liposomes stabilized with TEL (46). The immunological properties of these liposomal systems are tested in rabbits, and intestinal absorption of various compounds from TEL liposomes

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is studied in these animal models. These experiments are to be carried out at the University of Indonesia, Jakarta, and the Primate Study Centre of the Agricultural University in Bogor, Indonesia.

ACKNOWLEDGMENTS Most of the work was done in the Gustav-Embden-Zentrum der Biologischen Chemie, Laboratorium fu¨r Mikrobiologische Chemie, Klinikum der Johann Wolfgang Goethe-Universita¨t Frankfurt am Main by the working group headed by Prof. Dr. K. Ring until 1988 and then by me until I went to the University of Indonesia in Jakarta. I thank all the members of the working group. Emmanouil Antonopoulos, Anette BettinBogutzki, Detlef Bloecher, Barbara Deisinger, Gregoria Efstathiou, Marc Engelhardt, Raimund Gutermann, Michael Hartmann, Sabine Hartmann, Birgit Henkel, Lutz Henkel, Sabine Janku, Gabriele John, Frauke Lehr, Eva Lengsfeld, Cornelia Neisser, Ulrich Lo¨benberg, Anton Oertl, Petra Rudolph, Oliver Schivelbusch, Heike Wiesner, Sigrid Winter, and also the other coworkers who were not especially working in the field of TEL liposomes. Prof. Dr. G. Zimmer kindly carried on my work until the end of 1997, when our last grant from the Deutsche Forschungsgemeinschaft ended; their (DFG) support over many years is gratefully acknowledged. Furthermore, KarlFranzens Universitaet, Graz, Austria should be mentioned where I was appointed Guest Professor in 1993/94 and was supported by the Jubilaeumsfond der Oesterreichischen Nationalbank, together with Prof. Dr. W. Mlekusch, Prof. Dr. G. Reibnegger, and Dr. Karl Oettl, all of whom contributed to the results of this review, as well as other cooperating scientists whose names can be found in the citations and references. In Indonesia, Dr. Anja Meryandini and her coworkers will carry on research on Thermoplasma acidophilum and Thermoplasma volcanium from nearby volcanoes in the microbiological laboratory at the Bogor Agricultural University, and liposome technology is presently established by Dr. Ernie H.P. in the Department of Clinical Pharmacy at the University of Indonesia in Jakarta.

REFERENCES 1. Darland G, Brock TD, Samsonoff W, Conti SF. A thermophilic, acidophilic mycoplasm isolated from a coal refuse pile. Science 1970; 170:1416–1418. 2. Segerer A, Langworthy TA, Stetter KO. Thermoplasma acidophilum and Thermoplasma volcanium sp. nov. from solfataria fields. Syst Appl Microbiol 1988; 10: 215–224.

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3. Freisleben H-J, Henkel L, Gutermann R, Rudolph P, John G, Sternberg B, Winter S, Ring K. Fermentor cultivation of Thermoplasma acidophilum for the production of cell mass and of the main phospholipid fraction. Appl Microbiol Biotechnol 1994; 40:745–752. 4. Langworthy TA. Lipids of archaebacteria. In: Woese C, Wolfe R, eds. The Bacteria: A Treatise on Structure and Function, Vol. VIII, Archaebacteria. New York: Academic Press, 1985:459–496. 5. Langworthy TA, Pond JL. Archaebacterial ether lipids and chemotaxonomy. Syst Appl Microbiol 1986; 7:253–257. 6. De Rosa M, Gambacorta A. Lipid biogenesis in archaebacteria. Syst Appl Microbiol 1986; 7:278–285. 7. Langworthy TA. Lipids of thermoplasma. Methods Enzymol 1982; 88:396– 406. 8. Antonopoulos M. Extraktion, Reinigung und chemische Modifizierung von Tetraetherlipiden aus Thermoplasma acidophilum. PhD Dissertation, JWG-University, Frankfurt, Germany, 1999. 9. Ernst M, Freisleben HJ, Antonopoulos E, Henkel L, Mlekusch W, Reibnegger G. Calorimetry of archaeal tetraether lipid: indication of a novel metastable thermotropic phase in the main phospholipid from Thermoplasma acidophilum cultured at 59°C. Chem Phys Lipids 1998; 94:1–12. 10. Bakowski U, Rothe U, Antonopoulos E, Rietz R, Henkel L, Freisleben HJ. Monomolecular organization of the main tetraether lipid from Thermoplasma acidophilum at the water-air interface and transfer of the monofilms to solid surfaces. Chem Phys Lipids, in press. 11. Strobl C, Six L, Heckmann K, Henkel B, Ring K. Physico-chemical characterization of tetraether lipids from Thermoplasma acidophilum. II. Film balance studies on the monomolecular organisation of the main glycophospholipid in monofilms. Z Naturforsch 1985; 40c:219–222. 12. Freisleben HJ, Antonopoulos E, Rothe U, Bakowsky U. Tetraetherlipide und diese enthaltende Liposome sowie deren Verwendung. Patents P 19607722.2, 1996; PCT/ EP 97/01011, 1997. 13. Swain M, Brisson JR, Sprott GD, Cooper FP, Patel CB. Identification of β-L-gulose as the sugar moiety of the main polar lipid Thermoplasma acidophilum. Biochim Biophys Acta 1997; 1345:56–64. 14. Blo¨cher D, Gutermann R, Henkel B, Ring K. Physico-chemical characterization of tetraether lipids from Thermoplasma acidophilum. Differential scanning calorimetry studies on glycolipids and glycophospholipids. Biochim Biophys Acta 1984; 778: 74–80. 15. Stern J, Freisleben HJ, Janku S, Ring K. Black lipid membranes of tetraether lipids from Thermoplasma acidophilum. Biochim Biophys Acta 1992; 1128:227–236. 16. Ring K, Henkel B, Valenteijn A, Gutermann R. Studies on the permeability and stability of liposomes derived from membrane spanning bipolar archaebacterial tetraetherlipid. In: Schmidt KH, ed. Liposomes as Drug Carriers. Stuttgart: G.Thieme Verlag, 1986:100–123. 17. Blo¨cher D, Gutermann, R, Henkel B, Ring K. Physico-chemical characterization of tetraether lipids from Thermoplasma acidophilum. IV. Calorimetric studies on the

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19. 20.

21. 22. 23.

24.

25.

26. 27. 28.

29.

30.

31. 32.

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miscibility of tetraetherlipids with dipalmitoyl phosphatidylcholine and dipalmitoyl phosphatidylglycerol. Z Naturforsch 1985; 40c:606–611. Blo¨cher D, Gutermann R, Henkel B, Ring K. Physico-chemical characterization of tetraether lipids from Thermoplasma acidophilum. V. Evidence for the existence of a metastable state in lipids with uncyclizated hydrocarbon chains. Biochim Biophys Acta 1990; 1024:54–60. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ. Protein measurement with the Folin phenol reagent. J Biol Chem 1951; 193:265–275. Bradford MM. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 1976; 72:248–254. Chen PS Jr, Foribara TY, Warner H. Microdetermination of phosphorus. Anal Chem 1956; 28:1756. Bangham AD, Standish MM, Watkins JC. Diffusion of univalent ions across the lamellae of swollen phospholipids. J Mol Biol 1965; 13:238–252. Finer EG, Flock AG, Hauser H. Mechanism of sonication of aqueous egg yolk lecithin dispersions and nature of the resultant particles. Biochim Biophys Acta 1973; 298:1015–1019. Weder HG, Zumbu¨hl O. The preparation of variably sized homogeneous liposomes for laboratory, clinical, and industrial use by controlled detergent dialysis. In: Gregoriadis G, ed. Liposome Technology, Vol. I. Boca Raton, FL: CRC Press, 1984: 79–108. MacDonald RC, MacDonald RI, Menco BPM, Takeshita K, Subbarao NK, Hu L. Small-volume extrusion apparatus for preparation of large, unilamellar vesicles. Biochim Biophys Acta 1991; 1061:297–303. Barenholtz Y, Amselem S, Lichtenberg D. A new method for preparation of phospholipid vesicles (liposomes)—French press. FEBS Lett 1979; 99:210–214. New RRC. Liposomes: A Practical Approach. Oxford: IRL Press, 1990. Zimmer G, Freisleben HJ. Membrane fluidity determinations from viscosimetry. In: Aloia RC, Curtain CC, Gordon LM, eds. Advances in Membrane Fluidity: Methods for Studying Membrane Fluidity, Vol. 1. New York: Alan R. Liss, 1988:297–318. Weinstein JN, Ralston E, Leserman LD, Klausner RD, Dragsten P, Henkart P, Blumenthal R. Self-quenching of carboxyfluorescein fluorescence: uses in studying liposome stability and liposome-cell interaction. In: Gregoriadis G, ed. Liposome Technology, Vol. III. Boca Raton, FL: CRC Press, 1984:183–204. Rudolph P, Wiesner H, Engelhardt M, Freisleben HJ. Liposomes of the main phospholipid (MPL) from the archaebacterium Thermoplasma acidophilum: size and stability [abstract]. Biol Chem Hoppe-Seyler 1993; 374:145. O’Connor CJ, Wallace RG, Iwamoto K, Taguchi T, Sunamoto J. Bile salt damage of egg phosphatidylcholine liposomes. Biochim Biophys Acta 1985; 817:95–102. Schivelbusch O. Der Einfluss von Gallensaeuren auf Groesse und Membranfluiditaet von Eilecithin-Liposomen und Liposomen aus dem Hauptphospholipid von Thermoplasma acidophilum. Eine ESR-spektroskopische Studie. MD Dissertation, JWG University, Frankfurt, Germany, 1996. Freisleben HJ, Zwicker K, Jezek P, John G, Bettin-Bogutzki A, Ring K, Nawroth T. Reconstitution of bacteriorhodopsin and ATP synthase from Micrococcus luteus

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9 Transfection of Eukaryotic Cells with Bipolar Cationic Derivatives of Tetraether Lipid Larissa A. Balakireva and Maxim Yu. Balakirev Russian Academy of Science, Novosibirsk, Russia

I. INTRODUCTION Gene therapy and genetic engineering require reliable and efficient systems for the delivery of exogenous genes into target cells. Among nonviral vectors, cationic lipids are most widely used as a delivery vehicle for nucleic acids (1,2). In aqueous solution, cationic lipids form lamellar aggregates that, on simple mixing with diluted plasmid DNA, condense the nucleic acid into smaller multimolecular particles coated with cationic lipid bilayer. The transfection efficiency of such DNA–lipid complexes depends significantly on their ultrastructure, which in turn depends on the condensing conditions and the chemical structure of the cationic lipid (3). A number of different cationic lipids have been synthesized, and their efficiency for DNA transfer has been estimated both in vitro and in vivo (1,2). Despite some differences, all these compounds have a common structure of eukaryotic lipids in which a charged polar headgroup is connected to two hydrophobic alkyl chains. This similarity might be a significant constraint in construction of the new DNA–lipid complexes. In this chapter we describe the synthesis of cationic lipids based on the lipids of Archaebacteria, which possess quite different bola-amphiphilic structures. These lipids consist of branched phytanyl chains ether-linked to a substituted glycerol instead of unbranched ester-linked chains of eubacteria and eukaryotic lipids. Furthermore, the lipids of extreme thermoacidophilic archaebacteria appear as bipolar molecules in which two substituted glycerol residues are ether153

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Figure 1 Typical structure of archaeal lipids.

linked via two C40 ω–ω′ biphytanyl chains into a closed cycle (Fig. 1). This cyclic structure, known as a tetraether (TE), has been used in the design of new cationic lipids. Three different derivatives, AF1, AF2, and AF⫹, have been synthesized that differ in the structure of their polar headgroups (diethylamino, amino, and triethylamino, respectively) at both ends of the TE cycle. All these lipids form liposomes in aqueous solution that are able to complex plasmid DNA. The observed 1:1 (lipid/phosphate) stoichiometry of DNA–AF complexes suggests that only one of the two AF cationic groups is involved in the interaction with nucleic acid. All derivatives were found to mediate the transient transfection of cultured cells via an endocytosis-dependent pathway. The efficiency of AF-mediated transfection was found to be comparable to that of lipofectamine and could be further augmented by their mixing with dioleylphosphatidylethanolamine (DOPE).

II.

EXPERIMENTAL

A.

Synthesis of Archefects

Synthesis of the different tetraether lipids has been carried out according to Scheme 1 using the crude lipid extract of Thermoplasma acidophilum cells as the starting material. The procedure included acid hydrolysis of the total lipid components to produce tetraether (TE), subsequent oxidation of the tetraether to the diacid compound (DA), production of diacid dichloroanhydride (DA-Cl), and its coupling with different amines, resulting in the formation of cationic bipolar lipids called archefects (AF1, AF2, AF⫹). The structures of all synthesized compounds (except DA-Cl) has been proved by infrared spectroscopy, mass spectroscopy, and elemental analysis (C, H, N ⬍ ⫾0.4). 1.

Tetraether

A 750 mg portion of the crude lipid fraction of Thermoplasma acidophilum cells (see chapter by Freisleben in this book) was dissolved in 800 mL of 1 M HCl

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Scheme 1 Chemical synthesis of bipolar cationic lipids. TE, tetraether; DA, diacid; DACl, diacid dichloroanhydride.

solution in methanol and refluxed for 10 h. The solution was evaporated, the residue was redissolved in 100 mL of CHCl3 and washed with water (5 ⫻ 100 mL). The chloroform fraction was evaporated and chromatographed on silica gel (eluant: CHCl3). Fractions were collected and analyzed on thin-layer chromatography (TLC) plates (eluant CHCl3 /methanol 9:1); fractions that contained tetraether were evaporated, giving 390 mg of TE.

2. Diacid Compound Tetraether (80 mg) was dissolved under stirring in 2 mL CH2Cl2 solution of 4methoxy-TEMPO radical (1 mg), and tetraoctylammonium bromide (1 mg) at 0 °C. Sodium hypochlorite (0.35 M, pH 8.6, 5 mL) was added to this solution, and the mixture was stirred vigorously at 0 °C. Five minutes later, chloroform (30 mL) was added and the organic layer was washed with 0.25 M HCl (5 ⫻ 30 mL). The organic fraction was evaporated and chromatographed on silica gel (eluant: CHCl3 /methanol/acetic acid 100: 5:0.1). Fractions were collected and analyzed on TLC plates (using the same eluant); fractions that contained diacid compound (DA) were evaporated, giving 47 mg of DA (yield ⬃ 50%).

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3.

Diacid Dichloroanhydride

Diacid compound (45 mg) was dissolved in the solution of thionyl chloride (100 µL) in dry CH2Cl2 (5 mL) and refluxed. After 6 h the solution was evaporated, and crude diacid dichloroanhydride (DA-Cl) was used for archefect synthesis without further purification. 4.

Archefects AF1, AF2

3-Dimethylaminopropylamine (50 µL) was added to the solution of diacid dichloroanhydride (DA-Cl) (⬃45 mg) in dry CH2Cl2 (5 mL). After 5 min the solution was washed with water (5 ⫻ 30 mL), evaporated, and chromatographed on silica gel (eluant: CHCl3 /methanol/acetic acid 80: 20: 0.5). Collected fractions were analyzed on TLC plates (the same eluant), and the fractions containing compound AF1 were evaporated, giving 35 mg of archefect (yield ⬃ 75%). AF2 was synthesized by the same manner except that 100 µL of 1,3-diaminopropane was used instead of 3-dimethylaminopropylamine. 5.

Archefect ⫹ (AF⫹)

Archefect 1 (10 mg) was dissolved in the solution of dimethyl sulfate (10 µL) in dry CH2Cl2 (5 mL). After 20 h the solution was evaporated, and the residue was redissolved in 10 mL of CHCl3 and washed with 0.1 M HCl (3 ⫻ 20 mL). The organic solution was evaporated, giving 10 mg of AF⫹ (yield ⬃ 95%). 6.

Rhodamine Archefect (Rh-AF)

Rhodamine isothiocyanate (2 mg) was added to the solution of AF2 (2 mg) and triethylamine (5 µL) in dry CH2Cl2 (2 mL). After 15 h the solution was washed with water (5 ⫻ 30 mL), evaporated, and chromatographed on silica gel (eluant: CHCl3 /methanol/acetic acid 80:20:0.5). Fractions were collected and analyzed on TLC plates; fractions that contained fluorescent lipid were evaporated, giving 2 mg of Rh-AF.

B. Preparation of Liposomes To prepare lipid film, a solution of archefect (500 µL of 2 mg/mL) in chloroform/ methanol (1 :1) was slowly evaporated under rotation and dried in vacuum (24 h, ⬍0.05 mm). Lipid film was hydrated in 1 mL of 150 mM NaCl, 50 mM HEPES at pH 7.4 for 48 h at room temperature, sonicated in a Branson 1210 bath (15 min, RT), and finally sonicated with sonifier Branson B15 (10 min, RT).

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C. Preparation of DNA–Archefect Complex and Transfection of Cultured Cells Transfection was performed with pSV-lacZ plasmid DNA (Promega). Cells were plated on six-well dishes (about 105 cells/well) and were used for transfection 24 h later (⬃50% of confluent). The following protocol was used: 1. 3–5 µL of the lipid suspension was diluted in 100 µL of FCS-free DME medium. 2. 1–2 µg DNA was diluted in 100 µL of FCS-free DME medium. 3. Lipid and DNA solutions were gently mixed, then incubated for 15 min at room temperature to obtain a DNA–lipid complex. Usually, the formation of complexes was not accompanied by the appearance of aggregates. 4. At the end of incubation the DME medium (800 µL) was added to give 1 mL final volume. 5. Cells were washed with FCS-free DME medium, then 1 mL of the DNA–lipid complex solution was added. 6. Five hours later, DNA-containing medium was replaced by DME medium with 10% FCS. D.

Cell Assay for β-Galactosidase Expression In Situ

Twenty hours after transfection, cells were washed with PBS, fixed with ice-cold methanol, and colored overnight in the PBS solution of 4 mM ferrocyanide, 4 mM ferricyanide, 1 mM MgCl2 , and 0.1% X-gal, pH 7.4. The efficiency of transfection was estimated as the percentage of blue-colored cells. E.

Fluorescent Microscopy

To prepare Rh-AF-labeled liposomes, 1 mg of AF/Rh-AF mixture (100:1 molar ratio) was sonicated in 1 mL of 150 mM NaCl, 50 mM HEPES at pH 7.4. Cells were incubated with Rh-AF liposomes or DNA–liposome complex at lipid concentrations ranging from 10 to 100 µg/mL for 1 h at 37 °C, washed three times with PBS, and studied under a reverse fluorescence microscope.

III. RESULTS A.

Archefect Synthesis and Preparation of the Liposomes

The synthesis of tetraether cationic lipids is shown in Scheme 1. The advantage of the tetraether structure is its high stability to acid hydrolysis and oxidation. Since membranes of Th. acidophilum are mainly composed of tetraether lipids

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(4), one can produce more than 50 wt% of pure TE by simple HCl hydrolysis of total bacterial lipids. Attempts to modify TE using the various alkylating and acylating reagents were unsuccessful, presumably due to the low reactivity of hindered OH groups. We therefore tried to convert TE alcohol groups into carboxylic acids. Several oxidation conditions [different organic peracids (5) and catalysis with ruthenium salts (6,7)] were tested. The best results were achieved using nitroxyl-catalyzed heterophase oxidation with hypochlorite (8). The yield of DA compound obtained by this method can be as high as 50%, depending significantly of the purity of TE. The reaction of DA with thionyl chloride resulted in DA dichloroanhydride formation, which was coupled to different amines to produce cationic tetraether lipids with various headgroups (called archefects) (Scheme 1). Any of the synthesized cationic lipids could be used to prepare an aqueous dispersion. Sonication of the hydrated archefect film in NaCl–HEPES buffer at pH 7.4 resulted in the formation of liposomes of different sizes, which were shown by electron microscopy (Fig. 2A). Sometimes, along with the vesicles, the presence of a cubic lipid phase characteristic of bipolar archaeal lipids was observed (Fig. 2B). The archefect liposomes prepared by sonication were used for the formation of a complex with plasmid DNA (pDNA) and transfection. They were found to be remarkably stable, showing no lipid degradation or aggregation during 1 month of storage at room temperature. (A)

(B)

Figure 2 Electron microscopy of negatively stained archefect samples. (A) Monolamellar AF⫹ liposomes (5 mg/mL of lipid) prepared by 25 min sonication of the lipid film. (B) Formation of the cubic phase in AF⫹ dispersion (5 mg/mL) after 10 min of sonication. Bar represents 200 nm.

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B. Archefect–DNA Complex Formation Two methods have been used to study the interaction of archefects with pDNA. First, the formation of AF-pDNA complexes was shown fluorometrically using a DNA-specific fluorescent probe, ethidium bromide (EB). It is known that DNA binding to cationic lipids prevents EB intercalation into DNA, thus decreasing its fluorescence (9). The decrease in EB fluorescence can therefore be used to determine the amount of lipid-bound DNA. Complexes of archefects with pSVlacZ plasmid DNA were preformed at different AF/DNA ratios in 1 mL of 150 mM NaCl, 50 mM HEPES, pH 7.4 buffer following by EB addition. For all archefects a typical sigmoidal dependence of EB fluorescence on lipid concentration was observed (Fig. 3A). An increase in pDNA concentration resulted in a shift of the curve to the higher values of AF concentration. The data in Figure 3A show that the formation of the complex occurs at an archefect/pDNA phosphate molar ratio of 1:1. Taking into account two polar groups on each archefect molecule, it seems likely that only one of two AF cationic groups is involved in the interaction with nucleic acid.

Figure 3 Formation of lipid–DNA complexes followed by ethidium bromide–DNA fluorescence and DNA gel electrophoresis. DNA–lipid complexes were preformed as described in Section II. (A) Fluorescence of 0.1 µM EB (λext 518 nm, λem 605 nm) after addition to pDNA in the presence of different concentrations of cationic lipid (AF1), (■) 2.1 µM DNA; (䉱) 4.2 µM DNA; (䊉) 6.3 µM DNA. (B) pDNA retardation on 1% agarose gel induced by AF1 at different phosphate/lipid molar ratios.

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Formation of the AF–pDNA complex was further proved by DNA retardation assay using 1% agarose gel (Fig. 3B). Increasing the concentration of archefects resulted in AF–pDNA complex formation and pDNA retardation due to the masking of the negative charge. Here again, the optimal condition for the complex formation was a 1: 1 AF/pDNA phosphate molar ratio. At higher concentration of archefects we observed an apparent decrease in the visible amount of DNA on the gel. This could be explained by the inhibition of EB staining of pDNA in the complex with cationic lipid. C. Penetration of Archefect and AF–DNA Complexes Inside the Cells To study whether synthesized positively charged lipids and their complexes with pDNA are capable of penetrating cells, we synthesized the rhodamine-labeled archefect analog Rh-AF. Archefect liposomes labeled with Rh-AF were added to a monolayer of baby hamster kidney (BHK) cells, and intracellular distribution of the fluorescent probe was studied by fluorescence microscopy (Fig. 4). After 1 h of incubation, strong diffusive fluorescent staining of the cells was observed with all types of archefect liposomes (AF1, AF2, and AF⫹), suggesting the ability of AF liposomes to fuse with cellular membranes (Figs. 4A,4B). By contrast, when the fluorescent AF liposomes were mixed with pDNA before addition to the cells, comparatively little transfer of the labeled lipid to the cells was observed (Figs. 4C,4D). The data in Figure 4 suggest, in agreement with some previous proposals (10,11), that the lipid mixing and fusion between the liposomes and cell membranes depend on the net positive charge of the liposomes and is significantly inhibited by the formation of a liposome–DNA complex. D.

Archefect-Mediated Transfection of the Cells in Culture

In the next series of experiments, different archefects were studied for their ability to mediate the transfection of BHK cells in culture. The pSV-lacZ plasmid bearing the β-galactosidase reporter gene was used to enable measurement of transfection efficiency by determination of the number of transfected blue cells (Fig. 5A). As shown in Figure 5, all the synthesized archefects were able to mediate the efficient transfection of BHK cells. At the optimal conditions, up to 12% of the cells can be transfected with AF–pDNA complexes. AF-mediated transfection was also observed with another plasmid, pGL3-CMV, containing the luciferase reporter gene as well as with other cell lines (HeLa, CV-1). Transfection efficiency with archefects showed a sharp maximum at the pDNA/lipid molar ratio of ⬃1.5 (Fig. 5B). Since this value is higher than that obtained for optimal AF– pDNA complex formation (Fig. 2), it seems likely that a certain excess of positive charge is required for efficient interaction of the complex with a negatively

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Figure 4 Penetration of rhodamine-labeled archefect liposomes and archefect–pDNA complexes into BHK cells. (A) Phase-contrast and (B) fluorescence images of BHK cells treated with Rh-labeled liposomes (20 µg of AF1, 1 mol% of Rh-AF) for 90 min. (C,D) The same conditions as in (A,B) but the liposomes were mixed with pDNA (1: 1 AF1/ phosphate ratio) before the incubation with BHK cells.

charged cellular surface (3). Like other known cationic lipids (13), the transfection with archefect–pDNA complexes was transient: Maximum expression of βgalactosidase was observed 20 h after transfection, followed by a gradual decrease in the number of transfected cells (Fig. 6A). The presence of fetal calf serum during the transfection procedure did not decrease the efficiency of AFmediated transfection, whereas it did significantly inhibit transfection with the control positive lipid lipofectamin (data not shown). This may be due to the rigid

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Figure 5 Transfection efficiency of different archefects. Standard transfection procedure described in Section II was used. (A) X-gal staining of BHK cells transfected with 2 µg of pSV-lacZ plasmid DNA in the complex with 5 µg of AF1. (B) Transfection efficiency of lipid–pDNA complexes formed with 2 µg of DNA and indicated concentrations of lipids. (䊉) AF1; (䉱) AF2; (■) AF⫹.

structure of tetraether-formed membranes that hinder protein interaction with the cell surface and prevent protein incorporation into the lipid bilayer (12). The demonstrated ability of archefect lipids to mediate DNA entry into cells can not be explained by the fusion mechanism alone, since lipid mixing between the archefect–pDNA complex and cellular membranes was found to be inhibited (Figs. 4C,4D). To obtain more information about the mechanism of AFmediated transfection, we studied the effect of endocytosis inhibitors on transfection efficiency (Fig. 6B). Although it was shown that the transfection efficiency of cationic lipids can be augmented by lysosomotrophic agents (chloroquine, monensin, NH4Cl) (14,15), only a decrease in the number of AF–pDNAtransfected cells was observed with these agents. In fact, the presence of 100 µM chloroquine during the incubation of the AF–pDNA complex with the cells resulted in more than threefold reduction in transfection efficiency (Fig. 6B). This finding suggests that, as in the case of pH-sensitive liposomes (16), the endosomal/lysosomal pathway plays an important role in the entry of AF-pDNA complex into the cells. E.

Effect of Phospholipids on the Efficiency of Cell Transfection with AF–pDNA Complex

It is generally accepted that the efficiency of lipofection depends on the phase state of the lipid component in the complex with DNA (1–3). The presence of

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lipids capable of existing in a hexagonal phase can significantly enhance the transfection competence of the DNA–lipid complexes (17,18). While the tetraether lipids display a rich polymorphism similar to that of eukaryotic lipids, they easily adopt an unusual cubic phase instead of the lamellar phase (19). We have seen a cubic phase impurity in some liposome samples of archefects and MPL (major phospholipid of Thermoplasma acidophilum; see Freisleben, this volume) that appears as a bicontinuous lipid lattice in electron micrographs (Fig. 2B). Prolonged sonication (5–15 min) was necessary to prepare the homogeneous samples of unilamellar vesicles (Fig. 2A). The exact folding of the archefects in the complex with DNA was difficult to determine. To study the effect of the lipid phase state on transfection competence of the AF-pDNA complex, we mixed archefects with different phospholipids of certain phase preferences: dioleoyl phosphatidylethanolamine (DOPE, hexagonal phase), egg yolk phosphatidylcholine (PC, lamellar phase), and MPL (lipid polymorphism of Archaea). In agreement with previous studies (17,18), only DOPE could significantly augment the transfection efficiency of the AF-pDNA complex (Fig. 7)

Figure 6 Characteristics of AF-mediated transfection. (A) The time course of β-galactosidase expression in BHK cells transfected with AF–pDNA complex. (B) Effect of the inhibitor of endosomal acidification, chloroquine (100 µM), on the efficiency of AF-mediated transfection.

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Figure 7 Effect of the phospholipids on transfection efficiency of archefect. (A) BHK cells were transfected with pDNA (2 µg) and liposomes, prepared from the mixture of AF with different phospholipids at the indicated molar ratios (5 µg). (B) Transfection of BHK cells with 2 µg DNA and different concentrations of the mixed liposomes (1 : 1 AF1/ phospholipid molar ratio). (䊊) AF1; (䊉) AF1 ⫹ DOPE; (■) AF1 ⫹ PC.

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suggesting the crucial role of hexagonal phase formation in the lipofection mechanism. The inhibition of transfection by negatively charged MPL arises mainly from neutralization of the archefect charge rather than from the phase effect. By contrast, the inhibition of AF-mediated transfection by PC was quite unexpected. Although the reason for such an inhibitory effect is unclear, it seems likely that PC can induce changes in archefect membrane folding, thus weakening the lipid– DNA complex or inhibiting its entry into the cells.

IV. CONCLUSION In this chapter we have described archefects, positively charged bipolar lipid structure. The synthesis protocol has been elaborated, allowing the preparation of different cationic tetraether derivatives from the total lipids of Thermoplasma acidophilum. All synthesized archefects were found to be able to penetrate the cells, to complex with pDNA, and to mediate the efficient transfection of eukaryotic cells. Our study suggests that tetraether cationic lipids can be useful for gene therapy and for structural studies of DNA–lipid interaction.

ACKNOWLEDGMENTS We are indebted to our colleagues E. Antonopoulos and L. Henkel for assistance in lipid synthesis, to Dr. R. Gropp for help in plasmid DNA preparation, and to Dr. E. Kiseleva for an introduction to electron microscopy. We are grateful to Dr. F. Gropp, Dr. H.-J. Freisleben, and Dr. K. Hartmann for fruitful discussions. We wish to thank especially Prof. Dr. G. Zimmer for his constant encouragement and help. The assistance of A. Hochberger in manuscript preparation is acknowledged. We thank the Deutsche Forschungsgemeinschaft for the support of this study, Zi 80/26-2.

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10 Cholera Toxin Conformational Changes Associated with Changes in Membrane Structure Jameson A. McCann St. Louis University, St. Louis, Missouri

William D. Picking University of Kansas, Lawrence, Kansas

I. INTRODUCTION A.

Overview of Cholera Toxin

The enterotoxin of Vibrio cholerae (CT) is responsible for the major manifestations of cholera, including severe abdominal cramping and the discharge of voluminous amounts of ‘‘rice-water’’ diarrhea (1). Left untreated, the loss of fluids and electrolytes in cholera results in severe dehydration and death within hours of onset. Cholera is contracted from drinking water contaminated with human fecal waste, which provides a vehicle for rapid dissemination of this diarrheal disease (2). Although V. cholerae infection can be treated with antibiotics and its symptoms countered by rehydration therapy and electrolyte replenishment, cholera continues to be a serious problem in developing nations having poor sanitation systems and inadequate medical facilities (3). One of the problems encountered when combating cholera is the absence of an effective vaccine to provide long-term protection against the illness (2). The difficulty in developing a cholera vaccine is the need for a long-lived mucosal immunity that neutralizes the pathogen and its toxin (4). Generation of a specific, long-term mucosal (secretory) immune response has also proven to be an obstacle in combating other enteric pathogens such as Shigellae and Salmonellae. Ironically, while production of an effective vaccine with long-lived protection against cholera remains elusive, CT and its pentameric binding subunit (CTB) have been 167

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found to be potent mucosal adjuvants that promote a secretory IgA response to many coadministered antigens that would otherwise not elicit such responses (5–7). CTB’s behavior as a mucosal antigen and adjuvant is probably related to its ability to interact with cellular membranes at mucosal surfaces (8,9). The observation that CT is a better adjuvant than CTB (8,10), however, may stem from endosomal processing of the holotoxin following its uptake by target cells. Internalization and processing within endosomal compartments is required for delivery of the A1 polypeptide of CT (CTA1) to the target cell cytoplasm, where it carries out its biochemical function. In any case, it is clear that the interaction between CTB and the host cell membrane has an integral role in both the cytotoxicity and adjuvancy of CT. Moreover, the observation that recombinant CTB (completely devoid of CTA) exhibits adjuvancy (11) implies that the CTB–membrane interaction contributes substantially to the immunomodulatory behavior of CT. Exploration of the membrane-influencing properties of CTB is essential for a better understanding of the full nature of cholera intoxication and the action of the best mucosal adjuvant reported to date. B. Structural Features of Cholera Toxin A pentamer of identical protein monomers (58 kDa total), CTB is responsible for CT binding to the carbohydrate moiety of ganglioside GM1 receptors on target cell surfaces (1). Although CTB binds up to five equivalents of GM1, composite assemblies consisting of various combinations of active and inactive B monomers have been used to demonstrate that pentavalent binding is not a prerequisite for cellular intoxication (12). Binding of CTB to the cell surface is not sufficient for eliciting CT toxicity, because it is the A subunit (CTA) that possesses the ADP-ribosylase activity responsible for cellular intoxication (1). During its synthesis by V. cholerae, CTA (27 kDa) is proteolytically cleaved to generate two polypeptides, CTA2 (5 kDa) and CTA1 (22 kDa), which remain linked via a single disulfide bridge (1). CTA2 is held noncovalently within the hydrophilic central pore of CTB, where it serves as a tether for maintaining CTA1 as part of CT holotoxin (1). In the absence of CTB, CTA1 is nontoxic to cells because it is unable to associate with or enter target cells (1). Therefore, CTB is essential for the toxin’s recognition of target cells and thus for endosomal processing of CT. At the same time, the interaction of CTB with host cell membranes is an important aspect of CT’s behavior as a mucosal adjuvant. C. Processing of CT and the Mechanism of CT Action After gaining access to the target cell cytoplasm, CTA1 binds NAD⫹ and catalyzes the transfer of ADP-ribose to Gsα, the GTP-binding subunit of a heterotrimeric G protein complex (reviewed in Ref. 1). ADP-ribosylation of Gsα by

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CTA1 causes it to favor the binding of GTP over GDP, thereby leading to constitutive activation of adenylate cyclase with concomitant overproduction of cAMP in the target cell. The consequence of this series of events is an efflux of ions from the intestinal epithelium. The resulting osmotic imbalance in the intestine is compensated for by the massive export of water into the intestinal lumen. Cholera toxin binds to the surface of target cells with CTA1 oriented away from the cytoplasmic membrane (13). From this position, the holotoxin is taken up by the host cell and processed within endosomal compartments prior to development of overt cellular sequelae (14–16). Endosomal processing appears to be a prerequisite for (1) CTA1 entry into the host cell cytoplasm and (2) movement of the toxin from its initial point of entry (the apical face of enterocytes) to the site at which its intracellular targets are located (the basal side of these cells) (14–17). Endosomal processing is sensitive to compounds such as brefeldin A that interfere with events occurring in those intracellular compartments related to Golgi function (16). To explore the role of CTB in cellular intoxication by CT (with implications pertaining to this toxin’s behavior as a mucosal adjuvant), it is important to understand the interactions that occur between GM1-containing membranes and CTB throughout the intoxication process.

II.

LOW pH ELICITS STRUCTURAL CHANGES IN THE CTB-GM1 COMPLEX

A.

Using Fluorescence Spectroscopy to Monitor Protein Conformation

Based on the prolonged presence of CT within endosomal compartments (17), one could argue that low pH is an important environmental factor that must be considered when exploring the intracellular processing of the toxin. In our laboratory, fluorescence resonance energy transfer (FRET) has been a particularly useful tool for monitoring conformational changes in CTB as a function of environmental pH (18). FRET is the passage of excitation energy from a donor fluorophore (d ) to an acceptor molecule (a) whose absorption overlaps the emission of d (19). FRET efficiency (E ) is dependent upon (1) the spectral overlap of d emission and a absorption, (2) the relative orientation of d and a dipoles, and (3) the distance separating d and a. Therefore, FRET is a useful molecular ruler for calculating the distance from d to a when the spectral overlap and dipole orientation are known. The spectral overlap integral (J ) is calculated from spectral measurements, whereas the dipole orientation factor (κ2) is typically taken as 2/3, which assumes a dynamically averaged orientation over the fluorescence lifetime of d (19). Although the latter can introduce uncertainty into the resulting distance determination, dos Remedios and Moens (20) convincingly argue that assuming a random orientation factor is appropriate for most biological applications of FRET. Moreover, an inverse sixth power relationship between the dis-

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tance separating the probes and FRET efficiency makes this method extremely sensitive to structural changes that influence the distance between d and a within a macromolecular complex (19). The theory and application of FRET analysis for analyzing protein structure are well documented (21). FRET is measured from E ⫽ 1 ⫺ (Fda /Fd ), where Fda is the fluorescence intensity of d in the presence of a and Fd is the fluorescence intensity of d in the absence of an acceptor probe. E can be used to calculate the distance separating d and a when each occupies a specific position within a macromolecular complex according to E ⫽ R 60 /(R 60 ⫹ r 6), where R0 is the theoretical distance that would give 50% E and r is the actual distance separating the probes. R0 takes into account J, κ2, and other spectral factors (such as the quantum yield of d ), which makes it unique for each set of probes and conditions used (19). Numerous reactive fluorescent probes are available to label proteins for subsequent FRET analyses. In our laboratory, fluorescein, coumarin, and stilbene probes have all been used as amine-reactive extrinsic probes for examining CTB structure (18). The naturally occurring tryptophan residue present at position 88 within each CTB monomer (W88) has also been exploited as a probe for fluorescence analysis in our laboratory (18,22). Additional probes used in these analyses are fluorescent lipids incorporated into artificial phospholipid vesicles consisting of phosphatidylcholine (PC). Figure 1 provides a general description of the positions assumed by these different fluorophores in the complex resulting from CTB association with membrane-embedded GM1. B. FRET-Based Detection of pH-Dependent Changes in CTB Structure From its published crystal structure, W88 is located within the GM1 binding site of CTB (23). Using FRET-based analyses, the distance from W88 on CTB to a pyrene acceptor covalently attached to the nonpolar tail of GM1 (pyreneGM1) is about 3.2 nm, which is approximately the distance expected to separate these two groups (18) (see Fig. 1). As a component of the GM1 binding site, W88 should maintain a fixed position relative to the ganglioside in CTB–GM1 complexes as conditions are encountered that would otherwise alter CTB structure. This, in fact, is the case for pH as indicated by FRET-based distance determinations where pH is used as the environmental variable (Fig. 2A). The fact that low pH does not influence the distance from W88 to the probe on pyreneGM1 also shows that low pH does not disrupt the CTB–GM1 association. This has proven to be an important control for additional experiments designed to explore the influence of low pH on the structure of the CTB–GM1 complex. To better determine if low pH affects the structure of the CTB–GM1 complex, amine-reactive fluorescent probes were attached to CTB at available lysine residues so that energy transfer between pyreneGM1 and these sites could be

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Figure 1 The relative locations of fluorescent probes present on a single monomer of CTB and within phosphatidylcholine (PC) vesicles into which derivatives of ganglioside GM1 and/or phosphatidylethanolamine have been incorporated. On the CTB monomer, A denotes the position of tryptophan 88 (W88) located within the GM1 binding site and B marks the position of lysine 69 (K69) on the central helix of CTB, where the majority of extrinsic label is covalently attached. Another population of extrinsic fluorescent label is covalently bound at K91 located near the edge of the GM1-binding site on CTB. Within the membrane, C marks the position of the pyrene probe linked to the acyl tail of GM1 and D denotes the general position of the NBD probes bound to the polar headgroup of phosphatidylethanolamine. (From Ref. 18.)

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Figure 2 (A) Energy transfer is used to calculate the distance (shown in nm) from W88 (0.15 µM CTB) to the tail of a pyreneGM1 acceptor (5 µM incorporated into 0.1 mg/mL PC vesicles) as a function of pH. W88 was excited at 282 nm with fluorescence intensity measured at its peak of emission (334 nm). Fd was measured using 5 µM nonfluorescent GM1 in place of pyreneGM1. (B) The distance from the pyrene linked to the acyl tail of GM1 to (䉱) coumarin, (䊊) stilbene, or (䊉) fluorescein probes covalently attached to CTB increases as the pH is lowered. PyreneGM1 was present at 0.15 µM, and CTB (labeled and unlabeled) was used at 0.3 µM. PyreneGM1 was excited at 330 nm with emission measured at 398 nm for each acceptor probe. (From Ref. 18.)

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measured. CTB was labeled with isothiocyanate derivatives of coumarin, stilbene, or fluorescein probes. The coumarin and stilbene probes labeled to a stoichiometry of slightly less than one probe per CTB monomer, while fluorescein labeled at just over one probe per CTB monomer (18). The difference in labeling stoichiometry was probably a function of the relative solubility of each probe in aqueous solution. Mass spectrometry indicated that fluorescein labeling yields products that are more than 80% monolabeled with minor amounts of unlabeled and dilabeled species (18). Subsequent peptide mapping of the fluorescein-labeled protein used here suggested that the most readily available site for fluorescein labeling was lysine 69 (K69), with significant labeling also occurring at lysine 91 (K91). According to the published crystal structure of CTB, the label on K69 should be about 1.7 nm from W88 on a given CTB monomer, while the label on K91 is near the edge of the GM1 binding site (about 1.0 nm from W88) (23). Labeling at K69 is perhaps fortuitous because this residue is located on the central αhelix residing within the hydrophilic central pore of the CTB pentamer. Coumarin labeling tends to be more heterogeneous than fluorescein labeling, and the stilbene-labeling site has not been precisely determined (18). All three forms of extrinsically labeled CTB can be used for FRET experiments with pyreneGM1 serving as the donor species. The distance separating extrinsic probes on CTB and the pyrene attached to GM1 can be calculated as a function of pH based on the spectral properties of each probe and the subsequent FRET measurements for individual d–a pairs (Fig. 2B). In each case, there is a well-defined increase (up to 1.0 nm) in the distance between the extrinsic probes on CTB and the pyrene on GM1 when the pH is lowered to below 5.0. Differences observed in the calculated distances using CTB labeled with fluorescein versus that labeled with stilbene or coumarin may represent slight differences in labeling position, labeling stoichiometry, and the effect of pH on the spectral overlap between pyreneGM1 and the individual probes attached to CTB. In any case, CTB labeled with each probe provides evidence that pH has a major effect on the structure of the pyreneGM1–CTB complex. These distance changes are not due to release of CTB from the membrane surface as indicated by the data presented in Figure 2A. Moreover, because there is no change in the fluorescence properties of pyreneGM1 bound to CTB (data not shown), it is unlikely that the observed distance changes are due to extraction of GM1 from the membrane. C. Low pH Alters the Distance from Sites on CTB to the Membrane Surface Because the distance from probes on CTB to the tail of pyreneGM1 (a point located within the membrane interior) increases as a function of low pH, it is

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important to find out whether there is a general movement of CTB relative to the membrane. For this, FRET from coumarin and stilbene probes linked to CTB to an NBD probe linked to the polar headgroup of phosphatidylethanolamine (giving NBD-PE) can be monitored. First, NBD-PE is incorporated into PC vesicles at a density that gives a FRET efficiency (E ) near 50% at neutral pH. This allows easy detection of distance changes as the pH is changed. Unfortunately, because positioning of NBD-PE in the fluid membrane is random, it is impossible to use this FRET determination to measure the precise distances from sites on CTB to the membrane surface. Instead, E can be used to determine the general position of the probes on CTB relative to an average density of probes on the surface of the membrane. In these experiments FRET increases rapidly as the pH is lowered to 5.5 or less (Fig. 3), suggesting that the probes on CTB are approaching the membrane rather than moving away as appears to occur when pyreneGM1 is the membrane-associated probe used in the FRET experiments.

Figure 3 The effect of low pH on FRET efficiency between probes on CTB and NBDPE incorporated into the outer face of PC vesicles. NBD-PE (4 µg/mL) was incorporated into 0.1 mg/mL PC vesicles along with 5 µM unlabeled GM1. FRET was measured from the probe on CTB to NBD as a function of pH. The open circles show the influence of pH on FRET efficiency from coumarin to NBD, and the filled circles show the same experiment using stilbene as the donor probe. Coumarin-labeled CTB was excited at 385 nm with emission measured at 465 nm. Stilbene-labeled CTB was excited at 336 nm with emission measured at 440 nm. FRET was also measured from W88 (0.33 µM CTB) to NBD-PE (4 µg/mL incorporated along with 5 µM nonfluorescent GM1 into 0.1 mg/mL PC vesicles) as a function of pH (䉱). W88 was excited at 282 nm with emission measured at 334 nm. In all cases, Fd was measured using PC vesicles that did not have NBD-PE incorporated into their outer face. (From Ref. 18.)

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To extend these experiments, the effect of pH on the FRET efficiency between W88 and NBD-PE can be monitored. In this case, W88 appears to move toward the membrane surface in small increments as a function of lowering the pH (Fig. 3). This graded movement, in contrast to the abrupt change in distance seen with the extrinsic probes linked to CTB, is interpreted to indicate that low pH does not induce movement of W88 relative to the membrane surface. Alternatively, low pH may induce conformational changes in CTB that result in some portions of the protein moving toward the membrane surface while others move away. W88 may lie at a position that does not allow it to move relative to the tail of pyreneGM1 but does allow it to show minor movement with respect to the membrane surface. In contrast, the extrinsic probes on CTB could move more dramatically relative to the membrane because of their relative positions on the protein. Taken together, the data suggest that low pH induces structural changes in CTB–GM1 complexes that may, in turn, influence the packing of the phospholipid membrane to which CTB is associated.

III. THE EFFECT OF CTB BINDING ON MEMBRANE STRUCTURE The fluorescence properties of pyreneGM1 are greatly influenced by the microenvironment of the pyrene probe (24,25). At low concentrations in aqueous solutions, pyreneGM1 exists with its nonpolar pyrene moiety sequestered at the interior of micelles, where the close proximity of these groups results in excimer formation (25). PyreneGM1 excimers are characterized by self-quenching of fluorescence at the major fluorescence peaks of emission (380, 398, and 415 nm) accompanied by increased fluorescence at 480 nm (25). PyreneGM1 excimers are eliminated when the pyrene moieties are dispersed as would occur in organic solvent or when it is incorporated into PC vesicles (24,25). Similarly, a reduction in the excimer properties of pyreneGM1 micelles is observed when they are incubated with an excess of CTB, suggesting that CTB binding to GM1 influences the ability of this lipid to pack with other lipids (26). To follow up this observation, pyreneGM1 was incorporated into PC vesicles and changes in the quenching of pyrene by the small polar quenching agent acrylamide were monitored in the presence and absence of CTB. Steady-state fluorescence can be used to monitor the relative accessibility of a fluorescent probe to a particular quenching agent according to the method of Stern and Volmer (27). The Stern–Volmer quenching constant (KQ) for pyreneGM1 with acrylamide is calculated by measuring the fluorescence spectrum of the sample upon adding increasing amounts of acrylamide. Fluorescence quenching is plotted as F0 /F versus the concentration of quenching agent according to F0 /F ⫽ 1 ⫹ KQ[Q], where F0 is the fluorescence intensity in the absence of quenching agent, F is the fluorescence intensity at a given concentration

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of quenching agent, and [Q] is the concentration of quenching agent (27). The slope of this plot gives KQ . Incorporation of pyreneGM1 into the outer face of PC vesicles results in a dramatic decrease in the quenching of pyrene fluorescence by acrylamide. Micellar pyreneGM1 has a KQ of 2.1 M⫺1 , which decreases to 0.47 M⫺1 after incubation with a 360-fold molar excess of PC (as preformed lipid vesicles). Reduced quenching by acrylamide is caused by the integration of pyrene into the hydrophobic interior of the membrane where it is free to diffuse but is no longer accessible to polar quenching agents (24,25). Therefore, protection of pyreneGM1 from quenching by acrylamide accompanies the loss of pyreneGM1 excimer formation. Interestingly, the addition of CTB to the pyreneGM1-containing vesicles causes KQ to once again increase (Table 1), indicating that CTB binding to the modified GM1 elicits changes in its packing within the membrane. This altered packing provides better access for the small polar quenching agent acrylamide to the pyrene attached to the acyl tail of GM1, which is still located within the hydrophobic core of the phospholipid vesicles (Table 1).

Table 1 Effect of CTB Binding and pH on the Acrylamide Quenching of the Fluorescence from PyreneGM1 Incorporated into PC Vesiclesa Stern–Volmer quenching constant (M⫺1) pH

Without CTB

With CTB

7.0 6.0 5.0 4.0 3.0

0.47 0.49 0.53 0.50 0.57

1.09 1.13 1.24 0.76 0.56

PyreneGM1 (0.33 µM) was incubated for 1 h with preformed PC vesicles (0.12 mM). These composite vesicles were then incubated with 1.5 µM CTB at neutral pH for 30 min, and the pH of the samples was subsequently adjusted to the given pH. Acrylamide quenching of pyreneGM1 fluorescence was determined using an excitation wavelength of 330 nm with emission measured at 398 nm according to the method of Stern and Volmer (27). Source: Ref. 22.

a

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It has been reported that CTB binding to phospholipid vesicles allows the escape of small polar molecules trapped within the vesicles (28,29) and that environmental pH may profoundly influence this event (30). As shown in Table 1, as the pH is lowered, the ability of CTB to cause increased acrylamide quenching of pyreneGM1 is abolished (Table 1). There is no such pH effect on pyreneGM1 quenching in the absence of CTB. This is not an artifact of pH-dependent release of CTB from GM1, as it has already been shown that these conditions do not disrupt the CTB–GM1 interaction (Fig. 2A). Moreover, these data are not a result of extraction of pyreneGM1 from the PC vesicles because (1) low pH causes increased protection of the pyrene from acrylamide quenching and (2) there is no change in the fluorescence intensity of the pyrene as would be expected upon its movement from a nonpolar to a polar environment (data not shown). It should be noted that the pH-dependent change in pyrene quenching occurs in the same pH range needed to elicit the changes in CTB described above. To further explore the influence that pH-induced changes in CTB structure have on GM1-containing membranes, FRET was measured from the hydrophobic tail of pyreneGM1 to NBD probes (as NBD-PE) located on the surface of PC vesicles (18). In the absence of CTB, pH has no influence on FRET from the pyrene to the NBD (Fig. 4). In contrast, low pH causes an increase in FRET from pyreneGM1 to NBD-PE when CTB is bound to the pyreneGM1 prior to

Figure 4 Energy transfer efficiency from membrane-embedded pyreneGM1 (0.16 µM) to 4 µg/mL NBD-PE located on the surface of PC vesicles as a function of pH. Each lipid was incorporated into preformed 0.1 mg/mL PC vesicles, and FRET efficiency was measured in the (䊉) absence or (䊊) presence of CTB. In each case, pyreneGM1 was excited at 330 nm, with emission measured at 398 nm. Fd was measured using PC vesicles that did not have NBD-PE incorporated into their outer face. (From Ref. 18.)

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changing the pH (Fig. 4). These data imply that pH does not normally influence the position of pyreneGM1 incorporated into phospholipid membranes but that pH-dependent changes in CTB structure may be relayed to the membrane-bound ganglioside receptor, whose position within the membrane then becomes pHresponsive.

IV. DISCUSSION The major steps of cholera intoxication have been described, but the mechanism by which CTA1 enters the target cell cytoplasm and the nature of the CTB– membrane interaction remain poorly understood. Evidence that endosomal processing of CT is essential for cellular intoxication suggests that the response of this protein complex to low pH could provide an important contribution to the current understanding of the mechanism of CT intoxication. Moreover, the implication that low pH influences CT processing extends beyond simply providing information on the mechanism of CT intoxication, as both CT and CTB are now known to be potent mucosal adjuvants (6–9,11). The ability of CT and CTB to serve as mucosal adjuvants has been proposed to be an outcome of the interaction these protein complexes have with the membranes of cells at mucosal sites (8,9). The observation that CT may be a better adjuvant than CTB could also implicate CT’s toxigenic potential and perhaps its potential as a transmucosal carrier in its ability to modulate the host immune response to coadministered antigens at mucosal surfaces (10). The latter possibility has prompted study of the use of reconstituted CT holotoxin-like analogs for the delivery of novel organic compounds into mucosal epithelial cells or cultured cells (31). Furthermore, holotoxin-like protein complexes have been reconstituted in vitro that consist of the CTB pentamer and the A2 polypeptide (32). Such protein complexes could, by virtue of the chemically reactive sulfhydryl present on CTA2, be used to target any of a variety of compounds for delivery into host cells or for use as novel vaccines or as a pharmacological delivery system. The attractiveness of the latter compounds is that they need not be limited to the use of protein antigens, because CTB– CTA2 complexes have been reconstituted in vitro that possess a coumarin fluorescent probe attached to the free sulfhydryl of CTA2 (32). An important aspect of the adjuvant action of CT is probably the interaction that occurs between CTB and GM1-containing cellular membranes. We have shown that low pH, such as that which may be encountered as CT or a CT analog is processed within the endosomal compartments of target cells, induces structural changes in the CTB–GM1 complex. In turn these structural changes appear to influence the packing of the membrane phospholipids surrounding the GM1 receptor. By understanding the nature of the interaction between CTB and membranes in low pH environments, it may be possible to further dissect the mecha-

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nism of cholera intoxication while revealing those factors governing the properties of mucosal adjuvants.

ACKNOWLEDGMENTS This work was supported by the Beaumont Faculty Development Fund and a Faculty Summer Research Award to WDP from Saint Louis University.

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11 Parasite Mitochondrial Membrane Functions as Targets for Chemotherapy Akhil B. Vaidya, Michael T. McIntosh, and Indresh K. Srivastava MCP Hahnemann University, Philadelphia, Pennsylvania

I. INTRODUCTION It is a goal of antimicrobial drug discovery efforts to identify pathways that are unique, conserved, and essential in microbial pathogens, with the hope of developing agents with selective toxicity. For viruses and eukaryotic parasites, this goal becomes quite challenging, because the physiological processes used by these pathogens are often very similar to those of their hosts. In this chapter, we suggest that a significant number of essential physiological processes carried out by mitochondria of eukaryotic pathogens are likely to be sufficiently divergent from the mitochondria of their hosts and that these could be fruitfully explored in drug discovery endeavors. Eukaryotic pathogens, comprising a vast group of organisms including protozoa, fungi, and invertebrate animals, take an enormous toll on the health and economy of the world’s population. In general, we have fairly limited means available to counter these pathogens, often with drugs that have low therapeutic indices and high toxicity. While oxidative phosphorylation to generate ATP is the most obvious role for mitochondria, it is clear that these organelles play a critical role in many other metabolic processes such as calcium homeostasis, amino acid metabolism, isoprenoid synthesis, lipid metabolism, and heme synthesis. Indeed, there are cell type- and species-specific functions associated with mitochondria. Furthermore, the central role played by mitochondria in life and death decisions by the metazoan cells has become increasingly clear, resulting in a resurgence of interest in 183

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these organelles (1–3). The current view of mitochondrial evolution tends to favor a monophyletic origin of these organelles, that is, the view is that all extant eukaryotes share a common mitochondrial ancestor and that mitochondrial evolution is as ancient as eukaryotic cells themselves (4). This view would also suggest that vast evolutionary distances separate mitochondria from vastly divergent taxa. This level of divergence becomes apparent in light of unusual characteristics associated with mitochondria from some of the protists such as kinetoplastid (RNA editing, mitochondrial DNA organization) and apicomplexan parasites (scrambled ribosomal RNA organization, extreme reduction of mitochondrial genome). In addition to these relatively major differences, the evolutionary divergence also results in many subtle but significant changes, even in highly conserved mitochondrial functions such as the electron transfer chain. In this chapter, we describe one such example that underlies selective toxicity of a new antiparasite drug.

II.

MITOCHONDRIA OF APICOMPLEXAN PARASITES

The phylum Apicomplexa comprises thousands of known parasite species with sophisticated sexual stages (5). Among these are pathogens of great medical and veterinary importance such as Plasmodium, Cryptosporidium, Babesia, Theileria, Toxoplasma, and Eimeria species. Malaria alone is responsible for an estimated 500 million cases and 2.7 million fatalities each year (6). Mitochondria in malaria parasites were believed to be important mainly to serve as an electron sink for the enzyme dihydroorotate dehydrogenase (DHOD; see Ref. 7). This enzyme is critical in pyrimidine biosynthesis, and since the parasites cannot salvage pyrimidines and rely entirely on de novo synthesis (8), mitochondrial function was deemed essential. This view was derived from the observations that the parasite mitochondria lacked citric acid cycle enzymes (9) and had saclike morphology in the erythrocytic stages with few discernible cristae (10). In the 1980s, malaria parasites were found to contain a very unusual mitochondrial genome consisting of tandem arrays of DNA molecules with a unit length of 6 kb (11–17). Subsequently, similarly small mtDNA have been reported in other apicomplexan parasites such as Theileria (18) and Babesia (14,19) species, although the tandem arrangement appears to be limited to the Plasmodium species at this point. As can be seen in Figure 1, these smallest known mitochondrial genomes encode only three proteins: cytochrome b and subunits I and III of cytochrome c oxidase. In addition, the mtDNAs encode fragmented and scrambled ribosomal RNA molecules that appear to associate in trans to form very unusual mitochondrial ribosomes (13,15,17). It is clear that a vast majority of proteins required for mitochondrial functions are synthesized in the cytoplasm and imported into mitochondria (20).

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Figure 1 Schematic representation of the mitochondrial genomes of (a) Plasmodium and (b) Theileria. Both genomes are linear and appear to lack tRNA genes. Plasmodium elements are tandemly arrayed with a unit length of 6 kb (13,15), while Theileria genomes remain as 7.1 kb molecules with terminal inverted repeats (TIRs) (18). Protein coding regions encode portions of the electron transport system that include cytochrome c oxidase subunits I and III (COX I, COX III) and apocytochrome b (CYT B). Additional genes encode fragmented portions of mitochondrial rRNA that have lost their conventional order of transcription. Portions of large subunit (LS) and small subunit (SS) gene fragments are numbered in ascending order as they would appear 5′–3′ in conventional rRNA. The arrows indicate the direction of transcription for the upper and lower strands.

Transport of these as well as of a number of metabolites across the mitochondrial membranes is dependent upon the maintenance of an electrochemical gradient around the mitochondrial inner membrane. Proton-pumping electron transfer chain complexes are responsible for generating this gradient. In malaria parasites, only two of these complexes are present; ubiquinol–cytochrome c oxidoreductase (the bc1 complex) and cytochrome c oxidase. The first complex, NADH dehydrogenase, is absent from malaria parasites (21,22). This could be explained by the fact that these parasites do not contain a citric acid cycle (9) and thus do not produce much NADH inside the mitochondria. Instead, as seen in Figure 2, electrons are most likely provided by the three dehydrogenases—DHOD, succinate dehydrogenase, and glycerol 3-phosphate dehydrogenase—all of which generate ubiquinol (QH2). The bc1 complex oxidizes QH2, transferring the electrons to

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Figure 2 Representation of the electron transfer protein complexes within the inner mitochondrial membrane of Plasmodium species. Coenzyme Q (Q) is reduced to ubiquinol (QH2) by one of three dehydrogenases oxidizing succinate, glycerol-3-phosphate, or dihydroorotate. NADH dehydrogenase (complex I) appears to be absent from Plasmodium species. Ubiquinol is oxidized through the Q cycle (63) by the cytochrome bc1 complex (complex III), which in turn reduces cytochrome c (Cyt c) present in the intermembrane space. Atovaquone inhibits ubiquinol oxidation (bold arrow). Cytochrome c is oxidized and molecular oxygen reduced to H2O by cytochrome c oxidase complex (complex IV). Electron transfer reactions at the bc1 complex and cytochrome c oxidase are coupled to proton translocation from the matrix side of the membrane to the intermembrane space (vertical arrows). This results in the generation of an electrochemical gradient across the inner mitochondrial membrane. Electron transfer is likely to be coupled to the synthesis of ATP by the ATP synthase (complex V) via H⫹ translocation from the intermembrane space to the matrix; however, the components of complex V and mitochondrial ATP synthase activity remain undemonstrated in Plasmodium.

cytochrome c and protons across the inner membrane. It is this step in the electron transfer chain that is affected by the new antiparasitic drug atovaquone. III. ATOVAQUONE AS A BROAD-SPECTRUM ANTIPARASITIC DRUG In the 1940s Fieser and colleagues carried out extensive investigations on antiparasitic properties of a large number of hydroxynaphthoquinones (23). One of these

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compounds, lapinone, cured P. vivax-infected patients (24). Because this compound required parenteral administration in high doses, and because orally available potent antimalarial agents such as chloroquine became available, the interest in developing hydroxynaphthoquinones as antimalarials waned in the 1950s (see Refs. 25 and 26 for reviews). However, in the late 1970s, Wellcome Research Laboratories successfully developed an anti-Theileria hydroxynaphthoquinone called parvaquone (27), leading to a renewal of interest in designing this class of drugs with enhanced metabolic stability and broad-spectrum antiparasite activity. These efforts led to the synthesis of 2-[trans-4-(4′-chlorophenyl)cyclohexyl]-3hydroxy-1,4-naphthoquinone (initially called 566C80, later named atovaquone). Atovaquone demonstrated potent activity against P. falciparum in culture as well as in Aotus monkeys (28). Additional studies showed atovaquone to be effective against Toxoplasma (29) and Pneumocystis (30) as well. Clinical evaluation in uncomplicated P. falciparum malaria showed an excellent response but an unusually high rate of treatment failure (⬃30%) due to recrudescence (31,32). However, atovaquone’s effectiveness against Pneumocystis pneumonia and toxoplasmosis in AIDS patients kept this drug alive. The drug has been approved for treating Pneumocystis and toxoplasma infections. Canfield et al. (33) sought a way to use atovaquone in malaria by assessing a partner drug that could act in synergy with it, resulting in identification of an established antimalarial, proguanil, as one such compound. The atovaquone– proguanil combination has given encouraging results with an essentially 100% cure rate with no indication of recrudescence (32,24,35). This combination has also proved to be effective as a chemosuppressive antimalarial agent, and its use as a prophylactic drug for travelers can be anticipated.

IV. MECHANISM OF ATOVAQUONE EFFECTIVENESS A.

Antimitochondrial Activity

As early as the 1940s, studies on hydroxynaphthoquinones suggested that these compounds inhibited mitochondrial respiration (36–38). All of these early investigations, however, were done in heterologous mitochondria from yeast or mammalian tissues. Thus, these results, while suggestive of mitochondria as targets for hydroxynaphthoquinones, could not explain the apparent selective toxicity and therapeutic value of these compounds. Studies on parasite mitochondria were not carried out until recently. Successful isolation of malaria parasite mitochondria for biochemical studies was reported in 1991, although these mitochondria did not seem to be well coupled (21). Using the isolated organelles from P. falciparum and P. yoelii, Fry and Pudney (39) showed that atovaquone inhibited the bc1 complex with an EC50 of ⬃10⫺9 M, while the mammalian bc1 complex was inhibited with an EC50 of ⬃5 ⫻ 10⫺7 M. These results for the first time suggested

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the selective toxicity of an antimalarial hydroxynaphthoquinone through preferential inhibition of the parasite mitochondrial electron transfer chain. B. Mitochondrial Membrane Potential (∆Ψm) Collapse Because of the difficulties associated with isolating biochemically active mitochondria from malarial parasites, alternative approaches to investigating mitochondrial physiology in these organisms are needed. To this end, a flow cytometric assay to measure ∆Ψm in intact, live malaria parasites was developed (40). This assay uses a lipophilic cationic fluorescent dye, 3,3′-dihexyloxacarbocyanine iodide (DiOC6), as a probe in conjunction with flow cytometry to assess ∆Ψm . The use of this probe was carefully validated: At 2 nM concentration, DiOC6 rapidly partitions into energized mitochondria within the live parasites; at the concentration used, the probe did not perturb the parasite mitochondrial physiology and remained partitioned in the mitochondria for an extended period; the partitioning was dependent upon the maintenance of ∆Ψm , because inclusion of carbonyl cyanide m-chlorophenyl hydrazine (CCCP), a protonophore, abolished the probe accumulation (40). Thus, this assay permitted analysis of the effect of various compounds on the mitochondrial physiology of live parasites. The usual mitochondrial electron transfer chain inhibitors, myxothiazol, antimycin, and cyanide, collapsed the parasite ∆Ψm in a dose-dependent manner. Rotenone, a complex I inhibitor, did not affect ∆Ψm as expected due to the absence of this complex from the parasite mitochondria. Atovaquone collapsed ∆Ψm in a dose-dependent manner with an EC50 of ⬃15 nM. Measurement of respiration by the parasites under the influence of these compounds correlated with the effects on ∆Ψm . Other antimalarial drugs, such as chloroquine and tetracycline, did not affect parasite mitochondrial physiology under these conditions. Further, atovaquone did not have any significant effect on ∆Ψm of mammalian cells, again confirming its selective activity against parasite mitochondria (40). These results could be interpreted to suggest that ∆Ψm collapse is a consequence of electron transport inhibition by atovaquone. Under the conditions used in these experiments, mitochondria from most organisms will be able to maintain ∆Ψm even in the face of electron transfer chain inhibition by reversing the F0F1 ATP synthase, which will hydrolyze ATP and pump protons across the inner membrane. A lack of robust F0F1 ATP synthase in the parasite mitochondria would not permit such compensatory reestablishment of ∆Ψm . However, this interpretation is not settled yet in light of recent observations suggesting that atovaquone may act as a site-specific uncoupler of the ∆Ψm in malaria parasites (see Sec. V). C. Parasite Death In multicellular organisms, collapsed ∆Ψm has profound effects on cellular physiology (1–3). The collapsed ∆Ψm could be a cause or a consequence of mitochon-

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drial permeability transition in which a large pore forms within the mitochondrial inner membrane, resulting in the release of solutes up to 1500 Da in size from the mitochondrial matrix (41). In mammalian cells this also leads to the release of proteins, most notably cytochrome c present within the intermembrane space (42). A large body of evidence accumulated over the last few years shows that mitochondrial permeability transition is a central step in the activation of the programmed cell death pathway (1–3). Cytochrome c released in the cytoplasm has been shown to participate in activation of caspases that lead to apoptosis (42,43). While analogous apoptosis pathways have not been detected in unicellular organisms, programmed cell death has been hinted at in several organisms (44–47). It is conceivable that a programmed cell death pathway would be evolutionarily selected for in unicellular organisms inasmuch as it may provide a measure of protection for the neighboring members of the species when a cell dies. It clearly would be of interest to investigate such pathways, especially in parasitic pathogens. As for the malaria parasites with atovaquone-mediated ∆Ψm collapse, the demise may occur through the disruption of several processes. Lack of pyrimidine synthesis due to inhibition of DHOD is one such process. In addition, all metabolite and protein transport across the inner membrane will cease in parasites with collapsed ∆Ψm , the net result being a major disruption of the parasite metabolism. Death could thus ensue due to such disruption; the form of death, however, remains unclear. Do parasites lyse, being unable to maintain homeostasis, in a manner similar to necrotic cell death? Or do they undergo an ordered dismantling of their macromolecular organization in a manner reminiscent of apoptosis? Although we do not have much information about these processes, morphological observations of ‘‘crisis forms’’ of parasites are suggestive of an orderly demise. If correct, an understanding of the death process could prove quite useful.

V.

A MECHANISM FOR ATOVAQUONE–PROGUANIL SYNERGY

As mentioned earlier, the unacceptable level of treatment failures arising from the use of atovaquone as a single drug led to the inclusion of a synergistic partner, proguanil, in a formulation that has been registered as a new antimalarial agent in several countries (33–35). The success of proguanil as a synergistic agent was somewhat surprising, as resistance to proguanil alone was widespread in parts of the world where the combination was tested in clinical trials (32). Further, the antimalarial activity of proguanil is mediated by cycloguanil, a parasite dihydrofolate reductase (DHFR) inhibitor that is a metabolite of cytochrome P450-mediated cyclization of proguanil (48,49). About 20–30% of the Asian and African population are deficient for the P450 isoform believed to carry out this metabolic activation (50,51), yet the atovaquone–proguanil combination was quite effective

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in these populations. Thus, there was a suspicion that proguanil acted as a prodrug to provide synergy to atovaquone rather than as its metabolite cycloguanil. Proguanil was tested alone and in combination with atovaquone for its effect on parasite ∆Ψm as well as on electron transport (52). At pharmacologically relevant concentrations, proguanil by itself had no appreciable effect on parasite ∆Ψm or respiration (52). However, it significantly enhanced atovaquone-mediated ∆Ψm collapse at micromolar concentrations (Fig. 3). This enhancing effect was mediated by the prodrug itself, since neither cycloguanil nor pyrimethamine (another parasite DHFR inhibitor) were able to affect atovaquone-mediated ∆Ψm

Figure 3 Effects of atovaquone alone and in combination with proguanil on the mitochondrial membrane potential in live intact malaria parasites. Mitochondrial membrane potential was assayed by flow cytometry of DiOC6 fluorescence intensity as described by Srivastava et al. (40). Atovaquone collapsed ∆Ψm with an EC50 of ⬃15 nM, and the inclusion of proguanil at 3.5 µM decreased the EC50 to ⬃2 nM.

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collapse (52). Proguanil-mediated enhancement was specific for atovaquone, since effects of other inhibitors of mitochondrial electron transport such as myxothiazol and antimycin were not altered by inclusion of proguanil. It was rather surprising that proguanil did not enhance the ability of atovaquone to inhibit parasite electron transport (52). Thus, the synergy between atovaquone and proguanil appears to be mediated by the enhancement of ∆Ψm collapse at a lower concentration of atovaquone. It appears quite clear that proguanil acts to enhance the atovaquone effect as a biguanide rather than as its metabolite cycloguanil (52). Biguanides have a long history of being used as agents that affect cellular physiology (53). Drugs such as metformin are currently used as hypoglycemic agents for the treatment of insulin-independent diabetes (54). It was initially suggested that these compounds are uncouplers of oxidative phosphorylation (55); however, the uncoupling effect is seen at millimolar concentrations, which are pharmacologically not relevant. The puzzle of atovaquone–proguanil efficacy against malaria in areas where a large number of individuals do not metabolize proguanil and where proguanil-resistant parasites are widespread can also possibly be explained by these results. The slow uptake of atovaquone and its high lipophilicity (26) may result in a relatively prolonged period of parasite exposure to suboptimal concentrations of the drug when it is used as a single agent. Under these conditions, atovaquone-resistant parasites appear to emerge frequently (31,32). Inclusion of proguanil with atovaquone will effectively lower the in vivo IC50 of atovaquone, thereby resulting in parasite demise at atovaquone concentrations that otherwise would have been suboptimal. The net result will be a much lower incidence of treatment failure and resistance emergence, which is what has been observed in clinical trials (32,34). Pharmacokinetic studies have shown that proguanil concentrations required for the enhancement of the atovaquone effect are achieved in an adult within 3.5 h after an oral dose of 200 mg (49,56). Hence the dose of the atovaquone–proguanil combination recommended for treating adults (1000 mg atovaquone and 400 mg proguanil per day for 3 days) should be sufficient to achieve effective concentrations in plasma for its optimal antimalarial activity. The molecular basis for the proguanil enhancement is unclear. The fact that proguanil did not enhance the atovaquone-mediated electron transport inhibition suggests that the ∆Ψm collapse effect of atovaquone could be uncoupled from its electron transport inhibition effect. In this regard, the atovaquone–proguanil combination may act as a site-specific uncoupler of the mitochondrial membrane potential. Fidock and Wellems (57) suggested that proguanil may have intrinsic activity independent of its metabolic activation and DHFR inhibition. Indeed, at concentrations higher than 12 µM, proguanil does seem to have an effect on ∆Ψm in malaria parasites, although specificity of this effect is unclear, as a number of compounds can affect mitochondrial physiology at high concentrations. The concentrations of proguanil used by Fidock and Wellems (57) are significantly higher (IC50 of 50–75 µM) than those achieved in vivo (49,56). Thus, the rele-

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vance of the intrinsic in vitro activity of proguanil to the clinical situation remains unclear. The synergistic activity of proguanil in experiments described in Figure 3 was observed at pharmacologically achievable drug concentrations. Nevertheless, it would be important to determine whether similar mechanisms apply to both the intrinsic in vitro activity and the atovaquone synergy shown by proguanil.

VI. RESISTANT MUTANTS DEFINE ATOVAQUONE BINDING REGION A.

Atovaquone-Resistant Mutants

To understand the molecular basis of atovaquone action it is important to identify the site at which the drug binds in a selective manner, and an investigation of the basis for resistance mutation could prove quite useful. Toward that end, a series of atovaquone-resistant P. yoelii parasites were derived by suboptimal treatment of infected mice with atovaquone. Resistance in this rodent malaria system arose in a single step and quite readily, reflecting the observation of 30% recrudescence during the clinical trials when atovaquone was used as a single agent. A total of nine independent mutants were derived through this regimen (58). The atovaquone-resistant malaria parasites were also resistant to the collapse of ∆Ψm as well as respiration inhibition by atovaquone (58). The IC50 ranged approximately 1000-fold higher than that for the parental parasites, ranging from 10,000 to 25,000 nM compared to ⬃ 15 nM for the parental parasites. Thus, development of atovaquone resistance in malaria parasites is accompanied by concomitant resistance to atovaquone-mediated ∆Ψm collapse and electron transport inhibition. The ∆Ψm in the resistant parasites in the presence of myxothiazol, a standard cytochrome bc1 complex inhibitor, was also examined. The EC50 for myxothiazol in the parental parasites is about 180 nM, and in the resistant parasites the EC50 values increased only two- to fourfold. Hence, the level of crossresistance for this inhibitor appeared to be much lower than for atovaquone (58). It was of interest to see if proguanil could continue to have synergism even in atovaquone-resistant parasites. However, for atovaquone-resistant parasites inclusion of proguanil did not affect the ∆Ψm collapse profile for atovaquone (58). This suggests that inclusion of proguanil will have no synergism with atovaquone once atovaquone resistance has emerged. B. Structural Basis for Atovaquone Resistance The above observations strongly suggest the possibility of structural changes within the atovaquone-binding site in malaria parasites. In a number of different organisms, essentially all naturally arising mutants resistant to inhibitors of the

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cytochrome bc1 complex carry mutations in the cytochrome b gene (59). Hence, the cytochrome b genes of the 6 kb mitochondrial genome from each of the nine independent atovaquone-resistant P. yoelii lines were examined. Mutations leading to amino acid changes were seen to cluster around a region of the ubiquinol oxidation (QO) site (58). As summarized in Figure 4b, three independent atovaquone-resistant lines had identical two-base-pair changes resulting in L271V and K272R mutations. Three others had identical one-base-pair changes resulting in I258M mutation; two lines had a one-base-pair change giving Y268C mutation; and one had a single change resulting in F267I mutation. For one of the atovaquone-resistant lines, the entire mtDNA was cloned and completely sequenced. This revealed total sequence identity with the parental mtDNA except for the two-base-pair mutations in the cytochrome b gene noted above (58). The mtDNA in malaria parasites is remarkably well conserved, with its apparent evolutionary drift being much slower than that for the nuclear genome (60). Furthermore, there is little sequence variation within a parasite species as judged from essentially complete sequence identity of the 6 kb mtDNA sequenced from geographically isolated clones; indeed, only a single nucleotide difference was observed over the entire 5966 bp mtDNA sequenced from two geographically distant P. falciparum isolates (60,61). Therefore, the rapid sequence changes observed in atovaquone-resistant parasites indicate a great selective advantage provided by these mutations in the presence of this drug. The mitochondrial DNA in malaria parasites appears to replicate through a rolling circle mechanism followed by extensive recombination and strand invasions (62). We have suggested that this mode of DNA replication may result in a high level of copy correction of the tandemly arrayed mitochondrial genome; while this would limit sequence divergence under normal conditions, rapid dissemination of an advantageous mutation would ensue once it does arise. The parental cytochrome b sequence was undetectable in atovaquone-resistant P. yoelii through a PCR-based assay. There are approximately 100 copies of mtDNA molecules per parasite in P. yoelii (11); it appears that in the resistant parasites none of these molecules displays the parental cytochrome b sequence. This indicates that, in very short order, the entire mtDNA repertoire has been converted to the resistant sequence, suggesting a rapid gene conversion rate for the parasite mtDNA. Inhibitors of cytochrome bc1 complex are believed to act mainly as ubiquinone/ubiquinol antagonists by interfering with ubiquinol oxidation or ubiquinone reduction steps of the proton-motive Q cycle (63). Crystallographic evidence has now localized myxothiazol, stigmatellin, and antimycin in their respective sites, which correspond very well with the locations of the mutations providing resistance in various systems (64–67). Atovaquone resistance mutations also localized to the general vicinity of the ubiquinol oxidation region of cytochrome b (Fig. 4a). They mapped within a highly conserved 15 amino acid region within the e–f loop of this subunit (Fig. 4b), which contains the universal PEWY sequence found in all cytochrome b. Of the five amino acid changes

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conferring atovaquone resistance, three (I258, Y268, and L271) involved residues that are absolutely conserved in all cytochrome b, while the other two (F267 and K272) involved residues that were different between the parasite and vertebrate proteins. Toxoplasma parasites resistant to atovaquone have been examined, and one mutant was found to contain an amino acid change (I254L) corresponding to the I258M change seen in Plasmodium; in another mutant a change involved amino acid M129L within the c–d loop (see Fig. 4b) (D. McFadden and J. C. Boothroyd, personal communication). In Plasmodium, mutations involving the c–d loop have not been seen as yet. In the cavity defined by the atovaquone resistance associated mutations (Fig. 4a), two amino acid residues that bear different side chains in the parasite and vertebrate cytochrome b seem to acquire a special significance with respect to the selectivity of atovaquone binding. These are F267 and K272 in malaria parasites, corresponding to A278 and R283 in the chicken or human, respectively (Fig. 4b). These positions are also conserved in Toxoplasma and Theileria cytochrome b (in Toxoplasma there is a tyrosine instead of phenylalanine at position 267). Hence, we would suggest that an aromatic side chain at position 267 in combination with lysine at 272 is required for sensitivity to atovaquone. If so, then in the vertebrate cytochrome b, alanine at position 278 in combination with the larger arginine at position 283 would appear to be responsible for resistance to atovaquone. This then would explain the selective toxicity of atovaquone toward parasites without affecting the vertebrate mitochondrial functions. Consistent with this proposal is the observation that other atovaquone resistance associated amino acid changes also subtly alter the hydrophobicity or volume of this cavity

Figure 4 Location of amino acid residues known to be involved in atovaquone resistance or susceptibility in Plasmodium yoelii and Toxoplasma gondii. (a) Coordinates from the X-ray diffraction study of beef heart bc1 complex (67) were obtained from the PDB, and the cytochrome b component was modeled using RASMOL. Positions of conserved residues in which alterations are known to confer resistance to atovaquone in Plasmodium yoelii or Toxoplasma gondii are highlighted against a dark background. Important residues were found to reside within the quinol oxidation (QO) pocket formed by the loops c–d and e–f linking transmembrane helices c, d and e, f, respectively. Five residues were identified in the e–f loop and one in the c–d loop. This pocket is proximal to the ironsulfur cluster (Fe2S2) of the Rieske protein (not shown) and to the low potential heme moiety of cytochrome b (heme bL). (b) Alignment of c–d and e–f loops of the cytochrome b from eight different organisms. Positions of conserved residues in which alterations are known to confer resistance to atovaquone in Plasmodium yoelii (AtqR line) or Toxoplasma gondii (arrow) are indicated. Concurrent mutations leading to AtqR resistance in Plasmodium are marked with a double line. Sequences were obtained from GenBank, and conserved residues were highlighted using BOXSHADE. Four genera of Apicomplexan parasites display a four amino acid deletion in the c–d loop.

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at absolutely conserved positions (Fig. 4b). This may greatly diminish the accessibility and/or affinity of atovaquone for this cavity without significantly affecting that of ubiquinol and effective electron transfer to the iron-sulfur protein. For efficient electron transport, ubiquinone has to move in and out of the catalytic sites of the bc1 complex. The flexible polyisoprenyl side chain at the carbon-2 position could aid this movement, whereas the relatively inflexible cyclohexylchlorophenyl side chain at the 2-position in atovaquone may hinder it by possibly interacting directly with the resistance-associated residues of cytochrome b.

VII.

PERSPECTIVES

This chapter has focused mainly on one aspect of rather subtle differences between the parasite and host bc1 complex that appear to be the basis for selective toxicity of a new class of antiparasitic drugs. Structural features of atovaquone binding site revealed here may assist in further development of selectively toxic drugs against structurally distinct cytochrome bc1 complexes of mitochondriacontaining eukaryotes. Such organisms form a vast group, ranging from protists and fungi to nematodes, and extract a major toll on health and the global economy. Rapid emergence of resistance, however, is likely to be a major problem if such compounds are used as single agents. To counter this, it is conceivable that combination therapy with multiple hydroxynaphthoquinones with subtle differences in their side chains may provide means to restrict rapid emergence of resistance. Acquisition of resistance to such drug combinations would require multiple mutations; such mutations would be rare and may be incompatible with the functional integrity of the parasite cytochrome bc1 complex. As we learn more about the unusual physiological processes associated with the parasite mitochondria, it is quite conceivable that these could form the basis for efforts at drug development. Examples of such parasite-specific features include mtDNA replication, transcription, and translation within mitochondria as well as the machinery involved in import of the entire complement of tRNAs for mitochondrial use. As the malaria parasite genome project comes to completion, additional targets among these processes are likely to be identified. The challenge will be to validate such targets, and, more important, to marshall the will as well as the resources to develop compounds based on these insights.

ACKNOWLEDGMENTS We deeply appreciate the grant support (AI28398) provided by the National Institutes of Health to enable our work on malaria parasite mitochondria. We also thank Dr. Fevzi Daldal for many stimulating discussions.

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12 Hepatic Cell Function in Liver Fluke Infection Linda M. Lenton and Carolyn A. Behm Australian National University, Canberra, Australia

I. INTRODUCTION Infection with the Fasciola species of liver fluke (fasciolosis) compromises growth and productivity in grazing animals [reviewed in Dargie (1)]. Predisposition to other diseases and sudden death may also be the outcome. Liver fluke infection presents a serious problem in many parts of the world; sheep, cattle, goats, and pigs are affected as well as many species of wildlife. In some countries where aquatic plants such as watercress are consumed as part of the diet, fasciolosis is also a significant human problem. Fasciola hepatica, a parasite of temperate climates, and its tropical counterpart Fasciola gigantica together are estimated to infect in excess of 2 million people, 200 million sheep, and 300 million cattle worldwide. F. hepatica is the more widely studied of these two species and is the subject of this chapter. The antihelminthic triclabendazole has proven to be the most effective in the treatment of Fasciola in animals and humans (2–4), though the cost of the drug restricts its application in developing countries. Problems of long-term residues in animals used in the meat and dairy industries and the development of drug resistance are also drawbacks in the application of such flukicidal compounds (5). At present there is no effective vaccine to prevent this infection. Fasciola antigens with the ability to lower fluke burden in cattle show some promise, but there are many problems to be overcome if this is to lead to a successful vaccine (6).

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II.

LIFE CYCLE OF THE PARASITE

Infection with F. hepatica commences when the mammalian host ingests the larval stages, metacercariae, encysted on vegetation. The metacercariae excyst in the small intestine, and the juvenile flukes, which are just visible to the naked eye, pass through the intestinal wall into the peritoneal cavity and thence penetrate the liver. They move through the liver, creating burrows and tracks and feeding on hepatic cells. The juvenile flukes grow very rapidly and eventually (after about 4–6 weeks in the rat and 8 weeks in the sheep) penetrate the walls of the main bile ducts, enter the bile ducts, become sexually mature, and start to release eggs into the bile. Adult flukes are leaf-shaped and attain a final length of 1–2 cm. They remain in the bile ducts, feeding on blood, for the duration of the infection, which varies between host species but can be as long as the lifetime of the host. The eggs pass in the bile to the intestine to be voided in the feces. In moist conditions, the eggs hatch and infect the intermediate host, a snail of the genus Lymnaea, in which the remainder of the life cycle is completed. More detailed information on the parasite and its biology can be found in Boray (7), Pantelouris (8), and Schmidt and Roberts (9). Rats, mice, guinea pigs, and rabbits have been used extensively as hosts in experimental liver fluke infections.

III. THE ROLE OF THE LIVER The liver plays a particularly vital role in the metabolism and physiology of all higher animals. Liver function is especially critical in ruminants because the liver is responsible for synthesizing and secreting into the blood substrates such as glucose and ketone bodies from precursors derived from the diet. The liver damage caused by flukes is substantial and has a significant impact on hepatic function. Major metabolic lesions are observed in the liver early in the infection of rats, mice, and sheep. Many hosts become severely ill and may die during this stage of the infection.

IV. PATHOLOGY Fasciola hepatica damages the liver directly by mechanical injury that occurs mainly during the tissue migration phase of the juvenile flukes (10) and by indirect means either as a result of secretions and excretions of the flukes (in the migratory phase or as adults in the bile ducts) or by eliciting cellular immune reactions and their pathological consequences (discussed below). During the bile duct stage of the infection, liver flukes also cause severe anemia due to blood

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loss into the bile (11) and a generalized edema that is due to loss of plasma proteins (12). In the rat, the first evidence of liver damage is the appearance of yellowwhite migratory tracks at the periphery of the liver at about 2 weeks postinfection (13). These become extensive as the infection progresses and are often hemorrhagic. Infiltration of these tracks with leukocytes, of which eosinophils form a prominent component, has been observed from 2 weeks post infection (14). Patches of necrosis appear, and the liver becomes enlarged, fibrosed, and in the long run cirrhotic. Enlargement and thickening of the bile ducts is apparent well in advance of the entry by the flukes (15). Maximal damage is inflicted on the liver by about 4 weeks postinfection, the period immediately prior to the flukes entering the bile duct. Despite the extensive literature on liver fluke infection, detailed study of functional changes in the liver of the host that might influence survival, growth, and productivity has been limited. The metabolism of infected livers has been studied at a number of functional levels, namely, whole organ in a perfused liver system, or cellular, using isolated hepatocytes, and subcellular, using standard fractionation techniques, in rats, mice, and sheep. The majority of this experimental work on liver function during fluke infection has been carried out using rats as a model host for the parasite. Strategic experiments and hypotheses have been tested in infected sheep, because they are an economically important host. Major liver functions that are compromised during liver fluke infection include energy metabolism (16), cytochrome P450-based xenobiotic metabolism (17), and albumin synthesis (18). There is also evidence that the liver is under oxidative stress during infection (19). All of these changes are most severe during the tissue migratory stage.

V.

MAJOR LESIONS IN HOST LIVER BIOENERGETIC METABOLISM IN THE RAT

A.

Uncoupling of Mitochondria

The observation that rat liver mitochondria are uncoupled during liver fluke infection was first made by Van den Bossche et al. (20). They observed uncoupling in isolated mitochondria as well as increased histochemical staining for mitochondrial ATPase activity in infected liver slices, indicative of a loss of respiratory control in situ (21). Using improved methods to isolate mitochondria, this uncoupling was confirmed by Rule et al. (16). At 2, 4, and 6 weeks postinfection, mitochondria isolated from infected livers exhibited acceptor control ratios of 1.0 and ADP/O ratios approached zero with both site I (pyruvate/malate) and site II (succinate) substrates, though uncoupling was more severe for site I. (See also Chap. 7.) This implies a loss of ability to maintain a proton gradient across

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the inner mitochondrial membrane. Respiration showed no response to the chemical uncoupler carbonyl cyanide m-chlorophenylhydrazone (CCCP) or the F0F1ATPase inhibitor oligomycin. At 2 and 6 weeks postinfection, state IV respiration rates of mitochondria were high, similar to those of control mitochondria in the presence of CCCP. However, at 4 weeks postinfection, when infected livers exhibit maximum physical damage, respiratory rates were severely attenuated at all three energy-conserving sites, suggesting damage to the mitochondrial electron transport chain. The integrity of the mitochondrial inner membrane was also questioned in the light of findings that mitochondria from infected livers showed enhanced respiration in the presence of NADH, to which mitochondria are normally impermeable. (See also Chap. 6.) Mitochondria from 4 week infected livers were subsequently shown to be unable to synthesize ATP in vitro (22). It is noteworthy that uncoupling was evident as early as 2 weeks postinfection, when the flukes are very small and little visible damage has occurred to the liver, and that uncoupling persisted into the bile duct stage (until at least 21 weeks postinfection), suggesting that the effect of the parasite on the liver may be an indirect one (16). By 11 weeks postinfection, some recovery was apparent, with a burden of at least three flukes resident in the bile duct being necessary for the persistence of uncoupling. B. F0 F1-ATPase ATPase activity in mitochondria isolated at 3 and 4 weeks postinfection was first found by Rule et al. (16) to be insensitive to oligomycin. This was confirmed by Lenton et al. (23), who extended the inhibitor study to include the F0F1-ATPase inhibitors N,N′-dicyclohexylcarbodiimide (DCCD) and diethylstilbestrol (DES). A pattern of insensitivity similar to that found for oligomycin was observed, with the ATPase complex regaining its inhibitor sensitivity by 6 weeks postinfection when the flukes were leaving the liver parenchyma for the bile duct. All of these inhibitors bind to the F0 moiety of the ATPase complex. Though it is thought that the oligomycin and DCCD binding sites are interrelated, DES is considered to bind at a distinct site. Therefore, lack of sensitivity to all of these inhibitors most likely represents a general problem in communication of inhibitor binding to the F1 portion of the complex, rather than specific damage at inhibitor binding sites. Purification of the F0 F1-ATPase from infected liver mitochondria produced an intact oligomycin-insensitive complex, but at a very low yield relative to control preparations (23). There was a failure to enrich specific activity during purification from infected material, indicating that the complex was structurally unstable. Leakage of ATPase activity from membranes into the supernatant fraction was evident during preparation of inner membrane vesicles. This was corrobo-

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rated by electron micrographs showing fewer F1 components projecting from inner membranes prepared from infected livers (Lenton et al., unpublished results). It therefore appears that the connection between the F0 and F1 moieties is compromised during infection. The yield of mitochondria from infected livers was less than 50% of that from control livers, suggesting instability of the organelle during isolation, though it was not ruled out that fewer mitochondria were present in vivo in infected livers (23). The specific activity of the mitochondrial enzyme citrate synthase increased in the cytosolic fraction from a control value of 3.0 nmol/(min.mg protein) to 11.3 nmol/(min.mg protein) (p ⱕ 0.002) at 4 weeks postinfection, also signifying structural damage to the mitochondrial membranes (Lenton et al., unpublished results). Citrate synthase specific activity in the mitochondrial fraction from infected livers did not differ significantly from control values. There was no apparent increase in cytosolic citrate synthase activity at 2 weeks postinfection. In isolated mitochondria, latent ATPase activity did not change throughout infection and was the same in mitochondria from control and infected livers. This is consistent with the above findings, as, unlike sensitivity to inhibitors, the ability of F1 to hydrolyze ATP is not dependent on attachment to F0. C. The Role of Nonesterified Fatty Acids and Altered Phospholipid Composition The concentration of nonesterified fatty acids (NEFAs), which are well-known uncoupling agents, is abnormally high in mitochondria isolated from infected rat livers (24). At 2 weeks postinfection the rate of state IV respiration (elevated during infection) of individual preparations correlated strongly with NEFA content and uncoupling could be largely reversed in vitro by the addition of defatted bovine serum albumin (BSA), which sequesters NEFA. By 3 weeks postinfection the concentration of NEFA in the mitochondria had increased further. The attenuated rate of electron transport typical of this stage could be relieved in vitro by the addition of excess BSA, but respiration remained uncoupled. Small volumes of mitochondria from 3 week infected livers caused uncoupling of control mitochondria in vitro, an effect that could be reversed by adding BSA but not by including the phospholipase inhibitor dibucaine in the incubation. Overall, these observations show that mitochondrial uncoupling has a reversible (early) component and an irreversible (late) component and that the reversible component could be a result of high concentrations of NEFA present in the mitochondria. Similarly, attenuation of electron transport also had a component that could be reversed with BSA. NEFAs at high concentrations are known to inhibit mitochondrial respiration (25). An increase in mitochondrial inner membrane permeability leading to uncoupling of oxidative phosphorylation can be brought about by the presence of

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an endogenous proton-translocating agent such as the NEFA discussed above, as well as by structural damage to the membrane resulting in ‘‘leakiness.’’ (See also Chaps. 6 and 7.) Analysis by 31P NMR spectroscopy of the phospholipid composition of mitochondria isolated from infected rat livers revealed that there was a significant decline in the relative concentration of phospholipids, to 40% of controls at 3 weeks postinfection (24). Significant decreases were observed in phosphatidylcholine, phosphatidylethanolamine, and phosphatidylinositol, accompanied by increases in a variety of degradation products such as lysophosphatidylethanolamine, glycerophosphocholine, glycerophosphoethanolamine, and phosphocholine. There was no apparent change in phosphatidylserine, cardiolipin, and sphingomyelin. When the phospholipid composition of inner mitochondrial membranes was analyzed, however, there was found to be a 50% loss of cardiolipin in the infected state. This, along with the loss of mitochondrial ATPase activity during the preparation of the inner membrane fraction (23), emphasizes the fragile nature of mitochondrial membranes from infected livers. That is, they were less able to withstand the harsher treatment involved in the preparation of inner membrane vesicles. The simultaneous loss of cardiolipin and ATPase activity may be related; many inner membrane proteins, including F0F1ATPase, are very tightly associated with cardiolipin (26). A 70% fall in total liver phospholipid content has also been reported (27). The changes in phospholipid profiles, coupled with the increased concentrations of NEFAs, implicate increased phospholipase activity plus possible derangement of phospholipid synthesis or acylation, as potential causes of the respiratory changes in mitochondria from infected livers. Further circumstantial evidence in support of increased phospholipase A2 activity is that the concentration of sphingomyelin was not altered by the infection (24); sphingomyelin is not a substrate for phospholipase A2 . A recent review of the role of phospholipases A2 in inflammation and their interaction with other inflammatory pathways can be found in Cirino (28). We also found a low level of lysophospholipids relative to phosphodiesters, indicating lysophospholipase activity. However, lysophospholipids still made up approximately 12% of the total mitochondrial phospholipids in infected livers compared with 1% in uninfected livers. Lysophospholipids may contribute to the mitochondrial pathology as they are capable of uncoupling oxidative phosphorylation, and at levels above their critical micellar concentration they can cause rearrangements of the membrane bilayer (29,30). As the concentration of NEFAs found in the mitochondria tallies with the amount of mitochondrial phospholipid lost, it is likely that the source of the NEFAs implicated in the mitochondrial uncoupling is the mitochondrial membranes. The elevated NEFAs are not the result of increased de novo hepatic fatty acid synthesis (Lenton et al., unpublished results). The liver is a complex organ

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composed of hepatocytes, Kupffer cells, stellate (Ito) cells, pit cells (resident natural killer cells), red blood cells, sinusoidal epithelial cells, and, in the inflamed state, recruited circulating leukocytes. With this in mind, there are many questions concerning changes in liver bioenergetic metabolism that are left unanswered. For example, it is not known in which cell population(s) the putative increase in phospholipase activity occurs [e.g., Kupffer cells are a rich source of mitochondrial phospholipase A2 (31)] or which cells contribute the NEFAs and lysophospholipids that give rise to uncoupled mitochondria.

VI. BIOENERGETIC FUNCTION IN VIVO In freeze-clamping experiments it has been shown that the ATP concentration of extracts of rat livers at 4 weeks postinfection was 75–85% that of matched control preparations (Millard et al., unpublished results); this shows that ATP synthesis is occurring in the infected liver but that there is a possibility of a shortage of ATP in the liver in vivo. It is therefore very unlikely that all the hepatic mitochondria are uncoupled in vivo. As the addition of a small amount of mitochondria from infected livers led to the uncoupling of control mitochondria in vitro (24), it is likely that a substance, such as NEFA, present in a subpopulation of uncoupled mitochondria in infected livers is uncoupling the remainder during or upon isolation or assay. The reversibility of uncoupling at 2 weeks postinfection by the addition of BSA supports this argument. However, preparation of mitochondria in the presence of BSA or the phospholipase inhibitor dibucaine did not prevent uncoupling in vitro at 2 or 3 weeks postinfection, showing that the damage, though possibly limited to only a subpopulation of the liver mitochondria, is probably present in the intact liver. In support of the hypothesized in vivo changes in mitochondrial bioenergetics discussed above, respiration in hepatocytes isolated from livers at 3 weeks postinfection was found to be abnormal (22). Like isolated mitochondria, which frequently exhibited attenuated respiration rates at this stage of the infection, hepatocyte respiration rates did not increase in the presence of CCCP. Also, at least some of the cells isolated from infected livers appeared to be permeable to succinate. This is symptomatic of the plasma membrane being breached and implies that perturbations to membrane structure during infection may not be confined to the mitochondria (see also Sec. VIII). Other indirect evidence supporting the concept of deranged bioenergetic function in vivo includes the demonstration of attenuated gluconeogenesis and abnormal Ca2⫹ fluxes in perfused infected rat livers following administration of glycogenolytic hormones (32). Calcium ion uptake by mitochondria is an important part of this process (33).

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VII.

HEPATIC BIOENERGETIC METABOLISM IN OTHER EXPERIMENTAL HOSTS

In sheep, abnormalities in the respiration of isolated mitochondria were observed from 4 to 15 weeks postinfection (34). That is, in common with the rat model, these changes persisted throughout the parenchymal stage and into the bile duct stage of the infection. But in many respects these abnormalities were different from those observed in the rat and were not accompanied by an increase in NEFA content or changes in phospholipid composition (35). For example, in the sheep, respiration in the presence of ADP (state III respiration) was attenuated, as was the response to CCCP, but there was no change from control values in the state IV respiration rate and the F0F1-ATPase retained its sensitivity to inhibitors. All changes in mitochondrial metabolism were restricted to regions of the liver that had been penetrated by the flukes. The site of these lesion(s) was assigned to the mitochondrial electron transport chain. Thus the characteristics of hepatic mitochondrial respiration and its effectors differ markedly between infected sheep and rats. This was also the case in mice, where uncoupling and attenuation of electron transport in mitochondria isolated from Balb/c and CBA mice was observed only at 4 weeks postinfection when liver damage was maximal and significant mortality occurred (36). These changes did not take place in Swiss outbred mice.

VIII. MICROSOMES The main focus of investigation into changes at the level of hepatic microsomal membranes during infection with F. hepatica has concerned host drug biotransformation systems, ranging from in vivo studies of drug pharmokinetics to in vitro microsomal drug metabolism. The principal effects of infection are a decrease in total cytochrome P450 content. The decrease is specific for isoforms of P450, which discounts a nonspecific inhibition of microsomal protein synthesis (37,38). This topic has been comprehensively reviewed elsewhere (39). Decreases in cytochrome P450 content and the implicit deficiencies in xenobiotic metabolism during fasciolosis are unique among the hepatic metabolic lesions examined to date in that they appear to be common across species. Studies have been conducted in sheep (40), cattle (41), rats (42), and mice (36). Whether the etiology of this lesion is common to these species has yet to be investigated. In the rat, the phospholipid composition of the microsomal fraction is also altered during infection and displays a pattern of change similar to that of the mitochondrial phospholipids (24). Loss of microsomal phospholipids (60%) exceeded that of the mitochondria (40%), with a concomitant massive increase in NEFA. As with mitochondria, such disruption to the lipid environment in the

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endoplasmic reticulum would be expected to have deleterious effects on the activity of integral membrane proteins such as P450, though this does not explain the isoform specificity mentioned above. The activity of the microsomal calcium ATPase has been shown to be dramatically reduced during the parenchymal phase of infection and was associated with a decrease in microsomal calcium uptake, a decline in microsomal membrane fluidity, and a rise in cytoplasmic Ca2⫹ concentration (38,43). While using enzyme markers to establish the levels of cross-contamination between isolated subcellular fractions, it came to our attention that glucose-6phosphatase activity could not be detected in homogenates or microsomes from infected livers (24). As hepatic glucose-6-phosphatase is a key enzyme in the maintenance of blood glucose homeostasis, this observation appeared to contradict findings that blood glucose levels in infected rats did not differ from controls under fed or starved conditions (Millard, Lenton, Bygrave, and Behm, unpublished results). Histochemical staining of liver sections for glucose-6-phosphatase activity revealed that substantial areas of infected liver were indeed deficient in activity, though some activity was present in all sections examined, with increased staining for this inducible enzyme apparent in both control and infected livers following fasting (Lenton et al., unpublished results). Glucose-6-phosphatase activity was clearly being lost during tissue fractionation. A factor present in part of the infected liver was responsible, as glucose-6-phosphatase activity from control microsomes was abolished upon incubation in the presence of microsomes from infected livers. Conditions in infected livers would certainly be conducive to inactivation of glucose-6-phosphatase during isolation, as this enzyme has been reported to be sensitive to inactivation in the presence of unsaturated fatty acids and products of lipid peroxidation as well as having a strong dependence on an appropriate phospholipid environment for activity (44,45). So far, evidence has been presented that several aspects of liver function are disrupted in vivo and in vitro during fasciolosis. It has now become clear that the nature of these changes, at least in the rat, predispose the infected liver to preparative artifacts upon subcellular fractionation, reinforcing the concept that observations in disrupted tissues need to be interpreted with caution.

IX.

OXIDATIVE STRESS

The excessive release of reactive oxygen species from sequestered and resident phagocytic cells of the immune response leads to oxidative stress to the liver in pathological states such as sepsis, endotoxemia, and ischemia-reperfusion. Liver pathology during liver fluke infection is considered to be initiated via a similar mechanism (19,46). A decrease in tissue glutathione (GSH) levels, which is indic-

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ative of oxidative stress, has been reported in rats during the tissue migratory phase (27,46,47). At the same time, concentrations of products of lipid peroxidation, malondialdehyde and conjugated dienes, are augmented (19,27). Though Galtier et al. (48) observed no change in GSH in rats, they did report a decline in GSH in the cytosolic fraction of infected lamb livers (37). When GSH was administered intraperitoneally to rats for the first 40 days of the infection, the result was normalization of glutathione and malondialdehyde levels, a reduction in the loss of phospholipids, and amelioration of the impairment of the drug-metabolizing capacity of the liver (27). As the liver is unable to take up GSH under normal circumstances, the authors proposed that the increase arose from an increase in the availability of precursor amino acids. Administration of uridine diphosphoglucose (UDPG) had previously produced similar results: an amelioration of the levels of hepatic GSH and malondialdehyde as well as P450 and associated enzyme activities (46). It was suggested that the mechanism of action of UDPG was possibly both direct and indirect, the latter involving increased NADPH production through the supply of substrate for glycolysis, the NADPH then being available for regeneration of GSH. The role of NADPH and the hexose monophosphate shunt during oxidative stress in the liver and in the nonparenchymal cells in particular has been reviewed recently (49). As has been proposed by Maffei Facino et al. (27,46), the administration of hepatoprotective agents that are able to prevent or counteract the metabolic damage during the acute stage of the infection has considerable merit. The maintenance of the liver’s detoxifying capacity would expedite the metabolism of flukicidal compounds, diminishing the associated problems of toxicity, tissue residues, and environmental contamination, all of which are exacerbated in animals with compromised xenobiotic metabolism. It has been shown in nutritional states that give rise to depletion of liver GSH that dietary supplementation with sulfur amino acids restores GSH levels (reviewed in Ref. 50). This, too, may have an application in fasciolosis.

X. THE ROLE OF THE HOST IMMUNE RESPONSE Administration of the antiinflammatory agent dexamethasone to rats at 48 h intervals for 8 days prior to assay either prevented or ameliorated many of the metabolic lesions discussed above (22,32). These included the uncoupling of mitochondria and their inability to synthesize ATP in vitro, impairment of perfused liver responses to glycogenolytic hormones, and loss of total cytochrome P450. Dexamethasone is pleiotropic in its actions. For example, it is known to kill T-lymphocytes and eosinophils, to inhibit the production of inflammatory

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cytokines and eicosanoids, and to prevent tumor necrosis factor-dependent activation of cytosolic and mitochondrial phospholipase A2 (51–56). Dexamethasone has also been reported to protect mitochondria directly from alterations to their functional integrity in vitro, though it is not clear if this would apply to the in vivo situation (57). To extend this study, we examined hepatic mitochondria from infected congenitally athymic (nude) rats. These mitochondria did not develop uncoupling or attenuation of electron transport, nor were elevated NEFA and changes in phospholipid composition evident in the mitochondria of these rats (22,24). They also showed levels of cytochrome P450 comparable with those of uninfected controls (58). As congenitally athymic and dexamethasone-treated rats share a deficiency of T-lymphocytes, these observations are evidence that many of the metabolic dysfunctions observed during fasciolosis, at least in the rat, are hostmediated and require T-lymphocytes. Importantly, in these experiments the livers of dexamethasone-treated and athymic rats were traversed with fluke tracks to an extent similar to those of normal infected rats but were notable for their lack of infiltration by leukocytes (22). Necrosis, too, was less evident. The flukes matured normally in both dexamethasone-treated and athymic rats, with eggs being recovered in the feces of dexamethasone-treated rats at 10 weeks postinfection. In contrast to these findings in the rat, administration of dexamethasone to sheep early in infection led to increased liver damage and more rapid parasite development (59). Further, infected athymic (nude) mice generally died during the liver migratory phase (60), as did rabbits treated with an antilymphocyte serum (61). Increased fluke burdens and larger flukes have been reported in rats treated with anti-inflammatory drugs and in athymic rats, respectively (62,63). So even though the immune response in the rat has been implicated as a major contributing factor in the development of various hepatic lesions, it appears also to have a protective role, particularly in other species.

XI.

CONCLUSION

The differences between species in their metabolic responses to fasciolosis have been highlighted. The immune response has emerged as a major player in the determination of the extent and type of injury sustained by the host liver in the rat, but whether this is the case in other host species needs to be investigated. As far as bioenergetic metabolism is concerned, the rat does not appear to be a suitable model for this infection in sheep. The lethality of the infection in mice, particularly immunocompromised strains, precludes their use in elucidating the role of the immune response in the hepatopathology of fasciolosis.

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REFERENCES 1. Dargie JD. The impact on production and mechanisms of pathogenesis of trematode infections in cattle and sheep. Int J Parasitol 1987; 17:453–643. 2. Boray JC, Crowfoot PD, Strong MB, Allison JR, Schellenbaum M, Von Orelli M, Sarasin G. Treatment of immature and mature Fasciola hepatica infections in sheep with triclabendazole. Vet Rec 1983; 113:315–317. 3. Suhardono, Widjajanti S, Stevenson P, Carmichael IH. Control of Fasciola gigantica with triclabendazole in Indonesian cattle. Trop Anim Health Product 1991; 23:217– 220. 4. Laird PP, Boray JC. Human fascioliasis successfully treated with triclabendazole. Aust NZ J Med 1992; 22:45–47. 5. Overend DJ, Bowen FL. Resistance of Fasciola hepatica to triclabendazole. Aust Vet J 1995; 72:275–276. 6. Spithill TW, Dalton JP. Progress in the development of liver fluke vaccines. Parasitol Today 1998; 14:224–228. 7. Boray JC. Experimental fascioliasis in Australia. In: Dawes B, ed. Advances in Parasitology. New York: Academic Press, 1969:96–210. 8. Pantelouris EM. The Common Liver Fluke. Oxford; UK: Pergamon Press, 1965. 9. Schmidt GD, Roberts LS. Foundations of Parasitology 3rd ed. Times Mirror/Mosby College Publishing, 1985: Chap 18. 10. Dawes B, Hughes DL. Fascioliasis: the invasive stages of Fasciola hepatica in mammalian hosts. Adv Parasitol 1964; 2:97–168. 11. Jennings FW. The anaemias of parasitic infections. In: Soulsby EJL, ed. Pathophysiology of Parasitic Infection. New York: Academic Press, 1976:41–67. 12. Dargie JD. Blood protein turnover in parasitized ruminants. Isotopes Radiat Parasitol 1981; 4:155–170. 13. Thorpe E. The pathology of experimental fascioliasis in the albino rat. J Compar Pathol 1965; 75:39–45. 14. Rajasekariah GR. Immunity to Fasciola hepatica in rats. PhD thesis, Zoology Department, Australian National University, Canberra, Australia, 1977. 15. Dawes B. Hyperplasia of the bile duct in fascioliasis and its relation to the problem of nutrition in the liver-fluke, Fasciola hepatica L. Parasitology 1963; 53:123–133. 16. Rule CJ, Behm CA, Bygrave FL. Aberrant energy-linked reactions in mitochondria isolated from the livers of rats infected with the liver fluke Fasciola hepatica. Biochem 1989; 260:517–523. 17. Galtier P, Battaglia A, More J, Franc M. Impairment of drug metabolism by the liver in experimental fascioliasis in the rat. J Pharm Pharmacol 1983; 35:729–733. 18. Dargie JD. The applications of radioisotope techniques to the study of red cell and plasma protein metabolism in helminth diseases of sheep. In: Taylor AER, Muller R, eds. Pathogenic Processes in Parasitic Infections. Oxford, UK: Blackwell Scientific, 1975:1–26. 19. Maffei Facino R, Carini M, Genchi C, Tofanetti O, Casciarri I. Participation of lipid peroxidation in the loss of hepatic drug-metabolizing activities in experimental fascioliasis in the rat. Pharmacol Res 1989; 21:549–559.

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20. Van den Bossche H, Verhoeven H, Lauwers H. Uncoupling of liver mitochondria associated with fascioliasis in rats. Normalization by closantel. In: Van den Bossche H, ed. The Host Invader Interplay. Amsterdam: Elsevier, 1980:699–704. 21. Van den Bossche H, Verheyen A, Verhoeven H, Arnouts D. Alterations in rat liver mitochondria caused by Fasciola hepatica. In: Gigase PL, Van Marck EAE, eds. From Parasite Infection to Parasitic Disease. Basel: Karger, 1983:30–38. 22. Hanisch MJE, Topfer F, Lenton LM, Behm CA, Bygrave FL. Restoration of mitochondrial energy-linked reactions following dexamethasone treatment of rats infected with the liver fluke Fasciola hepatica. Biochim Biophys Acta 1992; 1139: 196–202. 23. Lenton LM, Behm CA, Bygrave FL. Characterization of the oligomycin-sensitivity properties of the F1F0-ATPase in mitochondria from rats infected with the liver fluke Fasciola hepatica. Biochim Biophys Acta 1994; 1186:237–254. 24. Lenton LM, Behm CA, Bygrave FL. Aberrant mitochondrial respiration in the livers of rats infected with Fasciola hepatica: the role of elevated nonesterified fatty acids and altered phospholipid composition. Biochem J 1995; 307:425–431. 25. Rottenberg H, Hashimoto K. Fatty acid uncoupling of oxidative phosphorylation in rat liver mitochondria. Biochemistry 1986; 25:1747–1755. 26. Schlame M, Haldar D. Cardiolipin is synthesized on the matrix side of the inner membrane in rat liver mitochondria. J Biol Chem 1993; 268:74–79. 27. Maffei Facino R, Carini M, Aldini G, Ceserani R, Casciarri I, Cavalletti E, Verderio L. Efficacy of glutathione for treatment of fascioliasis. Arzneim-Forsch/Drug Res 1993; 43:455–460. 28. Cirino G. Multiple controls in inflammation. Extracellular and intracellular phospholipase A2, inducible and constitutive cyclooxygenase, and inducible nitric oxide synthase. Biochem Pharmacol 1998; 55:105–111. 29. Witter RF, Morrison A, Shephardson GR. Effect of lysolecithin on oxidative phosphorylation. Biochim Biophys Acta 1957; 26:120–129. 30. Prokazova NV, Zvezdina ND, Korotaeva AA. Effect of lysophosphatidylcholine on transmembrane signal transduction. Biochemistry (Moscow) 1998; 63:31–37. 31. Hatch GM, Vance DE, Wilton DC. Rat liver mitochondrial phospholipase A2 is an endotoxin-stimulated membrane-associated enzyme of Kupffer cells which is released during liver perfusion. Biochem J 1993; 293:143–150. 32. Hanisch MJE, Behm CA, Bygrave FL. Beneficial effect of dexamethasone on attenuated hormone-induced uptake of calcium and glycogenolysis by perfused liver of rats infected with Fasciola hepatica. FEBS Lett 1991; 285:94–96. 33. Altin JG, Bygrave FL. Second messengers and the regulation of Ca2⫹ fluxes by Ca2⫹mobilizing agonists in rat liver. Biol Rev 1988; 63:551–611. 34. Rule CJ, Hanisch MJE, Behm CA, Bygrave FL. Aberrant energy-linked reactions in mitochondria isolated from the livers of sheep infected with the liver fluke Fasciola hepatica. Int J Parasitol 1991; 21:353–355. 35. Lenton LM, Bygrave FL, Behm CA. Fasciola hepatica infection in sheep: changes in liver metabolism. Res Vet Sci 1996; 61:152–156. 36. Somerville AC, Bygrave FL, Behm CA. A study of hepatic mitochondrial respiration and microsomal cytochrome P450 content in mice infected with the liver fluke Fasciola hepatica. Int J Parasitol 1995; 25:667–672.

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37. Galtier P, Larrieu G, Tufenkji AE, Franc M. Incidence of experimental fascioliasis on the activity of drug-metabolizing enzymes in lamb liver. Drug Metab Dispos 1986; 14:137–141. 38. Galtier P, Cambon-Gros C, Fernandez Y, Deltour P, Eeckhoutte C, Hoellinger H. Fasciola hepatica: liver microsomal membrane functions in host rat. Exp Parasitol 1994; 78:175–182. 39. Behm CA, Sangster NC. Pathology, pathophysiology and clinical aspects. In: Dalton JP, ed. Fascioliosis. Wallingford, UK: CABI, 1999:185–224. 40. Galtier P, Larrieu G, Beaune P. Characterization of the microsomal cytochrome P450 species inhibited in rat liver in the course of fascioliasis. Biochem Pharmacol 1986; 35:4345–4347. 41. Maffei Facino R, Carini M, Genchi C. Impaired in vitro metabolism of the flukicidal agent nitroxynil by hepatic microsomal cytochrome P-450 in bovine fasciolosis. Toxicol Lett 1984; 20:231–236. 42. Maffei Facino R, Carini M, Bertuletti R, Genchi C, Malchiodi A. Decrease of the in vitro drug-metabolizing activity of the hepatic mixed function oxidase system in rats infected experimentally with Fasciola hepatica: pharmacological implications. Pharmacol Res Commun 1981; 13:731–741. 43. Galtier P, Eeckhoutte C, Larrieu G. Fasciola hepatica: liver enzymes in rats and interaction with chemical inducers. Exp Parasitol 1987; 63:189–194. 44. Mithieux G, Bordeto J-C, Minassian C, Ajzannay A, Mercier I, Riou J-P. Characteristics and specificity of the inhibition of glucose-6-phosphatase by arachidonic acid. Eur J Biochem 1993; 213:461–466. 45. Koster JF, Slee RG, Montfoort A, Lang J, Esterbauer H. Comparison of the inactivation of microsomal glucose-6-phosphatase by in situ lipid peroxidation-derived 4hydroxynonenal and exogenous 4-hydroxynonenal. Free Rad Res Commun 1986; 1:273–287. 46. Maffei Facino R, Carini M, Genchi C, Tofanetti O, Casciarri I, Bedoschi D. Antihepatotoxic properties of uridine-diphosphoglucose in liver fluke infection. ArzneimForsch/Drug Res 1990; 40:490–498. 47. Gonzalez P, Tunon MJ, Lopez P, Diez N, Gonzalez J. Hepatic disposition of organic anions in rats infested with Fasciola hepatica. Exp Parasitol 1991; 73:396–402. 48. Galtier P, Vandenberghe Y, Coecke S, Eeckhoutte C, Larrieu G, Vercruysse A. Differential inhibition of rat hepatic glutathione S-transferase isoenzymes in the course of fascioliasis. Mol Biochem Parasitol 1991; 44:255–260. 49. Spolarics Z. Endotoxaemia, pentose cycle, and the oxidant/antioxidant balance in the hepatic sinusoid. J Leukocyte Biol 1998; 63:534–541. 50. Bray TM, Taylor CG. Tissue glutathione, nutrition and oxidative stress. Can J Physiol Pharmacol 1993; 71:746–751. 51. Alnemrie S, Litwack G. Glucocorticoid-induced lymphocytosis is not mediated by an induced endonuclease. J Biol Chem 1989; 264:4104–4111. 52. Rolfe FG, Hughes JM, Armour CL, Sewell WA. Inhibition of interleukin-5 gene expression by dexamethasone. Immunology 1992; 77:494–499. 53. Moncada S, Palmer RMJ. Inhibition of nitric oxide synthase by glucocorticoids: yet another explanation for their anti-inflammatory effects? Trends Pharmacol Sci 1991; 12:130–131.

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54. Van den Bosch H, Schaljwijk C, Pfeilschifter J, Marki F. The induction of group II phospholipase A2 by cytokines and its prevention by dexamethasone. Adv Exp Med Biol 1992; 318:1–10. 55. Barnes PJ, Adcock I. Anti-inflammatory actions of steroids: molecular mechanisms. Trends Pharmacol Sci 1993; 14:436–441. 56. Hoeck WG, Ramesha CS, Chang DJ, Fan N, Heller RA. Cytoplasmic PLA2 activity and gene expression are stimulated by tumour necrosis factor: dexamethasone blocks the induced synthesis. Proc Natl Acad Sci USA 1993; 90:4475–4479. 57. Campbell PI, Al-Nasser IA. Dexamethasone inhibits inorganic phosphate stimulated Ca2⫹-dependent damage of isolated rat liver and renal cortex mitochondria. Comp Biochem Physiol C 1995; 111:221–225. 58. Topfer F, Lenton LM, Bygrave FL, Behm CA. Importance of T-cell-dependent inflammatory reactions in the decline of the microsomal cytochrome P450 concentration in livers of rats infected with Fasciola hepatica. Int J Parasitol 1994; 25:1259– 1262. 59. Sinclair KB. The effect of corticosteroid on the plasma protein of lambs infected with Fasciola hepatica. Res Vet Sci 1968; 9:181–183. 60. Erikson L. Influence of thymus function on the course of infection in mice. Nord Veterinaermed 1980; 32:493–500. 61. Dodd K, O’Nualla´in TO. Effect of antilymphocytic sera on the histopathology of Fasciola hepatica infestations in rabbits. J Pathol 1969; 99:335–337. 62. Doy TG, Hughes DL. Evidence for two distinct mechanisms of resistance in the rat to reinfection with Fasciola hepatica. Int J Parasitol 1982; 12:357–361. 63. Baeza E, Poitou I. Influence of anti-inflammatory treatments on experimental infection of rats with Fasciola hepatica: changes in serum levels of inflammatory markers during the early stages of fasciolosis. Res Vet Sci 1994; 57:172–179.

13 Lipopolysaccharide A Membrane-Forming and Inflammation-Inducing Bacterial Macromolecule Ulrich Seydel, Artur J. Ulmer, Stefan Uhlig, and Ernst Theodor Rietschel Research Center Borstel, Borstel, Germany

I. INTRODUCTION Endotoxins are formed by a certain group of bacteria, the gram-negative bacteria, which differ from other microorganisms by the unique architecture of their cell wall and thus their staining behavior as devised by Hans-Christian Gram (1). Endotoxins were originally described as heat-stable components of Vibrio cholerae by Richard Pfeiffer (2) and are known today to be generally present in the cell envelope of gram-negative bacteria. Here they are major and integral components of the outer membrane, being exclusively located in its outer leaflet facing the bacterial environment. Endotoxins participate in the physiological membrane functions and are therefore essential for bacterial growth and viability (3). At the same time, endotoxins represent a primary target for interaction with antibacterial drugs and components of the immune system of the host. They have therefore attracted the interest of microbiologists and bacterial geneticists, who initiated studies to understand the biosynthesis and the molecular basis of the vital function of endotoxins for bacteria. Further, endotoxins were recognized as potent toxins that, in higher organisms, elicit a broad spectrum of biological activities. They play an important role in the pathogenesis and manifestation of gram-negative infection in general and of septic shock in particular. As a result, endotoxins intrigued clinical and biological researchers, who set out to elucidate their mode of action and to devise strategies aiming at control of the detrimental endotoxin effects observed during severe 217

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bacterial infection and sepsis. In addition, endotoxins are capable of producing beneficial effects in higher organisms depending upon their amount and route of introduction (4). Thus, endotoxins belong to the most active stimulators of the mammalian defense system. They induce the formation of antibodies directed not only to themselves but also to unrelated antigens, and they activate immune cells, resulting in an enhancement of the body’s capacity to cope with microbial infections and also malignant tumors. It was found that harmful as well as beneficial host responses to LPS are mediated by immune modulators such as tumor necrosis factor α (TNFα), members of the interleukin family (e.g., IL-1, IL-6, IL-8, IL-12), interferons, reduced oxygen species, and lipids. These mediators are released by various types of host cells, e.g., monocytes/macrophages, polymorphonuclear cells, vascular cells, and T lymphocytes (5–10). Endotoxins have therefore become a major subject of immunological research. Finally, endotoxins possessing a complex primary structure and an even more complex three-dimensional architecture have always fascinated chemists and physicists, who wished to elucidate those structures that are responsible for endotoxin activity, such as membrane formation or mammalian toxicity and its function as a physiological membrane component and immunoactivator. This chapter summarizes recent advances made with regard to the chemical and physical structure of endotoxin and attempts to translate this knowledge into an understanding of the membrane-forming, cell activating, and proinflammatory activities of endotoxins in vitro and in vivo.

II.

MEMBRANE ASPECTS OF ENDOTOXIN ACTIVITY

Membranes, in general, constitute the border between a cell or a cell compartment and its environment. They are composed of (glyco)lipids and proteins, function as permeability barriers, maintain constant ion gradients across the membrane, and guarantee a steady state of fluxes in the cell. Furthermore, cell membranes carry recognition sites for components of the immune system and for the interaction with other cells. These functions, to work properly, require a particular lipid composition on each leaflet as well as a particular lipid distribution between the two leaflets of the lipid bilayer. Thus, a membrane is built up from a large variety of lipids, differing in their charge and fatty acid pattern (length and degree of saturation). These lipids are in a delicate equilibrium, providing a suitable environment for protein function and membrane permeability. By a complex interaction of passive and active transport processes—by diffusion through the lipid matrix or proteinaligned transmembrane channels and by energy-dependent ion pumps and transport proteins, respectively—ion gradients are built up that contribute, together

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with the charge distribution of the lipids on the two leaflets of the membrane, to a transmembrane potential. The biological activity of endotoxins may be discussed in the context of the interaction of LPS molecules with the membranes of host cells, because intercalation of endotoxin molecules is considered to be an important step in the initiation of biological effects (11,12). A prerequisite for normal cell functioning is the maintenance of a particular composition of the lipid matrix at given ambient conditions (13). Disturbances of this composition, e.g., by uptake of exogenous lipids that differ in their chemical structure (e.g., acylation pattern, headgroup conformation, net electric charge) from that of the normal constituents of the cell matrix, may lead to (1) alterations of membrane fluidity and/or permeability, (2) phase separation and domain formation, (3) disturbance of the lamellar membrane architecture, and even (4) internalization of the extraneous lipids. In general, the cell may be able to compensate for the changes by altering the composition of the lipid matrix [‘‘homoviscous adaptation’’ (14)]. If this is not possible (the process takes more than minutes), any one of these membrane alterations may cause severe dysfunctions of the cell. These may manifest themselves, for example, in transient or permanent alterations in the functioning of transmembrane proteins that might be involved in signal transduction. The membrane alterations and their influence on cell functioning will be the more severe the more the chemical structures and conformations of the constituent and the interacting lipids differ. As their amphiphilic character suggests, endotoxins should interact with cell membranes nonspecifically via hydrophobic interaction. This interaction has been postulated in various investigations (11,15,16) and should consist of a direct interaction of small endotoxin aggregates down to monomers. This mechanism may be assumed to be responsible for cell activation at high endotoxin concentrations. At low endotoxin concentrations another mechanism of interaction is proposed that proceeds via specific coupling either directly to a membrane bound receptor protein (mCD14, TLR2, CD11/CD18, CD55) (17–23) or indirectly to the acute-phase protein lipopolysaccharide-binding protein (LBP), which then transfers LPS to membrane-associated receptors such as mCD14 (24,25). Furthermore, it has been shown that in the case of CD14-negative endothelial and epithelial cells, a soluble form of CD14 (sCD14) mediates LPS binding (26). Because mCD14 is anchored in the membrane by glycosylphosphatidylinositol (GPI) and therefore lacks a transmembrane domain, it may be hypothesized that none of these pathways leads directly to cell activation and that further steps are involved in the initiation of cell activation and signaling. One early step could be binding (directly or CD14-mediated) to a transmembrane protein such as TLR2, CD11/ CD18, or an ion channel (27), another the internalization of endotoxin (28), and both processes may subsequently be involved in triggering an intracellular signaling pathway. The discussion of the activation pathways will be taken up in more

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detail in a later section of this chapter. However, independent of the detailed mechanism of endotoxin–cell interaction it may be expected that unspecific intercalation as well as specific binding of endotoxin should depend on its physical parameters like the critical aggregate concentration [CAC; also termed critical micelle concentration (CMC)], phase state of the acyl chains, and the size and conformation of the hydrophobic moiety of LPS, i.e., of lipid A, its endotoxic principle.

III. CHEMICAL STRUCTURE OF ENDOTOXIN According to their chemical nature, endotoxins constitute lipopolysaccharides (LPS), and in the case of Enterobacteriaceae they consist of three structural units, i.e., the O-specific chain, the core region, and the lipid A component (Fig. 1). The O-specific chain is unique and characteristic for a given LPS serotype and is therefore highly variable among LPS molecules of different serotypic origin. The structure of the O-specific chain is characterized by a polymer of repeating oligosaccharide units that carry the epitopes for O-specific (serotype-specific) antibodies and phages. The core portion is structurally less variable and may be divided into the O-chain proximal outer core and the lipid A proximal inner core. The variability of the outer core is due to different locations and linkages of its constituents. In Enterobacteriaceae, common elements present in the outer core

Figure 1 Schematic representation of the architecture of an enterobacterial wild-type lipopolysaccharide.

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comprise the pyranosidic hexoses d-glucose, d-galactose, 2-amino-2-deoxy-dglucose (GlcN), and 2-amino-2-deoxy-d-galactose (GalN). The inner core is composed of 3-deoxy-d-mannooctonic acid (dOclA, also termed 2-keto-3-deoxyoctulosonic acid, Kdo) and heptose residues, in general of the l-glycero-d-manno configuration, the latter being often phosphorylated. Lipopolysaccharides expressing the complete core and the O-specific chain are termed wild-type or smooth (S-form) LPS, those lacking the O-chain, rough mutant LPS (R-LPS). LPS carrying only the Kdo units bound to lipid A are classified as deep rough mutant LPS (Re-LPS). Lipid A has been identified as the membrane-forming and inflammationinducing principle of LPS, thus constituting the minimal structure that expresses many of the biological activities and functions of intact LPS (29). The chemical structure of lipid A of most gram-negative bacteria studied consists of a bisphosphorylated β1,6-linked d-GlcN disaccharide that carries up to six acyl groups (in a few cases even up to seven, e.g., Salmonella enterica spp.). This structure is highly conserved among bioactive LPSs (30). As an example, Figure 2 shows the chemical structure of the lipid A component of Escherichia coli. Six fatty acids are nonsymmetrically distributed over the two GlcN residues. An esterlinked (R)-3-hydroxymyristic acid at C-3′ carrying myristic acid at its 3-OH group and an amide-linked (R)-3-hydroxymyristic acid that is acylated by lauric acid at its 3-OH group are attached to the nonreducing GlcN residue (GlcN II). The reducing GlcN residue (GlcN I) carries 2 mol of (R)-3-hydroxymyristic acid, one in ester (position 3) and one in amide (position 2) linkage. Dissimilarities among lipid A structures of different bacterial origin are due to variations in the

Figure 2 Chemical structures of agonistic lipid A of LPS from Escherichia coli (corresponding to the synthetic compound 506) and of the LPS antagonists precursor Ia (compound 406) and synthetic analog of lipid A from Rhodobacter capsulatus LPS (compound E5531).

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number, location, and chain length of fatty acids, the substitution of phosphate groups, and the presence of 2,3-diamino-2,3-dideoxy-d-glucose (GlcN3N) instead of GlcN (30).

IV. STRUCTURAL REQUIREMENTS FOR ENDOTOXIN BIOACTIVITY In recent years, we and others have investigated R- and S-form LPSs of various bacterial origins and different lipid A and lipid A partial structures with our aim being to characterize the bioactive regions of LPSs in more detail. In particular, the successful chemical synthesis of lipid A and corresponding lipid A partial structures, such as E. coli-type lipid A (termed compound 506 or LA-15-PP) or precursor Ia (compound 406, LA-14-PP, or lipid IVa) carrying only four (R)-3hydroxymyristic acid residues in symmetrical distribution has provided the experimental basis for these investigations (31–34). The chemical structures of bioactive (agonistic) lipid A from E. coli LPS (compound 506) and two prominent antagonistic lipid A partial structures or analogs (compounds 406 and E5531) are shown in Figure 2. It was found that full biological activity is expressed by a lipid A molecule consisting of a hexaacylated, bisphosphorylated β1,6-linked d-GlcN disaccharide, i.e., an E. coli-type lipid A (or compound 506, see Fig. 3) (34). Lipid A partial structures deficient in one of these elements are less active or even nonactive regarding the induction of monokines in human monocytes. For instance, the 1-dephospho (compound 504) and the 4′-dephospho (compound 505) synthetic lipid A partial structures were less active than compound 506, highlighting the significance of the phosphoryl groups (or the charges per se) for the biological activity of lipid A. The lipid A precursor Ia (compound 406) is completely inactive in inducing IL-1, IL-6, and TNF release in human monocytes (31,33,35– 37) and in activating human T lymphocytes (9). The heptaacylated lipid A component of S. enterica vs. Minnesota spp. (compound 516) shows lower bioactivity than compound 506. Also, the location of the secondary acyl residues is of importance, as shown by the low bioactivity of compound LA-22-PP, having, in contrast to compound 506, a symmetrical distribution of the fatty acids. As it is known that target cell membranes contain various LPS-binding proteins (e.g., CD14), it seemed reasonable to test lipid A-like structures or partial structures for their ability to block LPS-induced activation. To this end, a variety of lipid A analogs and nonbioactive synthetic partial structures have been tested with respect to LPS-induced reactions in vitro and in vivo. We and others could show that among these the lipid A precursor Ia (compound 406) is a very effective LPS-antagonist, inhibiting monokine production (35–39). Similar antagonistic effects were described for the pentaacyl bisphosphoryl lipid A isolated from Rhodobacter sphaeroides (37,40) and Rhodobacter capsula-

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Figure 3 Molecular conformations (shapes) of lipid A monomers (left) in relation to supramolecular structures (right).

tus (41). These observations led to the development of an antagonistic pentaacyl lipid A analog, the synthetic compound E5531, the structure of which is based on R. capsulatus lipid A. This lipid A analog has been analyzed for therapeutic application in in vivo models and was, indeed, found to protect mice from endotoxin-induced lethality and, when administered simultaneously with an antibiotic, from the lethal outcome of an E. coli-induced peritonitis (42). Concerning the mechanism of the inhibitory action of the LPS antagonists, Lineweaver–Burk plot analyses suggested a competitive inhibition of LPS binding to its receptor on human monocytes by lipid A analogs (43). However, at very low concentrations precursor Ia is able to block cytokine release in the human monocytic cell line THP1 under conditions where binding of LPS was not affected (44). In this case, therefore, a noncompetitive mechanism is likely to be operative. Further experimental evidence in favor of this has been provided by Thie´blemont et al. (45), who found that the antagonistic LPS from R. sphaeroides, when presented as a monomeric complex with soluble CD14, remained at the cell membrane of polymorphonuclear cells and monocytes and, in contrast to

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inflammatory active LPS, was not transported to cytoplasmic vesicles and, furthermore, that LPS from R. sphaeroides inhibited the intracellular transport of bioactive LPS.

V.

PHYSICAL STRUCTURE OF ENDOTOXIN

As early as the 1960s, physicochemical parameters of LPS were considered to be important for an understanding of its bioactivity. The importance of physicochemical parameters refers in the first place to the critical aggregate concentration (CAC), the phase state (fluidity of the endotoxin acyl chains), which is a property of the bulk lipid, and the molecular conformation (shape), which is a property of the individual molecules, the molecular shape being deduced from the type of supramolecular aggregate structure built up from a large number of identical endotoxin molecules. Since lipid A as the endotoxic principle of LPS constitutes the molecular entity that primarily interacts with the host cell membrane, the determination of its conformation should be of utmost importance for an understanding of biological action. The role of the sugar moiety of LPS should be restricted to a modulation of lipid A bioactivity, mainly due to its influence on the hydrophobicity of endotoxin molecules and on their CAC but also on the fluidity of the lipid A acyl chains. Thus, a more detailed physicochemical characterization of the above-mentioned parameters, mainly of lipid A but also of LPS, remains an important issue in the characterization of the parameters governing endotoxicity and in answering the question as to why some endotoxin structures are biologically highly active but others are inactive (for an extensive review, see Ref. 46). A.

Phase States

For amphiphilic compounds like lipid A and LPS, an endothermic transition between a highly ordered gel phase (β phase) and a less ordered liquid crystalline phase (α phase) takes place upon temperature variations. The transition temperature Tc depends on properties of the hydrophilic headgroup (charge, size, conformation, water-binding capacity) as well as of the hydrophobic moiety (number, length, and degree of saturation of the acyl chains). A transition between the different phase states is usually accompanied by changes in the geometry of the molecules involved, i.e., of the geometrical cross sections of the molecules, and may therefore have an impact on the structure of the supramolecular assemblies. We found a very characteristic dependence of Tc on the chemical structure of the sugar region of LPS (47–51). Thus, for enterobacterial strains, Tc was highest for free lipid A (around 45°C), lowest for deep rough mutant LPS (around 30°C), and, with increasing length of the polysaccharide portion toward comple-

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tion of the O-chain (wild-type LPS), it increased again up to 37–40 °C. The phase transition temperatures of various nonenterobacterial LPSs and lipid A were found to be significantly lower than those of Enterobacteriaceae (48,52). This observation correlates with the fact that the former contain shorter acyl chains with a higher degree of unsaturation (53). The addition of divalent cations or a lowering of pH caused a significant rigidification of the acyl chains of free lipid A and LPS preparations and, in part, also led to an increase in Tc . B. Critical Aggregate Concentration, Supramolecular Structure, and Molecular Conformation Lipopolysaccharide and lipid A represent amphiphiles and therefore form aggregates in aqueous medium above their CAC. So far, no exact value for the CAC of lipid A could be determined because of extreme experimental difficulties in the low concentration range. A rough estimation on the basis of a limited number of available data for other lipids leads us to propose a value of 10⫺7 –10⫺10 M, depending on the ambient salt concentration. The three-dimensional supramolecular structure of the aggregates depends on the molecular conformation (shape) of the contributing individual molecules, which is determined by their primary chemical structure and ambient conditions. Thus, the molecular conformation can be derived from the aggregate structure. The correlation between different molecular shapes of lipid A or lipid A part structures and the corresponding supramolecular structures is schematically outlined in Figure 3. Briefly, our results on the three-dimensional structure of various lipid A preparations at physiological conditions can be summarized as follows. In pure lipid–water systems and at physiological water concentrations, free lipid A of the LPS of E. coli and S. enterica vs. Minnesota aggregate into nonlamellar cubic structures below Tc . With the beginning of the acyl chain melting process, free lipid A assumes nonlamellar cubic structures. With the completion of chain melting, the cubic structures change into inverted hexagonal HII. In further studies, measurements on the aggregate structure were extended to other lipid A samples, i.e., enterobacterial LPS and lipid A in different salt forms, monophosphoryl lipid A, and lipid A of nonenterobacterial sources like those of R. capsulatus, Rhodopseudomonas viridis, Rhodocyclus gelatinosus, Rhodospirillum fulvum, Campylobacter jejuni, and Chromobacterium violaceum (52,54,55). The measurements were performed exclusively under near physiological conditions to directly correlate the results to data from biological analyses. It was found that the type of counterions (endotoxins in different salt forms) influenced the aggregate structure and that the different nonenterobacterial lipid A samples showed a variety of aggregate structures: lipid A from Rc. gelatinosus adopted HII structures; that of C. jejuni and monophosphoryl lipid A of enterobac-

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terial strains, mixed cubic/lamellar; and those of C. violaceum, R. capsulatus, R. viridis, and R. fulvum, pure lamellar structures (52,54–56). Subsequently, it was shown that these different structural preferences are of importance for the expression of biological activity.

VI. RELATION OF PHYSICOCHEMICAL PARAMETERS TO BIOLOGICAL ACTIVITY Assuming that the phase behavior and the three-dimensional structure determined under near physiological conditions and outlined here with emphasis on lipid A reliably describe the structural polymorphism of these substances, there are some striking correlations between the phase transition temperatures of endotoxins as well as of their supramolecular structures and various biological effects. However, since the reversible chain melting transition frequently goes along with a transition between different structures, it is, in many cases, difficult to decide whether the phase states of the acyl chains or the structural behavior of the whole lipid assembly, or a combination of the two, is the governing process. Nevertheless, there are various biological effects that can be correlated with the value of Tc or the state of order of the acyl chains at 37 °C. Here, only one example is given for the induction of monokine secretion by LPS. The strong cation-induced decrease of fluidity of LPS at 37 °C has been correlated to an enhanced monokine secretion (TNFα and IL-1β) of monocytes (57). The increasing order of the hydrocarbon chains brought about by Zn2⫹ could facilitate a stronger bond between LPS and LBP, thus enhancing the transport of LPS to the target membrane. [For a more complete presentation of the influence of the acyl chain fluidity, the reader is referred to a previous review (46).] A most striking correlation was found between the biological activity of lipid A from different bacterial species and their preference to adopt particular supramolecular structures. Lipid A samples that adopted lamellar structures (individual lipid A molecules having a cylindrical conformation) like those from Rb. capsulatus and C. violaceum were completely inactive, those that assumed mixed lamellar/cubic structures (lipid A monomers possessing partly conical conformation) had intermediate activity, and those samples that preferred pure nonlamellar (Q,HII ) structures (conical conformation) were highly active (Fig. 4). In this context also, the consequences of the structural characteristics of the synthetic lipid A analogs and partial structures (see above) for their bioactivity can be understood: all samples with less than six acyl chains are almost inactive (58), which is again apparently correlated with their preference for lamellar structures. As stated earlier, the lipid A phosphate groups, or possibly other negatively charged groups at the lipid A backbone, are of great importance for biological

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Figure 4 Correlation between supramolecular structure or molecular conformation and biological activity of various lipids A at 37 °C (Adapted from Ref. 145.)

activity. Thus, we recently showed that phosphate groups strongly influence the conformation of lipid A and thus its biological activity (59). In these investigations, we could also demonstrate that a lipid A analog with the 1-phosphate substituted by a carboxymethyl group had an IL-6-inducing capacity comparable to that of the native lipid A. A very interesting fact concerning the induction of biological activity was described by Tahri-Jouti and Chaby (60). It was found that LPS (of Bordetella pertussis, E. coli J5, and S. minnesota R595) bound to mouse peritoneal macrophages both by specific and nonspecific interactions. The nonspecific interactions most probably occurred as a result of the insertion of LPS into the lipid layer of the cellular membrane. These investigations suggest that LPS–LPS associations may also contribute to the nonspecific binding and that the nonspecific binding is less temperature-dependent than specific binding. Above 22 °C (and particularly at 37 °C), specific binding was completely obscured. The authors concluded that the latter observation could be explained by increased nonspecific interactions with the lipid layer resulting from a modification of the fluidity of the cellular membrane at this temperature. Of course, the fluidity of the LPS aggregates is also temperature-dependent in a characteristic way (see above).

VII.

ENDOTOXIN TARGET CELLS

Various types of cells have been found to respond to endotoxin, and it appears that the most important endotoxin target cells belong to the innate immune sys-

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tem. Different types of cells react in typical ways, these reactions including phagocytosis, proliferation, differentiation, mediator production, and apoptosis. A.

Monocytes/Macrophages

The most investigated cell population responding to endotoxin are the cells of the monocyte/macrophage lineage. The prominent role of these cells in endotoxemia is demonstrated by experiments showing that LPS-sensitive macrophages are able to mediate endotoxin shock in LPS-low responder C3H/HeJ mice (61). Upon stimulation by LPS, monocytes/macrophages form and release a variety of cytokines, including interleukin 1 (IL-1), IL-6, IL-8, IL-10, IL-12, and TNFα. In addition to these cytokines, reduced oxygen species [superoxide anion, hydrogen peroxide, hydroxyl radicals, and nitric oxide (NO)] are produced. Bioactive metabolites of arachidonic acid (prostaglandins, thromboxane, and leukotrienes) and of linoleic acid (S-13-hydroxylinoleic acid) are lipidic products of activated monocytes/macrophages in addition to platelet-activating factor (PAF). B. Polymorphonuclear Leukocytes Polymorphonuclear leukocytes (PMN) represent the first host barrier against invading microorganisms. These cells phagocytose and kill bacteria and take up bacterial fragments, reactions that are enhanced by LPS. PMN possess enzymes (acyloxyacyl hydrolase and phosphatases) to degrade LPS and lipid A to nontoxic partial structures (62,63). Furthermore, PMN contain and release bactericidal permeability-increasing protein (BPI) (64,65). This cationic 55 kDa protein binds LPS and neutralizes its bioactivity. Activated PMN contribute to inflammatory reactions by causing damage to endothelial cells and by destroying the blood vessel lining. After penetrating the vessel wall into the tissue, PMN may contribute substantially to local inflammatory reactions. C. B and T Lymphocytes B-lymphocytes of murine origin (but not human B-lymphocytes) are known to proliferate, differentiate, and secrete immunoglobulins in response to stimulation by LPS (66). Polyclonally induced, the resulting immunoglobulins may contribute to an early defense against invading microorganisms by providing antibodies against numerous bacterial specificities. Stimulation of T lymphocytes by LPS is a less known phenomenon, which is now reliably established. Human T cells (CD4- as well as CD8-positive ones) proliferate after LPS stimulation and produce Th1-type lymphokines (9,10). The activation of T cells by LPS was shown to be monocyte-dependent, to require direct cell-to-cell contact and B7-CD28 recognition, but to be MHC-unrestricted

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(see Fig. 5). Furthermore, the presence of IL-12 is obligatory. In addition to monocytes, it has now become evident that CD34⫹ hematopoietic stem cells are also essential for activation of T cells by LPS. Induction of cytokine production and proliferation of T lymphocytes by LPS needs a low number of CD34⫹ stem cells (67). The mechanism, however, by which these CD34⫹ stem cells exert their accessory cell function has to be clarified (see Fig. 5). Murine T cells have also been reported to proliferate in vitro in response to LPS (68,69). In addition, LPSstimulated CD8⫹ /CD4⫺ murine T-lymphocytes are described to suppress the humoral immune response to bacterial polysaccharides (70). Based on these findings, one may assume that indeed T lymphocytes are of great relevance during infection with gram-negative bacteria. D.

Vascular Cells and Epithelial Cells

In addition to immune-competent cells, vascular cells (endothelial cells, smooth muscle cells) and epithelial cells also take part in inflammatory reactions of the host during infection (71,72). It is now well established that activation of these cells by LPS and various other stimuli results in the production of IL-1, IL-6, and/or IL-8 but also of other mediators such as PGI2, NO, PAF, and/or interferons (7,26,73). In addition, the expression of adhesion molecules is a further result of

Figure 5 Schematic representation of cellular and humoral parameters involved in activation of T lymphocytes by LPS.

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the activation of these cells during infection and inflammation. The stimulation by cytokines like IL-1 or TNFα is most effective, but LPS alone also has a profound stimulatory capacity (7). It was found that during infection with gram-negative bacteria one of the most fatal reactions of endotoxin in mice is the induction of apoptosis in the endothelial cells of the colon (74). This ultimately fatal reaction is largely mediated by TNFα, but direct cytotoxic activity of endotoxin in this system was also observed.

VIII. LPS-BINDING PROTEINS/RECEPTORS AND POSSIBLE ACTIVATION MECHANISMS Details of the mechanisms of the interaction of endotoxin with host cell membranes finally leading to their activation are only partly understood. Nevertheless, a number of LPS receptors/LPS-binding membrane proteins have been described. CD14 has unequivocally been shown to represent a prominent LPS-binding structure on monocytes/macrophages. Interaction of LPS with CD14 is necessary for specific binding of LPS and activation of human monocytes or murine macrophages (17,75). Binding of LPS to CD14 is facilitated by the LPS-binding protein LBP (17,24,76), which forms a complex with LPS. Therefore, LBP appears to be responsible for the fact that only minute amounts of LPS are necessary to stimulate monocytes/macrophages. The importance of LBP for the pathophysiological reaction during LPS-induced and bacteria-induced shock is supported by experiments involving mice whose LBP gene has been destroyed. Using these mice it was found that LBP is required for the induction of inflammatory reactions in response to LPS as well as gram-negative bacteria (77). Similarly, CD14 ‘‘knock-out’’ mice were found to be resistant to lethal shock induced by either live gram-negative bacteria or isolated LPS, demonstrating the essential role of CD14 in endotoxicity in vivo (78). In addition to membrane-bound CD14, a soluble form of CD14 (sCD14) has been described (79). sCD14 is present in normal serum, and its level is enhanced during acute-phase reaction. It is either released by activated cells or enzymatically cleaved from the membrane of CD14-expressing cells (80). sCD14 is of importance for the LPS-mediated activation of CD14-negative cells such as endothelial, epithelial, or smooth muscle cells (26). These cells can be stimulated by LPS only in the presence of serum, and it has been shown that the serum proteins that mediate the activation are LBP and sCD14. LPS forms complexes with sCD14, and complex formation is again enhanced by LBP. Binding of sCD14 to nonmyeloid cells was observed in the presence of LPS and LBP (81). It is therefore likely that endothelial cells and smooth muscle cells carry a surface receptor for LPS–sCD14 complexes (26). However, sCD14 may also express

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stimulatory activity by itself, i.e., in the absence of LPS, as has recently been shown by induction of TNFα in human monocytes or monocytic cell lines (82). Both serum proteins LBP and CD14, which in physiological concentrations enhance the activation of cells by LPS, have been shown to attenuate cell activation when present in high concentration (83–85). CD18 represents another LPS-binding membrane protein that is present on leukocytes and that recognizes E. coli bacteria via LPS (86). It is noncovalently linked to either CD11a (LFA-1), CD11b (iC3b receptor), or CD11c (87). It has been demonstrated that CD11c/CD18 functions as a transmembrane signaling receptor for LPS in the absence of CD14 (20). However, according to most recent findings, the transmembrane domain of CD11c/CD18 is not involved in signal transduction (21). Membrane-bound mCD14 is GPI-anchored to the membrane (88) and is thus lacking a transmembrane domain. Therefore, this protein is not capable of transmitting a signal to the cellular interior, and it has been assumed that in addition to a receptor-mediated activation the insertion of endotoxin molecules into the lipid matrix could be a prerequisite for the activation of host cells (89,90). This intercalation could be achieved in various ways: (1) via direct intercalation by hydrophobic interaction of a priori existing endotoxin monomers that are present in sufficient numbers for those chemotypes with longer sugar chains (91) or (2) via the intercalation of monomers or smaller endotoxin aggregates that are produced by the disaggregating properties of LBP and transported to the membrane through a complex interplay of sCD14, mCD14, and LBP (92,93). One further point deserves attention. LBP, initially defined as lipopolysaccharide-binding protein, has recently been shown not to be LPS-specific but rather to interact with and transport other negatively charged lipids. Thus, LBP seems to be a lipid transfer protein in a more general sense (27,92). The mere intercalation of endotoxin molecules into the lipid matrix would, however, not be sufficient for activation, and two basically different fates for the molecule are conceivable. One possible mechanism is the internalization of endotoxin, which is described in several investigations (28,94,95) and could be associated either with cell activation or with clearance and detoxification. The other mechanism constitutes the direct or CD14-mediated interaction of endotoxin with other membraneassociated or membrane-spanning proteins (96,97), and several candidates have been proposed. One of these is an LPS-binding protein found on human monocytes and endothelial cells that has recently been characterized to represent CD55 (98). This protein binds LPS and lipid A only in the presence of soluble CD14 and LBP. Furthermore, chemical cross-linking experiments revealed an association of sCD14/LPS complexes with a 216 kDa membrane protein (99). Other candidates include a purine receptor (100) and ion channels (27,101). Very recently, TLR2, a member of the family of Toll-like receptors, has been shown to be essential in LPS-mediated activation of cells (19,102). The

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Figure 6 Proposed model of cell activation by endotoxin. Endotoxin molecules bind (secondary to an interaction with surface molecules such as mCD14 and TLR2) via their lipid A component to a membrane-spanning signal-transducing molecule, perhaps an ion channel. Binding is facilitated via hydrogen bonding. This requires the existence of hydroxyl or other H-bonding groups in the lipid A part. A further prerequisite for activation is a particular conformation of lipid A.

association of LPS with TLR2 is augmented by LBP and dependent on CD14. It appears that TLR2 is responsible for transmitting an LPS-initiated signal to the cellular interior, resulting in NF-κB activation and the production of endogenous mediators. The correlation between the conformation of endotoxin molecules and their bioactivity is in favor of the latter mode, the existence of a transmembrane signal-transducing protein that is triggered by binding of endotoxin molecules. The triggering signal requires a particular conformation of the lipid A component of endotoxin (Fig. 6). Thus, only those endotoxins will be active that possess a lipid A portion leading, in the isolated form, to nonlamellar aggregate structures. We furthermore postulate that for binding to the signaling protein, the existence of a sufficient number of hydroxy fatty acids in lipid A is necessary, allowing the formation of hydrogen bonds. This conformational concept would readily explain the antagonistic action of biologically inactive endotoxins. In these cases, the binding sites of the transmembrane protein would be occupied by the inactive molecules, thus inhibiting the interaction of the active structures. The fact that compound 406 (synthetic lipid A precursor Ia) and E5531 (synthetic compound related to lipid A of R. capsulatus) express antagonistic activity in humans, but the former not in mice and hamsters and the latter not in hamsters (37,103), could be explained in this model by assuming that the conformation and/or the binding

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sites of the membrane-spanning signaling molecule or other components of the signaling cascade vary in the different systems. If the transmembrane protein were an ion channel, the interaction of endotoxin with the channel protein could lead to mismatches in intracellular ion concentrations, provoking the activation of the subsequent signaling cascade. In our model, the above-mentioned types of endotoxin intercalation should express different efficiencies. Thus, the direct intercalation of monomers and the LBP-mediated process will lead to a random intercalation in the lipid matrix, whereas the mCD14-mediated process will associate endotoxin directly to the signaling protein, assuming that mCD14 is located in the direct vicinity of the signal transducer. This asumption is backed by the observation that mCD14mediated activation can be blocked by anti-CD14 antibodies. At high endotoxin concentration the blockade by anti-CD14 antibodies can be overcome (104), and obviously in that case the CD14-independent activation pathway is operative.

IX.

ENDOTOXIN EFFECTS IN VIVO EXEMPLIFIED FOR LUNG TISSUE

Endotoxins exert a wide variety of effects in vivo. As an example and to illustrate the proinflammatory properties of LPS in vivo, we focus here on the lung and discuss the pulmonary consequences of systemic or airborne LPS exposure. This topic is of utmost clinical relevance, since endotoxemia is an important risk factor for the development of septic shock and the acute respiratory distress syndrome (ARDS) (105), while airborne LPS is involved in the development of byssinosis (farmer’s lung). An extensive overview on endotoxin and the lungs appeared in 1994 (106). Here we reiterate only the most important principles and discuss some more recent aspects, with emphasis on the early effects of LPS. A.

Distribution of LPS in Lung Tissue

Several studies have traced the fate of intravenously applied endotoxin in vivo. Most of the circulating LPS is bound to high-density lipoprotein (107) and is removed by the liver (108,109) and the lungs (only about 2%) (109). It should be noted, however, that the specific activity expressed in micrograms per gram of tissue is comparable in liver and lung (108). Only 1 h after intravenous injection of LPS into rats, LPS is found inside some (3%) alveolar macrophages (110), indicating that LPS can rapidly penetrate through the lung tissue. After 2.5 h, 20%, and after 3 days all alveolar macrophages are positive for LPS (110). In line with this, in isolated mouse lungs perfused with LPS (50 ng/mL) for 1 h, the LPS concentration in the hypophase is about 1.5 ng/mL, i.e., 3% (von Bethmann and Uhlig, unpublished data). Moreover, in the alveolar space of LPS-

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treated rats, dense complexes similar to those obtained by mixing pulmonary surfactant with LPS in vitro (111) are found (112). After 18 h of injection, LPS can also be found in pulmonary endothelial and pulmonary epithelial cells (110). These findings indicate that diffusion or distribution of LPS in lung tissue occurs, but only to a limited extent. Thus, intravenous administration of LPS results in much lower LPS, and hence, much lower TNFα, levels in the alveolar space than are attained by giving the same dose intratracheally (113,114). This conclusion is further corroborated by the observation that the pulmonary sequestration of leukocytes is more pronounced when LPS is administered via the airways compared to systemic application (113,115). B. Pulmonary Sequestration of Leukocytes Even under normal conditions the lung harbors a large pool of marginated leukocytes. These leukocytes, however, accumulate largely due to mechanical forces owing to the fact that leukocytes are larger than capillaries. Hence, during their transit through the lungs, the leukocytes are more retarded than red blood cells (116). Thus, the margination of leukocytes differs from the accumulation in inflamed lungs in terms of both mechanism and location. A hallmark of inflamed tissue is the presence of leukocytes, and this also holds true for lungs from ARDS patients and from LPS-treated animals. In lung tissue from LPS-treated animals, predominantly neutrophilic granulocytes are found. The process of pulmonary leukocyte sequestration starts 15 min after injection of endotoxin and has been impressively visualized by the use of 111In and external scintigraphy (117). This process depends on the expression of adhesion molecules on leukocytes and endothelial cells as well as on the topical release of chemokines (116), which finally leads to invasion of neutrophils into the lung tissue. The expression of adhesion molecules probably results from a direct (CD14-mediated) interaction of LPS with leukocytes and endothelial cells, since these responses are also observed in cell culture (118). In addition, LPS is a potent stimulus for the release of chemokines from a wide variety of cells (119). The fact that intratracheal administration of LPS causes a stronger pulmonary inflammation than intravenous application (113,115) provides evidence that this response relies on the release of chemokines from parenchymal lung cells. C. Consequences of Pulmonary Leukocyte Sequestration: Edema Pulmonary sequestration of neutrophils is not harmful to the lungs per se (120) but may become so if the sequestered leukocytes are activated. Activated neutrophils can release a broad spectrum of deleterious agents such as reactive oxygen species, proteases, and proinflammatory mediators. The major consequence of release of this injurious cocktail in lungs from LPS-treated animals appears to

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be pulmonary edema. Accordingly, four different strategies have been used to ameliorate LPS-induced pulmonary edema: (1) prevention of pulmonary neutrophil sequestration, e.g., by blocking chemotactic factors such as LTB4 (121) or MIP-1α (122); (2) antioxidant strategies such as treatment with superoxide dismutase (123) or N-acetylcysteine (124); (3) inhibition of proteases such as elastase (125); and (4) blocking of inflammatory mediators derived from neutrophils such as platelet-activating factor (126). However, the role of neutrophils in the induction of edema in endotoxic animals is not exclusive, since neutropenic animals also develop edema in response to LPS (127). Moreover, preventing pulmonary neutrophil sequestration with ICAM-1 antisense oligonucleotides had no effect on the development of pulmonary edema (128). In line with this, edema formation occurs in neutropenic ARDS patients (129,130). Obviously, cells different from neutrophils are also capable of releasing edematogenous substances. For example, reactive oxygen species may be derived from endothelial xanthine oxidase (131,132), and PAF is also formed in the lungs of neutropenic rats (126). Alternatively, LPS could damage pulmonary vascular endothelial cells directly (133), probably by inducing apoptosis. However, since components of the extracellular matrix as well as integrins prevent this response (134,135), the direct effect of LPS on endothelial cells may be of only limited importance in vivo. It has been shown that LPS causes TNFα- and ceramide-dependent apoptosis of endothelial cells in vivo (74). Whether this mechanism will result in altered vascular permeability is presently unknown. Two proinflammatory mediators that are found in lungs from LPS-treated animals are TNFα and IL-1. Applied in high doses, both cytokines cause pulmonary neutrophil sequestration and pulmonary edema (136,137). Whether these two cytokines contribute to the LPS-induced lung edema has only rarely been investigated. In LPS-treated animals, TNFα antibodies were ineffective in preventing edema in mice (138). If LPS is instilled in the airways, then edema formation is reduced in animals pretreated with IL-1 receptor antagonist (139,140). Thus, the role for both cytokines in LPS-induced edema formation needs further clarification. The most consistent results have been obtained with PAF antagonists in that all PAF antagonists studied so far have alleviated LPS-induced pulmonary edema. D.

Bronchoconstriction

Many studies relevant to bronchoconstriction have been performed in sheep or pigs. Both species are sensitive to endotoxin (5 µg/kg) and are large enough to allow continuous measurements of physiological lung parameters in whole animals. In both species, intravenous administration of LPS causes bronchoconstriction that is maximal after 60 min and is mediated by thromboxane (141–143).

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This response can be reproduced in perfused rat lungs (144) and does not depend on the presence of neutrophils. In keeping with this, lungs from neutropenic rats still produce thromboxane when challenged with LPS (145). The formation of thromboxane depends on the induction of cyclooxygenase-2 (146). This is in contrast to the PAF-induced bronchoconstriction, which also depends on thromboxane (147) but where PGH2 is supplied by cyclooxygenase-1 (146). The source of thromboxane in lungs is unclear. However, since incubation of isolated pneumocytes with LPS or PAF does not enhance thromboxane synthesis, it seems likely that the thromboxane production in the whole organ depends on the interaction of at least two different cell types (148). A similar phenomenon is known for leukotriene synthesis from endothelial and epithelial cells that lack lipoxygenase and that receive their substrate leukotriene A4 from neutrophils (149,150). E.

Pulmonary Hypertension

In the first phase, i.e., during the first 2 h, the vascular response in LPS-exposed animals follows the airway response and is also caused by thromboxane (141). However, in isolated blood-free lungs exposed to LPS, the formed thromboxane causes only bronchoconstriction and not vasoconstriction (144,146). There are a number of possible explanations for this discrepancy between observations made in whole animals and perfused lungs. 1. The thromboxane that is responsible for vasoconstriction in vivo is derived from blood cells or, alternatively, blood cells may inactivate a vasodilator that is released simultaneously to thromboxane. 2. Vascular obstruction in vivo may occur by thrombosis. 3. In the whole animal, leukocytes and platelets may plug the vessels. The role of cyclooxygenase-1 versus cyclooxygenase-2 for the LPS-induced pulmonary hypertension has not yet been clarified. The response of the pulmonary vasculature to LPS is biphasic. The first, thromboxane-dependent, peak subsides after 2 h, when a secondary increase in vascular resistance sets in. The nature of this second phase has been unknown for a long time, but recent evidence suggests that it is mediated by endothelin. Endothelin is a powerful spasmogen for airway and vascular smooth muscles (151), and the endothelin receptor antagonist bosentan largely prevented the second, but not the first, increase in pulmonary vascular resistance (152). Another important alteration in the pulmonary vasculature in septic animals and patients is the altered vascular reactivity. Of particular interest is the diminished hypoxic vasoconstriction that is observed in LPS-treated lungs (153,154). Hypoxic vasoconstriction is an important regulatory mechanism in healthy lungs as it redirects blood away from nonventilated lung segments. Therefore, the loss of hypoxic vasoconstriction probably contributes to the mismatch in ventilation/

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perfusion seen in LPS-treated animals as well as in humans with ARDS. The mechanism of this diminished reactivity is still unclear. F. Alterations in Surfactant Homeostasis Pulmonary surfactant is a unique product of epithelial alveolar type II cells. Surfactant consists of phospholipids (90%) and proteins (10%). Its major function is to reduce the surface tension inside the alveoli, but it also has immunomodulating functions (155). Deterioration of the pulmonary surfactant system results in decreased lung compliance and is probably a critical pathogenic factor for the development of hypoxemia in LPS-treated animals and in ARDS patients (156). Consequently, various (though not all) studies have shown that treatment of ARDS patients with surfactant is beneficial. There are several interactions between LPS and surfactant: (1) LPS directly and indirectly inactivates surfactant function, (2) LPS disturbs surfactant homeostasis, and (3) surfactant itself affects LPS-induced immune responses. These are discussed below. 1. Mixing of surfactant with gram-negative bacteria (157) or LPS (158), but not with gram-positive bacteria (158), increases surface tension. This is probably due to complex formation between surfactant and LPS (111). Similar complexes can be observed in the alveolar space from LPS-treated rats (112). These complexes may be of functional relevance, because in vitro (158) as well as in the whole organ (112) LPS reduces the hysteresis area of surfactant, indicating that the surfactant becomes less compressible. The biochemical properties that are responsible for complex formation between endotoxin and surfactant have not been defined yet, but it is known that LPS interacts with surfactant proteins SP-A (159) and SP-D (160). LPS may also indirectly lead to inactivation of surfactant by inducing alveolar edema. Since blood proteins such as fibrin inhibit surfactant function (161), alveolar pulmonary edema will also compromise surfactant function. 2. Our view of the effects of LPS on surfactant homeostasis is summarized in Figure 7. As early as 2 h after LPS treatment, giant (surfactant-storing) lamellar bodies accumulate in type II cells (112,144). These giant lamellar bodies probably originate from fusion of several smaller lamellar bodies as a result of impaired surfactant secretion (112). Thus, in the alveolar space there is less phospholipid (112,162,163). In addition, alterations in the biochemical and structural composition of surfactant in the alveolar space are observed. Biochemically, there is a decrease in phosphatidylcholine and a concomitant increase in sphingomyelin (162,163). Structurally, there is loss of tubular myelin, which is important for the formation of the bioactive surfactant monolayer, and more multilamellar vesicles are present, which represent used-up surfactant (112,164). These alterations that occur within 2 h after LPS treatment continue to worsen for at least the next 10 h and may finally lead to a decrease in lung compliance (164). Because until

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recently edema proteins were thought to play the major role in surfactant inactivation in endotoxic lungs (156,161), it is noteworthy that these changes in surfactant homeostasis occur before and are independent of edema formation (112,144,164). The relative importance of each of these two mechanisms is not known, but obviously they could occur together. Finally, it should be noted that similar structural and biochemical changes, i.e., giant lamellar bodies and loss of tubular myelin, have also been reported after ozone treatment (165–167). The mechanism of the deterioration of surfactant homeostasis as described above is not known. However, some studies have attempted to characterize the effects of LPS on alveolar type II cells. Incubation of type II cells with 10–200 µg/mL LPS results in unspecific high affinity binding of LPS that leads to increased microviscosity of cell membranes (168). LPS (500 µg/mL) first accumulates in the microvilli (25 min) and later in the cytoplasm (40 min), mitochondria (90 min), and nucleus (120 min) (169). Treatment of type II cells with LPS suppressed [1–10 µg/mL LPS (170)] or stimulated [10–200 µg/mL (168,171)] surfactant synthesis as assessed by the altered incorporation of radiolabeled choline. The former condition with the lower LPS concentration is likely to be more relevant for the in vivo situation, since type II cells from rats treated with a shock dose of LPS in vivo also showed diminished incorporation of choline (172) and no alterations in membrane fluidity (173). 3. So far, the effects of LPS on surfactant have been discussed; however, this interaction is mutual. Thus, surfactant inhibits endotoxin-stimulated cytokine secretion from alveolar macrophages (174). The surfactant proteins SP-A and SP-D both bind and interact with LPS and may therefore represent a first line of defense against bacterial infection of the lower airways. SP-A recognizes the

Figure 7 Schematic representation of the effects of LPS on surfactant homeostasis. 1. In alveolar type II cells (T-II), vesicles (V) that are derived either from the Golgi body or from reuptake of surfactant from lamellar bodies (LB). In lungs from LPS-treated animals, surfactant synthesis may be reduced. 2. In LPS-exposed lungs, several lamellar bodies fuse to form giant lamellar bodies (GLB). 3. Surfactant is secreted as lamellar body–like surfactant (not shown). Surfactant secretion is decreased by LPS. 4. Lamellar body–like surfactant is transformed into tubular myelin (TM), the precursor for the surface-active monolayer (ML). In the alveolar space of LPS-treated animals, less tubular myelin is found. Complexing of surfactant with LPS may also decrease surfactant bioactivity. 5. Used-up surfactant is transformed into unilamellar (not shown) and multilamellar vesicles (MV). The latter accumulate in the alveolar space of LPS-treated animals. 6. Multilamellar vesicles are either removed by alveolar macrophages (AM) or recycled by reuptake into type II cells. 7. Edema fluid may additionally reduce surfactant activity. For further details see text.

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lipid A region of LPS (159). SP-D can bind to various forms of LPS, but gross agglutination is achieved only with R-form LPS (160). The interaction between SP-D and LPS has been postulated to involve terminal glucose and/or heptose residues of the LPS core region (160). In a number of studies, surfactant or surfactant proteins have been shown to modulate LPS-induced cytokine secretion under various conditions. However, at present the results are partly contradictory, and the reader is referred to two reviews on this topic (175,176).

X. FINAL REMARKS In earlier decades, endotoxin fascinated scientists and clinicians because of its suspected association with severe infections and gram-negative sepsis. The desire to understand the molecular basis of endotoxin activity was thus disease-driven and has ultimately led to the characterization of the toxic region of LPS, the elucidation of its primary chemical structure, the delineation of biosynthetic pathways, the identification of host target cells as well as humoral and cellular receptors, the description of signal transduction cascades associated with the production of proinflammatory mediators, and understanding of the mode of action of mediators and endotoxins causing organ dysfunction or failure. In these studies, however, the physics of endotoxin were largely neglected, and it was only recently that the conformational and membrane-forming properties of LPS have been addressed. Despite considerable progress in this field, much remains to be learned. As lipid A has not yet been crystallized, the models proposed here for its molecular conformation await further proof. Also, the parameters governing the membrane-forming properties of LPS are not fully understood in submolecular terms. It is our conviction that only a deep knowledge of the physicochemical properties of LPS will allow the precise definition and understanding of the interaction of LPS with its humoral and cellular receptors and thus the initial events in endotoxemia. The characterization of the first steps in endotoxin–host interaction, however, is a prerequisite for the rational development of preventive or therapeutic strategies in order to control endotoxemia and endotoxin-based diseases. In the years to come, therefore, the circle in endotoxin research is likely to close: Interest will focus again on the role of endotoxin in diseases, this time, however, on scientifically firm grounds.

ACKNOWLEDGMENTS Part of the studies reported in this review were supported by the Deutsche Forschungsgemeinschaft [SFB 367, projects B8 (U.S.), C5 (A.J.U.), and B2 (E.Th.R.); SFB 470, projects B4 (E.Th.R.) and B5 (U.S.)], by the Federal Minister

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of Education, Science, Research, and Technology [BMBF grant No. 01KI9471/ A6 (U.S., E.Th.R.) and 01KI9471/A1 (A.J.U.)], and by the Fonds der Chemischen Industrie (E.Th.R.).

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14 Effect of Cholestasis on Biomembranes Su¨krettin Gu¨ldu¨tuna and Ulrich Leuschner Johann Wolfgang Goethe University, Frankfurt, Germany

I. PATHOPHYSIOLOGY OF CHOLESTASIS The term ‘‘cholestasis’’ means the stasis of bile and is derived from the histological appearance of bile plugs within the lumen of the bile canaliculus. Ultrastructural findings characteristically demonstrate dilatation of the canalicular space, loss of microvilli on the luminal membrane, thickening of the microfilaments in the pericanalicular space, and adjacent accumulation of vesicular structures, many of which are filled with lipid and bile pigment. Characteristically, substances normally secreted into bile, such as bilirubin, bile acids, and cholesterol, are elevated in the serum. In addition, enzymes and protein found in bile, such as alkaline phosphatase, 5-nucleotidase, γ-glutamyltranspeptidase, leucinaminopeptidase, and immunoglobulin A, generally are increased in the serum of patients with cholestasis (1,2). Cholestasis results from either failure of bile formation or obstruction to bile flow. Intrahepatic cholestasis results in most instances from the inability of the hepatocyte to form osmotic gradients within the canalicular lumen that generates the forward movement of bile (2). The mechanisms of bile formation are complex, and thus the process may be impaired at many subcellular steps and is still imperfectly understood. The subject has been extensively reviewed elsewhere (3). The formation of osmotic gradients within the small canalicular channel depends on a highly coordinated series of transport phenomena, beginning with membrane transport systems (Na-K-ATPase) on the basolateral membrane that generate the driving forces, that regulate the secondary active transport systems for the translocation of bile acids across the sinusoidal and canalicular mem253

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254 Table 1 Mechanisms of Cholestasis Cholestatic agent Ethinyl estradiol Bile salts

Protoporphyrin Endotoxin Chlorpromazine

Mechanism Altered membrane fluidity; leaky tight junctions Altered membrane fluidity and membrane pumps; increased cytosolic calcium Altered membrane fluidity and membrane pumps Altered membrane fluidity and membrane pumps Altered membrane fluidity and membrane pumps; impaired membrane turnover

Clinical correlate Jaundice of pregnancy Bile sale toxicity

Protoporphyria Cholestasis of sepsis Drug cholestasis

brane domains. These and other membrane transport systems are maintained within these domains in a highly polarized manner (4). The canalicular membrane appears to be formed by the transcytotic movement of vesicles from the sinusoidal membrane, where lateral diffusion of transport proteins is prevented by the tight junctions that also function as sealing elements that tighten the canalicular lumen between hepatocytes and enable osmotic gradients in bile to be established (4,5). While it remains unclear how the hepatocyte establishes the secretory polarity, cholestatic liver injury may impair this process at many steps. However, the major subcellular alterations may be divided into several categories that are summarized in Table 1 where the mechanisms and possible clinical correlates are summarized.

II.

BILE ACIDS AND THEIR ROLE IN CHOLESTASIS

Bile acids are derived from cholesterol and have a steroid nucleus from which hydroxy groups project into space. One surface of the steroid ring always has more ionizable hydroxy groups and therefore is more hydrophilic than the other (6). Thus each bile salt molecule has a relatively hydrophilic (water-soluble) surface and a hydrophobic (lipid-soluble) surface. Also, bile salts with a greater number of hydroxy groups are more hydrophilic. Conjugation of bile acids to taurine or glycine in the liver, forming sodium salts of the conjugates, markedly increases their water solubility (7,8). With increasing bile salt concentrations, aggregates are formed in which the hydrophobic surfaces of individual molecules face each other, leaving the hydrophilic surfaces facing the aqueous environment. Bile salts also partition into lipid cell membranes and cause membrane-to-micelle

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movement of lipids. The ability of a bile salt to disrupt lipid membranes increases with its concentration and hydrophobicity. Thus, bile salt interactions with lipid cell membranes have both major structural and functional consequences (9–12). The apolar α-dihydroxy bile salts are cytotoxic and induce cholestasis and necrosis of the liver (13–17). In liver perfusion studies with chenodeoxycholate, an α-dihydroxy bile salt, progressive damage to the liver was observed. Following perfusion with 0.01 mmol/L chenodeoxycholic acid (CDC), liver plasma membranes were isolated and inhibition of the Na-K-ATPase was detected (Fig. 1). Na-K-ATPase is localized in the basolateral domain of the plasma membrane. Alterations in the lipid structure could induce inhibition of this enzyme accompanied by cholestasis (18,19). Concentrations of 0.3 mmol/L CDC induced morphological alterations of the plasma membranes (Fig. 2). Electron microscopy revealed bleb formation at the basolateral membrane and destruction of the canalicular domain. Increasing the CDC concentration to 0.5 mmol/L led to damage of cell organelles, e.g., swelling of mitochondria (Fig. 3). Ursodeoxycholate,

Figure 1 Activity of Na-K-ATPase in isolated liver plasma membranes after perfusion of the liver with different concentrations of CDC for 90 min. (*) p ⬍ 0.05; (**) p ⬍ 0.01 versus control.

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Figure 2 Morphological alterations of the plasma membranes detected by electron microscopy after perfusion with 0.3 mM CDC for 90 min. Notice the destruction at the canalicular domain (marked with arrow).

Figure 3 Morphological alterations of the liver detected by electron microscopy after perfusion with 0.5 mM CDC for 90 min. Notice the swelling of mitochondria.

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a more polar bile salt, has no toxic effects on the liver, whereas, in contrast, UDC has beneficial effects in chronic cholestatic liver diseases (20–22). Since the initial effects of bile salt toxicity were observed on hepatocyte plasma membranes, investigations were made into alterations in the plasma membrane.

III. MECHANISM OF LIPOSOMAL DAMAGE BY BILE SALTS Large unilamellar vesicles (LUVs) are an excellent model for studying membrane disturbances induced by asymmetrical bile salt binding. Moreover, with LUVs the transitions of bile salt lipid mixed bilayers to mixed micelles can be easily correlated with the binding data. Cabral et al. (23) determined in 13C animal studies very low rates for transbilayer movement (flip-flop) of several hours at low concentrations and a depronated state of bile salts. The results of the study indicate that bile salts are absorbed exclusively by the outer membrane monolayer. This initial membrane binding site of bile salts can be described by the interaction of bile salts and membrane lipids. The influence of bile salts on membrane lipid structure is due to their binding to the membrane and depends on the hydrophobicity of the bile salts. The greater the hydrophobicity, the stronger the binding to the membrane. Schubert et al. (11,12) investigated the binding equilibria of bile salts to model membranes (LUV) by means of Scatchard plots. The highest association constant (Ka) is found with CDC, followed by UDC and cholate (C). The association constant depends on the lipid composition of the membrane. Increasing the membrane cholesterol content decreases the association constant. Thus cholesterol hampered the binding of bile salts to phospholipid membranes. The number of lipid molecules that are associated with one bile salt molecule at low bile salt concentrations could also be calculated by means of Scatchard plots. Cholate is associated with six lipid molecules, CDC with three, UDC with 16. The number of lipid molecules per bile acid molecule increases with increasing cholesterol content of membrane. After binding to membranes, bile salts pass through the membrane. The transbilayer movement of bile salts has been measured with pyranine, a pH-sensitive fluorescent molecule (24). Passing the membrane, bile salts decrease the pH, e.g., deoxycholate induces a decrease in pH from 7.4 to 6.9. Thus the pK of bile salts is different in the outer and inner leaflets of the membrane. The diffusion time of CDC and DC is very short, t1/2 ⬍ 1 s, whereas that of cholate, a more polar bile salt, is 10–15 times as long. Nuclear magnetic resonance (NMR) studies have shown that the diffusion time of C is more than 50 times longer than that of α-dihydroxy bile salts; conjugated bile salts do not pass through the membrane (23).

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Membrane disturbances start when bile salts in the outer monolayer are forced to share increasingly more lipids involved in bile salt binding. Efflux of carboxyfluorescein (CF) or inulin from vesicles made out of model membranes are standard methods to investigate membrane permeability. CF efflux was induced even by concentrations of 0.05 mmol/L CDC, and similar results were obtained for inulin release (Fig. 4). The CF efflux depends on the cholesterol content of the membrane. In order to induce the same CF efflux in membranes with increased cholesterol content, the CDC concentration has to be increased. However, cholesterol contents of more than 30% have no additional stabilizing effect against CDC-induced permeability changes. Concentrations of 50% cholesterol destabilize the membrane, and the permeability is increased compared to LUVs without cholesterol. Thus, the stabilizing effect of cholesterol is limited, and more than 40% destabilizes the membranes. Membrane defects and changes in membrane elasticity induced by bile salts can serve as starting points for budding of the membrane or fusion to multilamellar aggregates. In vesicles with increasing amounts of bile salts, therefore, hydro-

Figure 4 Carboxyfluorescein efflux of large unilamellar vesicles with different cholesterol concentrations induced by CDC; incubation time 60 s.

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Figure 5 Membrane diffusion of bile salts showing the induction of pore formation in the membrane. (a) Insertion of CDC into the outer leaflet of a model membrane, increasing the polarity and decreasing the order of membrane lipid. (b) Diffusion of CDC to the inner leaflet of a model membrane.

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philic interactions of bile salts with lipids gradually are replaced by contact of the hydrophobic hemisphere of the bile salt steroid backbone with the hydrocarbon part of the phospholipids. Upon entering the outer leaflet of the membrane, bile salts increase the polarity and decrease the order by forming polar compartments in the outer leaflet. The increase in the energetic potential ∆G between the outer and inner leaflets of the membrane increases the diffusion potential. This could induce a redistribution of bile salt molecules, leading to the formation of transient pores in the membrane, which increases the permeability of the membrane (Fig. 5). Increasing the bile salt concentration of α-dihydroxy bile salts to the critical micellar concentration (CMC) induced solubilization of model membranes into bile salt–lipid mixed disk micelles (Fig. 6). Prior to onset of membrane solubilization, vesicle size increased, indicating that the bile salt–lipid ratio increased (9). The membrane solubilization induced by CDC occurs at concentrations of 1.25 mmol/L in 100% egg yolk lecithin vesicles. With increased membrane cholesterol content the CDC concentration has to be increased. UDC, a hydrophilic 3α, 7β-dihydroxy bile salt, induced vesicle fusion above the CMC to multilamellar aggregates larger than 1 µm, probably by forming apolar clusters of UDC on the surface of the vesicles similar to cholesterol. In carboxyfluorescein

Figure 6 Effect of CDC (1,2) and UDC (3,4) on the size (measured by laser light scattering) of large unilamellar vesicles with different cholesterol contents.

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Figure 7 Release of CF from LUV with 100% EYL. 1, Control; 2, incubation with 0.1 mM UDC; 3, incubation with 0.25 mM UDC and 0.1 mM CDC; 4, incubation with 0.1 mM UDC and 0.1 mM CDC; 5, incubation with 0.1 mM CDC.

efflux measurements, continued addition of UDC induces a protective effect (Fig. 7), whereas the common bile salts with 7α-hydroxy configuration lead to local disorder; these data could be confirmed by EPR studies (24).

IV. EFFECT OF BILE SALTS ON ERYTHROCYTE AND HEPATOCYTE MEMBRANES Alterations in the molecular structure of membranes can be detected by using sensitive methods such as electron paramagnetic resonance (EPR) spectroscopy. We therefore investigated alterations in the lipid structure of the membrane with this technique. Specimens were labeled with nitroxide-stable radical attached to a stearic acid molecule. The spin labels 16-DSA and 5-DSA provide information on the polarity (aN ) and molecular motion in their environments (Fig. 8). 16DSA reports on the apolar domain, and 5-DSA on the interface of the membrane (24–27) (Fig. 9).

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Figure 8 EPR spectra of (a) 16-DAS and (b) 5-DAS incorporated into erythrocyte membranes. Polarity estimations: aN assesses the water penetration into biological membranes. The ratio h⫺1P/h⫺1H is a sensitive parameter for measuring the relative amounts of spin label molecules in polar and apolar environments.

Figure 9 Localization of 5-DSA and 16-DSA in the bilayer.

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Apolar bile salts increased the polarity of the membrane. Figure 10 shows results for three different plasma membranes: erythrocyte, canalicular, and basolateral liver plasma membranes. Addition of CDC induced an increase in the polarity of the membrane lipid domain. The extent of the change in the polarity depends on the cholesterol concentration of the membrane. In membranes low in cholesterol the polarity increased, whereas membranes with higher cholesterol content were not affected. However, higher concentrations of CDC also induced an increase in the polarity of erythrocyte (ERM) and canalicular liver plasma membranes (cLPM), which have higher cholesterol content than basolateral liver plasma membranes. Thus, membranes with low cholesterol concentration are more sensitive to apolar bile salts; similar data could be obtained with model membranes. Moreover, EPR studies with erythrocyte and hepatocyte membranes have shown that an increase in membrane polarity depends on the hydrophobicity of the bile salt molecule. The greatest increase in membrane polarity is induced by deoxycholate (DC), followed by chenodeoxycholate (Fig. 11). The conjugates had a milder effect on membrane polarity. UDC decreased the polarity in the hydrophobic region of the membrane, whereas its conjugates did not affect the apolar domain (24,28). An increase in membrane polarity is due to a perturbation in membrane structure that permits water to permeate deeper into the membrane

Figure 10 Correlation of cholesterol concentration (mg/mg membrane protein) in different membrane fractions and sensitivity to CDC damage, shown by changes in membrane polarity aN (plotted inversely). Membranes were labeled with 16-DAS. (*) p ⬍ 0.001 versus control.

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Figure 11 Effect of different bile salts (µmol)/mg membrane protein on the polarity of erythrocyte membranes. The polarity (a N) of the apolar domain of the membranes was detected with 16-DAS. *p ⬍ 0.01 versus control.

interior. The increase in membrane polarity caused by toxic bile salts is accompanied by an increase in membrane permeability, as shown by carboxyfluorescein and inulin efflux methods (9,11,12).

V.

THERAPEUTIC EFFECT OF BILE SALTS

Ursodeoxycholate (UDC) has been successfully used in the therapy of liver diseases (20,21). In in vitro and in vivo experiments UDC prevented the damage induced by apolar bile salts (24,29–34). Increased membrane permeability induced by chenodeoxycholate was reduced by simultaneous incubation with UDC (see Fig. 7) (9). In EPR experiments UDC prevented the increase in membrane polarity caused by apolar bile salts (24). Since this protective effect could be observed in both erythrocyte and basolateral liver plasma membranes, this phenomenon is not specific for hepatocyte membranes. A possible mode of protection of UDC and UDC conjugates is polar interaction with CDC. Another possible protective mechanism of UDC and its conjugates is that they are bound to plasma membranes. The interaction of UDC with membrane lipids could be confirmed by experiments with labeled bile salts, by EPR experiments with biomembranes, and by equilibrium binding data of bile salts and model membranes by Scatchard plots (11,12,24).

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The membrane stabilization of UDC appears to be a direct effect on the membrane. In preincubation experiments non-membrane-bound UDC was removed and membranes were then postincubated with CDC, but UDC still prevented the increase in membrane polarity (24). The possible arrangements of bile salt molecules and the membrane bilayers are shown in Figure 12. NMR experiments have shown that α-dihydroxy bile salts are inserted as dimers into the membrane (35), whereas cholate molecules lie flat on the membrane surface (36). In EPR experiments with different lipid labels, UDC induced a decrease of polarity in the apolar part of the membrane and the conjugates induced a decrease of polarity in the membrane interface. These findings indicate that the steroid nucleus of UDC after insertion into the membrane is located in the apolar domain and that of the conjugates, in the interface. The binding site of UDC is similar to that of cholesterol. In some ways UDC mimics the effect of cholesterol on phospholipid membranes, e.g., UDC decreases the polarity of the membranes as does cholesterol. Cholesterol interacts with membranes and affects their thermotropic properties by inserting into the hydrophobic part of the membrane (37,38). This leads to a broadening, reduction, or elimination of the phase transition without altering the phase transition temperature. The thermal behavior of a lipid is an important factor in the permeability, fusion, and protein binding of lipid membranes (39,40). Experiments with differ-

Figure 12 Scheme indicating the different localizations of bile salt molecules in the lipid bilayer. (From Ref. 43.)

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ential scanning calorimetry (DSC) revealed an effect of UDC on the phase transition that was comparable to that of cholesterol (9). Addition of UDC to egg yolk lecithin resulted in phase transition behavior very similar to that of cholesterol (12). Enthalpy was reduced by about 50% when the UDC/phospholipid or cholesterol/phospholipid ratio in the model membranes was as low as 1:5 (Fig. 13). In experiments with dimiristoylphosphatidylcholine, the inhibitory effect of UDC on the enthalpy of the phase transition could be observed, even at concentrations of 0.1% (9). In DSC studies with taurocholate and taurodeoxycholate the transition temperature of DPPC was changed, indicating that micellar structures affect the cooperativity of the lipid. In contrast, under experimental conditions UDC did not change the transition temperature of the lipid. The similar effects of UDC and cholesterol on the phase transition therefore indicate that these substances may interact with lipids in a similar fashion. UDC interacts with membrane lipids, altering the chains, and decreases their motion, as does cholesterol. Cholesterol is incorporated into the membrane with its hydroxy group oriented to the aqueous surface and the aliphatic chain aligned parallel to the acyl chains in the center of the bilayers. The steroid nucleus is positioned

Figure 13 Changes of enthalpy (∆H ) in EYL during the phase transition determined by cooling scans (40 to ⫺40 °C) with different percentages of (1) cholesterol or (2) ursodeoxycholate. Data were correlated with the peak broadening ∆D induced by (3) cholesterol and by (4) ursodeoxycholate.

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Figure 14 Localization of UDC and tauroursodeoxycholate (TUDC) in the plasma membrane as detected by EPR technique.

along the first 10 carbons of the phospholipid chain. EPR studies suggest that the steroid nucleus of UDC is located a few atoms deeper than that of cholesterol, but in principle in the same domain (Fig. 14). In experiments with LUVs, the membrane-protective effect of UDC is inversely correlated to the cholesterol concentration of the vesicles inhibiting the insertion of UDC. Thus, membrane-bound UDC impedes the penetration of cholesterol into the membrane. The similarities between UDC and cholesterol concerning the interaction with phospholipid membranes may be of relevance to the therapeutic effect of UDC. In cholestatic liver diseases, liver plasma membranes (especially canalicular hepatocyte membranes) may change in lipid composition, since during cholestasis the more hydrophobic toxic bile salts seem to accumulate around the bile capillaries (41,42). Since bile secretion in these patients is reduced and the more apolar, toxic bile salts into blood are retained, similar alterations may occur at basolateral hepatocyte membranes. One of the therapeutic effects of UDC could be the maintenance of membrane structure, stability, and function.

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2. Fallon MB, Anderson JM, Boyer JL. Intrahepatic cholestasis. In: Schiff L, Schiff ER, eds. Diseases of the Liver. 7th ed. Philadelphia: Lippincott, 1993:343–361. 3. Sellinger M, Boyer JL. Physiology of bile secretion and cholestasis. In: Popper H, Schaffner F, eds. Progress in Liver Diseases. New York: Grune and Stratton, 1990: 237–259. 4. Hubbard AL, Barr VA, Scott LJ. Hepatocyte surface polarity: In: Arias IM, Boyer JL, Fausto N, Jakoby WB, Schachter DA, Shafritz DA, eds. The Liver. Biology and Pathobiology. 3rd ed. New York: Raven Press, 1994:189–213. 5. Stieger B, Landmann L. Effect of cholestasis on membrane flow and surface polarity in hepatocytes. J Hepatol 1996; 24(suppl 1):128–134. 6. Carey MC, Duane WC. Enterohepatic circulation. In: Arias IM, Boyer JL, Fausto N, Jakoby WB, Schachter DA, Shafritz DA, eds. The Liver. Biology and Pathobiology. 3rd ed. New York: Raven Press, 1994:719–767. 7. Small DM. The physical-chemistry of cholanic acids in the bile acids. Chemistry, physiology and metabolism. In: Nair PP, Kritchevsky D, eds. Chemistry, Vol 1. New York: Plenum Press, 1971:249–356. 8. Armstrong MJ, Carey MC. The hydrophobic-hydrophilic balance of bile salts. Inverse correlation between reverse high performance liquid chromatographic mobilities and micellar cholesterol-solubilizing capacities. J Lipid Res 1982; 23:70–80. 9. Gu¨ldu¨tuna S, Deisinger B, Weiss A, Freisleben H-J, Zimmer G, Sipos P, Leuschner U. UDC stabilizes phospholipid-rich membranes and mimics the effect of cholesterol: investigations on large unilammelar vesicles. Biochim Biophys Acta 1997; 1326:265–274. 10. Gu¨ldu¨tuna S, Zimmer G, Imhof M, Bhatti S, You T, Leuschner U. Molecular aspects of membrane stabilization by ursodeoxycholate. Gastroenterology 1993; 104:1736– 1744. 11. Schubert R, Schmidt KH. Structural changes in vesicle membranes and mixed micelles of various lipid compositions after binding of different bile salts. Biochemistry 1988; 27:8787–8794. 12. Schubert R, Beyer K, Wolburg H, Schmidt KH. Structural changes in membranes of large unilamellar vesicles after binding of sodium cholate. Biochemistry 1986; 25:5263–5269. 13. Greim H, Tru¨lzsch D, Czygan P, Rudick J, Hutterer F, Schaffner F, Popper H. Mechanism of cholestasis. 6. Bile acids in human livers with or without biliary obstruction. Gastroenterology 1972; 63:846–850. 14. Attili AF, Angelico M, Cantafora A, Alvaro D, Capocaccia L. Bile acid induced liver toxicity: relation to the hydrophobic balance of bile acids. Med Hypoth 1986; 19:57–69. 15. Drew R, Priestly BG. Choloretic and cholestatic effects of infused bile salts in the rat. Experientia 1978; 35:809. 16. Dryska H, Chen T, Salen G, Mosbach EH. Toxicity of chenodeoxycholic acid in the rhesus monkey. Gastroenterology 1975; 69:333–337. 17. Miyai K, Price VM, Fisher MM. Bile acid metabolism in mammals. Ultrastructural studies on intrahepatic cholestasis induced by lithocholic and chenodeoxycholic acids in the rat. Lab Invest 1971; 24:292–302. 18. Keeffe EB, Scharschmidt BF, Blankenship NB, Ockner RK. Studies on relationship

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35. Saito H, Sugimoto Y, Tabeta R, Suzuki S, Izumi G, Kodama M, Toyoshima S, Nagata C. Incorporation of bile acids of low concentration into model and biological membranes studied by 2H and 31P NMR. J Biochem (Tokyo) 1983; 94:1877–1887. 36. Ulmius J, Lindblom G, Wennerstro¨m H, Johansson LB, Fontell K, So¨dermann O, Arvidson G. Molecular organization in the liquid-crystalline phases of lecithin-sodium cholate-water systems studied by nuclear magnetic resonance. Biochemistry 1982; 21:1553–1560. 37. Yeagle PL. Cholesterol and the cell membrane. Biochim Biophys Acta 1985; 822: 267–287. 38. El-Sayed MY, Guion TA, Fayer MD. Effect of cholesterol on viscoelastic properties of dipalmitoylphosphatidylcholine multibilayers as measured by laser-induced ultrasonic probe. Biochemistry 1986; 25:4825–4830. 39. Ladbroke BD, William RM, Chapman D. Studies on lecithin-cholesterol-water interactions by DSC and x-ray diffraction. Biochim Biophys Acta 1968; 150:333–340. 40. Demel RA, De Kreuyff B. The function of sterols in membranes. Biochim Biophys Acta 1976; 457:109–132. 41. Okishio T, Nair PP. Studies of bile acids. Some observations on the intracellular localization of major bile acids in rat liver. Biochemistry 1966; 5:3662–3668. 42. Barnwell SG, Lowe PJ, Coleman R. Effect of taurochenodeoxycholate or tauroursodeoxycholate upon biliary outputs of phospholipids and plasma membrane enzymes, and the extent of cell damage, in isolated perfused rat livers. Biochem J 1983; 216: 107–111. 43. Hofmann AF. The Liver. New York: Raven, 1994. 44. Owen JS. Extrahepatic cell membrane lipid abnormalities and cellular dysfunction in liver diseases. Drugs 1990; 40(suppl):73–83.

15 3,5-Diiodothyronine Binds to Subunit Va of Cytochrome c Oxidase Possible Mechanism of Short-Term Effects of Thyroid Hormones Bernhard Kadenbach Philipps-University, Marburg, Germany

Susanne Arnold Children’s Hospital and Harvard Medical School, Boston, Massachusetts

I. THYROID HORMONES AND ENERGY METABOLISM The most obvious effect of thyroid hormones in animals consists of the increase of basal metabolic rate (the ‘‘calorigenic effect’’). The original assumption that thyroid hormones uncouple oxidative phosphorylation of mitochondria (1,2) was later withdrawn (3,4). The stimulation of cell respiration by thyroxin (T4) or its physiologically active derivative 3,3′,5-triiodo-l-thyronine (T3) was explained by turning on the expression of modified enzyme patterns, particularly in mitochondria (5,6) via binding of the hormone to the nuclear thyroid hormone receptor (7). Thyroid hormones have a dual effect on metabolism of vertebrates (for review, see Refs. 8 and 9). Short-term effects occur via direct binding to extranuclear proteins (enzymes), accompanied by their activation, whereas long-term effects occur via binding to nuclear receptors and subsequent stimulation of specific protein expression. The thyroid hormone receptors belong to a large family of zinc finger transcription factors (10,11), which form homo- or heterodimers, allowing complex interactions with numerous genes (12–15). The identity of extranuclear binding proteins is still not clear. Plasma membrane binding sites could mediate hormone transport into the cell but may also influence calcium, amino acid, and sugar uptake, whereas cytosolic binding pro271

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teins (CTBPs) are assumed to transport the thyroid hormones into the nucleus. Mitochondrial binding sites have been repeatedly described (for review, see Ref. 8). Specific binding sites for T3 were identified in the inner mitochondrial membrane by Sterling et al. (16,17) and were later confirmed by others (18,19). An immediate stimulation of the respiration of isolated rat liver mitochondria by T3 was shown (20), but a specific mitochondrial receptor for T3 could not be identified (21). As the result of a growing number of studies, diiodo-l-thyronines (3,5T2 and 3,3′-T2), instead of T3, have been identified as the active hormones that immediately stimulate mitochondrial oxidative capacity and respiration, independent of nuclear transcription, in humans and rats (22–27) as well as in fish (28). The diiodothyronines are formed from T4 or T3 by deiodinases (29,30). A direct stimulation of cytochrome c oxidase activity by diiodothyronines could be shown in rat liver homogenates (31) and for the isolated enzyme (32). Recently one binding site for 3,5-T2 could be identified at subunit Va of cytochrome c oxidase (33). II.

THYROID HORMONES STIMULATE UNCOUPLED MITOCHONDRIAL RESPIRATION

Mitochondria isolated from hypothyroid rats are characterized by decreased state 3 and state 4 respiration rates relative to mitochondria from euthyroid rats, while mitochondria from hyperthyroid rats show elevated state 3 and state 4 respiration rates (34). In contrast, the mitochondrial proton motive force ∆p decreased in the order hypothyroid ⬎ euthyroid ⬎ hyperthyroid (35). This means that mitochondrial energy coupling is highest under hypothyroid and lowest under hyperthyroid conditions. These results, measured on isolated mitochondria of hypo-, eu-, and hyperthyroid rats, were corroborated with isolated rat hepatocytes (36,37). The molecular mechanism of thyroid hormone action, however, remained a matter of discussion. Brand and coworkers (35,36) measured enlarged mitochondrial proton conductance (proton leak) as a consequence of hormone action in vivo and related it to an increase in the ratio of area of the inner membrane to protein mass and to an elevated permeability of the phospholipid bilayer (38). More recently a direct stimulation of cytochrome c oxidase, and of cytochrome c reducing components of the respiratory chain, was found 1 h after injection of 3,5-T2 by ‘‘top-down’’ elasticity analysis of isolated rat liver mitochondria (39). III. THYROID HORMONES STIMULATE EXPRESSION OF UNCOUPLING PROTEINS In addition to their direct effect on mitochondrial respiration and energy coupling, thyroid hormones could also indirectly increase the uncoupled respiration, and

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thus thermogenesis and the basal metabolic rate, by stimulating the expression of uncoupling proteins (UCPs). The uncoupling protein (UCP-1) was originally identified in brown adipose tissue and was shown to dissipate the energy of the mitochondrial proton gradient as heat instead of being used for ATP production (40,41). Recently two further uncoupling proteins were identified. While UCP2 is expressed in most tissues (42), UCP-3 is expressed only in brown adipose tissue and skeletal muscle (43). In rats, T3 stimulates the expression of UCP-1 in brown adipose tissue (44) and that of UCP-3 in skeletal muscle and brown adipose tissue (45,46). Also the expression of UCP-2 was increased after T3 administration to rats in white adipose tissue and skeletal muscle (47) and in heart and skeletal muscle (48).

IV. CONTROL OF CYTOCHROME c OXIDASE BY NUCLEOTIDES The reduction of oxygen to water by cytochrome c oxidase, the terminal enzyme of the mitochondrial respiratory chain, represents the irreversible step of the pathway. Therefore this step is expected to be regulated (49). Mammalian cytochrome c oxidase is composed of three mitochondrial and 10 nuclear coded subunits, three of which (subunits VIa, VIIa, and VIII) occur in tissue-specific isoforms (50). Recently seven high-affinity binding sites for ATP or ADP and three additional only for ADP were identified at the isolated bovine heart enzyme by equilibrium dialysis (51–53). The 10 nucleotide-binding sites were corroborated by 10 tightly bound cholate molecules, which are structurally very similar to ADP, at the enzyme crystals (52,54). One binding site is located at the matrix domain of the transmembranous (54) subunit VIaH (heart-type, expressed only in heart and skeletal muscle). Exchange of bound ADP by ATP decreases the H⫹ /e⫺ stoichiometry of the reconstituted bovine heart enzyme from 1 to 0.5 (55). This mechanism was suggested to participate in thermogenesis of skeletal muscle at rest (49,56). The reconstituted enzymes from bovine liver and kidney revealed H⫹ /e⫺ stoichiometries of 0.5, which were independent of the intraliposomal ATP/ ADP ratios (57). The lower H⫹ /e⫺ ratios of the enzymes from nonskeletal muscle tissues were postulated to participate permanently in mammalian thermogenesis. A further binding site for ATP or ADP is located at the matrix domain of the transmembranous (54) subunit IV. Exchange of bound ADP by ATP at high intramitochondrial ATP/ADP ratios decreases the enzyme activity accompanied by a change from hyperbolic to sigmoidal kinetics (58). Half-maximal decrease of enzyme activity was obtained at an ATP/ADP ratio of 30 in proteoliposomes (59). This allosteric inhibition of respiration at high ATP/ADP ratios, which is independent of the mitochondrial membrane potential (59), represents the ‘‘second mechanism of respiratory control’’ according to the utilization of ATP in eukaryotes, but not in prokaryotes (60). The generally known ‘‘first mechanism of respiratory control’’ is based on

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the mitochondrial proton motive force ∆p (61). The physiological significance of the ‘‘second mechanism of respiratory control’’ by the intramitochondrial ATP/ADP ratio, via binding to subunit IV of cytochrome c oxidase, could be to maintain a low mitochondrial membrane potential ∆Ψ, the major component of ∆p. Control of respiration by ∆p alone would allow ∆Ψ to rise to high levels of 160–200 mV as measured with isolated mitochondria (35). High mitochondrial membrane potentials, however, support the formation of reactive oxygen species such as the superoxide radical anion (62,63). In addition, the unspecific and energetically unproductive proton leak of biological membranes increases exponentially with the membrane potential (64). In fact, in perfused rat hearts, a low membrane potential of about 100–130 mV was measured that was little affected by varying the rate of respiration by a factor of 4 (65). Cytochrome c oxidase could limit the mitochondrial respiration rate only if it represents the rate-limiting step of the respiratory chain. From application of metabolic control analysis (66), a manifold excess of cytochrome c oxidase capacity over the amount required to support the endogenous respiration of isolated mitochondria was concluded (67–69). A limitation in the metabolic control analysis applied to isolated mitochondria, compared to the in vivo situation, however, is the significant alteration of the situation due to the possible loss of essential metabolites and to the absence of the cytosolic substrate and coenzyme environment. Recent metabolic control analysis of respiration in intact cultured cells revealed tight control of cell respiration by cytochrome c oxidase, exceeding its capacity above the endogenous respiratory activity by only 7–25% (70,71). Also in saponin-permeabilized muscle fibers a higher control strength for cytochrome c oxidase was found than in isolated mitochondria, in particular at lower oxygen pressure (72).

V.

EFFECT OF 3,5-DIIODOTHYRONINE ON CYTOCHROME c OXIDASE

Recently the specific binding of [3-125I]5-diiodo-l-thyronine to subunit Va of isolated cytochrome c oxidase from bovine heart was shown. Binding of the hormone completely abolished the allosteric inhibition of activity by high ATP/ADP ratios (33). The specific binding of the hormone to subunit Va was confirmed by the action of a monoclonal antibody against subunit Va, which prevents the effect of 3,5-T2. Figure 1 shows the ascorbate respiration of cytochrome c oxidase from bovine heart, reconstituted into liposomes containing asolectin and 5% cardiolipin in the presence of 5 mM ATP. Titration of ascorbate respiration with increasing concentrations of cytochrome c reveals a sigmoidal curve (control), which was not changed after reconstitution in the presence of a monoclonal antibody against subunit Va (AbVa). Complete inhibition of activity is observed up

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Figure 1 3,5-Diiodo-l-thyronine abolishes the allosteric inhibition of cytochrome c oxidase activity by intraliposomal ATP, and this effect is prevented by a monoclonal antibody against subunit Va. Cytochrome c oxidase from bovine heart (3 µM) was reconstituted into liposomes containing asolectin and 5% cardiolipin in 10 mM K-HEPES, pH 7.4, 40 mM KCl in the presence of 5 mM ATP, as previously described (33). The ascorbate respiration of proteoliposomes was measured polarographically at the indicated concentrations of cytochrome c [TN ⫽ turnover number (mol 1/4 O2) (Mol heme aa3)⫺1 s⫺1]. Reconstitution of the enzyme was performed without further additions (control) or in the presence of 10⫺6M 3,5-T2 (T2) and 6 µM of monoclonal antibodies (73) against subunit IV (AbIV) or subunit Va (AbVa), as indicated.

to 2 µM cytochrome c, followed by a slight increase. Reconstitution in the presence of 10⫺6 M 3,5-T2 completely abolished the ATP inhibition and resulted in a higher activity curve (T2) corresponding to that obtained after reconstitution of the enzyme in the presence of 5 mM ADP instead of ATP (not shown). This activity is not affected by a monoclonal antibody against subunit IV (T2 ⫹ AbIV). The monoclonal antibody against subunit IV alone, however, completely reversed the allosteric ATP inhibition (58). 3,5-T2 has no effect when the enzyme is reconstituted in the presence of 3,5-T2 and the monoclonal antibody against subunit Va (T2 ⫹ AbVa).

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Reversal of ATP inhibition by 3,5-T2 at high ATP/ADP ratios would not necessarily increase mitochondrial respiration, owing to the control of proton pumps by ∆p. Stimulation of the rate of electron transfer by 3,5-T2 also increases proton pumping and thus ∆p if the H⫹ /e⫺ stoichiometry remains constant. Increased ∆p would decrease the electron transfer activity of proton pumps if the rate of ATP utilization remained unchanged. We found, however, a second effect of 3,5-T2 on reconstituted cytochrome c oxidase from bovine heart, as shown in Figure 2. In addition to reversing ATP inhibition, the hormone also decreases the respiratory control ratio based on ∆p (RCR) in the presence of intraliposomal ATP but not intraliposomal ADP. In other words, under conditions of low energy utilization (high ATP/ADP ratios), or at rest, the hormone induces a proton leak

Figure 2 3,5-Diiodothyronine decreases the respiratory control ratio of reconstituted cytochrome c oxidase from bovine heart. Cytochrome c oxidase was reconstituted into liposomes containing asolectin and 5% cardiolipin in 10 mM K-HEPES, pH 7.4, 40 mM KCl, in the presence of 5 mM ADP (gray bars) or 5 mM ATP (black bars) with 10⫺6 M of either thyronine (T0), 3,5-diiodo-l-thyronine (T2), triiodo-l-thyronine (T3), or thyroxine (T4) or without additions, as indicated, in 10 mM K-HEPES, pH 7.4, 40 mM KCl, as previously described (33). After reconstitution the proteoliposomes were dialyzed overnight against 10 mM K-HEPES, pH 7.4, 40 mM KCl. The ascorbate respiration of proteoliposomes (38 nM heme aa3) was measured polarographically in 10 mM K-HEPES, pH 7.4, 40 mM KCl, 18.5 mM ascorbate, 25 µM cytochrome c in the absence or presence of 1 µM valinomycin and 3 µM CCCP. The respiratory control ratio (RCR) represents the ratio of respiration in the presence and absence of the uncouplers. (From Ref. 76.)

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(or slip) in cytochrome c oxidase, thus abolishing partly the respiratory control based on ∆p. The proton leak, measured by Brand and Murphy (34) in mitochondria from rats after thyroid hormone treatment, could thus be explained as a changed proton permeability within cytochrome c oxidase and possibly also within other proton pumps of the respiratory chain. From our results we conclude that 3,5-T2 increases coupled as well as uncoupled respiration of cytochrome c oxidase, which would increase basal metabolic rate and thermogenesis in vivo. This conclusion is supported by the calorigenic effect of 3,5-T2 in vivo (74). The cold tolerance of hypothyroid rats increased after injection of 3,5-T2, owing to increased energy expenditure of the whole animals. In addition, 3,5T2 stimulated the oxidative capacity and cytochrome c oxidase activity in rat heart, liver, skeletal muscle, and brown adipose tissue (75).

ACKNOWLEDGMENTS Preparation of this chapter was supported by the Deutsche Forschungsgemeinschaft (Ka 192/28-4) and Fonds der Chemischen Industrie.

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16 Regulation of Glucose Transport by Insulin in Muscle and Fat Cells Translocation and Activation of Glucose Transporters Romel Somwar and Karen Yaworsky The Hospital for Sick Children and The University of Toronto, Toronto, Ontario, Canada

Gary Sweeney, Toolsie Ramlal, Peter Tong, Zayna Khayat, and Amira Klip The Hospital for Sick Children, Toronto, Ontario, Canada

I. INTRODUCTION The stimulation of glucose transport across the plasma membrane by insulin in muscle and fat tissue, the main sites of postprandial glucose utilization, is crucial to the maintenance of glucose homeostasis (1,2). The first insights into this intricately regulated membrane transport process were provided in 1980 when it was demonstrated that the stimulation of glucose transport by insulin in isolated adipocytes was accompanied by the translocation of presynthesized glucose transporters to the plasma membrane from a poorly defined intracellular locale (3,4). It was later discovered that this transporter, now referred to as glucose transporter 4 (GLUT4), constitutes the main, but not the only, insulin-responsive glucose transporter in muscle (5), fat (6), and other insulin-responsive cell lines (7,8). In addition to GLUT4, insulin recruits GLUT1, the principal mediator of basal glucose uptake, to the plasma membrane in insulin-responsive tissues and cell lines (1,2,9). Although necessary, it remained unclear whether increased plasma membrane content of GLUT1 and GLUT4 suffices to achieve stimulation of glucose uptake by insulin. Recent work from this laboratory has uncoupled the translocation of glucose transporters from the activation of newly arrived transporters 283

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in two different insulin-responsive cell lines (10). In this chapter we discuss recent studies indicating that there are at least two distinct signaling pathways that are indispensable for maximal stimulation of glucose transport: one controlling the translocation of glucose transporters to the plasma membrane and the other controlling the intrinsic activity of the newly arrived glucose transporters.

II.

SIGNALING PATHWAY REGULATING THE TRANSLOCATION OF GLUCOSE TRANSPORTERS TO THE PLASMA MEMBRANE

A.

Insulin Receptor Substrates

Activation of the insulin receptor tyrosine kinase is the first biochemical consequence of the hormone binding to its receptor (11) and is the first critical step in the stimulation of glucose transport (12). In 1982 it was reported that the insulin receptor is a tyrosine kinase that phosphorylates itself and several cellular substrates (13). Today, these substrates include a family of proteins referred to as insulin receptor substrates (IRSs), of which four isoforms have been identified (IRS-1–IRS-4) (11). IRS-1 and IRS-2 are ubiquitously expressed (11), while IRS3 is highly expressed in adipocytes (14). It is unknown whether IRS-4 is a physiologically relevant mediator of insulin action as it is mainly expressed in a human embryonic kidney (HEK) cell line (15). IRS proteins couple to the insulin receptor via their phosphotyrosine binding (PTB) domain, which interacts with an NPXY motif in the juxtamembrane region of the receptor (11). In addition, the pleckstrin homology (PH) domains of the IRS proteins are also important for transmitting the insulin signal (16). Upon binding to the activated insulin receptor, the IRS proteins become tyrosine-phosphorylated on multiple sites (11). The IRS proteins can then serve as ‘‘docking proteins’’ that recruit and modulate the regulatory or catalytic activity of a wide variety of signaling proteins (16). Downstream, signaling molecules interact with the IRS molecules by binding to the phosphotyrosine residues found within YXXM or YMXM motifs (Y ⫽ Tyr, X ⫽ any amino acid, and M ⫽ methionine). It is not known with certainty which of the IRS proteins are involved in the stimulation of glucose transporter translocation by insulin. IRS-1 was initially thought to be the principal mediator of GLUT4 translocation and the stimulation of glucose transport based on observations that (1) overexpression of IRS-1 in rat adipocytes increased basal GLUT4 content in the plasma membrane (17) and (2) ablation of IRS-1 with antisense ribozyme in rat adipocytes reduced insulin sensitivity of GLUT4 translocation. However, the ability to translocate GLUT4 and achieve maximum stimulation of glucose transport by insulin was unaltered. In contrast, several lines of evidence now indicate that IRS-1 may not play a crucial role in insulin-stimulated GLUT4 translocation. First, interfering with

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IRS-1 function by microinjecting anti-IRS-1 antibodies, PTB domain constructs, or NPX-phosphotyrosine peptides into 3T3-L1 adipocytes had no effect on insulin-stimulated GLUT4 translocation (18). Moreover, transgenic mice lacking IRS-1 retained a significant capacity to regulate blood glucose levels in response to insulin (19). This latter discovery provided the impetus to search for other substrates of the insulin receptor that might mediate the translocation of glucose transporters and the stimulation of glucose transport. The existence of the highly homologous IRS-2 may explain the inconsistency of results regarding the participation of IRS-1 in insulin-stimulated glucose transport. Indeed, IRS-2 has emerged as the insulin receptor substrate that is most likely to mediate insulin-stimulated glucose transport (20), because genetic ablation of IRS-2 caused diabetes in mice (21). These mice show progressive deterioration in glucose homeostasis that ultimately results in severe diabetes due to liver and skeletal muscle insulin resistance (21). This is distinctly different from the consequences of IRS-1 disruption in mice, which results in growth retardation and mild insulin resistance but not diabetes (19). In addition, overexpression of IRS-2 in rat adipocytes stimulated the translocation of GLUT4 in the absence of insulin (22). Whether other insulin receptor substrates participate in the stimulation of glucose transport is unknown at present. It is possible that they do, as it has been demonstrated that all four isoforms of the IRS proteins engage the same signaling molecules, some of which are important mediators of the stimulation of glucose transport by insulin. B. Phosphatidylinositol 3-Kinase The members of the phosphatidylinositol 3-kinase (PI 3-kinase) family are dual specific enzymes that possess lipid and serine kinase activities (23) and interact with all the known IRS proteins (16,23). In vivo, PI 3-kinases phosphorylate phosphatidylinositol 4,5-bisphosphate (PI 4,5-P2) to produce phosphatidylinositol 3,4,5-trisphosphate (PI 3,4,5-P3), which may then be converted to PI 3,4-P2 by the action of a phosphatase (23). Although insulin is able to stimulate several different classes of PI 3-kinase, it is currently believed that class IA PI 3-kinases participate in the translocation of glucose transporters and the stimulation of glucose transport (24). The functional class IA enzymes consist of a regulatory subunit that is constitutively associated with a catalytic subunit (23). Following insulin treatment, the two Src homology 2 (SH2) domains of the regulatory subunit (p85α) interact with phosphotyrosine residues within a YMXM or YXXM motif (see above) on the IRS molecules. This results in a marked increase in the lipid kinase activity of the associated catalytic subunit (23). To date, three isoforms of the class IA catalytic subunits have been identified; only two (p110α and p110β) are believed to participate in insulin signal transduction (23,24). Of the five regulatory subunits that couple to the class IA catalytic subunits (p85α, p85β,

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p50α, p55α, and p55γ) it is currently believed that only p85α is important for the translocation of glucose transporters and the stimulation of glucose transport by insulin (23). Insulin stimulates class IA PI 3-kinases in a similar time frame and with a dose dependence similar to its stimulation of glucose transport (25). Pharmacological inhibitors of PI 3-kinase provided the initial insights that this enzyme was one of the long-sought-after mediators of insulin-stimulated glucose transport. Wortmannin blocks the stimulation of glucose transport and the translocation of GLUT1 and GLUT4 in rat adipocytes (26), 3T3-L1 adipocytes (27), L6 rat skeletal muscle cells (28), and rat skeletal muscle (29). The results obtained using wortmannin in 3T3-L1 adipocytes and skeletal muscle were confirmed using LY294002 (30,31), a structurally distinct inhibitor of PI 3-kinase. Several other approaches have been used to show that PI 3-kinase participates in the stimulation of glucose transporter translocation and glucose transport by insulin (12,24): (1) Overexpression of constitutively active forms of PI 3-kinase can stimulate glucose transport and redistribution of GLUT1 and GLUT4 to the cell surface, albeit not to the same extent as insulin; (2) overexpression of a dominant negative mutant of p85α prevents the stimulation of glucose transport by insulin; and (3) microinjection of a mutant p85α or a glutathione S-transferase-p85 fusion protein interferes with the normal insulin-induced translocation of GLUT4 in 3T3-L1 adipocytes. Although necessary, activation of endogenous PI 3-kinase may not be sufficient to stimulate GLUT4 translocation. Several growth factors activate PI 3kinase to an extent comparable to insulin but fail to stimulate glucose transport in 3T3-L1 adipocytes (32) and L6 muscle cells (33). In addition, PI 3-kinase is stimulated to levels similar to that obtained with insulin by treatment of 3T3L1 adipocytes with a thiophosphotyrosine peptide derived from IRS-1 without a comparable stimulation of glucose transport (34). Several mechanisms may contribute to the specificity of insulin to stimulate glucose transport and the translocation of glucose transporters (12,24,35). Insulin might activate a targeted pool of PI 3-kinase that is physically distinct from those that other growth factors target. Alternatively, insulin may be able to direct PI 3-kinase to a distinct subcellular location following its activation. This latter model is supported by the observation that although insulin and PDGF caused robust activation of PI 3-kinase measured in cell extracts of 3T3-L1 adipocytes, only insulin was able to cause an accumulation of PI 3-kinase activity in low density microsomes (LDM) (24). This fraction represents a subpopulation of internal membranes from which the intracellular pools of glucose transporters are isolated and contains cytoskeletal elements as well (see later). We observed that in L6 myotubes, a 20 min insulin treatment also resulted in accumulation of PI 3-kinase in the LDM fraction, with no detectable amount of PI 3-kinase partitioning with the plasma membrane (Fig. 1A). In addition, insulin increased the amount of IRS-1-associated PI 3-kinase

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Figure 1 Subcellular redistribution of PI 3-kinase by insulin. Protein from plasma membranes (PM) or low density microsomes (LDM) isolated from control or insulin-treated (20 min) L6 muscle cells were resolved by 7.5% SDS-PAGE and immunoblotted with anti-p85 (regulatory subunit of PI 3-kinase) antibody. Representative immunoblots are shown. (A) The amount of p85 in the LDM was quantitated using the computer software NIH Image and is presented in the lower graph (relative units). (*) Significantly different from control, p ⬍ 0.005. (B) Total LDM isolated from control or insulin-treated (10 min) L6 cells was solubilized in 1% Triton X-100 and then centrifuged to obtain a detergentsoluble supernatant (S) and a detergent-insoluble pellet (P). Proteins were then separated and immunoblotted for PI 3-kinase as indicated in (A).

activity detected on GLUT4-containing vesicles isolated from 3T3-L1 adipocytes (36). Recent evidence suggests that insulin may also activate a second parallel pathway that is also required for maximal stimulation of glucose transport. This hypothesis is supported by two observations: 1. Although overexpression of the catalytic subunit of PI 3-kinase can significantly increase GLUT4 translocation and in some studies glucose transport (37), the stimulation is submaximal at best.

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2. Treatment of intact 3T3-L1 adipocytes with cell-permeable PI 3,4,5P3 could rescue the inhibition of insulin-stimulated glucose transport by wortmannin (38). However, cell-permeable PI 3,4,5-P3 alone did not elevate glucose uptake. These results suggest that activation of PI 3-kinase alone is not sufficient to fully stimulate glucose transport and that insulin may activate another parallel pathway that synergizes with PI 3-kinase to fully stimulate the translocation of glucose transporters and glucose transport (see Sec. III). C. Protein Kinase B/Akt Protein kinase B (PKB), also known as Akt, is a serine/threonine kinase (39). To date, three isoforms have been identified and are referred to as PKBα, -β, and -γ or Akt1–Akt3 (39). These isoforms are differentially regulated by insulin in a tissue-specific manner. Akt1 is the major isoform activated by insulin in liver and skeletal muscle (40). Akt1 and Akt3 are activated to similar extents in L6 muscle cells in culture (40). In isolated rat adipocytes and 3T3-L1 adipocytes in culture, Akt2 is the major insulin-responsive isoform (40). Interestingly, Akt2 is not activated to any appreciable extent by insulin in skeletal muscle or L6 muscle cells (40). Activation of all Akt isoforms by insulin is contingent upon prior activation of PI 3-kinase, regardless of the cellular background. This realization led to the suggestion that Akt may be a protein kinase that functions downstream of PI 3kinase in the regulation of glucose uptake by insulin. Indeed, a constitutively active Akt1, generated by adding a membrane-targeting motif (Src myristoylation sequence) to various Akt1 constructs, is sufficient to elicit maximal stimulation of glucose uptake and translocation of GLUT4 in 3T3-L1 adipocytes, independently of insulin (41). In addition, overexpression of wild-type or other constitutively active Akt1 mutants resulted in elevated GLUT4 translocation in rat adipocytes (42) and L6 muscle cells (43). Overexpression of a conditionally active Akt1 in 3T3-L1 adipocytes was also shown to stimulate glucose transport and the translocation of GLUT4 (44). It is apparent from these studies that activation of Akt1 may be sufficient to stimulate glucose transport to the extent achieved by insulin. However, they do not prove that Akt1 is necessary for the translocation of GLUT4 or the stimulation of glucose transport by insulin. Ablation of Akt1 activity is needed to prove that Akt1 is necessary for the translocation of GLUT4. In the absence of a pharmacological inhibitor, Akt1 mutants that may act as dominant negative inhibitors of endogenous Akt1 have been used. Cong et al. (42) overexpressed a kinase-inactive mutant of Akt1 (K179A) in rat adipocytes; this construct reduced insulin-induced GLUT4 translocation by 20%. Unfortunately, due to experimental limitations, it could not be demonstrated whether the endogenous Akt1 was inhibited by the kinase-inactive

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Akt mutant. Wang et al. (45) recently reported that an Akt1 construct (Akt1-AAA) with three mutations (K179A, T308A, and S473A) could successfully prevent activation of Akt1 by insulin in L6 muscle cells, suggesting that it acts in a dominant negative manner. When transiently expressed in L6 cells that stably express a GLUT4 cDNA harboring an exofacial myc epitope (L6 mycGLUT4 cells), Akt1-AAA prevented insulin-induced GLUT4 translocation. This implies that Akt1 is required for GLUT4 translocation in muscle cells (45). In contrast to our findings, overexpression of a different Akt1 construct (T308A, S473A), which inhibited more than 80% of endogenous Akt activity in 3T3-L1 adipocytes, was without effect on insulin-induced glucose transport or GLUT4 translocation (46). It remains to be determined whether inhibition of Akt2 will have more of an effect on insulin-induced GLUT4 translocation in rat adipocytes than inhibition of Akt1. Hence, Akt1 may play a physiological role downstream of PI 3-kinase leading to GLUT4 translocation in a cell-type-specific manner.

III. PARTICIPATION OF THE ACTIN CYTOSKELETON IN THE TRANSLOCATION OF GLUCOSE TRANSPORTERS Concomitant with the stimulation of glucose transporter translocation, insulin induces bundling of actin below the plasma membrane, a process referred to as membrane ruffling (47). It has long been recognized that the cytoskeleton plays a permissive role in the movement of vesicles within cells. With these concepts in mind, we hypothesized that the cytoskeleton may participate in the translocation of glucose transporters to the plasma membrane in response to insulin. Actin filaments continuously add monomers at their barbed end and lose monomers at their pointed end. Cytochalasin D prevents addition of monomers, leading in time to the persistence of short ‘‘disrupted’’ filaments. As shown in Table 1, inhibition of actin filament polymerization with cytochalasin D prevented insulin-dependent translocation of GLUT1, GLUT3, and GLUT4 and the stimulation of glucose transport in L6 muscle cells (47,48). In 3T3-L1 adipocytes, cytochalasin D reduced insulin-dependent GLUT1 translocation by 75% and GLUT4 translocation by 40% (36). The effect of cytochalasin D on insulin-stimulated glucose transport in 3T3-L1 adipocytes mirrors its effect on GLUT4 translocation, i.e., a 40% reduction (Table 1). Latrunculin B is another agent that prevents the continuous formation of actin filaments by scavenging actin monomers. Insulin-stimulated glucose transport was abolished in L6 muscle cells and reduced by 48% in 3T3L1 adipocytes by latrunculin B (Table 1) (36), confirming a role for the actin network in the stimulation of glucose transport. The extent of actin filament disruption by cytochalasin D or latrunculin B in muscle and fat cells might differ. Sufficient filaments may remain in the 3T3-L1 adipocytes to sustain partial trans-

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Table 1 Effects of Cytochalasin D and Latrunculin B on 2-Deoxyglucose Uptakea 2-Deoxyglucose uptake 3T3-L1 adipocytes Condition

Cytochalasin D (48)

Control Drug Insulin Insulin ⫹ drug

1.0 0.9 7.3 4.5

⫾ ⫾ ⫾ ⫾

0.00 0.06 0.50 0.30b

L6 cells

Latrunculin B (36) 1.0 1.03 5.9 3.07

⫾ ⫾ ⫾ ⫾

0.00 0.04 0.17 0.11b

Cytochalasin D (48) 1.0 1.03 2.05 1.18

⫾ ⫾ ⫾ ⫾

0.00 0.09 0.21 0.17b

Cells that were deprived of serum were incubated for 2 h with or without 1 µM (L6 cells) or 2 µM (3T3-L1) cytochalasin B or with 2 µM latrunculin B. During the final 30 min cells were stimulated with 100 nM insulin. 2-Deoxyglucose uptake was then determined over a 5 min period. Results are expressed relative to untreated control and are the mean ⫾ SE of three to five experiments in which there were at least three replicates of each condition. b Significantly different from corresponding insulin-treated cells in the absence of drug, p ⬍ 0.001. Source: Refs. 36 and 48, as noted. a

location of GLUT4. Alternatively, the contribution of the cytoskeleton to the translocation of glucose transporters and the stimulation of glucose transport by insulin may be variable, depending on cell type. IV. ROLE OF THE ACTIN CYTOSKELETON IN THE TRANSMISSION OF INSULIN-DEPENDENT SIGNALS REQUIRED FOR GLUT4 TRANSLOCATION Our current hypothesis is that the actin cytoskeleton may facilitate delivery of insulin signals to required cellular loci. Treatment of either L6 muscle cells (47) or 3T3-L1 adipocytes (36) with cytochalasin D did not affect insulin-induced tyrosine phosphorylation of IRS-1. In addition, the insulin-induced association of PI 3-kinase with IRS-1 or the insulin-stimulated lipid kinase activity of PI 3-kinase were not affected by cytochalasin D treatment (36,47). However, actin filaments may be important for the insulin-dependent relocalization of PI 3-kinase. In Figure 1A we demonstrate that PI 3-kinase (p85 regulatory subunit) was recovered with the LDM in response to insulin. To determine if PI 3-kinase in the LDM associates with the cytoskeleton, we treated LDM isolated from control and insulin-treated L6 cells with Triton X-100. The Triton X-100-insoluble fraction contains most of the cytoskeletal elements. We found that PI 3-kinase was present only in the detergent-insoluble fraction of the LDM (Fig. 1B). Following insulin treatment (10 min), PI 3-kinase increased only in this detergentinsoluble fraction. Further analysis of the cellular localization of PI 3-kinase is

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illustrated in Figure 2. This figure shows the distribution of actin filaments and of the p85 subunit of PI 3-kinase in L6 mycGLUT4 cells. Insulin treatment (10 min) resulted in a reorganization of actin filaments into a filamentous mesh (left panels). PI 3-kinase staining colocalized with this mesh of actin filaments (right panels). This colocalization was lost upon pretreatment of cells with cytochalasin D (results not shown). The actin filament-dependent relocalization of PI 3-kinase may also facilitate the propagation of some PI 3-kinase-dependent signals that are necessary for the translocation of glucose transporters. Figure 3 illustrates the effect of cytochalasin D and latrunculin B on insulin-dependent activation of Akt1 in L6 muscle cells. Both drugs reduced insulin-stimulated Akt1 activity by approximately 40%. Given that cytochalasin D completely prevents the stimulation of glucose transport by insulin in L6 muscle cells, it is tempting to speculate that the reduction in Akt1 activity represents 100% inhibition of a pool of Akt1 that is specifically involved in the stimulation of glucose transport. It is plausible that the role of actin filaments is to bring Akt1 proximal to the production of PI 3,4,5P3 and/or PI 3,4-P2 so that Akt1 could receive the PI 3-kinase-dependent signal required for full activation. Fully activated Akt1 would then be able to fulfill its

Figure 2 Colocalization of PI 3-kinase with actin filamentous mesh. L6 myotubes stably expressing a GLUT4 cDNA harboring an exofacial myc epitope grown on glass cover slips were treated for 10 min with or without 100 nM insulin. Cells were then washed, fixed, permeabilized, and dual-labeled with fluorescein phalloidin against filamentous actin (F-actin, left panels) or polyclonal rabbit antibody against the regulatory subunit of PI 3-kinase (p85, right panels). The specimen was examined using a Leica inverted immunofluorescence microscope with a 40⫻ objective lens.

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Figure 3 Role of the actin cytoskeleton in the activation of Akt1 by insulin. L6 myotubes that were deprived of serum for 5 h were treated for 2 h with 1 µM cytochalasin D (CD) or latrunculin B (LB). During the final 20 min of incubation, cells were stimulated with 100 nM insulin. Cells were lysed, Akt1 immunoprecipitated, and an in vitro kinase assay used to determine Akt1 kinase activity. Results represent the mean ⫾ SE of three or four experiments. Kinase activity under untreated conditions was assigned a value of 1, and all other values are expressed relative to this. (*) Significantly different from corresponding control, p ⬍ 0.01. (#) Significantly different from insulin-treated control, p ⬍ 0.05.

unidentified role in GLUT4 (and possibly GLUT1) translocation. Indeed, in L6 muscle cells, insulin causes a relocalization of both PI 3-kinase (Fig. 1A) and Akt1 (unpublished results) to the low density microsomes. It is important to note that not all PI 3-kinase-dependent signals are prevented by disruption of the actin network. Recently, we demonstrated that cytochalasin D treatment of L6 muscle cells does not affect the PI 3-kinase-dependent activation of the 70 kDa ribosomal protein S6 kinase (p70 S6 kinase) (49).

V.

SIGNALING PATHWAY REGULATING THE ACTIVATION OF GLUCOSE TRANSPORTERS BY INSULIN

A.

Role of p38 MAP Kinase

As discussed in Section I, it has long been advocated that insulin may also stimulate the intrinsic activity of glucose transporters (50). Although no clear evidence

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Table 2 Effect of SB203580 on 2-Deoxyglucose Uptake and GLUT4 Translocationa 2-Deoxyglucose uptake Condition Control SB203580 Insulin Insulin ⫹ SB203580

3T3-L1 adipocytes

L6 cells

GLUT4 Translocation L6 cells

⫾ ⫾ ⫾ ⫾

1.0 ⫾ 0.03 0.91 ⫾ 0.03 1.67 ⫾ 0.05 1.35 ⫾ 0.09

1.0 ⫾ 0.04 1.03 ⫾ 0.01 2.04 ⫾ 0.05 2.08 ⫾ 0.03

1.00 0.99 4.20 2.30

0.00 0.04 0.60 0.24

a

3T3-L1 adipocytes or L6 myotubes stably expressing a GLUT4 cDNA harboring an exofacial myc epitope (L6 mycGLUT4) were treated for 20 min with or without 10 µM SB203580 prior to stimulation for 30 min with 100 nM insulin. 2-Deoxyglucose uptake or translocation of GLUT4 to the plasma membrane was then determined. Results are expressed relative to untreated control and represent mean ⫾ SE of three individual experiments within which each point was assayed in triplicate. Source: Ref. 10.

has been presented to date, there is precedence for this phenomenon in response to other stimuli (51). For example, anisomycin stimulates glucose transport in 3T3-L1 adipocytes exclusively by increasing the intrinsic activity of GLUT1 (51). Recently, we demonstrated that SB203580, a potent inhibitor of p38 mitogen-activated protein kinase (p38 MAPK), prevents the stimulation of glucose transport by insulin in muscle and fat cells in culture (10). We were surprised to find that in 3T3-L1 adipocytes, normal insulin-dependent translocation of GLUT1 and GLUT4 to the cell surface occurred (measured by immunofluorescent labeling of GLUTs on plasma membrane lawns) without an increase in glucose uptake. We considered the possibility that SB203580 might have simply prevented the proper insertion of the glucose transporters into the plasma membrane such that they were not functionally competent. To address this, we used L6 mycGLUT4 cells where translocation of GLUT4 can be assessed in intact cells. In this system, the exofacial myc epitope on GLUT4 can be detected on the cell surface only if the mycGLUT4 is fully inserted into the plasma membrane. Using this approach we demonstrated that the insulin-dependent translocation and insertion of mycGLUT4 into the plasma membrane was unaffected by SB203580, yet insulin-stimulated glucose transport was reduced by the inhibitor (Table 2). We interpreted these results to indicate that following the translocation of GLUTs to the cell surface, they must be activated by a p38 MAPK-dependent signaling pathway. p38 MAPK may be a component of the second, parallel signaling pathway that is required for the full stimulation of glucose transport (12,35).

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VI. TYPE 2 DIABETES: DEFECTS IN INSULIN SIGNAL TRANSDUCTION AND GLUCOSE TRANSPORTER TRANSLOCATION Normal glucose homeostasis is tightly coordinated by insulin secretion from the pancreatic β cells in response to changes in blood glucose concentration. Type 1 diabetes (insulin-dependent diabetes mellitus) is a consequence of insulin deficiency arising from autoimmune destruction of the β cells. Type 2 diabetes (noninsulin-dependent diabetes mellitus) is characterized by (1) resistance of peripheral tissues, especially skeletal muscle and adipocytes, to the action of insulin on glucose uptake; (2) augmented hepatic gluconeogenesis and glucose output; and (3) dysregulated insulin secretion (20,53). One hundred million people worldwide suffer from Type 2 diabetes, yet, despite intense research, the primary lesion(s) responsible for the disease state remain unknown. The hallmark of Type 1 and Type 2 diabetes is hyperglycemia (54). In the latter disease, hyperglycemia may be a direct consequence of insulin resistance of the muscle and liver. In turn, hyperglycemia induces or exacerbates insulin resistance by affecting muscle, liver, and β-cell function. The reduction in glucose uptake by the diabetic muscle could be due to either a defect in the translocation or activation of glucose transporters or a reduction in the total amount of glucose transporters. Evidence supporting these two phenomena have been demonstrated in animal models of Type 2 diabetes such as the db/db and ob/ob mice, Zuker and Goto-Kakizaki rats, and in Type 2 diabetic humans. GLUT4 protein is markedly affected in adipose tissues of animal models of Type 2 diabetes and Type 2 diabetic humans. In an animal model of Type 2 diabetes created by transgenic ablation of brown adipose tissue in mice, total GLUT4 protein in adipocytes was diminished (1). GLUT4 expression in adipocytes also drops as diabetes develops in older Zuker rats (55). Similarly, adipose cells taken from humans with Type 2 diabetes show a reduction in GLUT4 content (56). This change in GLUT4 levels is notably restricted to adipose tissue, as normal expression of GLUT4 is observed in muscle of db/db mice and Zuker rats (57,58). Muscle biopsies taken from humans with Type 2 diabetes also show normal skeletal muscle GLUT4 content (59). The diminished insulin-mediated glucose uptake observed in muscle of Type 2 diabetes patients and animal models of the disease is associated with lower insulin-induced translocation of glucose transporters to the plasma membrane (58,60). Several possibilities could explain the reduced translocation of GLUT4 to the cell surface in both skeletal muscle and adipocytes: (1) altered signaling emanating from the insulin receptor; (2) impaired translocation machinery; (3) inability of the transporters to functionally incorporate into the plasma membrane upon arrival; (4) impaired activation of glucose transporters. In animal models of the disease there is considerable evidence for alterations in the early

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steps of insulin action. For example, there is an approximately 50% reduction in insulin receptor phosphorylation and an 80% drop in IRS-1 phosphorylation in liver and skeletal muscle of ob/ob mice (61). This is associated with ⬎90% decrease in insulin-stimulated PI 3-kinase activity associated with IRS-1 and no detectable stimulation of total PI 3-kinase activity. In addition, insulin-stimulated Akt1 kinase activity in skeletal muscle of the lean diabetic Goto-Kakizaki rat is reduced by 70% (62). Skeletal muscle isolated from individuals with Type 2 diabetes also shows defects at the level of the insulin receptor tyrosine kinase activity, IRS-1 expression and phosphorylation, and IRS-1 associated PI 3-kinase activity (63). Reductions in IRS-1 expression (by 70%) and IRS-1-associated PI 3-kinase activity have also been reported in adipose cells isolated from humans with Type 2 diabetes (56). Thus, in this disease, there are defects at four early steps of insulin action. Whether there are also defects distal to the initial signaling events that contribute to impaired translocation of GLUT4 remains to be determined. In addition to alterations in the expression and regulation of signaling molecules in Type 2 diabetes, the pattern of signaling also changes. In adipose cells isolated from humans with Type 2 diabetes IRS-2 becomes the main docking protein for PI 3-kinase and adapter proteins in response to insulin (56). In fact, expression of IRS-2 increases such that this protein predominates as the main insulin receptor substrate in liver of mice lacking IRS-1 (19). The upregulation of IRS-2 expression in IRS-1-deficient mice and in cells isolated from diabetic animals suggests that IRS-2 could substitute for IRS-1 in an adaptive response. Hence, it remains to be determined if the main role of IRS-2 is to mediate insulin signals in peripheral tissues or, as suggested, its main function is the regulation of insulin secretion in the β cells of the pancreas (20). VII.

CONCLUSIONS

There are two components to the stimulation of glucose transport by insulin in muscle and fat cells: translocation of glucose transporters to the plasma membrane, and activation of the transporters following their arrival at the plasma membrane (Fig. 4). These two processes are controlled by independent signaling pathways. The PI 3-kinase/Akt signaling pathway is important in the translocation of GLUT4 in muscle cells and adipocytes. An intact actin network appears to be required for the translocation process, perhaps by allowing signaling molecules to meet their effectors. Activation of glucose transporters is mediated by a p38 MAPK-dependent signaling pathway. Given that the stimulation of glucose transport by insulin is regulated in such a complex fashion, there are multiple levels at which defects can and do occur. In Type 2 diabetes there are defects at the level of glucose transporter expression and at the level of intracellular

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Figure 4 Our current hypothesis of the stimulation of glucose transport by insulin in muscle cells. Insulin binds to its receptor, resulting in the upregulation of the receptor’s intrinsic tyrosine kinase activity. Tyrosine phosphorylation of a family of insulin receptor substrate (IRS) proteins occurs, resulting in the formation of binding sites for phosphatidylinositol 3-kinase (PI 3-kinase). Occupation of the two Src homology 2 (SH2) domains of PI 3-kinase by phosphotyrosine residues within specific motifs results in elevation of the lipid kinase activity of the enzyme. The lipid products of PI 3-kinase have direct targets such as 3-phosphoinositide-dependent kinase (PDK)-1, PDK-2, and Akt1. Insulindependent activation of Akt1 results from the direct interaction of Akt1 with PI 3,4-P2 and/or PI 3,4,5-P3 and phosphorylation by PDK-1 and PDK-2. Actin filaments play an unidentified role in the activation of Akt1 by insulin. Active PI 3-kinase and Akt1 accumulate in low density microsomes (LDM, the subcellular membrane compartment that generates GLUT4-containing vesicles). GLUT4-containing vesicles are released and translocate to the plasma membrane. Once fully inserted into the plasma membrane, the intrinsic activity of the transporters is increased by a signaling pathway that involves p38 mitogenactivated protein kinase (p38 MAPK). Steps downstream of Akt1 and p38 MAPK remain to be defined. Known upstream activators of p38 MAPK include MAPK kinase kinase (MKK)3 and MKK6.

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signaling required for the proper translocation of glucose transporters. It remains to be determined whether there are any defects in intracellular signaling that mediate the activation of glucose transporters, such as the expression or activation of p38 MAPK or any of its upstream regulators in Type 2 diabetes.

ACKNOWLEDGMENTS We are grateful to Theodorus Tsakadiris and Qinghua Wang for their contribution to some of the experiments discussed. Financial support (to Amira Klip) was provided by the Medical Research Council of Canada and the Canadian Diabetes Association. Romel Somwar and Karen Yaworsky are supported by studentships from the Hospital for Sick Children Research Training Centre. Zayna Khayat is supported by a Doctoral Research Award from the Medical Research Council of Canada. Gary Sweeney is supported by a joint postdoctoral fellowship from Novo Nordisk and the Banting and Best Diabetes Centre at the University of Toronto. We also thank Dr. M. Birnbaum (University of Pennsylvania) for the supply of anti-p85 antibody.

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17 Glucose-6-Phosphatase A Member of the Newly Identified HPP Superfamily That Consists of Histidine Phosphatases and Vanadium-Containing Peroxidases and Consequences for Membrane Topology, Active Site, and Reaction Mechanism Wieger Hemrika, Rokus Renirie, and Ron Wever E. C. Slater Institute, University of Amsterdam, Amsterdam, The Netherlands

I. INTRODUCTION Our main research efforts were directed in the past to a peculiar class of enzymes carrying vanadate as a prosthetic group, the vanadium-containing haloperoxidases. We recently established a surprising homology between this group of enzymes and several families of acid phosphatases that were formerly considered unrelated. These families of acid phosphatases include glucose-6-phosphatase, lipid phosphatases from bacteria and yeast, lipid phosphatases from higher eukaryotes that are involved in signal transduction, and secreted bacterial phosphatases. Some excellent reviews within recent years describe these enzymes and their (putative) physiological roles (1–10). In this chapter we describe the homology and some of the important implications, especially for glucose-6-phosphatase, the enzyme involved in von Gierke disease. For more extensive treatment of the groups of enzymes mentioned in this chapter, the reader is directed to Refs. 1–10.

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II.

GLUCOSE-6-PHOSPHATASE

A.

Function and Role in Glycogen Storage Disease Type 1 (von Gierke Disease)

The liver plays an important role in the regulation of blood glucose levels. When bllod glucose is high the liver takes up glucose, which can than be used in glucose-consuming processes such as glycolysis or stored for future use in the form of glycogen. On the other hand, when the body is in need of glucose the liver supplies this sugar, produced via gluconeogenesis or glycogenolysis, to the blood, which transports it to other tissues that can use it. Glucose-6-phosphatase (G-6-Pase) catalyzes the hydrolysis of glucose-6phosphate to glucose and inorganic phosphate, which is the last step in both gluconeogenesis and glycogenolysis. The enzyme is present in high levels only in liver, kidney, and pancreatic islet β-cells (6), and, being the only enzyme in the body capable of producing significant amounts of glucose (6), G-6-Pase has a key role in glucose homeostasis. The enzymes catalyzing the reactions of gluconeogenesis and glycogenolysis are almost exclusively located in the cytosol. This made the finding in 1951 (11) that G-6-Pase activity was located in the endoplasmic reticulum (ER) a very surprising one that led to the insight that G-6-Pase activity has to be accompanied by transport functions for both its substrates and products. To date two opposing models explain the localization and kinetic behavior of the enzyme. The first and most prevalent model is called the substrate transport or catalytic subunit model (12,13). In this model it is proposed that G-6-Pase is part of a multicomponent system in which the hydrolytic enzyme is accompanied by accessory proteins (14) that are responsible for transport of the substrate(s) and the product(s) and for stabilization of the catalytic unit. One transport protein, named T1 (13), is required for the translocation of glucose-6-phosphate from the cytosol to the ER lumen (15). G-6-Pase, the hydrolytic subunit, is a relatively nonspecific phosphohydrolase with its active site facing the ER lumen. In addition to glucose-6-phosphate the enzyme hydrolyzes substrates such as pyrophosphate and carbamoyl phosphate (4). Furthermore, G-6-Pase has been shown to carry out a number of phosphotransferase reactions (6). Two other transport proteins (T2 and T3) are responsible for transport of phosphate (16) and glucose from the lumen back to the cytosol. Recently a candidate for T3, the glucose transporter, was identified and named GLUT7 (17–19). The second model, the combined conformational flexibility substrate transport model, originally proposed by Schulze et al. (20) and modified to its present signature by Berteloot et al. (21,22), describes G-6-Pase as a single membranespanning protein that functions in hydrolysis and substrate transport. According to this model the enzyme forms a water-filled space around the catalytic site that

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is deeply buried in the membrane (21). Using this model all the kinetic observations are explained by assuming that the protein can be found in several active conformations, each one possessing particular properties (21–26). The effects that accompany the absence of G-6-Pase activity exemplify the importance of this enzyme in regulating blood glucose levels (6). It was demonstrated in 1952 (27) that a disease first described by von Gierke in 1929 (28) was the result of the absence of G-6-Pase activity. This disease, glycogen storage disease type 1 (GSD 1), also known as von Gierke disease, is an autosomal recessive disorder that has an incidence of about 1 in 200,000. The disorder manifests itself by hypoglycemia, growth retardation, hepatomegaly, kidney enlargement, hyperlipidemia, hyperuricemia, and lactic acidosis (6). Without dietary treatment patients suffering from this disease die early, usually in their teens, of liver and kidney complications (29). Not all patients diagnosed with GSD lack the hydrolytic capacity of G-6Pase, and cases are described in which G-6-Pase activity could be restored after detergent treatment of microsomes of patients diagnosed with GSD type 1. Later it was demonstrated (30) that these patients were deficient in microsomal membrane transport for glucose-6-phosphate. Several GSD type 1 subtypes are described to account for such findings. According to the substrate transport model the subtypes of GSD type 1 are linked to the defects in the hydrolytic subunit or proteins associated with that. GSD 1a is then the result of a defect in the hydrolytic enzyme itself. GSD 1b is linked to defects in the (putative) glucose-6-phosphate transporter (30), type 1c results from defects in the transport of phosphate (31), and type 1d, linked to defects in microsomal glucose transport, has also been described (6). It should be noted that the mentioned GSD type 1 subtypes are also explained (although less elegantly) within the framework of the combined conformational flexibility substrate transport model when the effects of one or several types of inhibitors such as fatty acyl CoA esters (32), are taken into account (5).

B. Cloning of the G-6-Pase Gene and Structure–Function Analysis In 1993 research on G-6-Pase took an important step forward with the cloning of the murine cDNA encoding this enzyme (33) and the subsequent cloning of the G-6-Pase genes from rat, human, dog, and recently two fish species (34,35). The sequence database also contains a mouse G-6-Pase isoform cloned from the islets of Langerhans for which additional information is unfortunately absent. Analysis of the protein sequences reveals a C-terminal ER retention signal, and based on the hydropathy profile a model was predicted (Fig. 1) in which the

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Figure 1 Six transmembrane helix topology model adapted from Ref. 42. Residues predicted to be on the membrane interface are depicted in the open ovals on the helices (depicted as cylinders). Arg83 (R83) and the conserved histidines are depicted as open ovals on the loops. H ⫽ His, R ⫽ Arg, F ⫽ Phe, L ⫽ Leu, I ⫽ Ile, P ⫽ Pro, V ⫽ Val. This model was later corrected to the one depicted in Figure 8.

enzyme is anchored in the ER membrane via six transmembrane helices (33,35). However, we show later that this topology model is in need of correction. All G-6-Pases are highly similar. Overall similarity between the mammalian liver enzymes is higher than 85%, while the human and the fish enzyme show 51% identity. Identity between the mouse liver and pancreatic isoforms is 49%. Analysis of the G-6-Pase genes from GSD type 1a patients revealed at least 29 different mutations (36–40), and, importantly, it was shown that, in favor of the substrate transport model, patients diagnosed with GSD type 1b or type

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1c did not have mutations in their genes coding for G-6-Pase (41). It was found that more than 40% of GSD type 1a patients carry a mutation that affects Arg83 of G-6-Pase and changes it into a cysteine or a histidine (40,41). In a near-saturation mutagenesis study it was shown that Arg83 is indispensable for G-6-Pase activity (42). Based on the role of an arginine residue in alkaline phosphatases (43,44) it was proposed that Arg83 in G-6-Pase is involved in positioning of the phosphate group (42). Since it was already shown (45,46) that a histidine functions as a phosphate acceptor in G-6-Pase catalysis, Lei et al. (42) mutated all four conserved histidines (see Fig. 1) that were predicted to reside on the same (luminal) side of the ER membrane as Arg83. It was shown by this approach that from the four mutated histidines only His119 was absolutely required for G-6-Pase activity. It was therefore proposed that His119 functions as the acceptor of the phosphoryl moiety. The cloning of the G-6-Pase gene also enabled transcription regulation studies. Although historically it was assumed that there is no short-term regulation of G-6-Pase activity other than via its substrate concentration (6), it has now become clear that G-6-Pase gene expression is highly regulated. Glucocorticoids, cAMP, glucose, and fatty acids have all been shown to increase G-6-Pase mRNA (34,47–51). On the other hand, G-6-Pase gene expression is shown to be repressed by insulin, tumor necrosis factor-α, and interleukin-6 (34,47,52–54). Overexpression of G-6-Pase is proposed to play a role in some forms of diabetes (55). Research on G-6-Pase has largely benefited from the cloning of the gene encoding this enzyme from healthy individuals or from GSD type 1a patients. Analysis of the primary sequence and mapping of the mutations in patients enabled structure–function relation studies that led to a secondary structure model and to proposed active-site residues. However, to obtain good insight into the structure of the active site and the role of the active-site residues, information about the tertiary structure of the enzyme is essential. For G-6-Pase, which is a highly hydrophobic protein, obtaining such information is anticipated to be a difficult task. Very recently such information was provided from an unsuspected site, indicating, among other things, that His119 is not the phosphate acceptor.

III. VANADIUM-CONTAINING HALOPEROXIDASES A.

Discovery and Physiological Function

Haloperoxidases are enzymes that catalyze the two-electron oxidation of a halide to the corresponding hypohalous acid at the expense of hydrogen peroxide according to the reaction H2O2 ⫹ H⫹ ⫹ X⫺ → HOX ⫹ H2O

(1)

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When a suitable nucleophilic acceptor is present, a subsequent reaction will occur, giving rise to a variety of halogenated compounds. HOX ⫹ AH → AX ⫹ H2O

(2)

Haloperoxidases are named after the most electronegative halide they can oxidize, implying that a chloroperoxidase can oxidize chloride, bromide, and iodide while a bromoperoxidase can oxidize only bromide and iodide. Three groups of haloperoxidases are currently known. One of these groups consists of enzymes without a prosthetic group, and such enzymes have been detected in bacteria (56,57). The remaining two groups are enzymes that contain either heme or vanadate as a prosthetic group. Heme-containing haloperoxidases form the most studied group of haloperoxidases and are exemplified by the chloroperoxidase (CPO) from the fungus Caldariomyces fumago (58) or myeloperoxidase, which is found in white blood cells (59). Vanadium-containing haloperoxidases were first discovered in seaweeds about a decade ago (60–62) but since then have also been isolated from fungi (63) and from a lichen (64). Vanadium-containing bromoperoxidases (V-BPO) have thus far been isolated only from seaweeds and diatoms (65–67), and these enzymes are thought to be involved in the production of many halogenated compounds. V-BPOs are, in fact, believed to be responsible for the formation of large amounts of bromoform and halomethanes by seaweeds and phytoplankton (65). Their release into the atmosphere occurs when oceans become saturated with them, and this has been suggested to influence the ozone levels in the troposphere and in the stratosphere (68–70). As for a physiological function, these compounds are believed to function in the defense system of the seaweeds as they have antifeeding, antifungal, and antibacterial properties. Vanadium-containing chloroperoxidases (V-CPO) have thus far been isolated only from fungi (63,71), and the enzyme from the fungus Curvularia inaequalis is the most thoroughly studied CPO (72–74). The physiological role of the fungal CPO is not known yet, and, unlike the situation for the V-BPOs, no natural halogenated compounds are specifically attributed to the activity of the V-CPOs. Recently, based on a number of observations a hypothesis for the physiological function of the fungal V-CPOs was put forward (75). These enzymes are present at the surface of the fungal hyphae but are also found in the growth media of the fungi and are most probably secreted enzymes (75). Expression of the C. inaequalis V-CPO was shown to be high in the idiophase, when nutrients become limiting, and to be subject to glucose repression (75,76). Many of the fungi expressing V-CPOs are known as plant saprophytes or pathogens, e.g., C. inaequalis is the causative agent of Curvularia leaf spot on sweet corn plants (Zea mays). According to the hypothesis the enzyme functions in degrading the lignocellulose barrier of the host plant cell walls by producing HOCl. This would

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give the growing fungus access to the plant cells and thus make its nutrients available. B. Properties of the Cofactor and Enzyme Kinetics It is well established that the V-HPOs lose their cofactor in low pH buffers containing phosphate and EDTA, resulting in the inactive apoenzyme (77,78). The apoenzyme, however, can easily be reactivated at neutral or alkaline pH by the addition of vanadate (79). Binding constants for the cofactor have been reported to be as low as 35–55 nM for the BPO (78,79), while for the V-CPO a KD of approximately 100 nM at pH 7 has been reported (72). These high affinities correlate well with the concentrations of vanadate found in nature—approximately 50 nM in seawater (80) and levels of 20–130 ppm in soils (81). Furthermore, no additional protein factors are needed to convert the apo-V-CPO into the active holoform. The first step in catalysis of heme-containing peroxidases, formation of compound I by hydrogen peroxide, leads to oxidation of the heme iron from Fe3⫹ to Fe4⫹. Such a mechanism is not possible for the V-HPOs, because vanadium is present in these enzymes as vanadate and thus is already in its highest oxidation state, 5⫹ (61,82). Furthermore, no changes in the oxidation state of the vanadate were observed during catalysis, whereas reduction of the vanadate group resulted in an inactive enzyme (83). Vanadate is therefore considered to function as a Lewis acid, binding and activating the substrates for catalysis to occur. The steady-state kinetics of the V-HPOs have been extensively studied, particularly the V-BPOs, and these studies revealed that the enzymes work according to a substrate-inhibited bi-bi ping-pong mechanism (60,72,74,84–86). H2O2 is the first substrate that binds, and the affinity for this substrate was shown to increase with increasing pH. For the V-CPO of C. inaequalis, Km values for H2O2 have been reported to vary from 0.5 mM at pH 3.2 to 10 µM at pH 5 (74). The kinetic analysis also suggested that a group with a pKa of 5.6–6.5 is involved in the binding of peroxide to the enzyme. These pKa values are in the range of that of a histidine residue, and it was suggested that protonation of an active-site histidine results in the inhibition of peroxide binding to the enzyme (74). The halide is the second substrate that binds to the enzyme, and it was established that the pH profile for halide binding is opposite that of peroxide binding. For the V-CPO from C. inaequalis, the Km for Cl⫺ was shown to vary from 0.25 mM at pH 4 to 116 mM at pH 8 (74). The kcat for this enzyme reached a maximum of 22 s⫺1 at pH 4.5. Halide is also an inhibitor of the enzymatic reaction, and at low pH a competitive type of inhibition was established, whereas at higher pH values inhibition was shown to be noncompetitive (74,84,86). Recently we also determined the kinetic parameters for bromide oxidation by VCPO. It was shown that the V-CPOs from C. inaequalis and Embellisia didy-

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mospora have very high affinities for Br⫺ and the Km for this substrate is as low as 5 µM at pH 5 while substrate inhibition is observed at bromide concentrations higher that 0.5 mM (87). With bromide as the substrate, a kcat of 210 s⫺1 was established. These kinetic parameters reveal, in fact, that V-CPOs and V-BPOs are very similar enzymes, with the V-CPOs having the strongest oxidative ability. For comparison, kinetic parameters at pH 5 for the V-BPO of Ascophyllum nodosum are a Km for H2O2 of 150 µM, a Km for Br⫺ of 5 mM, and a kcat of 160 s⫺1. In the oxidation of chloride a Km for Cl⫺ of 344 mM was determined with a kcat of approximately 0.3 s⫺1 . These differences in halide specificity resulting from the differences in oxidative ability motivate an important question in our research on V-HPOs that considers the specific changes in primary and tertiary structure directing the specificity and the activity toward the different halides (see below). C. Primary and Tertiary Structure of the V-CPO from C. inaequalis Like research on G-6-Pase, research on V-HPOs recently obtained a large impetus with the cloning of the gene encoding the V-CPO from C. inaequalis (76), but, unlike the situation for G-6-Pase, the 3D structures of the native enzyme (88) and the catalytic peroxide intermediate (89) have also recently been determined. The C. inaequalis V-CPO gene encodes a protein of 609 amino acids with a calculated molecular mass of 67.488 kDa and, remarkably, although the fungus secretes the enzyme, no N-terminal secretion signal is present. Two cysteine residues were shown to be present in the primary structure, but it was established that these are present as free thiols, indicating that no disulfide bridges are formed in the mature protein while other forms of posttranslational processing such as glycosylation were also shown to be absent. Figure 2 shows an overall ribbon-type representation of the 3D structure ˚ resolution (88). The molecule has an overall as it was recently determined at 2.1 A ˚ . The secondary struccylindrical shape and measures approximately 80 ⫻ 50 A ture is mainly α-helical with two four-helix bundles as main structural motifs of the tertiary structure. V-HPOs are very thermally and chemically stable enzymes, with midpoint temperatures reported to be as high as 82–90°C (74,77). They show high stability in many solvents and are able to withstand high concentrations of hydrogen peroxide and hypohalous acid (90), which is in sharp contrast to the situation for the heme peroxidases. Analysis of the protein structure reveals that this high stability mainly results from the tight packing of the α helices, which show a strong hydrophobic effect (88). The vanadium binding site is located on top of the second four-helix bundle, and the amino acid residues providing the binding site span a length of approximately 150 residues in the primary sequence. Figure 3 shows that the metal is

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Figure 2 Ribbon-type representation of the CPO molecule. (Adapted from Ref. 88.)

bound as hydrogen orthovanadate (HVO42⫺) in a trigonal bipyramidal fashion via one covalent bond only to Nε2 of His496 . Hydrogen bonds to three positively charged residues—Arg360 , Arg490 , and Lys353 —compensate for the negative charges of the vanadate oxygens while Ser402 and Gly403 donate hydrogen bonds ˚ between vanadium and the apical to these oxygens. The bond length of 1.93 A oxygen is in the range of an OH ligand, which is hydrogen-bonded to His404 in the native structure. As mentioned above, protonation of a group with a pKa higher than 5 inhibits binding of peroxide and, based on the crystal structure, His404 seems a likely candidate for such a group. Two acidic and two aromatic residues may also be involved in V-CPO catalysis. Nδ1 of His496 is hydrogen-bonded to

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Figure 3 Stereoscopic view of the vanadium active site (adapted from Ref. 88) using Swiss PDB viewer (134) and POV ray.

Oδ1 of Asp500 , and this hydrogen bond might influence the protonation of N⑀1 of His496 in the apoenzyme. Asp292 forms a salt bridge with Arg490 and is also in close proximity of His404 , but its role in catalysis is not known yet. Trp350 and Phe397 together with the imidazole ring of His404 could provide the hydrophobic environment that seems necessary to stabilize chloride binding (88). D.

Events During Catalysis: A Reaction Scheme

Knowledge of the structure of the V-CPO active site and that of the peroxide intermediate (89) together with the information obtained from steady-state kinetics and spectroscopical studies enables us to develop a minimal reaction mechanism for the peroxidative activity of the CPO. The mechanism given in Figure 4 is a combination and extension of the reaction mechanisms proposed previously (2,3,89). The native enzyme is shown in Figure 4A. The hydrogen bond between the apical hydroxide and His404 makes this hydroxide a good nucleophile that will extract a proton from the incoming hydrogen peroxide and subsequently leave the coordination sphere as a water molecule. The deprotonated peroxide takes the empty coordination site on the vanadate (Fig. 4B) and the more nucleophilic oxygen, hydrogen bonded to Lys353, abstracts the proton from the singly bound peroxide. The now negatively charged peroxide displaces this formed newly hydroxide (Fig. 4B) and becomes bound side-on, as observed in the structure of the peroxide intermediate (89). Binding of the halide is the next step in catalysis. Binding can occur directly at the vanadate prior to oxidation (89), or the halide can carry out a nucleophilic

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Figure 4 Reaction scheme for the oxidation of halides by V-CPO.

attack on one of the oxygens of the bound peroxide. Based on mechanistic studies with model compounds (3) and on the structure of the peroxide intermediate, which is polarized by the H bond with Lys353 (Figs. 4C and 4D), we favor the latter possibility. Binding of the halide to the partial positive peroxide oxygen breaks the peroxide bond, and the nucleophilic OX group is formed (Fig. 4D). The OX group will take up a proton from an incoming water, possibly activated by His404 (Fig. 4E), and leave the coordination sphere as hypohalous acid. The formed hydroxide resulting from the deprotonation of the water molecule can take the empty coordination site on the vanadium, and the native structure is reformed (Fig. 4F).

IV. HALOPEROXIDASES AND ACID PHOSPHATASES: A CONSERVED ACTIVE SITE A.

Establishing the Homology

Initial database searches did not provide proteins homologous to the V-CPO (76,88). Aligning the amino acid sequence of the C. inaequalis V-CPO against the partial amino acid sequence of the A. nodosum V-BPO, however, revealed three stretches of high similarity in the regions providing the metal anion-binding site whereas the similarity turned out to be very low in the intervening regions (88). Using these three stretches of high similarity as templates in database searches, we were able to show that similar stretches are present in a large number

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of acid phosphatases (including G-6-Pase) that were previously considered unrelated (91). Figure 5 shows an alignment of the V-CPOs from C. inaequalis and E. didymospora (87), the V-BPOs from Corallina pilulifera (92) and Fuchus distichus, of which the genes were recently cloned, and some representative members of the acid phosphatases found to contain similar stretches. From the alignment it is clear that all residues coordinating vanadate in V-CPO are conserved. This allows the recognition of three separate domains that are highly similar among the aligned proteins. However, these three domains are connected by regions that are highly variable, both in sequence and in length. We thus suggested (91) that the binding pocket for vanadate in the peroxidases is similar to the phosphate-binding site in the aligned phosphatases. This is in agreement with the structural resemblance of vanadate and phosphate, with the observations that V-HPOs rapidly lose their activity in phosphate-containing buffers and that reconstitution of apo-BPO by vanadate is inhibited in the presence of phosphate (78). Furthermore, vanadate is known to be a potent inhibitor for many different phosphatases (93), including G-6-Pase, for which a Ki of 2 µM was established (94). Since our initial database searches, much improvement has been achieved in the available search algorithms. Furthermore, the linking of the databases, as done by the Entrez browser provided by the National Center for Biotechnology Information (NCBI), enabled us to quickly identify more than 40 (putative) proteins also containing domains 1, 2, and 3 in our more recent database searches; see also Refs. 95 and 96. The conservation of the active sites among the presented enzymes, both in structure and in the residues binding the anions, prompted us to determine whether the apo-CPO from C. inaequalis could act as a phosphatase (91). B. Apo-CPO Exhibits Acid Phosphatase Activity Since it is difficult to obtain pure apoenzyme from C. inaequalis in sufficient amounts for our experiments, we decided to use recombinant enzyme produced from the C. inaequalis V-CPO gene in a newly developed Saccharomyces cerevisiae expression system. This enabled us to produce large quantities of pure recombinant apoenzyme (rCPO), which, after activation with vanadate, behaves kinetically very similar to the enzyme isolated from C. inaequalis. Figure 6 presents the results of an experiment in which the putative phosphatase activity of apo-rCPO was assayed by measuring the release of p-nitrophenol ( p-NP), from p-nitrophenol phosphate ( p-NPP), a commonly used phosphatase substrate. This figure shows that p-NPP hydrolysis correlates linearly with the amount of enzyme added, demonstrating that the apo-r-CPO has phosphatase activity.

Figure 5 Alignment of the V-CPOs and V-BPOs with the acid phosphatases proposed to contain structurally similar active sites. Residues identical in 50% or more of the sequences are given in the gray boxes. Residues binding vanadate in the C. inaequalis VCPO are given underneath the alignment. YodM (accession Z99114), gene product of the YodM gene of B. subtilis and Methanococcus thermoautotophicum gene product (accession AE000858), which is similar to the bcrC gene product of Bacillus licheniformis (135). YG1P, hypothetical S. cerevisiae membrane protein (accession Z72821). Neutral phosphatase from Treponema denticola (136). phoN and phoC, nonspecific acid phosphatases from Salmonella typhimurium and Zymomonas mobilis (98,99). pgpB, lipid phosphatase from E. coli and Haemophillus influenzae (137,138); Wunen (100); phosphatidic acid phosphatase (PAP2a) (101); G-6-Pase of Haplochromis nubilis (accession AF008945) and human (35). BPO, bromoperoxidase of Fucus distichus (accession AF053411) and Corallina pilulifera (92). CPO, chloroperoxidase of C. inaequalis (76) and E. didymospora (87).

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Figure 6 Phosphatase activity as a function of apo-rCPO concentration. The enzyme was incubated with 1 mM p-NPP at 25°C in 100 mM citrate buffer at pH 5.0. After 4.5 h, reaction mixtures were quenched with NaOH and the production of p-NP was measured at 410 nm using ε410nm ⫽ 18.3 mM⫺1 cm⫺1 at pH 12. Each measurement was carried out in triplicate.

The presented data indicate that vanadate and p-NPP compete for the same binding site in the apoenzyme, implying that the phosphatase and peroxidase reactions are mutually exclusive. We have been able to show that this is indeed the case (91). Incubation of fully activated rCPO with 0.5 mM p-NPP at pH 5.0 resulted in a rapid decrease in CPO activity that was not observed in the absence of p-NPP. The decrease in CPO activity could be prevented by the addition of extra (100 µM) vanadate. We also determined the kinetic parameters of p-NPP hydrolysis catalyzed by apo-rCPO as a function of pH (91). This revealed that kcat varied by only a factor of 2 in the pH range 3.7–8.0, having an optimum of 1.7 min⫺1 at pH 5. This observation is in contrast with the case for haloperoxidase activity of VCPO, where kcat is affected much more within this pH range. The Km for p-NPP remained approximately 50 µM in the pH range 4.5–8.0 but increased strongly in the pH range 3.7–4.5 (Km-1.9 mM at pH 3.9), indicating that protonation of

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either a group on the free enzyme or the substrate itself is the cause of the reduced affinity for p-NPP.

C. Is V-BPO a Slightly Different Active Site? A close inspection of domain 1 in Figure 5 reveals one striking difference between the V-CPOs and the acid phosphatases on the one hand and the V-BPOs on the other. In domain 1 of V-CPO and the acid phosphatases there are six residues bridging the lysine (corresponding to the vanadate binding of Lys353 of V-CPO) and the arginine (corresponding to the vanadate binding of Arg360 of VCPO), while in the V-BPOs there are seven residues bridging these two amino acids. The presence of an extra residue in domain 1 of the V-BPOs may be a factor determining the enzyme’s low affinity for and low reactivity with chloride as a substrate. This is given weight by the fact that Lys353 of the C. inaequalis enzyme is the only one of the three equatorial basic residues (Arg360, Arg490, and Lys353) that forms a hydrogen bond with one of the oxygens of the bound peroxide in the catalytic peroxide intermediate (89). As mentioned, it is expected that this hydrogen bond activates the peroxide for nucleophilic attack by the halide. The slightly different geometry of the active site that will be the consequence of the additional amino acid in the domain of the V-BPOs might result in a less polarized peroxide intermediate. This in turn would result in its inability to oxidize the more electronegative chloride ion at physiological concentrations as in the marine environment.

V.

A CONSERVED ACTIVE SITE AND CONSEQUENCES FOR THE ACID PHOSPHATASES

A.

The HPP Superfamily

The common architecture of the active sites of the enzymes carrying domains 1, 2, and 3 may have important consequences for research in the acid phosphatase field since structural data are available for none of these enzymes. Figure 7 shows a dendrogram that is based on the alignment of the complete protein sequences. It should be noted that with this approach domains 1, 2, and 3 are not all aligned. This is because there is low sequence similarity outside these domains and also the positions of the domains in the proteins are variable. The method does enable us, however, to recognize the different protein families that together constitute the newly identified superfamily. Since a histidine is binding the vanadate in the peroxidases and, as suggested by the alignment, also the phosphoryl moiety in the acid phosphatases, we propose to call this superfamily the histidine phosphatase/peroxidase (HPP) superfamily.

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Figure 7 Dendrograms of proteins containing the conserved domains 1, 2, and 3. The dendrogram is based on a Clustal W alignment (default parameters) using the complete protein sequences.

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Inspection of Figure 7 reveals a bias toward membrane-bound proteins in the HPP superfamily. So far only two families—from the HPP superfamily— are identified that are soluble proteins; these are the vanadate-containing haloperoxidases and the class A nonspecific acid phosphatases (97–99). B. The LPP Family of Acid Phosphatases Together with the G-6-Pases, another (from an anthropocentric point of view) important and rapidly expanding protein family is that of which the Wunen protein (100) and the plasma membrane bound (type 2) phosphatidic acid phosphatase (PAP2a) (101) are members. Research on this family of enzymes, for which the name lipid phosphate phosphohydrolase (LPP) family was recently proposed (7), has also received an important stimulus. It is found that the acid phosphatases of the LPP family are not only involved in phospholipid and triacyglycerol biosynthesis but also play a role in signal transduction. This may occur via the balancing of the lipid second messengers diacylglycerol (DG) and phosphatidic acid (PA) (DG being the product of PA hydrolysis). Additionally, the ability of the LPPs to also hydrolyze lyso-PA, diacylglycerol pyrophosphate, ceramide-1-phosphate, and sphingosine-1-phosphate (102,103) may also indicate that the enzymes can attenuate signaling by these lipids while simultaneously generating other signals through the form of diacylglycerol, ceramide, and sphingosine (7,104). The interpretation of the regulatory functions of the LPPs in signal transduction is complicated by the occurrence of isoforms—four type LPP enzymes have been cloned in humans (7,101,105)—and the fact that all tissues examined express one or more of these LPPs at variable levels. PAP2a mRNA was recently shown to be expressed in 50 different human tissues, but expression levels were highest in the prostate and expression was stimulated by androgens (106). It was hypothesized that the ability of androgens to stimulate the expression of PAP2a may provide an important opportunity for cross-talk between signaling pathways involving lipid mediators and androgens (106). The importance of this group of proteins is exemplified by the functional analysis of the Wunen protein (100). This product of the Wun gene is a Drosophila protein of which the PA hydrolytic activity was only recently established (7). Activity of Wunen protein changes a permissive environment into a repulsive one, thereby guiding the germ cells in embryonic development from the lumen of the developing gut toward the mesoderm where they enter the gonads. Wunen shows high similarity to the protein encoded by the rat Dri42 gene (107). This protein was shown to be present in the ER membrane and was upregulated during development and differentiation of the epithelial cells of the intestinal mucosa. Members of the LPP family have almost superimposable hydrophilicity plots (7,100,107) and are predicted to have six transmembrane helices; the Dri42 protein was predicted to serve as a subunit for a channel protein (107). It should

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be noted that the predicted six transmembrane helix topology is in agreement with the proposal that the active site of these LPPs consists of the amino acid residues provided by domains 1, 2, and 3. Figure 7 also contains the LBP1 and LBP2 gene products from S. cerevisiae (108,109) that are, according to the dendrogram, more related to the G-6-Pases than to the enzymes of the LPP family. Strikingly, however, the LBP gene products were shown to be lipid phosphatases that modulate the stress response in yeast (110). C. A New Membrane Topology for G-6-Pase The knowledge that domains 1, 2, and 3 should be in close contact in order to constitute an active site can be an additional tool in determining the membrane topology of the membrane-bound enzymes contained in the HPP superfamily. An example of this was recently provided for G-6-Pase. As mentioned previously, the most common mutation in GSD type 1a affects Arg83, and this residue was proposed to be involved in positioning of the phosphate group (42). Such a role is confirmed by the alignment of Figure 5, since Arg83 of G-6-Pase aligns to Arg360 of V-CPO, which is one of the residues coordinating the vanadate. A subsequent mutagenesis study in which all four His residues of G-6-Pase predicted to reside at the same side as Arg83 were mutated revealed the importance of His119 of G-6-Pase, and this residue was proposed to be the acceptor of the phosphoryl moiety during catalysis (42). Our alignment, however, shows that His119 aligns to His404 of V-CPO, and this is not the histidine that covalently binds vanadate. The histidine residue that does covalently bind to vanadate is His496 of V-CPO, aligning to His176 of G-6-Pase. In the six transmembrane helix topology model (Fig. 1), His176 is positioned at the side opposite Arg83 and His119. Based on the conservation of the active sites of G-6-Pase and V-CPO we proposed that Arg83, His119, and His176 reside on the same side of the ER membrane and consequently that the six transmembrane helix topology model is in need of correction (91). Figure 8 shows our new model for the membrane topology (111) of G-6Pase, which is based on the presented evidence concerning the nature of the G6-Pase active site and on the results of two new algorithms for the prediction of membrane-spanning domains (112–114). These algorithms independently predict nine transmembrane helices in G-6-Pase, as opposed to the previous topology model. With this newly predicted topology all residues aligning to the activesite residues of V-CPO are situated on the same (luminal) side of the ER membrane. We propose therefore that helices II, III, IV, and V are in close contact and provide the glucose-6-phosphate binding and hydrolysis site of G-6-Pase. Experimental evidence for the new membrane topology was recently provided. Pan et al. (40) performed site-directed mutagenesis on His176 of G-6-Pase

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Figure 8 Newly proposed topology model for glucose-6-phosphatase. (Adapted from Ref. 111.) Residues predicted to be on the membrane interface are depicted in the open ovals on the helices (shown as cylinders). Residues corresponding to the active-site residues of V-CPO are depicted in the gray ovals. Possible N-linked glycosylation sites are indicated with the arrows. G ⫽ Gly, K ⫽ Lys, N ⫽ Asn, S ⫽ Ses, V ⫽ Val, Y ⫽ Tyr.

and were able to show that His176 (like His119) is indispensable for G-6-Pase activity, agreeing with its proposed function of binding the phosphoryl moiety. Furthermore, using N- and C-terminally tagged G-6-Pase, these authors were able to show that in intact liver microsomes the N-terminus is resistant to protease digestion, whereas the C-terminus is sensitive to such treatment. This indicated that G-6-Pase is anchored to the ER membrane via an odd number of transmem-

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brane helices, which is in agreement with the nine transmembrane helix topology model. Mutagenesis of potential N-linked glycosylation sites recently provided additional evidence for the nine transmembrane helix topology model (115). Analysis of mammalian multispan membrane proteins has indicated that N-linked glycosylation sites in these proteins are utilized when they are located on the luminal side of the ER and when they are at least 33 residues long (116–118). In accordance with the nine transmembrane helix topology model (see Fig. 8), Pan et al. (115) demonstrated that G-6-Pase was glycosylated at Asn96. Does the new G-6-Pase topology model have consequences for either the catalytic subunit model or the combined conformational flexibility substrate transport model? It is clear that the new topology model alone does not allow a decisive choice between these models. The proposed location of the active site, however, at the luminal surface of the ER membrane is not in agreement with the deeply buried active site proposed in the combined conformational flexibility substrate transport model (21). Furthermore, recent evidence from kinetic studies (119,120) and, even more important, genetic studies (121–124) identifying the putative glucose-6-phosphate transporter strongly support the catalytic subunit model.

D.

The Acid Phosphatases: A Reaction Scheme

Vanadium is coordinated in the V-CPO active site in a trigonal bipyramidal fashion (see Fig. 3). This coordination resembles the proposed transition state in the hydrolysis of the phosphate monoester by acid phosphatases in which a histidine acts as the nucleophile in the formation of the covalent enzyme–phosphate complex (125). On the basis of this resemblance and the role of the V-HPO activesite residues in haloperoxidase catalysis (see Fig. 4) it is possible to develop a reaction scheme for the acid phosphatase reaction of the apo-V-CPO and the homologous acid phosphatases (Fig. 9). The incoming phosphoester is positioned by the interaction of the phosphate oxygens with the basic Lys353, Arg360, and Arg490 or the corresponding basic residues in the acid phosphatases. A putative charge relay system (96) is formed between Asp500 and His496, which is likely to aid the nucleophilic attack of His496 on the incoming phosphate group (Fig. 9A). Simultaneously the alcohol leaving group moiety of the phosphoester interacts with a proton-donating group on the protein (group B in Fig. 9) and the SN2 transition state is formed (Fig. 9B). Upon formation of the covalent phosphoryl-histidine, the alcohol leaving group is released (Fig. 9C). The next step in catalysis, which again is likely to be aided by the charge relay system—hydrolysis of the phosphohistidine intermediate by an activated water molecule (Fig. 9D)—results in a noncovalent enzyme–phosphate

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Figure 9 Proposed reaction scheme of phosphatase activity of apo-CPO and the acid phosphatases.

complex (Fig. 9E). Subsequently, the phosphate is released and the native enzyme is re-formed (Fig. 9F). His404 and the corresponding histidines from the alignment are candidates for the proton-donating group B (see Fig. 9). However, according to the haloperoxidase reaction scheme (see Fig. 4) this histidine is unprotonated. Furthermore, in the reaction schemes of other acid phosphatases, such as the high molecular weight acid phosphatases that have a structurally very similar active site (126,127) but are not homologous to the enzymes of the HPP family, an aspartate is the proposed proton donor for the leaving group (128–131). Asp292 of V-CPO is a candidate for such a proton-donating group, and mutagenesis of this residue in V-CPO has revealed an important role in catalysis (Renirie and Hemrika, unpublished). However, no corresponding acidic residues are identified in the alignment of the V-HPOs and the homologous acid phosphatases. On the other hand, within each family of homologous acid phosphatases there are absolutely conserved aspartates that, based on the secondary structure predictions, could very well function as proton donors in the active sites of the acid phosphatases. Strikingly, G-6-Pase constitutes an exception to the HPP superfamily in that it does not contain the conserved aspartate corresponding to Asp500 of the

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C. inaequalis V-CPO (see Fig. 4) that is proposed to function in the charge relay network. It may be hypothesized, however, that Asp38 of G-6-Pase, which is positioned in helix 1 in the nine transmembrane helix topology model and is shown to be essential for enzyme activity (38,115), fulfills such a role in G-6-Pase. VI. EPILOGUE These are exciting times. The cloning of the G-6-Pase gene from healthy individuals and from GSD type 1 patients has been an important stimulus for research on this enzyme and has established the importance of specific amino acid residues for the catalysis and/or stability of this enzyme. Furthermore, the study of the enzyme’s regulation—at the levels of both gene transcription and enzymatic activity—has indicated that the role of G-6-Pase in glucose metabolism may be even more important than previously anticipated; see, e.g., Refs. 4 and 5. It is very possible that the inhibition of G-6-Pase activity by vanadate is an additional factor contributing to the insulin-mimicking effect of this compound (132,133). The phosphatases from the LPP family form another group of enzymes that are in the highlight of present-day research because they seem to play important roles in phospholipid-mediated signal transduction. It is anticipated that research in the G-6-Pases and LPPs will greatly benefit from the established homology with the V-CPO from C. inaequalis. The homology between vanadium-containing haloperoxidases and the acid phosphatases links families of enzymes that formerly seemed remote in the newly identified HPP superfamily of proteins. This would not have been possible without the knowledge of the 3D structure of the V-CPO, since overall sequence similarity between members of the different families contained in the HPP superfamily is very low. Furthermore, apart from their active sites, the enzymes of the HPP superfamily seem to share only their relatively low substrate specificity. Proteins from this superfamily have very different physiological functions and are found in species varying from E. coli to humans. However, vanadiumcontaining haloperoxidases have thus far been isolated only from eukaryotes and have not yet been detected in prokaryotes. Since phosphate and phosphate-metabolizing enzymes entered evolution at an early stage it seems likely that vanadate was coined by nature to become the prosthetic group in the vanadium-containing haloperoxidases at a more recent stage and thus that these enzymes have evolved from the phosphatases. ACKNOWLEDGMENTS This work was supported by the Netherlands Foundation for Chemical Research (SON) and was made possible by financial support from the Netherlands Organi-

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zation for Scientific Research, Netherlands Technology Foundation (STW), and the Netherlands Association of Biotechnology Centres (ABON).

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Alzheimer’s Amyloid β-PeptideAssociated Oxidative Stress Brain Membrane Lipid Peroxidation and Protein Oxidation D. Allan Butterfield University of Kentucky, Lexington, Kentucky

I. INTRODUCTION A.

Alzheimer’s Disease and Oxidative Stress

Alzheimer’s disease (AD) is the major dementing disorder of the elderly, affecting 4 million to 5 million persons in the United States currently and predicted to involve 14 million Americans early in the new millennium (1). AD patients present a progressive dementia characterized by cognitive and memory loss, and AD brain exhibits several characteristic pathological findings including loss of cortical neurons (with corresponding loss of synapses), the presence of neurofibrillary tangles composed mostly of hyperphosphorylated tau (a cytoskeletal protein), and senile (neuritic) plaques (SPs) composed of a central core of aggregated amyloid β-peptide (Aβ) surrounded by dystrophic neurites and other moieties (1). The molecular basis for the etiology and pathogenesis of AD remains uncertain, but recent research has provided strong evidence for the amyloid hypothesis for AD (2,3). This evidence, from many different perspectives, which supports the concept of a central role for Aβ in AD, is briefly summarized as follows: 1. Mutations in the amyloid precursor protein (APP), from which Aβ is derived, lead to excess deposition of Aβ and development of AD. 2. Persons with Down’s syndrome invariably develop AD after sufficient time, and APP is coded for on chromosome 21, the locus of trisomy in Down’s syndrome. 335

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3. Presenilin-1 and presenilin-2, two proteins coded for on chromosomes 14 and 1, respectively, may be involved in APP processing, and mutations in these proteins are associated with early-onset familial AD. 4. Mice overexpressing human APP develop a subset of pathology associated with AD. In all these cases, a common final pathway involves excess Aβ deposition, with consequent increased opportunity for oxidative stress and therefore death of neurons. There is considerable evidence of oxidative stress in AD brain (4). Markers of protein oxidation and lipid peroxidation, discussed in greater detail below, are significantly increased in AD brain (4,5). TBARS, an index of aldehydes formed from lipid peroxidation, are increased in AD brain (6). The lipid peroxidation product 4-hydroxy-2-trans-nonenal (HNE) also is increased in ventricular fluid and in frontal cortex in AD brain (7,8), as is free fatty acid release (9), consistent with increased lipid peroxidation in AD brain. Protein oxidation also is elevated in AD brain, in those areas in which amyloid-rich senile plaques (SPs) are found but not in SP-poor cerebellum (10,11). Consistent with Aβ-associated oxidative stress (5,10,12), APP-overexpressing mice, in which excess Aβ is deposited in brain, show evidence of brain-resident oxidative stress (13), and high-dose vitamin E treatment of AD patients is reported to significantly delay institutionalization (14). Implied in the above is the notion that a potential link between Aβ’s central role and oxidative stress in AD brain involes Aβ-associated free radical oxidative stress. Much evidence from many laboratories supports this concept, and the evidence suggesting Aβ-associated lipid peroxidation and protein oxidation is reviewed in this chapter. Reactive oxygen species (ROS) and reactive nitrogen species (RNS) are generally free radicals that react with important cellular components, resulting in altered structure and function (15). The term ROS encompasses H2O2, since this molecule, in the presence of Fe2⫹ or Cu⫹, can react to form hydroxyl radical (•OH). Oxidative stress is defined as a toxic condition characterized by chemical and/or functional alterations of biomolecules caused by an overproduction of free radicals, ROS, and/or RNS or resulting from a diminution of free radical defense mechanisms. Aging and age-related neurodegenerative disorders, such as AD, Parkinson’s disease, amyotrophic lateral sclerosis, stroke, diffuse Lewy body disease, and Wilson’s disease as well as traumatic brain injury, are associated with free radical oxidative stress (4,5,16–26). In addition, oxidative stress handling enzymes, including catalase, superoxide dismutase (SOD), and glutathione peroxidase, decline during aging (26).

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B. Major Markers of and Detection Methods for Oxidative Stress in Brain • Reactive oxygen species and reactive nitrogen species, such as •OH, O •⫺ 2 , HO 2, • • • ⫺ NO , ROO , RO , and ONOO , are formed in vivo chemically and biologically (15,16,26). The major markers of oxidative stress caused by ROS and RNS in brain cells are

1. Lipid peroxidation, detected by a number of methods such as formation of reactive aldehydic compounds (e.g., malondialdehyde) or alkenals (HNE) (27) or phospholipase A2-stimulated release of free fatty acids following oxidative stress (28,29), and by electron paramagnetic resonance (EPR) studies in which free radical-induced loss of signal intensity of lipid bilayer-resident stearic acid spin probes is followed (29– 31) 2. Protein oxidation, detected by increased protein carbonyl content (16,26) or by EPR in conjunction with protein-specific spin labels (10,32) 3. Formation of excess intracellular ROS, detected by ROS-sensitive fluorescence dyes (33) 4. Detection of 3-nitrotyrosine, a product of ONOO⫺ reaction with proteins (34) 5. Loss of activity of oxidatively prone enzymes such as glutamine synthetase (GS) and creatine kinase (CK) (10,16,35–40) 6. Increased message for and/or alterations in the activity of oxidative stress-handling enzymes, detected by RT-PCR message and/or activity analyses (6,41,42) 7. Oxidized DNA bases, usually detected as 8-hydroxyguanosine (43) 8. Prevention and/or significant modulation of these markers for oxidative stress and neuronal death by pretreatment of brain cells with appropriate free radical antioxidants (15) C. Aβ-Associated Free Radical Oxidative Stress and Neurotoxicity Free radicals can be detected by electron paramagnetic resonance (EPR) (44). In EPR spin-trapping studies, a nonparamagnetic species (the trap) in the presence of transient free radicals (the spin) forms a paramagnetic (EPR-detectable) moiety, usually a nitroxide (44,45). Using EPR spin trapping with the spin trap N-tertbutyl-α-phenylnitrone (PBN), several studies from different laboratories reported that Aβ(1-42), Aβ(1-40), or Aβ(25-35) yielded EPR-detectable spin adducts, evidencing the presence of a free radical (39,46–50). Others (51), using a variety of analytical methods, showed that Aβ(1-42) and, to a lesser extent, Aβ(1-40)

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donate an electron to substoichiometric amounts of peptide-bound Cu2⫹ , reducing the metal to Cu⫹ and forming a peptide free radical, i.e., confirming by non-EPR methods the existence of a peptide free radical. The reduced copper ion could then react with oxygen in a series of reactions to form H2O2 and other ROS. In the EPR studies, different times were required to detect free radicals from Aβ(1-42), Aβ(1-40), and Aβ(25-35), the times generally corresponding to the known different times for neurotoxicity associated with each of these peptides. Free radicals were not observed in solutions of the spin trap alone over these same time periods, indicating that free radicals were associated with the peptide. Further, when the spin trap finally did begin to break down in solution, the result was consistently a six-line EPR spectrum, in marked contrast to the three- or four-line EPR spectra of the peptides in PBN solutions. It is conceivable that trace contaminants in PBN led to the observed three- or four-line spectra. However, even though certain commercial lots of PBN do contain such contaminants, our studies, except the first, used PBN supplied by a pharmaceutical company whose products are based on nitrones or PBN we synthesized and purified ourselves, using recrystallization and sublimation methods, and in no case, even the first study, was a three- or four-line spectrum of the trap itself observed. Rather, as indicated above, when finally the spin trap began to decompose, a sixline spectrum was recorded. Highly purified PBN, even in the presence of 1 µM Fe2⫹ or Fe3⫹, gave no EPR spectrum over 6 h, again indicating that no contamination was present in the spin trap (Fig. 1). With Aβ(25-35), a four-line spectrum was observed after 24 h of incubation with purified PBN in the presence of chelator-treated buffer (Fig. 1). Thus, one can be confident that in Aβ studies if one uses highly purified PBN and buffers treated with metal chelators, the EPR spectra obtained emanate from peptide-associated free radicals. As reviewed below,

Figure 1 Spin trapping EPR spectra of PBN in the presence of Aβ(25-35) (top), purified PBN only (next to top), purified PBN plus 1 µM Fe2⫹ (next to bottom), and purified PBN plus Fe3⫹ (bottom). These spectra show that (1) Aβ(25-35) produces a free radical [we have also observed EPR spectra from spin trapping studies of Aβ(1-40) (39) and Aβ(142) (56), which give similar results] and (2) the PBN synthesized and purified in our laboratory appears to be contaminant-free; even redox metals are unable to form a spectrum in the time frame in which a spectrum is formed with Aβ.

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in ways inhibited by free radical antioxidants, Aβ induced lipid peroxidation (29– 31,52–54), caused brain cell protein oxidation (32,37,38,40,41,48,55–58), led to ROS formation (48,55), and resulted in neurotoxicity (32,48,55–61). On the basis of these and other findings, we postulated a central role for Aβ in the oxidative stress-induced neurotoxicity in AD brain (5,20,39). Figure 2 shows some of the many interrelationships of Aβ and oxidative stress and AD. As noted above, mutations in APP as well as mutations in presenilin proteins, both of which lead to familial AD (3,62,63), cause increased deposition of Aβ in brain. Aβ causes oxidative stress as manifested by increased lipid peroxidation and protein oxidation, which in turn causes synaptic damage and alterations in

Figure 2 Schematic diagram of Aβ-associated free radical oxidative stress and neurotoxicity in Alzheimer’s disease brain.

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cytoskeletal proteins and induces apoptosis, all of which lead to neuronal death. Lipid peroxidation and protein oxidation lead to altered Ca2⫹ homeostasis, resulting in significantly, and irreversibly, increased intracellular accumulation of Ca2⫹ , which in turn causes additional oxidative stress. Further, increased intracellular accumulation of Ca2⫹ leads to synaptic damage, alterations in cytoskeletal proteins, and apoptosis, indicating a large and complex feedback loop. Mitochondrial impairment, manifested, for example, by decreased activity of cytochrome oxidase and itself caused by oxidative stress, leads to free radical leakiness (15,26,64,65) and to increased intracellular accumulation of Ca2⫹ , which, as noted above, can cause neuronal death. Mitochondrial impairment also leads to leakiness of superoxide radical anion, which in the presence of NO forms peroxynitrite (34). Nitric oxide is formed by inducible-nitric oxide synthetase, which is stimulated by Aβ (66). ONOO⫺ is highly damaging to numerous tissues (34). Other pathways for neurotoxicity involving Aβ-associated oxidative stress are possible, including those associated with oxidative stress-induced transcription factors, e.g., Nfκ-B, inflammatory response, advanced glycation end products, and increased levels of redox-active trace metals (4). The cumulative data show that Aβ is a pro-oxidant. The mechanism(s) by which Aβ-associated free radicals form is still under investigation. However, methionine residue 35 may be involved (58,67). Methionine residue 35 is implicated in the mechanism of Aβinduced free radicals by the following (5,46,58,67): 1. Aβ(25-35) in solution forms a species that coelutes with Met sulfoxide, and this is prevented by a free radical scavenger. 2. Aβ(1-42) or Aβ(1-40), each substituted by norleucine for methionine at residue 35 (a CH2 in place of the S atom of methionine), do not yield an EPR spectrum in the presence of PBN. 3. Aβ(25-34), with no terminal methionine, likewise does not form an EPR-detectable spin adduct with PBN. The latter two peptides do not induce protein oxidation, and both are not toxic to neurons or glutamine synthetase, consistent with the spin trapping results. Studies with Met-substituted Aβ(1-42) expressed in transgenic animals are described below. Additional studies of the mechanism(s) by which Aβ-associated free radicals arise are ongoing. II.

Aβ-INDUCED LIPID PEROXIDATION

Membrane bilayer phospholipid unsaturated fatty acids are susceptible to oxidation. The brain’s low antioxidant activity and high oxygen consumption, coupled with high levels of polyunsaturated fatty acids (PUFAs), subject this organ to oxidative stress (15). Carbon-centered free radicals formed by free radical Hatom abstraction from unsaturated sites of phospholipid fatty acids immediately react with molecular oxygen to form lipid peroxyl radicals or hydroperoxides

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(15,26). Consequently, phospholipases are stimulated to cleave the oxidized fatty acids from the phospholipid backbone. The order and motion of phospholipids can change as a consequence of oxidation, and, because of the influence of membrane fluidity or oxidation on ion-motive ATPase function (68), increased intracellular calcium levels can result. Intracellular calcium also can cause activation of phospholipases (69), leading to even more intracellular Ca2⫹ . Phospholipase A2-induced release of arachidonic acid (AA) leads to a prostaglandin-mediated inflammatory response (28). Arachidonic acid also serves as a precursor for alkenals such as HNE (26–28). Several markers of lipid peroxidation are used to index free radical attack on phospholipids (27,70–74). In our laboratory, the methods commonly used to study lipid peroxidation include EPR with lipid-specific nitroxyl stearate spin labels, such as 5-NS and 12-NS, to examine changes in lipid bilayer order and motion (fluidity) and to monitor free-radical-caused loss of spin label paramagnetism, determination of conjugated dienes, analysis of free fatty acid release and phospholipid composition, and measurement of HNE levels. Cortical synaptosomal membranes treated with Aβ and incubated with the spin label 12-NS exhibited a 60% reduction in signal intensity of the EPR spectrum, indicating loss of paramagnetism of the spin label (30). No loss of the intensities of 12-NS in synaptosomal membranes was observed with the nontoxic reverse peptide, Aβ(35-25) (30). The antioxidant vitamin E inhibited the Aβinduced loss of paramagnetism in cortical synaptosomal membranes, consistent with the concept that Aβ is associated with free radical oxidative stress (29). This EPR paramagnetism reduction assay was used in another system to support this notion: PC-12 cells overexpressing Bcl-2, the gene product of which is thought to be an antioxidant, did not show Aβ-induced lipid peroxidation, in contrast to nearly 50% loss in signal following Aβ addition to PC-12 control cells (31). Aβ(25-35) incubated with synaptosomal membranes stimulated the release of phospholipid-resident fatty acids (29). The greatest release was for arachidonic acid, and this free fatty acid release, significantly, was inhibited by pretreatment of cortical synaptosomal membranes with vitamin E (29). Conjugated dienes were significantly elevated in brain membranes following Aβ additions (75). Aβstimulated lipid peroxidation also was reported by others, and the antioxidants vitamin E, melatonin, and others prevented this effect (52–54,76,77). The lipid peroxidation product HNE, produced following Aβ addition to hippocampal neurons (76), is neurotoxic (32,76,78,79). The membrane-damaging effects of Aβ are also produced by HNE: inhibition of ion-motive ATPase activity and loss of Ca2⫹ homeostasis (76,80); inhibition of glutamate transport (49,81); inhibition of glucose transport; and mitochondrial dysfunction in cultured neurons (82). The antioxidants propyl gallate and vitamin E protected against Aβ-induced lipid peroxidation (29) but not against HNE-induced alterations (32,76,82), consistent with HNE being a product of free radical–induced lipid peroxidation, not a free radical itself. Cortical synaptosomal membrane proteins bound HNE covalently

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and altered the conformation of membrane proteins (79), presumably accounting for HNE-induced inhibition of membrane transport and ion-motive ATPase functionalities. As noted above, Aβ stimulated i-NOS (65). Peroxynitrite-induced lipid peroxidation, measured by HNE-protein adducts and near-infrared alterations in lipid composition, is modulated by glutathione (83). The findings reviewed in this chapter strongly support the concept that amyloid β-peptide induces lipid peroxidation, consistent with the Aβ-associated free radical oxidative stress model for neurotoxicity in the brain of Alzheimer’s patients (Fig. 2). As mentioned above, there is considerable evidence for lipid peroxidation in AD brain. For example, thiobarbituric acid-reactive substances, principally malondialdehyde, are increased in AD brain (6), as are free (7) and protein-bound (8) HNE levels and free fatty acid release (9). Supporting this notion, high dose vitamin E treatment given to patients with moderate to severe AD delayed the progression of the disease (14), and addition of HNE to rat basal forebrain led to inhibition of choline acetyltransferase but not acetylcholinesterase and impaired spatial-temporal memory (84), common features of AD.

III. Aβ-INDUCED PROTEIN OXIDATION Several methods are employed to study free radical oxidative stress-induced protein damage. Some methods detect changes in the physical state of brain membrane or soluble proteins, while others detect chemical changes in oxidized proteins, i.e., increased protein carbonyls or changes in functionality manifested by loss of enzymatic activity (16,26). EPR, in conjunction with protein-specific spin labels, like 2,2,6,6-tetramethyl-4-maleimidopiperidin-1-oxyl (MAL-6) or (1oxyl-2,2,5,5-tetramethyl-delta3-pyrroline-3-methyl)methanethiosulfonate (MTS), is exquisitely sensitive to changes in the physical and conformational state of proteins (44). UV-Vis spectroscopy, immunochemistry, and histofluorescence methods are used to detect protein carbonyls (16,26). Kinetics studies measure changes in enzyme function (11,39). Each of these methods was used to demonstrate protein oxidation in AD brain and in Aβ-treated brain systems. Proteins are generally nonparamagnetic species. Consequently, proteinspecific spin labeling is required for EPR studies of oxidative modification of the physical state of proteins. The motion of the spin label is either relatively free or highly restricted, depending on the site of protein attachment. Surfacelocalized sulfhydryl (SH) residues spin labeled by MAL-6 reflect relatively unhindered spin label motion (weakly immobilized, W) with an associated resonance line that is relatively narrow (Fig. 3). In contrast, SH sites localized to narrow pockets or clefts within the protein spin labeled by MAL-6 reflect relatively hindered spin label motion (strongly immobilized, S) with an associated resonance line that is relatively broad and of low amplitude (Fig. 3) (44). The ratio of the low-field EPR signal amplitudes of the weakly (W) to strongly (S)

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Figure 3 Typical low-field EPR spectrum of MAL-6 spin-labeled synaptosomal membrane proteins. The W and S resonance lines are shown from which the W/S ratio is calculated. (See text.)

immobilized protein-bound spin labels (the W/S ratio) is a highly sensitive means of studying changes in the protein microenvironment (44). A decrease in the W/S ratio is an indirect measure of increased protein oxidation, resulting from increased protein cross-linking, increased protein–protein interactions, or changes in conformation or in accessibility to spin label binding sites (44). In several models of in vivo and in vitro oxidative stress leading to protein oxidation, such as hyperoxia (85,86), ischemia-reperfusion injury (21,87–89), accelerated senescence (90), and Fe2⫹ /H2O2-induced hydroxyl free radical–mediated oxidation (91), the W/S ratio of MAL-6-labeled synaptosomal membranes is decreased relative to controls. Synaptosomal membranes isolated from rodent brain and treated with Aβ resulted in a decreased W/S ratio of MAL-6, consistent with Aβ-induced protein oxidation (5,32). Synaptosomal membranes obtained from Aβ-rich hippocampus and inferior parietal lobule regions of AD brains following the University of Kentucky rapid autopsy protocol (2–4 h postmortem interval) showed decreased W/S ratios of MAL-6 relative to Aβ-poor cerebellum and to all areas of similarly obtained control brains (10). Antioxidants like vitamin E protected against the decrease in W/S ratios in rodent synaptosomal membranes treated with Aβ (5,32), consonant with the notion that free radicals and ROS are directly involved in the protein oxidation found in AD brain and in Aβ-mediated toxicity. That protein oxidation occurred in AD hippocampus and inferior parietal lobule brain regions or in rodent brain synaptosomal or astrocytic membranes treated with Aβ was confirmed by protein carbonyl measurements (see below) (5,32,36,48,55–58,81). Carbonyl groups, the most widely used markers of protein oxidation, can result from the direct oxidation of many amino acids such as histidine, lysine, arginine, proline, and threonine (16,26). Reaction of proteins with the lipid peroxidation product HNE or glycation or glycoxidation reactions of reducing sugars or their oxidation products with lysine can result in carbonyl formation (16,26). Aging results in an increase in brain protein carbonyls (16,26,90), and in several neurodegenerative disorders in addition to AD, such as amyotrophic lateral sclerosis, stroke, and Parkinson’s disease, an increase in brain protein carbonyls is observed (16,22,26).

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Protein carbonyls are detected by analysis of the resulting Schiff base formed by reaction of the carbonyls with a primary amine (reviewed in Refs. 16, 26, and 92). 2,4-Dinitrophenylhydrazine (DNPH) reacts with protein carbonyls to give a hydrazone derivative in acidic media (16,26,92): Protein–CCO ⫹ H2NENHE2,4-DNP → Protein–CCNENHE2,4-DNP ⫹ H2O The two principal means of detecting the hydrazone use spectroscopy (92) or immunochemical (37) methods. A different method for detecting protein carbonyls in hippocampal neuronal cell cultures is based on histofluorescence (48). This method uses biotin-4-amidobenzoic hydrazide to form the Schiff base with protein carbonyls followed by reaction with fluorescein-isothiocyanate-conjugated streptavidin. Digitization of the resulting fluorescence permits quantitation of the protein carbonyl levels. As noted above, cortical or hippocampal synaptosomes isolated from AD brain or from rodent brain incubated with Aβ peptides have increased protein carbonyls (5,10,11,32,55–58). Increased protein carbonyl levels also were observed in cultured hippocampal neurons incubated with Aβ(1-42), Aβ(1-40), and Aβ(25-35), and protection with the antioxidants propyl gallate or vitamin E was observed (5,32,48,55–58,81), consistent with Aβ-associated free radical damage to proteins and in agreement with the EPR studies noted above. Nontoxic reverse peptides did not cause protein oxidation. Maximum protein oxidation occurred at the time when, in separate spin trapping experiments, Aβ gave the most intense EPR spectrum (48,57). Protein oxidation in response to Aβ also has been demonstrated using oxidation-sensitive enzymes, particularly glutamine synthetase (GS) and creative kinase (CK) (16). The function of GS is to convert glutamate to glutamine, thereby preventing glutamate-activated NMDA receptor-mediated excitotoxicity. Oxidation of a single histidine residue found in the metal-binding region of the enzyme can lead to complete loss of enzyme activity (16). Hence, activity of GS is a useful marker of protein oxidation. CK plays a central role in energy transfer in cells with high energy requirements. Different CK isozymes, located in mitochondria and cytosol, are postulated to be necessary to establish an energy system that uses creatine as a phosphoryl carrier (93). Diminution of CK activity would have the effect of decreasing ATP availability with subsequent oxidative stress. Since CK is a sulfhydryl enzyme, this enzyme is susceptible to oxidative inactivation (94). Diminished GS activity occurs in AD brain (10,11). Aβ(1-40) or Aβ(2535) significantly decreased GS activity in cytosolic fractions of mammalian brain homogenates (35–39) and in cultured hippocampal neurons and astrocytes (48,55,81). Using purified GS, i.e., in the absence of any cellular components, GS was oxidatively inhibited by Aβ and protein carbonyls were incorporated in

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the enzyme (37), suggesting that Aβ-associated free radical oxidative stress occurs independently of cellular processes. In agreement with this suggestion, the free radical spin trap, sulfonated PBN, blocked the effects of Aβ on GS (37). Another study showing oxidation of GS by Aβ involved a kinetic investigation of the rate of covalent binding of the protein-specific spin label MTS (38). The rate of uptake of this SH-specific spin label was markedly reduced in oxidized sheep brain GS compared to control GS (38). Consistent with this finding and with oxidative stress in AD brain, highly purified GS isolated from AD brain showed a similar decreased rate of MTS binding relative to that from human control brain (38). A similar decrease in the rate of MTS uptake was observed when GS from sheep brain was treated with Aβ(1-40) (38). Decreased activity of CK in AD brain also was observed (10,11), and CK activity is inhibited by Aβ (39,55). The latter loss in CK activity is blocked by vitamin E, further supporting the concept of Aβ-associated free radical oxidative stress (55). Peroxynitrite also led to protein oxidation in cortical synaptosomal membranes, assessed by EPR spin labeling with MAL-6 and increased protein carbonyl levels (95). Pretreatment of synaptosomal membranes with glutathione prevented this effect (95). In vivo modulation of glutathione levels affected the action of ONOO⫺ : Injection of cyclohexene-1-one, known to decrease endogenous glutathione levels (89), led to significantly greater ONOO⫺-induced protein oxidation in subsequently isolated synaptosomal membranes than from peroxynitrite addition alone; in contrast, N-acetylcysteine, known to increase endogenous glutathione levels (96), partially protected subsequently isolated synaptosomal membranes from ONOO⫺-induced protein oxidation compared to peroxynitrite addition alone (97). As previously discussed, methionine residue 35 of Aβ may be important in the free radical oxidative stress and neurotoxicity associated with this peptide. We employed a unique in vivo model to test this idea. Caenorhabditis elegans transgenic animals expressing human Aβ(1-42) showed a 70% increase in protein carbonyls compared to vector-only control animals (58). In contrast, substitution of methionine residue 35 of Aβ(1-42) with cysteine resulted in healthy animals showing no increased oxidative stress assessed by protein carbonyl levels (98). This result is consistent with methionine substitution studies of Aβ(1-40) and Aβ(25-35) using EPR spin trapping, protein carbonyl, inhibition of GS, and hippocampal neurotoxicity (67). The finding of increased in vivo oxidative stress in Aβ(1-42)-expressing C. elegans transgenic animals is similar to increased oxidative stress in AD brain (4,5). Additional studies of this in vivo model of Aβassociated free radical oxidative stress are in progress. IV. CONCLUSIONS Physical and structural alterations of membrane proteins and the surrounding lipid bilayer can lead to changes in functionality of membrane-bound enzymes and ion transporters, resulting in ion imbalances. Inhibition of the activities of

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ion-motive ATPases, Na⫹ /K⫹ ATPase, and Ca⫹2 ATPase, as well as glucose and glutamate transporters, has been induced by Aβ and the lipid peroxidation product HNE (49,60,76,80,82). Several free radical antioxidants prevented Aβ-induced inhibition of ion-motive ATPases, consistent with a free radical process for this loss of protein functionality (5). Inhibition of the sodium pump can alter the cell potential, conceivably opening voltage-gated Ca2⫹ channels with consequent influx of Ca2⫹ . This increased intracellular Ca2⫹ can lead to further Ca2⫹ from intracellular stores, and because Aβ (or HNE) also inhibits the Ca2⫹ pump, the deleterious level of Ca2⫹ can not be extruded. Many destructive processes, such as proteolysis, breakdown of nuclear and mitochondrial DNA, and induction of apoptotic processes, result, leading to neurotoxicity (see Fig. 2). The many different approaches reviewed here strongly support the concept of Aβ-associated free radical protein oxidation. Taken together with lipid peroxidation reviewed above, with Aβ-induced ROS formation and neurotoxicity (55), with the prevention of all these deleterious effects by free radical scavengers (5), and with the central role of Aβ in Alzheimer’s disease, we suggest that Aβassociated free radical oxidative stress is important in neurotoxicity in AD brain. Further, these results suggest that brain-accessible free radical antioxidants should be evaluated for potential usefulness as AD therapeutics; indeed, the evidence reviewed here provides a rational basis for such a therapeutic approach, and early results with high-dose vitamin E treatment in AD (14) suggests that such a therapeutic strategy has merit. ACKNOWLEDGMENTS I thank my former and current graduate students and postdoctoral fellows for their excellent research and Drs. William Markesbery, Mark Mattson, and Christopher Link for useful discussions. This work was supported in part by NIH grants AG-05119 and AG-10836. NOTE ADDED IN PROOFS Others recently confirmed the four-line EPR spectrum of Aβ(1-42) in the presence of PBN (98). REFERENCES 1. Katzman R, Saitoh R. Advances in Alzheimer’s disease. FASEB J 1991; 4:278– 286. 2. Selkoe DJ. A central role of amyloid. J Neuropathol 1994; 53:438–447. 3. Selkoe DJ. Amyloid β-protein and genetics of Alzheimer’s disease. J Biol Chem 1996; 271:18295–18298.

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19 Membrane Orientation of the Alzheimer’s Disease–Associated Presenilins C. M. A. Boeve, P. Cupers, F. Van Leuven, W. Annaert, and Bart De Strooper Flanders Interuniversitary Institute, Gasthuisberg, and Catholic University Leuven, Leuven, Belgium

I. ALZHEIMER’S DISEASE Alzheimer’s disease (AD) is the most common neurodegenerative disease and is clinically characterized by a progressive dementia. All brains of patients with AD display cerebral amyloid plaques, which are extracellular deposits mainly consisting of the 40- and 42/43-residue β-amyloid peptide (βA); intraneuronal neurofibrillary tangles consisting of hyperphosphorylated tau; and extensive neuronal loss. In the large majority of cases, AD occurs sporadically with an age of onset of more than 65 years. Nonetheless, there are a number of families in which the disease has an early age of onset and a clear autosomal dominant inheritance pattern. The familial forms of AD (FAD) are associated with mutations in one of three genes. The first FAD gene identified is located on chromosome 21 and codes for the amyloid precursor protein (APP) (1). APP is proteolytically cleaved to give rise to βA. The amino terminus of βA is generated by β-secretase cleavage of APP. The resulting APP carboxy-terminal fragment is then further cleaved by γ-secretase(s) to yield the βA peptide, which consists of 40 or 42/43 amino acids. In 1995, two new FAD genes, presenilin 1 (PS1) on chromosome 14 and presenilin 2 (PS2) on chromosome 1, were identified (2–4). Mutations in these genes account for the majority of the autosomal dominant FAD pedigrees. PS1 and PS2 are highly homologous proteins, and more than 40 mutations in the PS1 353

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gene and two mutations in the PS2 gene have been identified (for a review, see Ref. 5). A first insight into the pathogenic mechanism of the presenilin mutations came from the observation that the levels of the βA(1-42/43) variant in plasma and in conditioned media from cultured fibroblasts derived from individuals with PS1 or PS2 FAD-linked mutations were significantly higher than those of control subjects (6). The longer (1-42/43) form of the peptide has a greater tendency to aggregate and thus acts as a seed in plaque formation (7). It is more toxic than the 1-40 form and is the predominant variant in AD plaques (8). Subsequent studies showed elevated levels of secreted βA(1-42/43) in cultured cells and in the brain of transgenic mice overexpressing mutant presenilins (9–14). Mutations in the presenilin genes thus cause an alteration in the processing of APP, probably at the level of the γ-secretase-mediated cleavage, leading to the formation of the more amyloidogenic form of βA. II.

CELL BIOLOGY OF THE PRESENILINS

Northern blot and in situ hybridization analyses show that presenilin mRNAs are widely expressed throughout the body and the brain (3,4,15,16). In the brain, the presenilins are predominantly expressed in neurons (15–17), where they are localized mainly in the cell body and dendrites (18,19). The levels of presenilin expression in mouse and rat brain are developmentally regulated with high levels in embryos and low levels in adult brains. In the latter, the presenilins appear to be primarily expressed in neurons of the hippocampus, the neocortex, and the cerebellum (16,20). The same pattern can be seen in primary cultures of rat hippocampal neurons, where the expression increases during neuronal differentiation until full polarization has been reached, and then declines (19). On the subcellular level, the presenilins are mainly located in the endoplasmic reticulum (ER) and to a lesser extent in the early Golgi (17,19,21,22; W. Annaert et al., unpublished results). There is no significant difference in the distribution of wild-type and mutant presenilin proteins (15,23). The presenilins are synthesized as an approximately 50 kDa precursor and undergo a physiological endoproteolytic cleavage to give rise to amino- and carboxy-terminal fragments of about 30 and 20 kDa, respectively (13,24–26). The cleavage is heterogeneous and occurs within the proximal portion of the hydrophilic loop region (27,28). The enzymatically generated fragments are the predominant presenilin forms observed in cultured cells and in brains. The levels of the fragments are highly regulated and saturable, since overexpression of presenilins in transfected cell lines or in transgenic mice results in the accumulation of full-length proteins without an increase in the level of the fragments (25,29). The amino- and carboxy-terminal fragments of either PS1 or PS2 are components of an oligomeric complex (30–33). Since the FAD-associated mutant PS1 lacking

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exon 10, which codes for a portion of the large loop domain including the cleavage site, elevates the level of βA(1-42/43) and is active in rescuing the egg-laying defect in Caenorhabditis elegans caused by mutations in sel-12 (see below), the proteolytic cleavage of the presenilins is not a prerequisite for their function (34,35). Moreover, FAD-linked missense mutant presenilin proteins seem to be normally cleaved (9,10,12,13). III. PRESENILIN HOMOLOGS The mammalian presenilins are homologous to the C. elegans gene product SEL12 that was identified by its ability to facilitate lin-12-mediated cell signaling (36). Lin-12 is a member of the Notch family of transmembrane receptors that are involved in the specification of cell fates during development (37). Another C. elegans homolog of the presenilins, HOP-1, was recently identified and shown to be functionally redundant with SEL-12 (38). Other presenilin homologs have been identified in Drosophila melanogaster and in Xenopus laevis (39,40). IV. MEMBRANE TOPOLOGY OF THE PRESENILINS The determination of the membrane topology of the presenilins is an important step toward the further understanding of their functions. The presenilins do not contain a secretory signal sequence. Using the indices of Kyte and Doolittle (41), hydropathy plot analyses of the presenilins reveal the presence of 10 hydrophobic regions in the polypeptide chain (Fig. 1). On the basis of this prediction, Sherring-

Figure 1 Hydrophobicity profile of PS1 using the Kyte–Doolittle algorithm (41) (window size ⫽ 15). The hydrophobic regions are numbered.

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ton and colleagues (4) proposed that PS1 is an integral membrane protein containing seven transmembrane domains. In their model, only the first six and the ninth hydrophobic regions were considered long enough to span the membrane lipid bilayer. In this theoretical model, the carboxy-terminal end of PS1 faces one side of the membrane while the amino-terminal part and the large acidic loop beyond the sixth hydrophobic region face the other side. Slunt et al. (42) used the newly developed TopPredII algorithm (43) to make predictions about the secondary structure of PS1. They proposed a ninetransmembrane domain model in which only the eighth hydrophobic region does not cross the lipid bilayer. In this model, the carboxy terminus of the protein and the large hydrophilic loop are positioned at the same side of the membrane while the amino-terminal end faces the other side. It was obvious that not predictions but experimental methods were needed to reveal the physiological topology of the presenilins. The most straightforward starting point was to determine the location of the hydrophilic loop and of both the carboxy and amino termini. These hydrophilic regions are not located in the lipid bilayer and are therefore available for interaction with antibodies and other proteins. Furthermore, as a first step toward the identification of those proteins, it is essential to determine whether these domains are oriented toward the cytoplasmic or luminal side of the endoplasmic reticulum membrane. We analyzed the orientation of the two most hydrophilic domains of PS1 using PS1 transfected COS cells treated with digitonin or saponin and immunostained with a set of antibodies against different regions of PS1 (22). Digitonintreated cells are permeable for antibodies only at the cell membrane and not at the ER membrane. The detergent saponin, on the other hand, permeabilizes the cell membrane and the intracellular ER membrane, exposing also the luminal side of the ER to the antibodies. This difference in the effect of the two detergents is a consequence of the difference in the cholesterol contents of the plasma membrane and the membrane of the ER (44). Digitonin-treated cells were immunoreactive with an antibody against the amino-terminal domain and an antibody against the hydrophilic loop, but not with an antibody against the second loop domain. Staining with the latter antibody was obtained only after treating the cells with saponin. Thus, both the amino-terminal end and the acidic loop after the sixth hydrophobic region are oriented toward the cytoplasmic face of the ER membrane. We concluded that at least the first six, but not the seventh, hydrophobic regions traverse the lipid bilayer of the ER (22). Doan and colleagues (45) essentially followed the same strategy but used streptolysin-O to selectively permeabilize the plasma membrane of cells. Using a panel of antibodies toward the different domains of presenilin, they came to the conclusion that both the amino- and carboxy-terminal ends as well as the hydrophilic loop region of PS1 are located in the cytoplasm. Furthermore, deleting exon 10, which codes for a portion of the large loop domain including a part

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of the seventh hydrophobic region, does not change the membrane topology of this domain. Assuming that the deletion of the 10 amino acid stretch of the seventh hydrophobic region would render this region too short to cross the membrane, this result suggests that the seventh hydrophobic region does indeed not span the membrane. In a couple of elegant studies, Li and Greenwald (46,47) studied the membrane topology of a series of β-galactosidase hybrid presenilin proteins in C. elegans. β-Galactosidase is active only in the cytoplasm of cells and not in the lumen of subcellular compartments. Fusion of LacZ (which codes for βgalactosidase) with the coding sequences for the second, fourth, sixth, seventh, ninth, or tenth hydrophobic region, respectively, resulted in transgenic lines displaying β-galactosidase activity, whereas no staining was detected when an additional synthetic transmembrane segment was inserted between the sel-12 sequences and LacZ. For the first, third, and fifth hydrophobic regions, the opposite was found: When LacZ was fused directly to those regions, β-galactosidase-negative transgenic lines were obtained, while including an additional transmembrane domain resulted in staining. Based on these observations, a membrane topology with at least six transmembrane regions was predicted, with the hydrophilic loop and both the amino and carboxy termini oriented toward the cytoplasm. The results obtained for the eighth hydrophobic region fusion proteins were not conclusive; neither the direct LacZ fusion construct nor the construct containing an additional transmembrane region yielded β-galactosidase-positive C. elegans strains. However, the β-galactosidase portion fused after the ninth hydrophobic region was found to reside at the opposite side of the membrane when compared to the same construct lacking the eighth hydrophobic region. The orientation of the ninth hydrophobic region seems to depend on the presence or absence of the eighth hydrophobic region, which implies that both the eighth and ninth hydrophobic regions are transmembrane. Moreover, a construct coding for a SEL-12 protein in which the eighth hydrophobic region was extended to obligatorily span the ER membrane was able to rescue the sel-12 mutant phenotype, indicating that the eighth hydrophobic region likely spans the membrane also in the wild-type PS1. These observations result in a model in which the presenilins have eight hydrophobic domains that span the membrane (Fig. 2). Using a different strategy, Lehmann et al. (48) proposed in contrast a six transmembrane domain structure. Those authors used reporter constructs containing glycosylation sites after each of the hydrophobic regions of PS1. The constructs were translated in vitro or expressed in COS cells, and glycosylation was analyzed. Only if the glycosylation sites were fused to the first, third, or fifth hydrophobic region was glycosylation seen, indicating that in those cases the reporter resided in the lumen of the ER. Further experiments analyzing the

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Figure 2 Predicted topology of PS1. The protein has six or eight transmembrane regions. The seventh and tenth hydrophobic regions probably do not cross the membrane.

protease sensitivity of the constructs confirmed these findings, leading to the conclusion that only the first six hydrophobic regions of PS1 effectively span the membrane, with the hydrophilic amino terminus and the long carboxy terminal region of the protein facing the cytoplasmic side of the membrane. Although the controversy around the question of whether the eighth and ninth hydrophobic regions of the presenilins span the membrane is not resolved yet, the above studies are consistent in their conclusion that the carboxy terminus, the amino terminus, and the hydrophilic loop of the presenilins reside in the cytosol. Dewji and Singer (49,50) postulated a totally different topology for the presenilins based on the observation that antibodies raised against the presenilin loop regions or the amino terminus of PS2 stained the cell surface of living transfected cells. Carboxy-terminal region antibodies were immunoreactive only when the cells were permeabilized. They proposed a seven transmembrane topology with both the amino terminus and the hydrophilic loop oriented extracellularly and not cytoplasmically. Several technical problems with this study have not been addressed until now. Most important, the specificity tests of the antibodies used were not conclusive, since they stained several other bands in addition to the presenilins in immunoblotting experiments (51). Although it was suggested that those proteins were aggregated or modified forms of the presenilins, no experimental evidence for this possibility was provided. In conclusion, the available experimental evidence indicates that PS1 and SEL-12 have most likely six or eight membrane-spanning domains, with the car-

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boxy terminus, the amino terminus, and the hydrophilic loop oriented toward the cytoplasm (Fig. 2). The problem in determining the topology of the eighth hydrophobic region can be explained by the fact that its short sequence has a relatively low hydrophobicity disturbing its topology in synthetic constructs. Furthermore, the available experiments do not address the possibility that the seventh and/or tenth hydrophobic regions are associated with or embedded in the membrane. The high degree of homology between the different presenilins (Fig. 3) suggests that they all have the same membrane topology. It cannot be ruled out that some of the experimental procedures used in the studies discussed above modify the interaction of the presenilins with the lipid bilayer, resulting in an artificial membrane topology. To definitively determine their membrane topology, electron microscopical studies in untransfected cells, using a large set of antibodies directed to different sequences of each of the loop domains, have to be performed.

V.

POSSIBLE FUNCTIONS OF THE PRESENILINS

Already more than 40 missense mutations in the PS1 gene and two missense mutations in the PS2 gene have been identified in patients affected with FAD (for a review, see Ref. 5). In addition, one splice-site mutation of PS1, resulting in the deletion of exon 10, coding for a portion of the loop domain including the cleavage site, segregates with the disease. The mutations are distributed throughout the protein, but most of the affected residues reside within one of the predicted transmembrane regions or in the hydrophilic loop, primarily amino-terminal to the cleavage site. Alignment of the sequences of the presenilins shows that most of the mutations occur at highly conserved residues, suggesting that they cause conformational or structural changes that directly affect presenilin function. Furthermore, it has been pointed out that the affected residues in the transmembrane segments of PS1 occur with a periodicity of three or four residues, suggesting that they are arranged along one side of an α-helix. In a theoretical three-dimensional model of PS1 using helical wheel analyses, most of the affected residues face the hydrophilic interior of a helical bundle formed by the transmembrane regions (52). The described model is compatible with a proposed function for the presenilins as ion channels, or more generally as transporters. It is clear that mutations causing amino acid substitutions at positions in the transmembrane regions would disturb the conformation of the channel formed by the cluster of helices. It should be noted that an α-helical configuration of the transmembrane regions is not necessarily needed to form a channel. For a number of pores and channels, like porin, it has been shown that the arrangement of transmembrane regions with a β-strand conformation in a so-called β-barrel can also create a molecule with a channel at its center (53). Until now, however, there has been

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Figure 3 Alignment of the protein sequences of human PS1, human PS2, C. elegans sel-12, and C. elegans HOP-1. Residues conserved in at least three members of this protein family are drawn on a black background. The 10 hydrophobic regions are overlined. As can be noted, these hydrophobic stretches are highly conserved.

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no experimental evidence to support the putative pore-forming properties of the presenilins. An interesting anlaogy has been drawn between the presenilins and the C. elegans protein SPE-4, involved in the spermatogenesis of the worm. SPE-4 is apparently needed for the formation of the fibrous body–membranous organelle complexes, which are Golgi-derived complexes with a role in the transport of proteins from spermatocytes to spermatids (54). The homology between the presenilins and SPE-4 suggests that they could play a role in the intracellular trafficking of proteins or docking and budding of vesicles. As the presenilins reside in the ER and Golgi, a functional role in the transport between these compartments is conceivable. More direct insight into the role of the presenilins comes from studies demonstrating that PS1 deficiency causes inhibition of the proteolytic cleavage of APP and Notch (55; Boeve et al., unpublished results). Interestingly, those changes occur close to or in the transmembrane regions of these proteins. For the Notch protein, this cleavage is essential for signaling; upon ligand binding, the cytoplasmic domain of Notch translocates to the nucleus, where several genes become activated (for a review, see Ref. 56). The proteolytic step releasing the cytoplasmic domain is inhibited in PS1-deficient mice, explaining some aspects of the phenotype of those mice. Indeed, a role for the presenilins in the Notch signaling pathway was suggested earlier on the basis of the finding that the mammalian presenilins are able to rescue the egg-laying defect caused by mutant sel12 in Caenorhabditis elegans (34,35). Furthermore, the phenotype of the PS1 knock-out mouse shows indeed some similarities with that of the Notch 1 knockout mouse (57,58). Cleavage of a protein at a site in the membrane is quite unusual. One wellstudied example is the sterol-regulatory element-binding protein (SREBP) (59). SREBP is an ER-localized transcription factor with two membrane-spanning regions. The cytoplasmically oriented amino-terminal fragment translocates to the nucleus after having been clipped from the membrane by the sequential action of two different proteases. The first cleavage, which occurs in the luminal loop, is regulated by the ER-localized protein SCAP (60). SCAP and SREBP are complexed through interaction of their carboxy-terminal cytoplasmic domains. The topology of SCAP is very similar to that of the presenilins: SCAP is an eighttransmembrane-spanning protein with both the amino and carboxy termini residing at the cytoplasmic face of the ER membrane (61). The structural homology of the presenilins and SCAP led to the hypothesis that the presenilins are involved in the regulation of the processing of APP and Notch by γ-secretase (55,56). The finding that PS1 may interact with APP (62,63) is consistent with this idea. PS1 could present APP and Notch to γ-secretase or could be involved in the transport of APP or Notch with the γ-secretase to the intracellular compartment where cleavage occurs.

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A more straightforward interpretation of the data is that presenilin itself is γ-secretase. Interestingly, the S2P proteinase that is actually responsible for the membrane domain cleavage of SREBP is also a polytopic membrane protein containing a consensus sequence specific for metalloproteases (64). Although the presenilins, in contrast to the S2P proteinase, do not contain a clear protease domain, the possibility that they are proteinases cannot be ruled out. In contrast, S2P is not γ-secretase, since the levels of βA(1-40) and βA(1-42/43) in S2P-deficient cultured cells were not different from those in wild-type cells (65,66). An important approach to further unraveling the biological function of the presenilins is to screen for interacting proteins. Using the yeast two-hybrid assay, coimmunoprecipitation assays, and affinity purification, a series of interacting proteins have been identified of which β-catenin (33,67–69), δ-catenin (69), filamin and its structurally related protein filamin homolog 1 (70), calsenilin (71), and APP (62,63) are the most promising. It was only recently that these protein interactions were demonstrated, and for the large majority of them the functional significance has still to be explored.

VI. CONCLUSION Since their discovery in 1995, much has been learned about the presenilins: Their metabolism, their expression pattern, and their subcellular localization have been documented. More than 40 FAD-linked missense mutations have been identified, and we start to understand how those mutations are involved in the development of AD. It is now essential to further elucidate the structure and membrane topology of these intriguing proteins to unravel further and in more precise detail their mechanisms of action and structure–function relations. Such detailed knowledge is instrumental for the rational development of drugs that hopefully can interfere with the devastating disease process of AD.

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20 Membrane Structure Analysis in Apoptotic Processes of Leukemic Blasts and Leukemia-Derived Cell Lines* Uwe Ebener, Sibylle Wehner, Christoph Rietschel, Hu¨lya Cakmak, Eckhard Niegemann, and Matthias Eishold Johann Wolfgang Goethe University, Frankfurt, Germany

I. INTRODUCTION A.

Electron Spin Resonance Spectroscopy

Electron spin resonance (ESR) spectroscopy is a powerful technique to study the properties of the cellular membrane of viable cells. The evaluation of ESR spectra enables the determination of various parameters such as membrane fluidity, polarity, and signal reduction (1–3). Equipped with a characeristic set of proteins, every cellular membrane is capable of performing highly specialized functions. Various techniques are available to specifically study such proteins. Basically every cellular membrane consists of the same phospholipid bilayer structure as described by Singer and Nicolson (4). The physical and structural properties of this cellular backbone are of great interest. The bilayer in its entirety exhibits properties of both a liquid and a gel-like substance. In this context, the term ‘‘fluidity’’ describes the quality of ease of molecular motion in a macromolecular environment. Fluidity is essential for cellular plasticity and seems to be crucially important

* Dedicated to Prof. Dr. Med. Dr. h.c. Bernhard Kornhu¨ber (Director of Clinic of Pediatrics–III), Johann Wolfgang Goethe University, Frankfurt, Germany.

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for the regulation of membrane proteins (5). Previous studies pointed out the possible role of changes in membrane fluidity in correlation to malignancy (6–8). To a great extent the fluidity of a cellular plasma membrane is determined by its lipid composition. Thus, we can constructively regard all the general allusions to fluidity to refer in part to some aspect of fluid expansion or molal excess volume, which describes the density of the lipid microenvironment (9). Accordingly, changes in fluidity are mainly caused by alterations of the phospholipid bilayer. Several studies have shown the influence of plasma membrane proteins on the fluidity to be negligible (10,11), although the percent by weight in eukaryotic cells amounts to 50% and more (5). This consideration, however, does not hold for intracellular membranes, where the amount of protein is over 80% (i.e., in mitochondria). In that case, the membrane exhibits much more fluidity than the ‘‘lipid-rich’’ plasma membrane (12). Another structural property of great interest is the polarity of the cellular membrane. Studies of the polarity may provide important information on lipid– protein interactions and different patterns of perturbation in the phospholipid bilayer. There also is a gradient of polarity, similar to membrane fluidity or order, decreasing with distance perpendicular to the membrane surface.

B. Apoptosis Cell death can be classified into three categories: (1) necrosis, which occurs as a result of massive tissue damage; (2) terminal differentiation of specialized tissues such as red blood, skin, and intestine; and (3) apoptosis, which is a process of active cellular self-destruction that requires the expression of a number of genes (13–16). Most cells from higher eukaryotes have the ability to self-destruct by activation of an intrinsic cellular suicide program when they are no longer needed or have become seriously damaged. This normal physiological process is referred to as programmed cell death (PCD) or apoptosis (16). In general, cells undergoing apoptosis display profound structural changes, including cell shrinkage, plasma membrane blebbing, chromatin condensation, and DNA fragmentation. The nuclear disintegration is associated with extensive damage to chromatin and DNA cleavage into oligonucleosomal length DNA fragments following activation of a calcium-dependent endogenous endonuclease (17,18). Biochemical studies of nuclear damage have shown that the cellular DNA is cleaved into multimers of about 180–200 base pairs that may be visualized as a distinct ladder of bands following agarose gel electrophoresis (19,20). The membrane-bound apoptotic bodies are readily phagocytosed and digested by macrophages without generating an inflammatory response. These changes distinguish apoptosis from cell death caused by necrosis (8,15,21).

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Figure 1 Chemical structure of camptothecin (C20H16N2O4).

C. Initiation of Apoptosis The CD95 (APO-1/Fas) receptor, which is able to induce apoptotic cell death upon oligomerization by its natural ligand or by the monoclonal antibody antiCD95, is broadly distributed in normal and malignant hematopoietic cells (22– 24). Different human leukemic cell lines are very sensitive to various apoptotic stimuli. One of the drugs best investigated in vitro is camptothecin (CAM). This indole alkaloid is isolated from a Chinese tree, Camptotheca acuminata, and has a wide spectrum of anticancer activity in vitro and in vivo, mainly through inhibitory effects on topoisomerase I (18,25–31) (Fig. 1). D.

Visualization of Apoptotic Cells by Immunological Detection Systems

Several flow cytometric methods have been described for analyzing apoptotic cells. Recently an assay that makes use of the capacity of the nuclear enzyme deoxynucleotidyltransferase (TdT) to label the 3′-OH termini of DNA breaks with biotinylated dUTP was developed (TUNEL method). Using this TdT-assay, apoptotic leukemic cells contaminating the peripheral blood of leukemic patients during therapy were identified (32). An early indicator of apoptosis in mammalian cells is loss of the phospholipid membrane asymmetry of the cell. This results in exposure of phosphatidylserine on the outer surface of the plasma membrane. This change in membrane asymmetry can be analyzed by using annexin V (33,34). II.

OBJECTIVE OF THE STUDY

The aim of this study is to systematically investigate and characterize physical and structural properties of the cellular membrane of intact vital cells obtained

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from hematological patients. Following the encouraging results of our previous investigations, our interest was focused on the process of apoptosis and its influence on the properties of the cellular membrane as observed by ESR spectroscopy (35,36). Attention was paid especially to the dynamics and the time dependence of alterations within the phospholipid bilayer and a comparison to the process of necrosis.

III. MATERIALS AND METHODS Due to the lack of experience in the systematic study of leukemic cell material by ESR, our first interst was to observe blast cells from the onset of childhood leukemia, where no specific drug effects would complicate the interpretation of the results. A.

Cell Culture

Human leukemia derived cell lines Molt-4, KG1, K562, HL-60, and U937 represent a suitable control for the group of malignantly transformed cells. Following optimal conditions during cultivation, they are available at any time. Their cellular membrane properties were studied frequently in order to rule out systematic errors. All of the cell lines we used were maintained in RPMI 1640 medium supplemented with 10% fetal calf serum (FCS), antibiotics (penicillin and streptomycin), and 2 mM l-glutamine in a humidified atmosphere (humidity 95% and 37 °C). Human lymphocytes (MNCs) isolated from healthy donors represented the control group (n ⫽ 23). The study comprised mononucleated cells (MNCs) isolated from bone marrow (BM) of children with newly diagnosed subtypes of acute leukemia from 25 pediatric patients suffering from acute lymphoblastic leukemia (ALL) or acute myeloid leukemia (AML) [(16 cases of common ALL (cALL), four cases of thymus-dependent ALL (T-ALL), and five cases of AML]. B. Cell Separation Cell separation was performed using a density-gradient centrifugation with Ficoll-Hypaque or Percoll (Pharmacia Biotech) according to Bøyum (37). This technique yielded a purity of mononuclear or polynuclear cells of more than 95%. For ESR analysis an amount of at least 107 cells per sample is required (35). After separation of freshly aspirated heparinized bone marrow from leukemic patients or cells from cultured cell lines, a cell suspension with a concentra-

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tion of 107 cells/mL was established, and portions of 1 mL were distributed to Eppendorf cups.

C. Induction of Apoptosis To study the structural properties of the intact cell membrane of apoptotic cell lines and leukemic blasts versus normal viable MNCs and necrotic cells, we used camptothecin (CAM) as a well-known inducing agent of apoptosis (26,29,30,38). For induction of apoptosis in cell culture, freshly isolated leukemic blasts from bone marrow as well as different cell lines and control cells were treated with the alkaloid CAM as previously described (36,39). Cells were incubated with CAM (50 µg/mL) for a period of 3–30 h. In comparison to the in vitro induction of apoptosis, we produced necrotic cells by increasing the temperature to 55 °C for 4 h following a cooling procedure with a refrigerator. After checking for cell death with the trypan blue exclusion test, necrotic cells were available for ESR analysis.

D.

Detection of Apoptosis

Apoptosis can be assessed by monitoring a number of physiological and morphological characteristics. One of the morphological traits associated with cell death is a loss of membrane integrity. These cells are then not able to retain, or exclude, certain dyes. For example, trypan blue is specifically excluded by viable cells but will stain cells that have lost membrane integrity. However, this method will not differentiate between necrotic cells and apoptotic cells undergoing secondary necrosis. Similarly, cells in the early stages of apoptosis do not lose membrane integrity and will not be stained. By contrast, acridine orange (AO) seems to be suitable to assess cellular integrity. Similarly, changes in membrane asymmetry and DNA fragmentation associated with apoptosis can also be assessed. To confirm successful induction of apoptosis, we performed several established procedures such as DNA fragmentation analysis, acridine orange (AO) staining, and the TUNEL test. 1. DNA Fragmentation Analysis For DNA fragmentation analysis, 107 MNCs were lysed in detergent containing proteinase K. Total cellular DNA was prepared from cell line HL60 using the QIAamp Blood Kit according to the manufacturer’s instructions (QIAGEN, Hilden, Germany) with several minor modifications. Gel electrophoresis was applied to visualize the DNA laddering.

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2.

Acridine Orange Staining

Acridine orange (AO) is a fluorescent dye intercalating into the DNA double helix (40). Following the staining procedure, cells are visualized with an Olympus BH-2 microscope with a fluorescence attachment. 3.

TUNEL Test

The TUNEL test is based on the TdT-mediated dUTP nick end labeling of apoptosis-initiated DNA strand breaks. Those DNA fragments can be identified by labeling free 3′-OH DNA ends with fluorescein (FITC) conjugated nucleotides in an enzymatic reaction catalyzed by the DNA polymerase terminal deoxynucleotidyl transferase (TdT). Incorporation of fluorescein can be detected by flow cytometry (FACS analysis) or by fluorescence microscopy (32). All cell samples were analyzed on a FACScan instrument (Becton Dickinson, Heidelberg, Germany) equipped with a 488-nm argon excitation laser. E.

Spin Labeling

Eppendorf cups including 107 cells were centrifuged for 2 min at approximately 300 ⫻ g. The cell pellet was then resuspended in 50 µL of phosphate-buffered saline (PBS). Spin labeling was performed by adding 1 µL of a 5 mmol solution of 16-doxylstearic acid (16-DSA) in absolute ethanol. The cell pellet was vigorously vortexed for 5–10 s, after which the resuspended cells were incubated with the spin label for an additional 3 min. The surplus 16-DSA was then washed out by the addition of 1 mL PBS, stirring, and centrifugation at 300 ⫻ g (Eppendorf minicentrifuge). The cell pellet was resuspended in 50 µL of PBS and transferred to a 50 µL capillary which was placed in the ESR spectrometer (35,39,41). F. ESR Spectroscopy Electron spin resonance spectroscopy is based on the detection of magnetic spin moments as presented by free electrons. Consequently, the ESR technique requires the presence of molecular structures with stable free electrons. Biological membranes usually do not fulfill this prerequisite. Therefore, spin label molecules have to be introduced into the cellular membrane. Those molecules provide a paramagnetic reporter group with a free electron (1,2). The similarity of the basic structure of these amphipathic molecules, consisting of an apolar fatty acid chain and a polar carboxyl head group, to the structure of the membrane phospholipids enables the spin label to be readily incorporated into the cellular membrane (Figs. 2 and 3). Subsequent to the labeling of the cells, the sample was placed in the ESR spectrometer and simultaneously exposed to a magnetic field of increasing

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Figure 2 Spin label 16-doxylstearic acid (16-DSA) with paramagnetic reporter group.

strength and microwave irradiation of constant frequency. Increasing the magnetic field strength over a well-defined narrow range leads to energy absorption by free electrons, resulting in a characteristic resonance spectrum (1,2,42). The characteristics of this spectrum mainly depend on the degree of mobility of the spin label molecules within the cellular membrane as well as the polarity of the latter (3,6,7,10,11,42). The mobility is related to the degree of order of the cellular membrane and its fluidity. An increase in the degree of order leads to a decrease in fluidity. Figure 4 shows an ESR spectrum with the characteristic triplet form, consisting of three distinct peaks (h⫹1 , h0 , h⫺1) as obtained by a solution of spin label 16-doxylstearic acid (16-DSA) only in phosphate-buffered saline (3,11). In this case, spin label molecules are mobile in all directions of space without any limitation. Figure 5 shows an ESR spectrum of a cell suspension. In contrast to Figure 4, spin label molecules are incorporated into the cellular membrane and their

Figure 3 Schematic model of biological membrane with incorporated spin label 16-DSA.

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Figure 4 Solution of spin label (16-DSA) in phosphate-buffered solution.

Figure 5 ESR spectrum of 16-DSA spin label in cell suspension. a ⫽ apolar; p ⫽ polar.

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mobility is restricted. Increasing restriction of mobility corresponds to broadening and flattening of the curves. ESR spectra allow the quantitative determination of various parameters (11): 1. Order parameter S is calculated from the values T′储 (parallel) and T′⊥ (perpendicular). S measures the degree of order of the cellular membrane. An increase in the degree of order corresponds to a decrease in fluidity. S ranges from 0 (completely fluid) to 1 (immobilized). 2. The partitioning coefficient f can be determined when splitting of the high field peak occurs. It is given by the ratio of the two components of the h⫺1 peak: f ⫽ h⫺1H /h⫺1P. f represents the degree of membrane polarity. It ranges from ⬎0 to less than about 50, the lowest polarity. IV. RESULTS A.

Differentiation Between Malignant and Nonmalignant Cells

The evaluation of the obtained ESR spectra confirmed our assumption that each type of cell investigated in the course of this study can be assigned to a group with characteristic properties of the cellular membrane. Among these, the differentiation between malignant and nonmalignant cells by means of the provisional order parameter S p (see Table 1) was most significant. Within the groups ‘‘malignant’’ and ‘‘nonmalignant,’’ the determination of the polarity shows significant differences between leukemic blast cells and cell lines. Table 1 shows the results of calculating the order parameter S p and parti-

Table 1 Values of Order Parameter Sp and Partitioning Coefficient f Cell source

n

Order parameter Sp a

Partitioning coefficient f

Healthy donors (PBL) Malignant cells (leukemic blast cells) Leukemia-derived cell lines Without treatment CAM treatment Necrosis

23 25 12

0.155–0.170 0.101–0.157 0.101–0.377

0.35–1.05 0.1–1.3 0.998–1.377

12 8

0.13–0.15 b 0.5–0.6

0.21–0.5b 0.798–0.879

PBL ⫽ peripheral blood lymphocytes; CAM ⫽ camptothecin. a According to Gaffney (43), T′储 can be derived from experimental spectra with high accuracy for all order parameters greater than 0.2. There are factors that influence the accuracy of measurements of T′储 . Therefore, data on S are provisional (Sp). b Time of induction with CAM was about 14–28 h.

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tioning coefficient f. The results indicate that the cellular membrane of nonmalignant cells reveals a higher degree of order and is less fluid than that of the malignant group. Within both groups, with the exception of cases of acute myeloid leukemia (AML), no significant differences were observed. A tendency to relatively low fluidity was seen in four out of five cases of AML. The cellular membrane of patients’ leukemic blast cells showed a significantly higher polarity than that of the cell lines (KG1, Molt-4, K562, HL60, U937). No significant differences in polarity could be observed in the group of nonmalignant cells. B. Cellular Membrane Characteristics of Apoptotic and Necrotic Cells Our results are summarized in Table 1 and shown in Figures 6–8 with ESR spectra exemplarily given for a control, an apoptotic, and a necrotic cell suspension. The interpretation of estimated values of characteristic spectra demonstrates that ESR technology enables one to discriminate malignant transformed and nonmalignant peripheral blood lymphocytes (PBLs) derived from healthy donors. As given by order parameter Sp, patients’ leukemic blast cells are more fluid than investigated PBLs of donors. The heterogeneity within the patients group is additionally described by an extensive range in the partitioning coefficient. While donors’ lymphocytes are regularly arrested in the G0 /G1 phase of the cell cycle and therefore are of nearly the same size and morphology, in contrast, leukemic blast cell populations, especially cell lines, seem to be more heterogeneous. The spectrum given in Figure 6, which was obtained with 16-DSA, shows a control sample in an undefined metabolic state. The third line reveals partitioning, which may differ over a range from ⬎0 to ⬍50. In the latter case the polar part of the third line is still just measurable. In different samples, the partitioning coefficient f ⫽ h⫺1H /h⫺1P (i.e., the apolar/polar ratio) may vary considerably due to differing metabolic conditions. Therefore, in viable cells the provisional order parameter Sp and h⫺1H /h⫺1P vary over large ranges of values (see Table 1). Interestingly, cells undergoing cell death (apoptosis and/or necrosis) can be characterized by restricted ranges of the order parameter Sp describing a very fixed, inflexible condition within the population. The 16-DSA spectrum in Figure 7 shows an apoptotic sample. The third line is partitioned much in favor of its polar part (compare Fig. 6 and control). Under these conditions, however, metabolic activity is abolished. Therefore Sp and h⫺1H /h⫺1P show much less variation (see Table 1). Moreover, in necrotic cells we find spectra superimposed by motionally much more restricted components compared to the control or apoptotic samples (Fig. 8; see Table 1 for S ap value). Partitioning of the third line resembles that

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Figure 6 ESR spectrum of 16-DSA spin label incorporated in plasma membrane of control cell suspension.

Figure 7 ESR spectrum of 16-DSA spin label incorporated in plasma membrane of apoptosis-induced cells (U937; CAM incubation for 20 h).

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Figure 8 ESR spectrum of 16-DSA spin label incorporated in plasma membrane of necrotic cell suspension.

found in the control (Figs. 5 and 6). Similar to Figure 7 (apoptosis) and due to abolished metabolism, variation of h⫺1H /h⫺1P is much reduced (see Table 1). The study of the process of apoptosis yielded further interesting results. As expected, cellular necrosis revealed early changes indicating a perturbation of the phospholipid bilayer integrity. This corresponds to a strong increase in cellular membrane polarity, whereas the membrane fluidity was not significantly altered. Incidentally, it is a remarkable phenomenon that in the hydrophobic core of the cellular membrane, necrosis exerts a significant influence on membrane fluidity. Thus, bilayer fragments obviously reveal a probable increase in the density of both lipids and proteins. In apoptosis-induced cells, the start of DNA fragmentation can be monitored by gel electrophoresis after 2 h of induction, and it is completed after approximately 8 h. Accordingly, applying acridine orange staining, apoptosis is clearly recognizable in more than 90% of the cells. ESR spectroscopic studies after 2–8 h do not show any significant changes concerning the cellular membrane. In more than 90% of the cells, the cellular membrane was found to be intact. Earliest significant changes observed by ESR were recognized after 12– 14 h. This resulted in a strong restriction of Sp ranges as well as a strong increase in plasma membrane polarity. Maxima were reached after 16–18 h. Continuing ESR spectroscopic observations led to additional interesting results. After ap-

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proximately 24 h fluidity decreased again, finally reaching a level that is characteristic of necrotic fragments. This result, of course, is not surprising, as the process of apoptosis finally leads to a pattern of damage that no longer allows one to differentiate between apoptosis and necrosis. In this state, the cellular membrane is disintegraed to a large extent.

V.

DISCUSSION

Programmed cell death is a complex phenomenon that can be influenced by a large number of gene products, including the members of the bcl-2 gene family, p53, stress-activated protein kinase, and various death effectors such as the interleukin-converting-enzyme (ICE)-like proteases (21). Important physiological changes that occur during apoptosis include oxidative stress and the depletion of cellular l-γ-glutamyl-l-cysteinylglycine (GSH), phosphatidylserine exposure, and the mitochondrial permeability transition. These functional changes appear to be fundamental regardless of the specific molecular processes that are controlling apoptosis (13,16,20,44–46). Modern techniques of cellular biology and immunology enable scientists to study a great variety of processes concerning living and dead cells. Among these, many processes involve the cellular membrane. In our study we used electron spin resonance (ESR) spectroscopy, a powerful technique to analyze the properties of the cellular membrane of intact cells under various conditions. The evaluation of ESR spectra allows the determination of various parameters such as membrane fluidity, polarity (10,11), and signal reduction (42). In the course of our ESR spectroscopic studies, we were able to reliably discriminate different cell types by means of characteristic properties of the cellular membrane. A clear differentiation between intact vital malignant and nonmalignant cells could be established, although so far the systematic study of cellular membrane properties within the group of leukemias has not yielded an obvious correlation to clinical or immunological features in the case of ALL (27,28,47– 50). The tendency of AML to show relatively high fluidity in the hydrophobic membrane core is subject to further confirmation (data not shown). The considerable increase in membrane polarity in this area observed during the process of apoptosis is an interesting phenomenon. Our results suggest that the fluidization of the cellular membrane is an effort that is bound to the integrity of the phospholipid bilayer. Polarity changes to the extent found in apoptosis have not yet been exhibited under other conditions. Our ESR data have shown that this technique is able to detect cell death at the level of the plasma membrane only at a late stage of apoptosis. In addition, this technique can discriminate to necrotic situation. Moreover, these findings

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indicate that fluidization of the hydrophobic core of the cellular membrane cannot simply be based on membrane damage. Future studies will focus on the changes in the properties of cellular membrane caused by drug treatment, especially by cytostatic drugs.

ACKNOWLEDGMENTS We are grateful to Miss Susanne Hakuba for technical assistance and to Professor Guido Zimmer for helpful discussions and review of the manuscript. We thank Dr. Klaus Zwicker for assistance in ESR technology. This investigation was supported by ‘‘Verein Hilfe fu¨r Krebskranke Kinder Frankfurt/e.V.’’ and by ‘‘Frankfurter Stiftung fu¨r Krebskranke Kinder.’’

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21 Use of Monoclonal Antibodies in Cloning and Identification of Membrane Antigens Marco Bestagno and Oscar Burrone International Centre for Genetic Engineering and Biotechnology, (ICGEB), Trieste, Italy

I. INTRODUCTION Since their discovery, monoclonal antibodies (mAbs) have been looked at as very promising tools to diagnose and possibly treat human tumors. For that reason many mAbs have been developed to identify antigens on the surface of cells that are specific for cancer or that are at least expressed in larger amounts on tumor cells than on normal cells. The identification of membrane-associated antigens may prove of great utility in different fields of study. It may help in the understanding of the mechanisms of cell differentiation and tumor progression, it can shed more light on the processes of cell signalling and cell–cell interactions, and it contributes to our knowledge of membrane structure and composition. Several strategies have been designed to develop mAbs specific for cancer cells and to identify and characterize their antigens. The simplest way is to immunize mice with tumor cells (1,2), or with membrane preparations from tumor cell lines (3,4). To increase the specificity of the immune response for cancer cells, the mice can be previously made tolerant to normal human cells by immunosuppressive treatments following injections of human normal cell membrane extracts (5). An original approach was introduced by Yamashita et al. (6), who raised mAbs against a chemically synthesized artificial carbohydrate antigen whose 387

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characteristics resembled those of cell surface glycoproteins. Of the mAbs obtained in this way, one was shown to be poorly expressed in normal tissues, whereas it reacted strongly with gastric, pancreatic, and colon cancers. The corresponding natural antigen has not been cloned yet but is presently under characterization (7). Once obtained, these tumor-specific mAbs may open the way to a number of different applications and further studies. Cloning of the variable region genes of the antitumor mAbs allows the production of many recombinant therapeutic agents, such as chimeric or humanized antibodies (8), immunotoxins (9,10), different variants of single-chain antibodies (11,12), or bispecific antibodies that can, for example, redirect T-cellmediated cytotoxicity toward tumor cells (13,14). As an example, the discovery of a hybridoma cell line producing an mAb against lung carcinoma (1) allowed the construction of a chimeric antibody and the analysis of the antibody binding site (15), followed by the molecular cloning of the antigen (16), which in turn was used as a target to reshape the antigen-binding site, to obtain higher affinity antibodies (17), and to clone the murine antigen counterpart in order to establish an animal model for that tumor (18). The isolation of another mAb specific for ovarian cancer (5) led to the cloning of the antigen (19) and to the construction of a recombinant antibody fragment (Fab) for immunotherapeutic purposes (20). Monoclonal antibodies specific for membrane-associated antigens can be used to study the interactions between the proteins and the membrane (21,22) or the associations between the tumor antigens and other membrane-bound molecules (23,24), which can lead to a better definition of the biological role of the antigens under investigation and possibly help in designing new strategies of antitumor intervention. Most studies on membrane-associated antigens involve cloning of their cDNAs; knowledge of the amino acid sequence of the cloned proteins is a useful source of information on their possible structures or on their hypothetical functional domains, according to homologies found with other known molecules. Sequence homologies may allow the assignment of a particular antigen to an already described family of proteins (25,26) or the identification of new protein families (2,27); proteins studied by different methods and known under different names may be identified as the same molecule (28,29). The availability of the cloned antigen may allow further studies, such as a fine epitope mapping, for a better definition of the antigen structure and membrane orientation (30), or to identify the regions interacting with B and T cells (31). In addition, cloned antigens can be used as recombinant vaccines to prevent tumors in subjects at risk or to arrest tumor growth in patients at early stages (32,33). An interesting possibility that has gained credence in recent years is represented by the identification of the idiotypes of membrane-bound immuno-

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globulins expressed by B-cell lymphomas as tumor-specific antigens. Cloning of the variable regions of these immunoglobulins allows the preparation of a specific vaccine to induce an anti-idiotypic response in the patient against his own lymphoma: Very promising results have been obtained by the so-called DNA immunization methodology, with plasmids containing the variable region genes of the lymphoma immunoglobulin (34,34a). Various techniques are available to achieve the molecular cloning of the antigen recognized by an mAb. One possibility is to utilize the mAb to purify the antigen. A partial amino acid sequence can then be determined from the purified protein; on the basis of this amino acid sequence, degenerated primers can be synthesized, which can be used to amplify, by reverse transcription and polymerase chain reaction (PCR), a cDNA for the antigen. The amplified fragment (or the primers themselves) may serve in turn as a probe to screen libraries in order to obtain the full-length cDNA or the complete gene (25,29, 35,36). Another possibility is to directly use the mAb as a probe to screen an expression library from λ-phage in bacteria (19,37,38). Even though some proteins are correctly folded in the bacterial environment and the screening procedure may in some cases be carried on in nondenaturing conditions, this approach is usually advisable when the antibody can recognize its antigen in the denatured form, as for example in Western blotting after SDS-PAGE, and does not require posttranslational modifications of the epitope, like glycosylation, which are not provided by the bacterial expression machinery. An alternative and widely used strategy is to transfect a cDNA library in the monkey kidney COS cell line and to select for surface expression of the antigen by panning with the specific mAb (2,16,27,39–41). This latter strategy may present an advantage when the mAb recognizes a conformational or glycosylated epitope, since the antigen is likely to be expressed by a eukaryotic cell in a better folded and posttranslationally modified form than in a bacterial system. To exemplify these techniques, we now report in more detail the cDNA cloning and the biochemical characterization of the antigen recognized by the mAb anti-breast cancer 6C6 (38). The 6C6 mAb was derived from a mouse immunized with a plasma membrane preparation of human breast cancer cells. In immunohistochemical studies it mainly reacted with human breast cancer tissue samples as well as with metastatic lymph nodes of breast cancer patients and, to a lesser extent, with tissue samples from several other human tumors, including fibroadenoma, gastrointestinal carcinoma, and liver carcinoma (3). In immunofluorescence on nonpermeabilized cells, the 6C6 antibody showed a strong cellsurface staining of the human breast cancer–derived cell line MCF7 as well as of cell lines derived from other kinds of human tumors, including the erythroleukemia cell line K-562.

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In Western blotting the 6C6 mAb recognized with high specificity a 28 kDa protein on cellular membrane extracts from the human cell lines MCF7 and K-562, while it was negative on mouse and hamster cellular extracts. The 6C6 antibody had already been used in vivo for the radiodetection of tumor lesions in animal models as well as in clinical trials in patients (42). Because of its high specificity in Western blotting, it was decided to use the 6C6 mAb as a probe to screen an expression cDNA library from K-562 cells in λgt11.

II.

MATERIALS AND METHODS

A.

Cell Lines and Media

Human breast adenocarcinoma cell line MCF7, human erythroleukemia cell line K-562, and Chinese hamster ovary (CHO) cells are all available from American Type Culture Collection, Rockville, MD. The mouse fibrosarcoma cell line T241 was provided by Dr. Zhou Ping, The Cleveland Clinic Foundation, Cleveland, OH. MCF7, K-562, and T241 cells were all grown and maintained in RPMI1640 medium supplemented with 10% fetal calf serum; CHO cells were grown in α-MEM supplemented with 10% fetal calf serum, 40 µM deoxynucleosides, and 40 µM ribonucleosides.

B. cDNA Library Screening An expression library of K-562 cDNA in λgt11 vector (Clontech) was screened with the 6C6 antibody. Plating bacteria were prepared by growing Y1090 E. coli strain overnight in LB medium containing 0.2% w/v maltose and 50 µg/mL ampicillin, and by resuspending bacteria in half the original volume of 10 mM MgSO4 ⫹ 0.2% maltose. Of these plating bacteria, 200 µL were infected for 20 min at 37 °C with 3 µL of bacteriophage λ library diluted to 105 plaque-forming units per microliter (PFU/µL) in SM buffer (50 mM Tris-HCl, pH 7.5; 0.1 M NaCl; 10 mM MgSO4 ; 2% w/v gelatin). Eight milliliters of molten top-agar (0.7% w/v agar in LB medium) containing 10 mM MgSO4 was added to each tube, and the contents of the tubes were then poured onto 15 cm LB-agar plates (plaque density was 3 ⫻ 105 per dish, about 1.7 ⫻ 103 cm⫺2 ). The infected plates were incubated for 4 h at 42 °C, then overlaid with nitrocellulose filters soaked in 10 mM IPTG (isopropylthio-β-d-galactoside) and incubated an additional 4 h at 42 °C. The filters were washed in TBST (10 mM Tris-HCl, pH 7.5; 150 mM NaCl; 0.05% Tween-20) for 20 min, blocked in 5% w/v nonfat dry milk in TBST, and incubated with 6C6-hybridoma supernatant for 2 h at room temperature; after

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washing in TBST, they were incubated 1 h with alkaline-phosphatase-conjugated antibody anti-mouse immunoglobulin, washed again, and revealed with 0.33 mg/ mL NBT (nitro-blue-tetrazolium chloride), 0.165 mg/mL BCIP (5-bromo-4chloro-3-indolyl phosphate) in 0.1 M Tris-HCl pH 9.5, 0.1 M NaCl, and 5 mM MgCl 2 .

C. Phage DNA Preparation and Sequencing Plating Y1090 bacteria (200 µL) were infected with 100 µL of positive phage stocks (about 105 –106 PFU), plated on 9 cm LB-agar dishes in 3.5 mL of topagarose (0.7% agarose), and incubated at 42 °C until plaques were almost confluent. To each plate 5 mL of SM buffer was added, and dishes were shaken overnight at 42 °C. To the collected SM buffer, 100 µL of chloroform was added, then tubes were shaken for 10 min at room temperature and spun down for 5 min at 2000 ⫻ g. To 1 mL of supernatant, DNase I was added at 20 µg/mL and incubated 15 min at room temperature, then 14 µL of 0.3% gelatin and 1 mL of a 70% DE-52 (Whatman) suspension in LB medium were added, shaken for 15 more minutes and centrifuged twice for 5 min at 5000 ⫻ g. One milliliter of supernatant was adjusted to 60 mM EDTA and digested with 40 µg/mL proteinase K for 15 min at 45 °C; 30 µL of a 5% w/v solution of CTAB (hexadecyl trimethylammonium bromide) in 0.5 M NaCl was added, and samples were heated for 3 min at 68 °C, chilled in ice, and centrifuged for 10 min at 8000 ⫻ g; the pellets were resuspended in 300 µL of 1.2 M NaCl and precipitated with ethanol. The cDNA inserts in λ DNA were excised and subcloned in pUC18 for sequencing with the classical dideoxy chain-termination method, using the T7DNA polymerase.

D.

In Vitro Transcription and Translation

The 6C6 antigen coding region from clone 8.4.1 was inserted between the EcoRI and Xbal sites in the eukaryotic expression vector pcDNA3 (Invitrogen). The resulting plasmid (pcDNA-8.4.1) was used for in vitro transcription experiments with the T7-RNA polymerase and for cell transfections. In vitro transcription and translation reactions were done using the TNT-T7 coupled reticulocyte lysate system (Promega), following manufacturer’s directions. A typical 25 µL reaction contained 1.5 µg of pcDNA-8.4.1 plasmid, 12.5 µL of rabbit reticulocyte lysate, 1 µL of reaction buffer, 0.5 µL (about 10 U) of T7-RNA polymerase, 0.5 µL of 1 mM amino acid mixture without methionine, 20 U of recombinant RNAsin (Promega), 20 µCi of [35S]l-methionine 1000 Ci/mmol. After 60 min of incuba-

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tion at 30 °C, 1 µL of the reaction mixture was directly loaded on a 12% SDSPAGE and the remaining was immunoprecipitated with the 6C6 mAb. E.

Cell Transfection

Chinese hamster ovary (CHO) cells and the mouse fibrosarcoma cell line T241 were transfected with the same pcDNA-8.4.1 plasmid used for in vitro transcription. 1.2 ⫻ 106 T241 and 1.4 ⫻ 106 CHO cells were resuspended in 0.5 mL of cold PBS and put in a cuvette for electroporation with an electrode gap of 0.4 cm; 10 µg of Bgl II-linearized plasmid was added to the cells, and electroporation was performed with a single pulse at 960 µF, 290 V, in a Bio-Rad gene pulser equipped with a capacitance extender. Afterward, cells were kept 5 min on ice, washed, resuspended in 30 mL of culture medium, and seeded in 10 cm dishes at a density of approximately 4 ⫻ 105 cells/dish. After 24 h, selective medium containing G-418 (Geneticin, Gibco-BRL) at a final concentration of 400 µg/mL was added. Selected clones were screened by Western blotting analysis on total cell lysates for the expression of the 6C6 antigen. F. Immunofluorescence Analysis MCF7 and the transfected T241 and CHO cells, grown on glass slides until they reached approximately 75% confluence, were washed twice with PBS and fixed for 20 min in 3% w/v paraformaldehyde in PBS. The fixed cells were blocked with 0.25% w/v gelatin in PBS and incubated for 1 h at room temperature with 6C6 hybridoma culture supernatant. After several washes with PBS-gelatin, cells were incubated for 1 h with FITC-conjugated antibody-anti-mouse immunoglobulins. The slides were mounted in fluorescent mounting medium (Kirkegaard & Perry Laboratories) for the microscopic observation. G. Metabolic Labeling and Immunoprecipitation Human breast cancer MCF7 cells were cultured in 60 mm dishes until they were approximately 70% confluent. After washing with PBS, RPMI medium without methionine was added, and cells were incubated for 20 min at 37 °C to deplete the intracellular pool of methionine, then [35S]l-methionine was added to a final concentration of 100 µCi/mL. In glycosylation-inhibition experiments, the Nglycosylation inhibitor tunicamycin was added to the culture medium at 10 µg/ mL. After 2 h labeling, cells were washed and chased with nonradioactive methionine for 3 h. Cells were then washed with cold PBS, lysed in 100 µL of TNN buffer [50 mM Tris-HCl, pH 8.0; 250 mM NaCl; 0.5% NP-40; 1 mM phenyl methyl sulfonyl fluoride (PMSF)] and centrifuged for 10 min at 10,000 ⫻ g at

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4 °C. The supernatants were mixed with 1 mL of 6C6 hybridoma supernatant and incubated for 2 h at 4 °C. In a different set of experiments, intact labeled cells were incubated with 6C6 supernatant for 2 h at 4 °C, then washed three times in PBS and lysed in TNN. In both cases, after antibody incubation, 25 µL of protein A-Sepharose (Pharmacia) was added, and after incubation for 1 h at 4 °C, beads were washed with 4 mL of TNN, 4 mL of TNNB (TNN ⫹ 1% w/v BSA), 4 mL of RIPA buffer (0.1 M Tris-HCl, pH 8.0; 0.1 M NaCl; 5 mM MgCl2; 1% NP-40; 1% deoxycholate; 0.1% SDS), and 4 mL of PBS. Immune complexes bound to protein A were eluted with 50 µL of reducing SDS sample buffer (30 mM TrisHCl, pH 6.8; 1.5% SDS; 2.5% β-mercaptoethanol; 10% glycerol; 0.1 mg/mL bromophenol blue) and loaded onto a 12% SDS-PAGE, which was subsequently dried and exposed for autoradiography.

H. Biotinylation of Membrane Proteins Surface membrane proteins were biotinylated with the Amersham ECL protein biotinylation system, following the directions provided by the manufacturer. MCF7 cells grown in 60 mm dishes were washed twice with cold PBS, then covered with 2 mL of ice-cold biotinylation buffer (40 mM sodium bicarbonate buffer, pH 8.6, 100 mM NaCl). Amersham biotinylation reagent (80 µL) was added, and cells were incubated for 30 min at 4 °C on an orbital shaker. After washing with cold PBS, the cells were lysed and biotinylated proteins were immunoprecipitated as described above, separated in SDS-PAGE, transferred to nitrocellulose, then detected with peroxidase-labeled streptavidin and the chemiluminescent reagent provided with the kit.

I. Epitope Mapping Deletion mutants of the 6C6 antigen were generated by PCR. An oligonucleotide primer complementary to the 3′ end of the coding sequence was used to introduce a HindIII restriction site after the stop codon, while three oligonucleotides hybridizing in different regions of the coding sequence were used to introduce BamHI sites before codons 118, 155, and 194. The three PCR products were digested with BamHI and HindIII and cloned into the BamHI/HindIII sites of the prokaryotic expression vector pUR-289, in frame with the β-galactosidase gene. The resulting plasmids were used to transform E. coli DH5/α cells. The transformed bacteria were grown until an O.D.550nm of 0.4, then expression of the recombinant proteins was induced with 10 mM IPTG. After 2 h induction, cells were collected and lysed in one-tenth of the original volume of reducing SDSsample buffer containing 6 M urea. Two milliliters of bacterial lysates were sepa-

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rated in SDS-PAGE on 7.5% gels and analyzed by Western blotting with the 6C6 mAb. J. mRNA Preparation and Northern Blotting Total RNA was extracted from cells according to the acid guanidinium thiocyanate/phenol/chloroform extraction method of Chomczynski and Sacchi (43). mRNA was prepared from total RNA on oligo-dT-cellulose, using the Pharmacia QuickPrep Micro mRNA purification kit. Approximately 2 µg of mRNA was run on a denaturing 2.2 M formaldehyde, 1.2% agarose gel in morpholino propane sulfonic acid (Mops)/acetate buffer (40 mM Mops, 8 mM CH3COONa, 1 mM EDTA, pH 7), transferred to a nylon membrane by capillary blotting in 20 ⫻ SSC (3 M NaCl, 0.3 M trisodium citrate, pH 7), and cross-linked by UV irradiation. The probe used for hybridization was the EcoRI/Xbal cDNA fragment from pUC18-8.4.1, containing the complete coding sequence of the 6C6 antigen. Standard hybridization and washing conditions were used (44). Filters were rehybridized with a β-actin probe as an internal control. III. RESULTS A.

Isolation of a cDNA Clone Encoding the Antigen Recognized by the 6C6 Antibody

The 6C6 mAb was originally obtained from a mouse immunized with a plasma membrane preparation from human breast cancer cells (3). Immunohistochemical studies on formalin-fixed tissue samples showed a strong reaction with breast tumor cells; reaction with normal tissue, although not completely negative, was much weaker than with tumor tissue, suggesting an overexpression of the 6C6 antigen in cancer cells. In Western blotting the 6C6 antibody recognized with high specificity a 28 kDa protein in cellular membrane extracts from the human breast cancer– derived cell line MCF7, as well as from the erythroleukemia cell line K-562, whereas it was negative on cellular extracts from different mouse and hamster cell lines. Besides human cells, in a cell line of monkey origin (MA104), the 6C6 mAb stained a band of the same 28 kDa molecular weight. Taking advantage of the high specificity shown in immunoblotting by the 6C6 mAb, a λgt11 cDNA expression library from K-562 cells was screened. The 6C6 mAb had an affinity high enough to allow screening of the filters directly with hybridoma culture supernatants, with no need to produce high titer ascitic fluids or to affinity purify the antibody. Twenty 15 cm plates at a density of 3 ⫻ 105 PFU/plate were screened. Five plaques out of 6 ⫻ 106 were found to be positive; they were taken and rescreened twice by reinfection of bacteria at a lower plaque density (about 104 PFU/plate) until two clones were isolated (clones

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8.4.1 and 10.7.2); the two cDNA inserts were cloned and sequenced. Clone 8.4.1 contained a 1008 bp insert, with an open reading frame (ORF) of 738 bp (Fig. 1a), from position 100 to position 837, encoding a protein of 246 amino acids with a predicted molecular weight of 27,991 Da. This finding is in good agreement with the observed molecular weight of the 6C6 antigen in Western blotting on cellular extracts from MCF7 and K-562 cells. Clone 10.7.2 contained a 511 bp insert that started at position 331 and ended at position 841 of the 8.4.1 sequence, just after the stop codon in the identified ORF. Sequence analysis of the predicted protein revealed two distinct regions: a mainly hydrophobic amino-terminal region and a hydrophilic carboxy-terminal region. In the amino-terminal region three highly hydrophobic sequences of 18, 17, and 21 amino acids are present (H1, H2, H3, respectively, in Fig. 1b) and could represent either transmembrane domains, or, alternatively, H1 may represent a signal leader peptide for translocation into the endoplasmic reticulum. In the hydrophilic carboxy-terminal region Asn181 was identified as a possible N-glycosylation site. B. Biochemical Characterization of the 6C6 Antigen To confirm that the cloned cDNA actually encoded the 6C6 antigen, the 8.4.1 cDNA was cloned into the eukaryotic expression vector pcDNA3. In this vector the inserted DNA is under the control of the cytomegalovirus (CMV) promoter, for expression in eukaryotic cells, and of the T7 prokaryotic promoter. Therefore, the same plasmid can be used for both in vitro transcription/translation and cell transfections. In the first experiment the pcDNA-8.4.1 plasmid was used for in vitro transcription and translation using T7-RNA polymerase and the rabbit reticulocyte lysate system. The in vitro translation mixture was then immunoprecipitated with the 6C6 mAb and analyzed on a reducing SDS-PAGE. Figure 2 (lanes 1 and 2) shows that a 28 kDa protein was synthesized and specifically immunoprecipitated by the 6C6 antibody. Moreover, the mobility of the immunoprecipitated in vitro translated protein is coincident with that of the protein immunoprecipitated from extracts of MCF7 cells labeled in vivo with [35S]methionine (Fig. 2, lane 3), thus confirming its identity with the 6C6 antigen. It was also investigated whether the 6C6 antigen was glycosylated at the putative N-glycosylation site at Asn181 . This was done by labeling MCF7 cells with [35S]methionine in the presence or absence of tunicamycin (a known inhibitor of N-glycosylation), followed by immunoprecipitation of the labeled products with the 6C6 mAb. As shown in Figure 2 (lanes 3 and 4), no difference in the molecular weights of the immunoprecipitated proteins with or without tunicamycin was obtained. Furthermore, when the 6C6 antigen was first immunopre-

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Figure 2 SDS-PAGE analysis of in vivo produced and in vitro translated 6C6 antigen. The cDNA clone 8.4.1 was subcloned in the vector pcDNA3, under the control of the T7-RNA polymerase promoter, and used for an in vitro transcription and translation experiment. Lanes 1 and 2 show the product of the in vitro translation before (lane 1) and after (lane 2) immunoprecipitation with 6C6 mAb. Lanes 3 and 4 show the proteins immunoprecipitated with 6C6 mAb from [35S]methionine-labeled extracts of MCF7 cells grown without (lane 3) or with (lane 4) the N-glycosylation inhibitor tunicamycin. In lane 5 the immunoprecipitated material shown in lane 3 was treated with the endoglycosidase PNGase-F before SDS-PAGE analysis. SDS-PAGE was in a 12% polyacrylamide gel. The positions of the molecular mass standards are indicated. (From Ref. 38.)

Figure 1 Sequence and hydrophobicity of 6C6 antigen. (a) The sequence of the cDNA isolated with the 6C6 mAb is shown, with the amino acidic translation of the identified ORF below. The three hydrophobic regions are underlined, and the putative N-glycosylation site at position 181 is circled. The sequence of the 6C6 antigen cDNA is available from the EMBL database under accession number X81109. (b) Hydrophobicity plot of the predicted 6C6 antigen protein, showing the three hydrophobic regions and putative transmembrane domains H1, H2, and H3. On the vertical axis are represented relative hydrophobic units (H.U.), according to Kyte and Doolittle (51). (Modified from Ref. 38.)

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cipitated from [35S]methionine-labeled MCF7 extracts and then digested with the enzyme PNGase-F, which hydrolyzes both high mannose and complex type Nglycan chains from glycoproteins (45), no differences were visible in the molecular weight of the 6C6 antigen (Fig. 2, lane 5). These results, together with those of the in vitro translation, which show no difference between the molecular weights of the in vitro translated and natural proteins, provided convincing evidence that the 6C6 antigen was not N-glycosylated. In addition, the identity of the in vivo and in vitro products of the cloned cDNA with the natural protein indicates that the first hydrophobic region (H1) is not a cleavable leader peptide and that it could represent a membrane signal-anchor sequence. The same pcDNA-8.4.1 plasmid was used to transfect Chinese hamster ovary cells and the mouse fibrosarcoma cell line T241, which were known to be negative for 6C6 antigen expression. After selection with G-418, several transfectant clones were obtained. In Western blotting analysis on total cellular lysates from those clones, the 6C6 antibody clearly recognized the same 28 kDa band detected in MCF7 and K-562 cell extracts (Fig. 3). The transfected cells also showed positive immunofluorescent staining when assayed with the 6C6 mAb

Figure 3 Western blot analysis of total cell extracts. Total extracts from the cell lines K-562, MCF7, T241, and CHO, as well as T241 and CHO clones transfected with the 6C6 antigen cDNA (T241/6C6 and CHO/6C6), were separated on 12% SDS-PAGE, blotted onto a nitrocellulose membrane, and reacted with 6C6 mAb. The positions of the molecular mass standards are indicated. (From Ref. 38.)

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Figure 4 Immunofluorescent detection of the 6C6 antigen. Cells from the MCF7 line (a) or the T241 and CHO clones transfected with the 6C6 antigen (b and c, respectively) were fixed in paraformaldehyde, which preserves membrane integrity, and reacted with 6C6 mAb and a fluoresceinated antimouse IgG antibody. The positive staining of the transfected clones indicates the presence of the newly synthesized 6C6 antigen on the cell surface. (Courtesy of Dr. Erqiu Li, Peking University, P.R. China.) (bar ⫽ 10 µm).

in nonpermeabilizing conditions (Fig. 4), thus confirming that the protein synthesized by the transfectants is correctly expressed on the cell surface.

C. Localization and Orientation of the 6C6 Antigen Positive immunofluorescence staining of nonpermeabilized MCF7 and transfected T241 cells suggested a membrane-specific localization of the 6C6 antigen, with the epitope recognized by the antibody exposed on the outer side of the plasma membrane. A better characterization of the antigen localization was obtained by two sets of immunoprecipitation experiments. In the first case, MCF7, K-562, T241, and 6C6 antigen-transfected T241 cells were surface-biotinylated, lysed, and immunoprecipitated with 6C6 mAb. In the second experiment, MCF7, T241, and transfected T241 cells, metabolically labeled with [35S]methionine, were incubated, while still intact, with 6C6 antibody and washed to remove unbound antibody. Cells were then lysed, and immunocomplexes were recovered by precipitation with protein A-Sepharose. As shown in Figure 5, the 6C6 mAb specifically immunoprecipitated a 28 kDa protein from either the surface-biotinylated or the [35S]methionine-labeled MCF7, K-562, and transfected T241 cells, but not from the original T241 cells. These results confirm that the 6C6 antigen is a component of the plasma mem-

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Figure 5 Detection of surface membrane protein. (a) The indicated cells were surface biotinylated, lysed, and immunoprecipitated with 6C6 mAb and protein A-Sepharose (⫹) or with protein A-Sepharose alone (⫺). Immunoprecipitates were analyzed by SDS-PAGE and blotted onto nitrocellulose, and biotinylated proteins were detected with peroxidaselabeled streptavidin and chemiluminescent reaction. (b) The indicated cell lines were labeled with [35S]methionine and reacted with 6C6 mAb (⫹) or with a nonimmune mouse serum (⫺). The cells were lysed, immunoprecipitated with protein A-Sepharose, and analyzed on 12% SDS-PAGE. The arrow on the left of each panel identifies the position of the 28 kDa band. The positions of the molecular mass standards are shown. (From Ref. 38.)

brane exposed on the cell surface. In addition, its correct transport to the cell membrane in transfected cells indicated that this is an intrinsic property of the 6C6 antigen protein. The reactivity of the λgt11 clone 10.7.2 with the 6C6 mAb, which contained the sequence of the 6C6 antigen cDNA starting from codon 78 (Fig. 6a), suggested that the epitope recognized by the 6C6 mAb resided in the carboxy-

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Figure 6 Epitope mapping of the 6C6 antigen. (a) Schematic representation of the coding sequences of the two λgt11 clones that showed a positive reaction with the 6C6 mAb. (b) Three fusion proteins were produced that contained three different truncated forms of the 6C6 antigen fused with β-galactosidase. The three fusion proteins were expressed in bacteria, and the bacterial extracts were separated in 7.5% SDS-PAGE and reacted in Western blot with the 6C6 mAb. Lane 1, β-gal-6C6-(119-246); lane 2, β-gal-6C6-(156246); lane 3, β-gal-6C6-(195-246). Unfused β-galactosidase was included as a control (lane 4). The positions of the molecular mass standards are indicated. (Modified from Ref. 38.)

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terminal end region of the protein. To precisely map its position, we constructed a series of β-galactosidase fusion proteins carrying deletions of different lengths in the amino-terminal portion of the 6C6 antigen protein. Proteins β-gal-6C6(119-246), β-gal-6C6-(156-246), and β-gal-6C6-(195-246), having deletions between residues 1–118, 1–155, and 1–194, respectively, were all positive in Western blotting when reacted with the 6C6 mAb (Fig. 6b). This allowed us to conclude that the 6C6 epitope is located in the carboxy-terminal hydrophilic domain of the protein, between residues 195 and 246. Since the hydrophilic C-terminal half of the molecule should be exposed on the cell surface, and since no cleavage of any of the hydrophobic domains takes place, the mature 6C6 antigen protein appears to be a multiple-spanning membrane protein, with either two (H1 and H2) or three (H1, H2, and H3) hydrophobic transmembrane domains, anchoring the protein to the cell surface, as illustrated in the models presented in Figure 7.

D.

Expression of the 6C6 Antigen

The expression of the 6C6 antigen mRNA was investigated in the tumoral cell lines MCF7 and K-562 as well as in a panel of normal human tissues (pancreas, kidney, skeletal muscle, liver, lung, placenta, brain, and heart) by Northern blot hybridizations. Figure 8 shows that in all cases a single band of about 1.5 kb is

Figure 7 Orientation of the 6C6 antigen protein in relation to the cell surface. The 6C6 antigen is presented with the extracellularly exposed carboxy-terminal half and the aminoterminal end either intracellular (a) or extracellular (b), depending on whether the third hydrophobic region (H3) is or is not used as a transmembrane domain. (From Ref. 38.)

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Figure 8 Northern blot hybridization of mRNAs from different human cell lines and normal tissues. mRNAs were extracted from the indicated human normal tissues and cell lines, separated in a denaturing agarose gel, and blotted to a nylon membrane. Upper panels show hybridizations with the cDNA probe of the 6C6 antigen (6C6-Ag); a single 1.5 kb band is observed in all samples. Lower panels show hybridizations of the same filters with a β-actin probe; heart and skeletal muscle contain a second β-actin isoform of 1.8 kb. From the relative intensity of the signals with the 6C6 antigen probe compared to β-actin, the 6C6 antigen mRNA seems expressed at a higher level in the breast cancer derived cell line MCF7 and in normal pancreas. (From Ref. 38.)

obtained. When compared with the β-actin control, higher expression levels of the 6C6 antigen mRNA can be observed in the breast cancer MCF7 cells and in the normal pancreatic tissue.

IV. DISCUSSION The isolated cDNA encoding the human cell surface antigen recognized by the 6C6 mAb contains an ORF of 738 bp that encodes a protein of 246 amino acids, with a predicted molecular weight of 28 kDa. This value is in good agreement with the observed molecular weight of the band recognized by the 6C6 mAb in Western blotting on cellular extracts from MCF7 and K-562 cells. The identity of the cDNA clone was confirmed by in vitro translation experiments, followed by immunoprecipitation of the synthesized protein by the 6C6 mAb. These experiments resulted in the isolation of a protein indistinguishable from the one immunoprecipitated from the MCF7 cell extract (Fig. 2). Furthermore, transfections of CHO cells and mouse fibrosarcoma cell line T241, which

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do not express the 6C6 antigen, resulted in transfected clones expressing a 28 kDa protein, recognized by the 6C6 antibody in Western blotting and immunoprecipitation (Fig. 3) as well as after immunofluorescent staining (Fig. 4). Sequence analysis of the predicted protein identified two distinct regions: a hydrophilic carboxy-terminal region and an amino-terminal one containing three well-defined hydrophobic sequences, which may correspond to three transmembrane domains. The first hydrophobic region was not excised from the mature protein after transport to the endoplasmic reticulum, as occurs with classical signal peptides of type I transmembrane proteins, suggesting that it can constitute a membrane signal-anchor sequence. Biotin labeling of plasma membrane proteins and immunoprecipitation with the 6C6 mAb demonstrated the surface membrane localization of the 6C6 antigen (Fig. 5). This association with the plasma membrane was also confirmed by additional evidence such as immunofluorescent staining of nonpermeabilized cells with the 6C6 antibody (Fig. 4) and enrichment of the 28 kDa band in the membrane fraction after subcellular fractionation. In addition, the 6C6 epitope was mapped to the last 50 carboxy-terminal amino acids by Western blot analysis of deletion mutant proteins (Fig. 6). All these experiments clearly indicated that the 6C6 antigen is a type II membrane protein with its carboxy-terminal end oriented to the outside of the cell. The observation that the first hydrophobic region is not cleaved from the mature protein is in agreement with this orientation, since type II membrane proteins do not have cleavable amino-terminal signal sequences (46). These observations and the identification in the protein sequence of three hydrophobic regions are suggestive of a model where the 6C6 antigen is oriented with the amino-terminal end inside the cell and the carboxy-terminal end outside, after three spans of the membrane (Fig. 7a). An alternative model, given the lower hydrophobicity of the third putative transmembrane domain H3, contains only two transmembrane domains (H1 and H2) and has both the amino- and carboxy-terminal ends oriented outside the cell (Fig. 7b). The model with three membrane-spanning domains in the amino-terminal region and a long extracellular carboxy-terminal domain is particularly suggestive because computer searching in protein sequence data banks identified some weak homologies with a family of transmembrane proteins to which several tumorassociated antigens belong. Members of this family are the human surface antigens CD37 (47), CD9 (35), and CD63 (also known as the melanoma-associated antigen ME491) (28,29,48) ; the lymphoma cell-surface protein, target of an antiproliferative antibody, TAPA-1/CD81 (2); the lymphocyte and monocyte cell surface marker lA4 (39); the motility-related protein MRP-1, found in several human tumors (37); the tumor-specific antigens L6 (16) and CO-029 (41); the rat leukocyte antigen MRC OX-44 (49); and the Schistosoma mansoni membrane antigen SM23 (50).

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All members of this family appear to have a similar structure, with three closely spaced amino-terminal hydrophobic domains followed by a hydrophilic extracellular domain of variable length and a fourth hydrophobic domain located at the carboxy-terminal end of the protein. Except for the fourth hydrophobic domain, these features are shared by the 6C6 antigen, which could be tentatively assigned to this family, as a distantly related member. Little is known about the possible functions of the 6C6 antigen; the protein has been found overexpressed in tumor tissues and in human cell lines. Interestingly, Northern blot experiments on normal human tissues revealed a comparable expression level of the 6C6 antigen mRNA in kidney, skeletal muscle, liver, lung, placenta, brain, heart, and K-562 cells, while a higher expression level can be observed in pancreas and in the MCF7 cells (Fig. 8). However, it remains to be determined whether the overexpression observed in tumors is due to increased transcription or transport and/or to the stability of the 6C6 antigen protein. In addition, the similarities found with a protein family whose members are involved in controlling cellular proliferation (such as CD37 or TAPA-1/CD81) or overexpressed in tumor cells (such as L6, CD63/ME491, and CO-029) suggest that the 6C6 antigen may also play a role in the control of cell proliferation. REFERENCES 1.

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22 Polycystins Membrane-Associated Proteins Involved in Autosomal Dominant Polycystic Kidney Disease Katherine W. Klinger and Oxana Ibraghimov-Beskrovnaya Genzyme Corporation, Framingham, Massachusetts

I. IDENTIFICATION OF THE PRIMARY DEFECT OF AUTOSOMAL DOMINANT POLYCYSTIC KIDNEY DISEASE A.

Clinical Aspects and Etiology of Autosomal Dominant Polycystic Kidney Disease

Autosomal dominant polycystic kidney disease (ADPKD) is one of the most common genetic disorders, with an estimated incidence of 1 in 1000, accounting for 8–10% of all cases of end-stage renal disease (ESRD) (1–3). This disease is generally a late-onset disorder, manifesting in the fourth decade. A variety of studies have shown that in affected individuals the mean and median ages at renal failure occur between 51 and 60 years (4–6). The disease is primarily characterized by the progressive development and enlargement of fluid-filled cysts in the kidney. The development of renal failure is thought to result from the progressive enlargement of cysts that cause compression and loss of the normal renal parenchyma (fibrosis) and functioning nephrons (7,8). ADPKD is a multisystem disorder that leads to the formation of cysts in other organs such as the liver, pancreas, and spleen. Associated cardiovascular abnormalities include valvular abnormalities, cardiac hypertrophy, and intracranial vascular aneurysms (9). Among the complications accompanying ADPKD are hypertension, hematuria, acute and chronic pain, and urinary tract infection.

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B. Genetic Heterogeneity of ADPKD Autosomal dominant polycystic kidney disease is phenotypically and genetically heterogeneous. Variation occurs in age of onset and rate of progression as well as severity within kindreds. For example, although primarily an adultonset disease with ESRD occurring in late middle age, there are still about 25% of affected individuals who retain adequate kidney function at that age. Similarly, while most individuals present with clinical symptoms as adults, a number of cases have been described as presenting clinical symptoms in utero (10). Genetic heterogeneity exists, with 85% of cases caused by mutations in the PKD1 gene, which maps to 16p13.3 (11). Most of the remaining ADPKD cases can be attributed to the PKD2 locus, which maps to 4q12-22 (12). Clinical manifestation of the two genetic defects are very similar, although there is a trend toward slower progression to ESRD in families with mutations in PKD2. There are a small number of clinical cases that are not linked to either the PKD1 or PKD2 locus, indicating the existence of a third locus for ADPKD that has not been mapped yet.

C. Structure of the PKD1 Gene 1.

PKD1 Gene Is Duplicated

Originally mapped to chromosome 16 in 1985 by Reeders et al. (11) using linkage analysis, the PKD1 gene-containing region was eventually narrowed down to a 700 kb interval and a long-range physical map was produced (13,14). The region surrounding the PKD1 locus was also cloned as a 700 kb P1 contig in an effort to identify the gene (15). This genomic area is extremely gene-rich: at least 20 genes are located in the interval. Analysis of a key mutation by the European Polycystic Disease Consortium, which involved a translocation that interrupted the gene, allowed unequivocal identification of the PKD1 gene (16) (Fig. 1). Initial efforts to reveal the complete cDNA sequence were complicated by the extremely complex organization of the PKD1 locus. Sequences related to the PKD1 transcript appeared to be duplicated at least three times on chromosome 16, proximal to the PKD1 locus (Fig. 1), so that only the 3′ ⬃3.8 kb of the cDNA are truly unique (16). The remainder of the transcript shares significant (⬎95%) homology with the duplicated transcripts (17). Thus, it was very important to identify the complete sequence of the PKD1 genomic locus in order to have a reference standard for putative cDNAs. Genomic sequence was reported independently by three different groups (18–20). According to these studies, the 53 kb PKD1 genomic region is organized into 46 exons encoding an ⬃14 kb mRNA whose 3′ end lies close to the 3′ end of the tuberous sclerosis (TSC2) gene (Fig. 1).

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Figure 1 Chromosome 16 location of the PKD1 gene. Unique regions of the PKD1 gene and TSC2 genes are indicated with arrows. The duplicated region of the PKD1 gene is shown by striped boxes. (Based on information from Ref. 16.)

2. A Large Polypyrimidine Tract in the PKD1 Gene Sequence analysis of the 53 kb PKD1 genomic region reveals a number of unusual features that pose various sequencing and presumably biological challenges (19). The overall GC content of the segment is 62.5% with a CpG/GpC ratio of 0.485, at least twice that observed for total human DNA. The frequency of CpG islands is ⬃1 per 5 kb, about 12 times that of the genome average (1 per 67 kb) (21). Twenty-three Alu repeats are clustered across ⬃53 kb, while all other middle repeats are absent. Several simple sequence repeats are present, including two dinucleotide repeats, one tetranucleotide, and 17 tandem copies of a perfect 27 bp repeat. Most interesting is the 2.5 kb GC-rich segment contained within intron 21, wherein the coding strand is ⬃97% CT (22). These polypyrimidine tracts are known to form an alternative DNA structure, H-DNA, composed of a triple helix and influenced by pH, supercoiling, or heavy metals (23). Triplex formation by homopyrimidine sequences can inhibit replication and transcription (24,25). For example, the autosomal recessive disease Friedreich ataxia results from the expansion of a GAA triplet repeat. Repeat expansion at this locus generates a homopurine/homopyrimidine repeat, which interferes with transcription in vitro (26). Presumably, the presence of this long 2.5 kb polypyrimidine tract in the PKD1 gene can interfere with replication, transcription, or RNA processing. Most known human homopurine-homopyrimidine sequences are relatively short, from 30 to 200 bp (27–29). The genomic fragment with this repeat is in the duplicated region of the PKD1 gene in 16p13.1. This region of chromosome 16 is known to participate in translocations and deletions, suggesting that repeat motifs may play a role in the relative chromosomal instability of this region.

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II.

PRIMARY STRUCTURE OF POLYCYSTIN-1 PREDICTS A LARGE TRANSMEMBRANE PROTEIN WITH CELL ADHESION MOTIFS

A.

Full-Length PKD1 cDNA

Although it is unusual to carry out complete genomic sequencing of a disease gene in advance of cDNA characterization, this was a necessary step in the case of the PKD1 transcript. Identification of cDNAs that are specific for the 5′ end of the PKD1 locus is particularly difficult, because multiple transcribed copies of these sequences are also present at 16p13.1 (16). Therefore, direct comparisons of potential PKD1 cDNAs and genomic sequence are required to definitively map a cDNA to the PKD1 locus. Several groups used different strategies to identify PKD1 transcripts (18– 20). Hughes et al. (18) described the use of sequences derived from cDNAs from the homologous loci to predict the likely positions of PKD1 exons and then, using primers from these exons, to generate cDNAs from a somatic cell hybrid cell line 145.19 that contains only the bona fide PKD1 locus. This ‘‘exon linking’’ strategy generated a number of partial cDNAs that, in aggregate, were likely to span the PKD1 locus; however, no contiguous cDNA clone was recovered. In a related manner, the IPKD Consortium used the sequence derived from three authentic PKD1 partial cDNAs, 14 partial cDNAs from homologous loci, and a corresponding PKD1 genomic sequence to predict the PKD1 cDNA sequence (20). The APKD Consortium analyzed the genomic sequence with the GRAIL2 program to predict the position of exons (19). Finally, with the genomic sequence as reference template, cDNA clones recovered by both cDNA library screening and RT-PCR were used to physically assemble a stable 14 kb full-length PKD1 cDNA (30). B. Predicted Structure of Polycystin-1 The PKD1 14 kb cDNA encodes a novel, large protein, polycystin, with a predicted molecular weight of at least 462 kDa. It contains a number of recognizable protein motifs and numerous sites for glycosylation (Fig. 2). The predicted extracellular region of the protein starts with a signal sequence followed by two leucine-rich repeats (LRRs), flanked by characteristic cysteine-rich regions, which have been shown to be involved in protein–protein interactions (31). There are also two motifs indicative of protein–protein/carbohydrate binding, a C-type lectin domain, and an LDL-A domain that is homologous to the low-density lipoprotein receptor (18). The large portion of the polycystin-1 extracellular domain is organized into 16 PKD domains similar to domains found in cell-adhesion molecules and receptors (17). In the polycystin-1 molecule, the first PKD domain is localized between the LRRs and the C-type lectin domain. The rest of the 15 PKD domains are clustered in the middle part of the molecule. Originally thought

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Figure 2 The predicted structure of polycystin-1. (Based on information from Refs. 18, 33, and 35.)

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to be members of the immunoglobulin (Ig) superfamily, more recent work suggests that while the PKD domains contain an Ig-like fold, they represent a novel family rather than being true members of the Ig superfamily (32). Downstream of the PKD domains four FNIII (fibronectin repeats) were originally described (18). These domains are often found in extracellular matrix proteins and cell adhesion molecules. However, more recent modeling has replaced the FNIII-like domain with a newly recognized module consisting of about 700 amino acids (33). This new domain, called REJ, has 20% sequence identity and 40% sequence similarity to a region in the sea urchin sperm egg jelly receptor (REJ). Analysis of hydrophobicity predicts several potential transmembrane regions (TMs) in the C-terminal region. Initially 11 such segments were described (13), where the Nterminal end of polycystin-1 is extracellular and the C-terminal tail is located inside the cell. Other groups have hypothesized that polycystin spans the membrane only three times, since a membrane-spanning segment requires 20 amino acids in α-helix conformation (34). To refine the structure of polycystin-1, comparative analysis of the Fugu and human PKD1 genes was performed (35). Key structural motifs, originally described for human polycystin-1, were maintained in Fugu, but the structure of the membrane-spanning region was refined. This revised model of polycystin also includes 11 TM domains but redefines positions of these regions (35). In the cytoplasmic tail, phosphorylation sites for both tyrosine kinase and protein kinase C are present (18) as well as a coiled coil domain (36). Interestingly, analysis of Fugu polycystin-1 also predicts a possible coiled coil domain in the corresponding region (35). Based on this unique arrangement of known structural motifs present in the polycystin-1, a likely function of polycystin is to mediate cell–cell/matrix interactions or possibly play a role in signal transduction.

III. INSIGHTS INTO NORMAL POLYCYSTIN FUNCTION A.

In Vitro Expression of Polycystin-1

The primary structure of polycystin predicts a large integral membrane protein with multiple cell recognition motifs, but its function remains unknown. Insight into polycystin’s normal function and its role in the development of ADPKD requires the assembly of an extensive collection of molecular reagents to examine its expression and to create model systems for functional studies. Development of these crucial reagents has been complicated due to the presence of transcriptionally active homologous loci as described above. To generate an authentic fulllength PKD1 transcript, several cDNAs were recovered from the 145.19 radiation hybrid cell line, which contains only the PKD1 locus but no homologous loci. In addition, PKD1 cDNAs were cloned from brain and kidney cDNA libraries by RT-PCR or library screening (30). By using appropriate portions of seven

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different cDNAs, a complete, full-length cDNA was constructed in our laboratory. To examine the protein product of the assembled PKD1 cDNA, polycystin was translated in vitro and shown to be of predicted molecular mass of ⬃400 kDa. Interestingly, several alternatively spliced PKD1 mRNAs with different exon combinations have been found in 145.19 cells. The physiological significance of these isoforms is not clear but is noteworthy, as distinct domains of polycystin (such as both LRRs and the C-type lectin domain) are encoded by separate exons. Such a correlation of structure and possible function suggests that differential functions of polycystin in different cell types or tissues could be regulated by alternative splicing. Shorter transcripts corresponding to PKD1 were detected by Northern blot analysis of brain, and alternatively spliced forms of the murine PKD1 transcript have been described (37). Determination of the potential function of these species awaits further study. B. Endogenous Expression of Polycystin-1 in Normal and Cystic Tissues Expression of PKD1 mRNA was detected in kidney, liver, and brain and in a wide variety of cell lines, being particularly abundant in astrocytoma and primary fibroblast cell lines (16). Antipeptide antibodies have been used by several groups to examine tissue expression of polycystin-1, producing somewhat conflicting reports (38–40). Localization to normal and cystic renal tubular epithelial cells appears consistent across groups but with differing patterns of distribution. Differences between frozen and fixed tissues are pronounced, and a consensus on the expression pattern of polycystin has not yet been reached. Successful cloning of an authentic cDNA allowed us to raise antibodies against functionally important domains of polycystin, such as the LRRs of the presumed extracellular domain as well as the C-terminal segments that may mediate intracellular interactions (30). In these experiments, fusion proteins rather than synthetic peptides were used as antigens. In accordance with previous reports, our laboratory has demonstrated polycystin expression in normal and cystic renal tubular epithelium using fixed tissues, but it is confined to the distal convoluted tubules (DCTs) and collecting ducts (CDs). No glomerular, proximal convoluted tubular (PCT), or interstitial staining is seen. Polycystin expression in fresh tissues confirms the restricted distribution of polycystin along the nephron and demonstrates glomerular and peritubular capillary endothelial cell staining in addition to that seen in normal arterioles. Expression in the proximal nephron is observed only after injury-induced cell proliferation. Polycystin expression is confined to ductal epithelium in liver, pancreas, and breast and restricted to astrocytes in normal brain. In all tissues examined, except brain, polycystin expression is confined mainly to epithelial cells. This

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cellular distribution supports a role for polycystin in the developmental regulation of epithelial cell differentiation and morphology. Interestingly, polycystin expression in PCT epithelial cells correlates with their proliferative activity in that expression is seen after PCT injury and in culture. In this tissue polycystin may have a dominant role in cellular differentiation rather than in maintenance of the mature phenotype. In fetal liver, expression is confined to the ductal plates from which mature biliary epithelium is derived and is not seen in developing or mature hepatocytes. This concurs with the localization to biliary epithelium in adult liver. Whether polycystin is required for initiation and/or maintenance of cellular differentiation will require more detailed studies. The role of polycystin in endothelial cells and astrocytes remains unknown, although it is possible that abnormal endothelial cell function may contribute to the development of vascular abnormalities such as hypertension and saccular aneurysms by interfering with normal endothelial cell matrix regulation in a manner analogous to the abnormal matrix seen in ADPKD kidneys (41). In all tissue sections examined in our laboratory, cytoplasmic staining was also accompanied by clear and consistent membrane accentuation. C. Evidence for Membrane Association of Polycystin-1 Using antibodies produced in our laboratories, subcellular polycystin-1 localization was examined in cultured tubular and endothelial cells (HUVECs) using confocal microscopy and image reconstruction in two planes (Fig. 3). A punctate linear pattern of staining at the cell borders was seen at the lateral cell junctions in both cell types using laser scanning confocal microscopy (Figs. 3c and 3d). No apical or basal staining was seen. In subconfluent cultures, staining was confined to points of cell–cell contact (Fig. 3c). Free cell borders did not express polycystin. Platelet endothelial cell adhesion molecule 1 (PECAM-1 or CD-31) is known to stain cell junctions in HUVECs (41). Polycystin staining colocalized with PECAMs, confirming the presence of polycystin at cellular junctions (Figs. 3e and 3f). Identical results were obtained with cells derived from renal CDs, which appeared to have the highest level of expression. The linear pattern of staining was completely abolished with preabsorption of the antibodies (Fig. 3b). The nonspecific diffuse cytoplasmic and nuclear staining observed was shown to be due to the secondary antibodies (Fig. 3a). Therefore, when cultured cells made cell–cell contact, polycystin was localized to the lateral membranes of cells in contact and not to free cell borders (30). These data support the structural predictions of polycystin being an integral membrane protein. Studies by other workers using immunofluorescence with confocal imaging and immunoelectron microscopy suggest discrete localization to small structures in the peripheral cytoplasm with apical accentuation and no surface membrane localization (40). Unfortunately, analysis of subcellular fractions by Western blot has proven difficult

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Figure 3 Polycystin-1 expression in cultured umbilical vein endothelial cells (HUVECs). (a) HUVECs stained with secondary antibody alone demonstrating the nonspecific pattern of nuclear and cytoplasmic staining. (b,c) HUVECs stained with anti-polycystin-1 antibody with or without preabsorption with immunizing polypeptide demonstrating the specific punctate linear expression pattern of polycystin. (e) Staining with PECAM—a lateral membrane marker showing colocalization with polycystin to the lateral membrane. (d,f) Vertical sections of (c) and (e) also show the colocalization of polycystin and PECAM to the lateral membranes with no apical or basal staining. (From Ref. 30.)

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because all reported antibodies detect a number of immunoreactive bands, indicating either high levels of degradation during sample preparation or the presence of multiple isoforms in cells.

IV. MOLECULAR MECHANISM OF CYSTOGENESIS: ROLE OF POLYCYSTIN A.

Evidence for ‘‘Loss of Function’’ Hypothesis

Polycystic kidney disease is an autosomal dominant disorder. In theory, all somatic cells should carry the same mutant allele as well as one normal allele. Several aspects of the natural history are difficult to reconcile with this assumption. First, kidney cysts originate in only 2–5% of nephrons along the renal tubules and then progressively increase in size and number until kidney function is compromised. In addition, clinical heterogeneity exists not only between affected individuals from different families but also within a single family, which should carry the same germline mutation. Furthermore, it has been suggested that anticipation might be an explanation for cases of childhood ADPKD in subsequent generations of families with an adult form of the disease (42). Taken together, these observations suggest that cystogenesis might be a two-step process. Qian et al. (43) suggested that the focal nature of cyst development may be due to a ‘‘second hit.’’ By analysis of polymorphic markers using DNA derived from epithelial cells from single renal cysts, those authors demonstrated that renal cysts in ADPKD are monoclonal. Most important, loss of heterozygosity was demonstrated within individual cysts for two closely linked polymorphic markers within the PKD1 gene, with the ‘‘second hit’’ occurring in the normal allele. This model is very attractive, because it explains the multiplicity of second hits and provides an explanation for the progressive development of new cysts in the affected individual throughout his or her lifetime. Similar results were obtained by another group (44), and subsequent studies suggested that at least some ‘‘second hits’’ arise by gene conversion (45). Authors have described the genetic basis of hepatic cystogenesis in ADPKD, suggesting that the two-hit mechanism may be responsible for a second focal manifestation of the disease (46). While the two-hit hypothesis is very attractive, and it seems clear that somatic mutations occur in the PKD1 gene at an appreciable frequency, a definitive causative role in cystogenesis has yet to be proven. In collaboration with Dr. Sanford’s laboratory, we have analyzed polycystin expression in cystic tissues by immunostaining (30). In both renal and hepatic cysts, staining was seen in the epithelial lining of the cyst. Although staining appeared to be more intense in cystic kidney than in normal kidney, it was variable even within sections and within cysts. Some cysts were entirely negative for polycystin staining. Normal nephrons in the cystic kidney sections displayed the same pattern of staining

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seen in normal tissue. In ADPKD cysts staining appeared to be predominantly cytoplasmic, but areas of membrane staining were also observed. One consistent feature in previously published studies is the overexpression of polycystin in ADPKD kidney, with all cysts showing strong epithelial staining (38,39). In contrast with these observations, we detected variable polycystin expression in cystic epithelia, ranging from levels more intense than in normal kidney up to the absence of staining in some cysts. As cysts in ADPKD arise from all regions of the nephron, it might be expected that all cysts from the same patient should stain either positive or negative, as they all carry the same germline mutation in the PKD1 gene. Our observation of variable staining suggests that in some cysts no polycystin expression occurs, whereas in others the normal allele is overexpressed. The former may be due to a ‘‘second hit’’ as has been proposed (43,44). Thus, the differential level of expression or absence of polycystin in different cysts suggests that loss of function is one mechanism of cystogenesis. On the other hand, strong immunoreactivity for polycystin in cystic epithelial cells of patients with known mutations, where the antibody epitope could be presented only by the normal allele, has led some workers to argue against the two-hit hypothesis as the molecular basis of cyst formation (47). (It should be noted that in this study the ‘‘normal’’ allele was not analyzed for the presence of somatic mutations that might have compromised function while maintaining the antibody epitope.) The authors (47) suggest that cells may be predisposed to a cystic phenotype because of the loss of protein from one PKD1 allele, resulting in a dosage effect, which in turn triggers a small proportion of cells to switch to a cystic pathway. Recently, Pritchard et al. (48) reported the generation of transgenic mice by the insertion of several copies of an ⬃120 kb human PKD1 genomic fragment into the mouse genome. Transgenic mice were shown to produce higher levels of human PKD1 mRNA than the endogenous murine level. Polycystin-1 was shown to be widely expressed in embryonic and neonatal tissues. Transgenic animals developed renal displasia and tubular and glomerular cysts, indicating that overexpression of polycystin can cause the disease phenotype; therefore, the level of polycystin expression may be relevant in human disease rather than, or as well as, loss of function (48). B. Generation of Mice with Targeted Pkd1 Mutation There are several naturally occurring mouse models of cystic disease, none of which maps to the murine Pkd1 locus. Lu et al. (49) generated mice with a targeted Pkd1 mutation, disrupting the gene at exon 34. Heterozygous Pkd⫹/⫺ mice were followed for several months but did not exhibit any cystic phenotype. Homozygous Pkd⫺/⫺mice showed an embryonic lethal phenotype at stage E18.5. Histological analysis showed that the kidney developed normally until

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E14.5, with cystic transformation of proximal tubules detected at E15.5, followed by cyst formation in collecting ducts. Interestingly, no cysts were detected in the liver, but pancreatic cysts were developed at E13.5, which is earlier than kidney cystogenesis. The early stages of tubulogenesis proceeded normally in Pkd⫺/⫺ embryos, indicating that polycystin-1 is not required for nephrogenic induction but is critical for establishment of normal tubular architecture. In addition, these data support the ‘‘two-hit’’ hypothesis of cystogenesis. However, generation of these mice did not provide new insights into molecular mechanisms of human ADPKD, wherein disease is manifested in the context of a heterozygous state of germline mutation. Perhaps the unique molecular events underlying the development of cystic disease in humans can be partially explained by unique features of the human PKD1 locus that are not described for the murine locus, including the presence of homologous loci, unusual repeats such as the long homopyrimidine tract, etc.

V.

ROLE OF POLYCYSTIN-2, A MEMBRANE-SPANNING PROTEIN WITH HOMOLOGY TO Ca 2ⴙ CHANNELS IN ADPKD

A.

Polycystins-1 and -2 May Interact Through Coiled Coil Domain

The PKD2 gene is located in 4q21-23 and when mutated is responsible for about 12–15% of ADPKD cases. Cloned in 1996 (50), the PKD2 gene encodes polycystin-2, which is predicted to be an integral membrane protein of 968 amino acids with six membrane-spanning segments and both N- and C-termini located intracellularly. Polycystin-2 has homology to a family of voltage-gated calcium channels. ADPKD patients with either PKD1 or PKD2 mutations present the same clinical phenotype, which suggests that polycystin-1 and polycystin-2 activities may be part of the same molecular pathway and possibly might directly interact with each other. To address this question the C-terminal domains of the two proteins were coexpressed in a yeast two-hybrid assay (36). It was demonstrated that the two protein fragments were capable of interacting through a probable coiled coil domain in this system (36,51). Although this observation is very interesting and may explain the similar clinical presentations for PKD1- and PKD2-linked diseases, no evidence of direct interaction between the two proteins in vivo has been demonstrated as yet. B. Targeted Disruption of PKD2 Gene Recapitulates Human ADPKD Mouse models of PKD2 disease that recapitulate the human phenotype have been generated (52). Different mutations disrupting the mouse homolog (Pkd2 gene)

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were produced. One such mutation (WS25) resulted in the insertion of the disrupted exon1 into the intron1 region of the Pkd2 gene in tandem with wild-type exon1. A cystic renal phenotype was observed for both heterozygous Pkd2⫹/ ws25 and homozygous ws25/ws25 animals aged up to 14 weeks. Homozygous mutant mice produced a more severe heterogeneous phenotype, with bilateral tubular cysts, originating mostly from distal nephrons. Both homozygous and heterozygous mice developed liver cysts, which is the most common extrarenal manifestation of human ADPKD. Kidneys with cystic phenotype displayed a complete absence of polycystin-2 protein in cysts by immunostaining; however, normal staining was observed in adjacent noncystic tubules. It is likely that a subset of tubular epithelial cells lose the ability to express polycystin-2, resulting in cystogenesis. Based on the genomic structure of the WS25 allele, the authors concluded that genomic rearrangement occurred due to homologous recombination between tandems of the wild-type and disrupted exon1. These findings suggested that heterozygous mice for a true null allele Pkd2⫹/⫺ should develop a more severe phenotype than Pkd2⫹/WS25 animals. Indeed, Pkd2⫹/⫺ mice showed kidney and liver cysts and Pkd2⫺/⫺ mice were not viable. Cysts were negative for polycystin-2 protein. Therefore, in heterozygous mice (Pkd2⫹/ws25 and Pkd2⫹/⫺), an inactivating mutation or gene conversion had to occur on the normal allele to initiate cyst formation. These data provide support for the ‘‘twohit’’ hypothesis of cystogenesis and suggests that like PKD1, PKD2 is also essential for the establishment of normal renal tubular architecture. VI. NOVEL MEMBERS OF POLYCYSTIN FAMILY Recently, a novel human gene (PKDL) was identified, encoding a new member of the family of polycystins, designated polycystin-L (53). This protein has 50% amino acid sequence identity to polycystin-2 and is also homologous to the alpha 1 subunit of Ca2⫹ channels. It is expressed at high levels in fetal kidney and liver and to a lesser extent in adult tissues. PKDL maps to 10q24 and is not linked to any known human cystic diseases. However, although ADPKD phenotypes have yet to be mapped to this locus, this gene could be a candidate for human and murine cystic diseases. Veldhuisen et al. (54) reported identification of two PKD2-like genes: PKD2L1, which maps to 10q24, and PKD2L2, which maps to 5q23.3. PKD2L1 appears to be identical to the gene described by Nomura et al. (53). Veldhuisen also reported the identification of a human PKD1 paralog, which maps to chromosome 22q. VII.

CONCLUSIONS

The two genes responsible for the majority of cases of ADPKD (PKD1 and PKD2) have been cloned and characterized. Polycystin-1, a protein product of

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the PKD1 gene, is predicted to be a large membrane-associated molecule that is likely to be involved in cell–cell/matrix interactions. This protein may be important in establishment and maintenance of normal renal tubular architecture as well as playing a similar role in other cell types. While much smaller than polycystin-1, polycystin-2, a protein product of the PKD2 gene, is also predicted to be an integral membrane protein. Polycystin-2 has homology to voltage-gated calcium channels and, at least in in vitro models, can interact with polycystin-1. The predicted domain structure of polycystin-1 is consistent with a protein whose role is to mediate cell–cell or cell–matrix interaction, sense changes in these interactions, and communicate via signaling pathways with the intracellular environment. Polycystin-2 may be a component of such a receptor complex. Interpreted in the light of the natural history of the disease as well as studies in both cultured cell lines and animal models, it seems likely that mutations in the polycystins result in increased cellular proliferation and an altered differentiation status. Significant progress toward understanding the initial stages of molecular events underlying cystic development has been made. Recent studies have provided new insights into the role of polycystin-1 in cystogenesis. Evidence for a clonal nature of renal cysts together with demonstration of loss of heterozygosity in some cysts suggests a somatic ‘‘second-hit’’ mechanism for cyst development. This hypothesis could explain certain clinical features of ADPKD including striking phenotypic variability and the focal nature of cyst formation. On the other hand, an alternative hypothesis suggests that cells may be predisposed to a cystic phenotype because of a loss of protein from one mutated PKD1 allele. In this model the resulting dosage effect triggers initiation of the cystic pathway. Interestingly, analysis of targeted disruption of the PKD2 gene suggests that cyst formation occurs through a cellular recessive mechanism. Further biochemical structure–function characterization of both polycystins should shed light on their normal function, while analysis of causative mutations (in particular, genotype/phenotype studies) should help in developing an understanding of cystic molecular pathways.

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37. Lohning C, Nowicka U, Frischauf AM. The mouse homolog of PKD1: sequence analysis and alternative splicing. Mammal Genome 1997; 8:307–311. 38. Ward CJ, Turley H, Ong ACM, Comley M, Biddolph S, Chetty R, Ratcliffe PJ, Gattner K, Harris PC. Polycystin, the polycystic kidney disease 1 protein, is expressed by epithelial cells in fetal, adult, and polycystic kidney. Proc Natl Acad Sci USA 1996; 93:1524–1528. 39. Peters DJM, Spruit L, Klingel R, Prins F, Baelde HJ, Giordano PC, Bernini LF, deHeer E, Breuning MH, Bruijn JA. Adult, fetal, and polycystic kidney expression of polycystin, the polycystic kidney disease-1 gene product. Lab Invest 1996; 75: 221–230. 40. Griffin MD, Torres VE, Grande JP, Kumar R. Immunolocalization of polycystin in human tissues and cultured cells. Proc Assoc Am Physicians 1996; 108:184–197. 41. Albelda SM, Oliver PD, Romer LH, Buck A. EndoCAM: a novel endothelial cellcell adhesion molecule. J Cell Biol 1990; 110:1227–1237. 42. Fick GM, Johnson AM, Gabow PA. Is there evidence for anticipation in autosomaldominant polycystic kidney disease? Kidney Int 1994; 45:1153–1162. 43. Qian F, Watnick TJ, Onuchic LF, Germino GG. The molecular basis of focal cyst formation in human autosomal dominant polycystic kidney disease type I. Cell 1996; 87:979–987. 44. Brasier JL, Henske EP. Loss of the polycystic kidney disease (PKD1) region of chromosome 16p13 in renal cyst cells supports a loss-of-function model for cyst pathogenesis. J Clin Invest 1997; 99:194–199. 45. Watnick TJ, Gandolph MA, Weber H, Neumann HPH, Germino GG. Gene conversion is a likely cause of mutation in PKD1. Human Mol Genet 1998; 7:1239– 1243. 46. Watnick TJ, Torres VE, Gandolph MA, Qian F, Onuchic LF, Klinger KW, Landes G, Germino GG. Somatic mutation in individual liver cysts supports a two-hit model of cystogenesis in autosomal dominant polycystic kidney disease. Mol Cell 1998; 2:247–251. 47. Ong ACM, Harris PC. Molecular basis of renal cyst formation—one hit or two? Lancet 1997; 349:1039–1040. 48. Pritchard L, Sloane-Stanley J, Aspinwall R, Sharpe J, Ong A, Wood W, Harris P. A human PKD1 transgene associated with a cystic phenotype. J Am Soc Nephrol 1998; 9:381A. 49. Lu W, Peissel B, Babakhanlou H, et al. Perinatal lethality with kidney and pancreas defects in mice with a targeted Pkd1 mutation. Nature Genet 1997; 17:179–181. 50. Mochizuki T, Wu G, Hayashi T, Xenophontos SL, Veldhuisen B, Saris JJ, Reynolds DM, Cai Y, Gabow PA, Pierides A, Kimberling WJ, Breuning MH, Deltas CC, Peters DJ, Somlo S. PKD2, a gene for polycystic kidney disease that encodes an integral membrane protein. Science 1996; 272:1339–1342. 51. Tsiokas L, Kim E, Arnould T, Sukhatme VP, Walz G. Homo- and heterodimeric interactions between the gene products of PKD1 and PKD2. Proc Natl Acad Sci USA 1997; 94:6965–6970. 52. Wu G, D’Agati V, Cai Y, Markowitz G, Park JH, Reynolds DM, Maeda Y, Le TC, Hou H, Jr, Kucherlapati R, Edelmann W, Somlo S. Somatic inactivation of Pkd2 results in polycystic kidney disease. Cell 1998; 93:177–188.

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53. Nomura H, Turco AE, Pei Y, et al. Identification of PKDL, a novel polycystic kidney disease 2-like gene whose murine homologue is deleted in mice with kidney and retinal defects. J Biol Chem 1998; 273:25967–25973. 54. Veldhuisen B, Dauwerse H, Coucke P, Breuning M, Peters D. A novel gene highly homologous to the autosomal dominant polycystic kidney disease 2 gene (PKD2). J Am Soc Nephrol 1998; 9:384A.

23

Action of β-Agonists Compared to Cromoglycate on Mononuclear Cell Membranes Stabilizing or Destabilizing? Guido Zimmer, Markus Bernho¨rster, Patrizius Pilz, and Jutta Schuchmann-Fix Johann Wolfgang Goethe University, Frankfurt, Germany

I. INTRODUCTION Nowadays, physicians are urged to treat inflammation in the airways and thereby reduce the bronchial hyperreactivity that is responsible for many of the symptoms of asthma. Numerous cell types play a role in mediating the inflammatory process. There are mast cells (1), neutrophils (2–4) as well as eosinophils (4,5), granulocytes, macrophages (6,7), thrombocytes, bronchial epithelial (8) cells, endothelial cells, and various types of lymphocytes (7,9). Many of these cells increase in number. Normally, lymphocytes make up only 10–20% of total cells recovered by bronchoalveolar lavage (BAL), but they are the predominant (10) cells in airway biopsy specimens (11). Oral prednisolone reduces the number of eosinophils, lymphocytes, and mast cells. There is a reduction of the expression of IL-4; interestingly, expression of IFN-γ, which is derived from T-lymphocytes, is augmented (12). It is known that interleukins 3 (13), 4, 5 (14,15), and 13 (16) together constitute important contributing factors in inflammation. No such changes occur when a standard bronchodilator, such as salbutamol, is given. There are reports, however, that βagonists may exhibit membrane-stabilizing properties. In this study we investigated mononuclear cells (mostly lymphocytes) that were separated from buffy coats. B-lymphocytes (plasma cells) activate mast 427

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cells via the IgE-antigen complex (IgE-X) (17). In the case of membrane stabilization, shedding of cytokines after activation would be prohibited.

II.

MATERIALS AND METHODS

A.

Separation of Mononuclear Cells

Buffy coats from the blood of healthy human individuals were obtained through the courtesy of Blutspendedienst Hessen, Frankfurt am Main, Germany. Thirty milliliters of blood was layered onto 20 mL of FicoLite H and centrifuged for 40 min at 620 ⫻ g. The layer containing the mononuclear cells was carefully pipetted into new centrifuge tubes, which were then filled with phosphate-buffered saline (PBS) up to 50 mL and centrifuged for 17 min at 154 ⫻ g. The supernatant was sucked off, and the pellet was resuspended with 30 mL PBS and layered onto 20 mL FicoLite. After another centrifugation at 620 ⫻ g for 40 min the layer with mononuclear cells was again separated and PBS was added to a volume of 50 mL. Subsequently, the cells were centrifuged at 93 ⫻ g for 15 min. The supernatant was discarded, and the pellet containing the separated washed mononuclear cells was filled with RPMI buffer to a volume of 10 mL. Portions of 10 µL were diluted 1:10 with RPMI, and cells were counted with an Olympus microscope. Cells were diluted (107 to 1 mL) with RPMI. Viability of the cells was tested by means of trypan blue staining. B. Electron Paramagnetic Resonance Spectroscopy and Spin Labeling One milliliter of RPMI, containing 107 cells, was centrifuged using an Eppendorf centrifuge for 3 min at 268 ⫻ g (2000 UPM). The supernatant was discarded and the pellet was resuspended with 50 µL RPMI. After addition of a few microliters containing the desired amount of β-agonist, an incubation was carried out for 10 min at 37 °C. Subsequently, 1 µL of spin label from 5 mM stock solution was added under vortexing, and then the cell slurry was transferred into a 50 µL capillary tube, sealed, and put into the EPR cavity. Established methodology of electron paramagnetic resonance (EPR) spin labeling was used. In particular, we employed the lipid spin labels 5- and 16doxylstearate (5- and 16-DSA). 5-DSA probes the polar interface of the membrane. 16-DSA reports on the hydrophobic membrane interior. From the spectra, order parameters S as well as information about polarity of the hydrophobic membrane core (aN values) were obtained. The following β-agonists were investigated at different concentrations: fenoterol, salbutamol, reproterol. We compared the results with those obtained with the compound cromoglycin (Fig. 1).

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Figure 1 Structural formulas.

III. RESULTS In this section we first describe the results obtained with cromoglycate, since this is a very well-known substance exhibiting membrane-stabilizing properties. At rising concentration of cromoglycate the order parameter of 5-DSA increases. The control value is about 0.658, and values between 0.73 and 0.74 are attained at 30 nmol/107 mononuclear cells (Table 1). This means that the order (tightness!) of the polar part of the membrane or its interface becomes significantly increased by cromoglycate. Using the 16-DSA spin label we observe a decrease of aN values from controls of about 15.24 to 15.08 at 30 nmol/107 cells. These findings indicate that, in particular at 30 nmol/107 cells, hydrophobicity of the spin label’s 16-DSA environment increases or, alternatively, that the

Table 1 Order Parameters for 5-DSA and aN Values for 16-DSA 5-DSA

1. 2. 3. 4.

Cromoglycate Fenoterol Salbutamol Reproterol

16-DSA 7

Control

30 nmol/10 cells

Control

30 nmol/107 cells

0.658 0.652 0.72 0.674

0.735 0.628 0.65 0.67

15.24 15.1 15.18 15.1

15.08 15.44 15.48 14.95

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amount of water within the hydrophobic environment or core of the membrane is decreased. Investigating fenoterol, the following results were obtained. With 5-DSA we observe a significant decrease of order of the control value of 0.652 to 0.628 at 30 nmol/107 mononuclear cells. Conversely, the aN values increased from controls of 15.1 to 15.44 at 30 nmol/107 cells. Such findings indicate that the order of the polar interface of the membrane becomes decreased and that there is an increase in polarity of the hydrophobic core. Using salbutamol a decrease of the order parameter (5-DSA) from 0.72 (controls) to 0.65 at 30 nmol/107 cells is obtained. (The control value of order with 5-DSA may vary within a range, depending on the different buffy coats used. Also with other buffy coats and different control values for 5-DSA, we obtained a decrease between 10 and 30 nmol/107 cells.) So we can state that there is a definite decrease of order at the membrane polar interface or a loosening of structure. The investigation of aN values revealed an increase from the controls of 15.18 to 15.48 at 30 nmol/107 cells. From such findings we can conclude that we obtain an increase in polarity of the hydrophobic membrane interior. Using reproterol, we observed different results. The order parameter of 5DSA at the very low concentration of 5 nmol/107 cells first somewhat decreases but then steeply increases toward the control value, attaining 0.665 at 10 nmol/ 107 cells and 0.67 at 30 nmol/107 cells. So, unlike both fenoterol and salbutamol, the order parameter with 5-DSA stays in the control range from 10 nmol/107 upward. Similarly, the aN values also do not follow a clear-cut increase/decrease pattern: from a control value of 15.1 we obtain an increase to around 15.4 at 5 nmol/107 cells followed by a steep decrease to 30 nmol/107 cells going down to around 14.95. The direction of changes in aN resembles that found with cromoglycate. In addition to the structural investigations of polar and apolar membrane regions of the mononuclear cells (with spin labeling) we investigated viability of the cells using trypan blue (Table 2). Using cromoglycate, we found that viability of the cells was in the range of that of the control cells, which appeared comparatively stable in the investigated series. By contrast, in the presence of fenoterol, the stability of the mononuclear cells was decisively reduced. The number of stained cells increased progressively at longer incubation times and higher concentrations of the drug. Furthermore, salbutamol did not stabilize cells. On the contrary, the number of stained cells was very much increased at longer incubation times with salbutamol. The result resembles that obtained in the presence of fenoterol. On the other hand, reproterol was found to generally behave like cromoglycate or better.

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Table 2 Cell Survival (%) After 22 h Measured by Trypan Bluea nmol/107 cells

1. 2. 3. 4.

Cromoglycate Fenoterol Salbutamol Reproterol

c*

10

20

30

100

96 96 96 93

97 84 86 96

96 82 82 95

94 80 80 94

87 70 72 95

c* ⫽ control. a Differences in order from polar interface to hydrophobic core are decreased by fenoterol, salbutamol. Differences in polarity from polar outside to apolar inside are also decreased by fenoterol and salbutamol. Thus, the nature of the border between polar, extracellular, and hydrophobic intramembranous environments is challenged or compromised. Differences are straightened. The border becomes leaky.

IV. DISCUSSION This chapter provides another proof of the interrelationship of spin label data and the structural integrity of the cellular/subcellular membrane. Earlier work had already established that spin label 16-DSA spectra, revealing the polar/apolar ratios of the partitioned third line as well as aN values, reports on membrane integrity (18–20). We found that membrane integrity, and thereby cellular viability, decreased in the case of decreasing hydrophobicity of the membrane interior, measured by 16-DSA. Moreover, the polar part of the membrane, or the interfacial region, including the phospholipid phosphate and carbonyl area, which can be analyzed with 5-DSA, was found to decrease in order (21). The latter data were obtained on isolated rat liver mitochondria. Nevertheless, our present results confirm those earlier observations. Irrespective of plasma membranes or intracellular membranes, in both cases a loosening of the membrane interface appears as a sign of decreased stability or viability. This is corroborated by the decreasing order parameters in the presence of increasing amounts of fenoterol or salbutamol, using 5-DSA in mononuclear cells. On the other hand, with fenoterol as well as with salbutamol the hydrophobicity of the membrane interior was found to decrease. From these two observations of changes at the polar and hydrophobic parts of the lymphocyte membranes we can conclude that, due to the decreased tightness of the polar membrane barrier, there is an increased leakiness that is transduced to the hydrophobic part of the bilayer of the plasma membrane. Here we observe a decreased hydrophobicity or, conversely, an increased polarity. These two observations result in an overall diminished membranous stability, since bor-

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ders of the polar interface and hydrophobic core lose sharpness or clarity. The leaky polar part allows entry of increasing amounts of polar substances into the hydrophobic core (Fig. 2). Stabilization is indicated from the EPR data of mononuclear cells in the presence of reproterol. Like cromoglycate, this drug, at certain concentrations tested, increased membrane internal hydrophobicity and also does not decrease polar order at a similar range of concentrations. The findings on cellular viability can be considered as a further confirmation of our EPR spectroscopic spin labeling results. Previous work of Xiang and Anderson (22) indicated that among the three characteristic regions of a lipid bilayer membrane (as a model for a plasma membrane) comprising (1) polar and highly ordered chain, and (2) highly disordered chain regions, the highly ordered chain region may exhibit a significant barrier to permeable substances. Thus, a free energy plot gave a quantitative indication of the barrier properties of the bilayer, being highest at the region of ordered chains. Such a bilayer structure, where a similar order extends to about the depth

Figure 2 Along the lines described by Gaffney and McConnell (24), Seelig and Seelig (23), McConnell (25), and Xiang and Anderson (22), this scheme exemplifies findings described in the text. (Left) Half of a bilayer is shown as a functional membrane barrier. There is a region of highly ordered alkyl chains extending into the bilayer to about C7, followed by another region of highly disordered chains. (Right) Another half of a bilayer represents a nonfunctional, leaky state. There is a region of intermediately ordered alkyl chains extending into the layer. Further, toward the center, there is another region of intermediately disordered alkyl chains. Thus the barrier between ordered and disordered regions appears flattened. Similarly, the gradient of polarity near the phospholipid headgroups and the apolar region toward the center of the layer appears flattened (not shown). Cromoglycate and reproterol stabilize the functional structure (left), whereas fenoterol and salbutamol drive the lipids into the nonfunctional, leaky stage (right). DG° symbolizes the energetic barrier.

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of carbon atom 7 (i.e., nearly half the depth of a layer of C16 –C18 fatty acids would belong to the highly ordered chain region) was early corroborated by 2 1H NMR investigations (23) and EPR spectroscopy (24,25). Clearly, the 5-DSA spin label analyzes the region of highly ordered fatty acid chains. According to Xiang et al. (22,26), chain ordering imposes an additional diffusional resistance and an entropy barrier to partitioning. Such a barrier would decrease together with a decrease of order in this region (Fig. 2). The order parameter is generally characterized by measuring the amplitude of motion about the long axis of the fatty acid chain (27). In the case of a biological membrane where proteins become included into the bilayer structure, things become somewhat more complicated. In that case, a decrease in the amplitude of motion of fatty acid spin labels does not necessarily imply an ordering effect (28). However, the results obtained in the lipid bilayer and in a biological membrane resemble each other: ‘‘The lipid packing may be more disordered in the presence compared to the absence of a protein and the bilayer may be more permeable’’ (28). Disorder means, in the latter case, a decrease in the amplitude of molecular motion, particularly in the headgroup region. Such a result, being important for lipid–protein interactions in the absence or presence of protein, does not apply in our case of a plasma membrane in the absence or presence of a β-agonist or cromoglycate. Here, an increase in angular amplitude of molecular motion measured with 5DSA is correlated with a decrease in the order parameter. Moving along the fatty alkyl chains from carbon 7 further toward the center of the bilayer, polarity decreases. This is most efficiently revealed by a decrease of the aN values as measured on the spectra obtained by 16-DSA. An increase in hydrocarbon core molecular volume during oxidative stress (29) should be connected with an increase of polarity within this region. According to Lands (30), fluidity refers in part to some aspect of fluid expansion or molal excess volume, which describe the density of the lipid microenvironment. With fenoterol as well as with salbutamol we observe a decrease in order at the highly ordered chain region and an increase in polarity in the relatively disordered chain region. The contrary was found with cromoglycate and reproterol. Clearly, a decrease of order in the region of high order and an increase of polarity in a region of low polarity can only be reasonably interpreted as indicating an increasing leakiness of the barrier properties of a membrane. Therefore, cell membranes of mononuclear cells become destabilized by fenoterol and salbutamol, whereas they become stabilized by cromoglycate and by reproterol at the range of concentrations studied. For salbutamol, however, previous findings had clearly indicated an inhibition of IgE-dependent histamine release from human dispersed mast cells (31). In that respect, salbutamol appeared 30,000-fold more potent than cromoglycate (32). Thus, mast cell activation becomes inhibited by salbutamol to an extent

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that is nearly not comparable to the action of cromoglycate, which is known to be a particular membrane stabilizer for mast cells (32). It should be considered that mast cells release histamine from intracellular vesicles by means of exocytosis due to extracellular signaling. This means there is a regular excretory mechanism. This is a physiological action of a specialized cell type that is prohibited by some unknown mechanism through salbutamol and other β2-agonists. Membrane stabilization as defined earlier in this chapter should be differentiated from such protection against histamine release by mast cells (Fig. 3). The stabilizing influence of cromoglycate may be dependent on its highly polar structure (Fig. 1). By interaction with the membrane surface it could prevent the structural change (decrease of order) at the interface. Besides its β2-agonist properties, reproterol (Fig. 1), because of its bivalent structure, can act as an adenosine receptor antagonist. Adenosine and AMP induce vasoconstriction via mast cell degranulation (33). It could be anticipated that reproterol might also become effective at this site (Fig. 3). Generally, β2-agonists appear to be active more on the indirectly mediated bronchoconstrictive effect by mast cell degranulation through AMP than on the bronchoconstriction caused by directly acting agents such as methacholine, histamine, and sodium metabisulfite (34,35).

Figure 3 A schematic diagram of the sites of action on the membrane of the investigated compounds. (A) Stabilization/destabilization on mononuclear cell membrane; (B) inhibition of the degranulation of mast cells.

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ACKNOWLEDGMENT We thank Annette Hochberger for providing the many drafts of this chapter. She also prepared the expert artwork.

REFERENCES 1. Hunt LW, Gleich JH, Butterfield JH. Mast cells in fatal asthma: visualization with an immunofluorescent stain to mast cell tryptase. Am Rev Respir Dis 1991; 143(4): A41. 2. Callerame ML, Condemi JJ, Bohrod MG, Vaughan JH. Immunological reactions of bronchial tissues in asthma. N Engl J Med 1971; 284:459. 3. Fujisawa T, Kephart GM, Gray BH, Gleich GJ. The neutrophil and chronic allergic inflammation:immunochemical localization of neutrophil elastase. Am Rev Respir Dis 1990; 141:689–697. 4. Sur S, Crotty TB, Kephart GM, Hyma BA, Colby TV, Reed CE, Hunt LW, Gleich GJ. Sudden-onset fatal asthma: a distinct entity with few eosinophils and relatively more neutrophils in the airway submucosa? Am Rev Respir Dis 1993; 148:713–719. 5. Unger L. Bronchial asthma. Prog Allergy 1952; 3:142–221. 6. Cutz E, Levison H, Cooper DM. Ultrastructure of airways in children with asthma. Thorax 1978; 8:207–213. 7. Azzawi M, Johnston PW, Majumdar S, Day AB, Jeffrey PK. T lymphocytes and activated eosinophils in airway mucosa in fatal asthma and cystic fibrosis. Am Rev Respir Dis 1992; 145:1477–1482. 8. Guo FH, De RH, Rice TW, Stuehr DJ, Thunnissen FB, Erzurun SC. Continuous nitric oxide synthesis by inducible nitric oxide synthase in normal human airway epithelium in vivo. Proc Natl Acad Sci USA 1995; 92:7809–7813. 9. Cohen S. Lymphokines and the Immune Response. Boca Raton, FL: CRC Press, 1990. 10. James AL, Carroll N. The pathology of fatal asthma. In: Hogate ST, Busse WW, eds. Inflammatory Mechanisms in Asthma. New York: Marcel Dekker, 1998:1–26. 11. Liu MC, Calhoun WJ. Bronchioalveolar lavage in studies of asthma. In: Holgate ST, Busse WW, eds. Inflammatory Mechanisms in Asthma. New York: Marcel Dekker, 1998:39–74. 12. Bentley AM, Hamid Q, Robinson DS, Schetman E, Meng Q, Assouti B, Kay AB, Durham SR. Prednisolone treatment in asthma: reduction in the numbers of eosinophils, T-cells, tryptase-only positive mast cells and modulation of IL-4, IL-5 and interferon-gamma cytokine gene expression within the bronchial mucosa. Am J Respir Crit Care Med 1996; 153:551–556. 13. Miyamjima A, Mui AL-F, Ogorocki T, Sakamaki K. Receptors for granulocytemacrophage colony-stimulating factor, interleukin-3, and interleukin-5. Blood 1993; 82:1960–1974. 14. Bradding P, Feather IH, Howarth PH, Mueller R, Roberts JA, Britten K, Bews JP, Hunt TC, Okayama Y, Heusser CH, Bullock GR, Church MK, Holgate ST. Interleu-

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24 Cystic Fibrosis Transmembrane Conductance Regulator A Chloride Channel Regulator of Ion Channels Makoto Sugita Hiroshima University School of Medicine, Hiroshima, Japan

J. Kevin Foskett University of Pennsylvania, Philadelphia, Pennsylvania

I. INTRODUCTION Although the spectrum of clinical manifestations is quite wide, it has been generally believed that an underlying basis of cystic fibrosis (CF) is abnormal regulation of epithelial ion and fluid transport (1). Primary sequence analysis of the CF gene suggests that its protein product, the cystic fibrosis transmembrane conductance regulator (CFTR), is an integral membrane protein containing structural domains similarly present in proteins with known transport functions (2,3). CFTR is a member of the traffic ATPase or ATP binding cassette (ABC transporter) superfamily of proteins. It is composed of two membrane-associated domains, each with six putative transmembrane helices, two nucleotide binding domains (NBDs), and a large cytoplasmic R (regulatory) domain that contains numerous consensus sequences for kinase phosphorylation (2,3). Cystic fibrosis is caused by mutations in CFTR. The disease is usually manifested as exocrine pancreatic insufficiency, an increase in sweat Cl⫺ concentration, and airway disease. Airway disease leads to progressive lung dysfunction, which is currently the major cause of morbidity and is responsible for 95% of CF mortality. Although all the functions of CFTR may not yet be established, a variety of evidence has demonstrated that CFTR is a cAMP-regulated Cl⫺ channel. First, expression of CFTR in nonepithelial cells confers a cAMP-regulated Cl⫺ conductance (4–6). Second, site-directed mutagenesis of CFTR alters channel properties 439

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associated with its expression (7). Finally, reconstitution of purified CFTR into planar lipid bilayers revealed cAMP-activated Cl⫺ channels with properties similar to those in CFTR-expressing cells (8). Regulation of the Cl⫺ channel activity of CFTR is remarkably complex (9). Phosphorylation of the R domain at multiple sites by cAMP-dependent protein kinase (PKA) activates the Cl⫺ conductance (10), presumably by inducing conformational changes or modulating electrostatic interactions of the R domain with other parts of the molecule (11). Protein kinase C (PKC) also activates the channel, and phosphorylation by PKC may be a prerequisite for PKA activation (12). The ABC transporters couple ATP hydrolysis at NBDs to the transport of a wide variety of molecules (13). However, CFTR conducts Cl⫺ ions down an electrochemical gradient, a process that has no obvious need for external energy. Nevertheless, CFTR hydrolyzes ATP (14), and when the R domain is phosphorylated, intracellular ATP interacts with the NBDs to regulate channel activity by nucleotide hydrolysis (15,16). Nonhydrolyzable ATP analogs fail to open phosphorylated CFTR that can subsequently be opened by ATP (16), and they cause CFTR channels to lock into conducting states of markedly prolonged duration (17,18). ATP hydrolysis therefore appears to contribute to both opening and closing of CFTR channels. Mutation analysis suggests that ATP hydrolysis at the two NBDs may have opposing functions, with NBD1 and NBD2 controlling channel opening and closing, respectively (19). Some evidence exists that changes in the configuration of the TMs are involved in gating the channel (20,21). The structural basis for the conduction pathway in CFTR is poorly understood. How the 12 membrane-spanning helices are arranged to form the pore remains unknown. It is likely that TM1, TM3, TM5, TM6, and TM12 contribute to the conduction pathway, as evidenced by the identification of water-accessible residues in the transmembrane segments using scanning cysteine accessibility methods (22,23), the halide permeability or conductance sequences of wild-type and mutant CFTRs (24–26), and the accessibilities of blockers to specific residues (27–29). Relative halide permeabilities, determined from shifts in reversal potentials, are consistent with a ‘‘weak site’’ ion–channel interaction, suggesting that permeability ratios depend highly on anion differences in anion–water interactions (24,30). Based on the presence of anomalous mole-fraction effects, the CFTR channel was inferred to have multiple anion-binding sites (25), although more recent results cast doubt on this conclusion (31). Thus, the structural basis for anion conduction by CFTR remains largely a mystery. In CF, the Cl⫺ channel function of CFTR is defective. Over 800 different mutations in the gene have been reported (L.-C. Tsui, personal communication). Electrophysiological and biochemical analyses of some of these mutant CFTRs have revealed that mutations can affect CFTR Cl⫺ channel function in several different ways, which have been catalogued (7) into discrete, not necessarily ex-

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clusive, classes: Class 1 mutations are associated with improper protein production due to premature termination signals in the gene. Class 2 mutations impede exit of the fully translated proteins from the endoplasmic reticulum (ER), where they are synthesized. This class includes the most common mutation, deletion of phenylalanine at residue 508 (∆F508). CFTR containing the ∆F508 mutation and other mutants in this class fail to mature to the fully glycosylated, Golgi-processed form and are subsequently degraded in an ER-associated compartment (32,33) by a process that appears to be mediated at least in part by ubiquitination of CFTR and subsequent proteolysis by the 26S proteosome complex (34,35). Because ∆F508 functions as a regulated Cl⫺ channel (36), these mutations appear to cause disease as a result of the absence or very low levels of the protein at the cell surface. Class 3 mutations cause the protein to be defectively regulated by phosphorylation or ATP binding and hydrolysis. Class 4 mutations affect the Cl⫺ conduction pathway. This list may expand as more is learned about the effects of specific mutations on the Cl⫺ channel function of CFTR and as other possible functions of CFTR are elucidated (see below). Immunolocalization of CFTR to the apical membranes of epithelial cells involved in CF, including lung, pancreatic ducts, reproductive tissues, bile ducts, and colonic crypts (37), has provided evidence consistent with a model in which fluid and salt transport by these epithelia involves Cl⫺ transport across the apical membrane through CFTR. A simple view of the role of CFTR in lung biology recognizes CFTR as an apical membrane-localized Cl⫺ channel that functions in airway transepithelial fluid transport, affecting airway fluid balance. Proper fluid balance is critical for mucociliary clearance of inflammatory particles, including bacteria (38,39); defective or absent CFTR, by disrupting this balance, is believed to cause airway mucus to become too viscous to be effectively cleared from the lungs by normal ciliary mechanisms (39). The resulting chronic inflammation and tissue damage in the lung are the cause of most mortality in CF (40). Some evidence supports such a model that places the apical membrane Cl⫺ channel function of CFTR at the center of lung pathology in CF. For example, there is a good correlation between the level of residual Cl⫺ channel activity and the severity of pancreatic disease in CF (41–43), suggesting that, at least in this organ, the Cl⫺ channel function of CFTR is critical to normal functioning of the tissue. Furthermore, in a mouse model of CF in which CFTR was disrupted, few manifestations of pathology were apparent in many of the organ systems that are affected in humans with CF (44). Electrophysiological analyses indicated that the mouse tissues may be protected from pathological consequences of the absence of CFTR by the presence of a different apical membrane Cl⫺ channel, one regulated by intracellular Ca2⫹ concentration (44). These data too suggest that the function of CFTR as an apical membrane Cl⫺ channel involved in transepithelial salt transport is critical to avoid the pathology observed in CF.

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DOES DYSFUNCTIONAL Clⴚ PERMEABILITY REALLY ACCOUNT FOR LUNG PATHOLOGY IN CF?

Although this model is attractive, several lines of evidence suggest that it may be too simplistic. First, it is not possible to know whether lack of plasma membrane Cl⫺ channel activity can account for airway pathology in CF, because the mechanisms that govern fluid balance in the normal lung are still poorly understood. Recent studies of CF mice indicate that the consequences of defective fluid transport are not apparent in mice reared in a pathogen-free environment (45). Only after repeated inflammatory insults by exposure to bacterial pathogens did it appear that the absence of Cl⫺ channel function contributed to lung pathology (45). These results may suggest that CFTR is involved in fluid secretion and mucus hydration only in response to inflammatory stress and is not primarily involved in these processes in the lung under normal conditions. Second, it is becoming more apparent that CFTR may have functions in addition to its Cl⫺ conductance role. The roles of these activities in CF pathophysiology are unknown. The cellular heterogeneity and morphological complexity of the lung have inhibited efforts to define the mechanisms that govern fluid homeostasis in this organ. It is generally believed that most fluid in the lung is derived from distal regions, likely the alveolus (46). The alveolar mechanisms that contribute to the water load on the lung are not clear but may involve active secretion and/or passive movements due to Starling forces (47). Because of the tremendous surface area amplification of the distal compared with the proximal regions of the lung, the more proximal regions are likely involved in ion and fluid reabsorption. In support of this notion, in vitro studies of airway cells from humans have demonstrated that the small and large airways are Na⫹-absorbing tissues (46,48). However, the proximal airway regions also contain submucosal glands composed of serous and mucous acini, and it seems likely that these structures secrete fluids (49,50). The possible role of CFTR in this process has not been established, but, importantly, CFTR is expressed at very high levels in the submucosal gland serous cells, in contrast to its low level of expression in the rest of the lung (51). Nevertheless, the relative contributions of serous glands to proximal airway fluid balance is unclear, and it is not known if fluid secretion by these glands is tonic or is activated only in response to specific cues, perhaps associated with inflammatory insults (48,50). Fluid reabsorption occurs in the rest of the human lung and appears to involve active Na⫹ transport, with Cl⫺ movement across the tissues being driven passively between cells by electrical forces (46,52). Although CFTR is expressed at low levels in airway cells, paradoxically it may not be involved as a pathway for transepithelial Cl⫺ absorption. First, Cl⫺ appears to be at electrochemical equilibrium across the apical membrane of airway cells (53), so there is no driving

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force for Cl⫺ movement in either the absorptive or secretory direction. Second, NaCl absorption is enhanced in CF, despite the absence of functional CFTR (46,54,55). Thus, a functional importance of the CFTR Cl⫺ channel activity in fluid transport is questionable in the majority of cells in the lung. The role that CFTR plays in airway cells therefore remains obscure, a conclusion that is at once both surprising and distressing, given the central contribution of lung pathology to mortality in CF. Importantly, CFTR has been implicated in the regulation of other ionic conductances (discussed below). It is possible therefore that these functions of CFTR, rather than its Cl⫺ channel function, are more relevant for the etiology of lung pathology in CF patients.

III. CFTR MAY HAVE FUNCTIONS IN ADDITION TO ITS ROLE AS A PLASMA MEMBRANE Clⴚ CONDUCTANCE Some recent observations suggest that CFTR may function to regulate other epithelial conductances by mechanisms that are seemingly independent of its Cl⫺ channel function. A.

Outwardly Rectifying Clⴚ Channel

Regulation by cAMP of another airway Cl⫺ channel, the outwardly rectifying Cl⫺ channel (ORCC), is also defective in CF (56). Although ORCC was originally believed to be the defective Cl⫺ channel basis of epithelial impermeability to Cl⫺ in CF (56) and the molecular identity of ORCC is not yet known, ORCC and CFTR are now appreciated as separate gene products (57). The biophysical properties of ORCC and CFTR Cl⫺ channels are distinct and well established. ORCCs have a nonlinear current–voltage (I/V ) relationship with a 20–40 pS single-channel conductance at hyperpolarizing voltages and a 60–80 pS conductance at depolarizing voltages (58,59). ORCCs are blocked by a wide variety of molecules, including DIDS and the calixarenes (60), and have a halide permeability sequence I⫺ ⬎ Cl⫺ ⬎ Br⫺ (58). ORCCs can be activated by PKA and PKC. Conversely, the CFTR Cl⫺ channel has a linear I/V relationship with a 7–14 pS single-channel conductance (8,61). Channel activity can be blocked by diphenylamine-2-carboxylic acid (DPC) (28) and glibenclamide (62,63) but not by external DIDS. The halide permeability sequence for CFTR is Br⫺ ⬎ Cl⫺ ⬎ I⫺ (24). Expression of CFTR in airway epithelial cells corrected defective ORCC regulation (56). The correction by CFTR appeared to be associated with the ability of intracellular ATP to activate the ORCC from the extracellular side of the membrane (64). A model has been proposed (64) in which ATP efflux through CFTR regulates the ORCC activity by making available extracellular ATP to a purinergic receptor, which in turn regulates ORCC activity by as yet unknown

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mechanisms. The observed CFTR-dependent PKA-mediated activation of ORCC has also been observed in bilayer studies (65,66) that indicated that phosphorylated CFTR is required in a mechanism that may impinge on G-protein regulation of ORCC. B. ATP Channels Cantiello and coworkers (67) and Guggino and coworkers (64) initially, and others later (68,69), provided evidence that CFTR can either conduct ATP or is very closely associated with an ATP channel. However, these surprising results are quite controversial because they have not been observed by all investigators (70– 72). To characterize ATP channels associated with CFTR, we analyzed Cl⫺ and ATP single-channel currents in excised inside-out membrane patches from MDCK epithelial cells transiently expressing CFTR (68). With 100 mM ATP in the pipette and 140 mM Cl⫺ in the bath, ATP channels were associated with CFTR Cl⫺ channels in two-thirds of patches that included CFTR. CFTR Cl⫺ channels and CFTR-associated ATP channels had slope conductances of 7.4 and 5.2 pS, respectively, and had distinct reversal potentials and sensitivities to channel blockers. CFTR-associated ATP channels exhibited slow gating kinetics that depended on the presence of protein kinase A and cytoplasmic ATP, similar to CFTR Cl⫺ channels. Gating kinetics of the ATP channels as well as the CFTR Cl⫺ channels were similarly affected by nonhydrolyzable ATP analogs and mutations in the CFTR R domain and NBDs. Our results indicated that phosphorylation- and nucleotide hydrolysis-dependent gating of CFTR is directly involved in gating of an associated ATP channel. However, the permeation pathways for Cl⫺ and ATP are distinct, and the ATP conduction pathway is not obligatorily associated with the expression of CFTR. A CFTR-associated ATP permeability has also been observed in only a subset of CFTR Cl⫺ conductance-expressing Xenopus oocytes (73). CFTR-modulated ATP release depended on both cAMP activation and a gradient change in the extracellular Cl⫺ concentration. Mutagenesis of CFTR demonstrated that Cl⫺ conductance and ATP release regulatory properties could be dissociated to different regions of the CFTR protein. Despite the lack of a requirement for Cl⫺ conductance through CFTR to modulate ATP release, alterations in CFTR channel pore residues R347 and R334 caused changes in the relative ability of different halides to activate ATP efflux (wt CFTR: Cl⫺ ⬎⬎ Br⫺; R347P: Cl⫺ ⬎⬎ Br⫺; R347E: Br⫺ ⬎⬎ Cl⫺; R334W: Cl⫺ ⫽ Br⫺), leading to the suggestion that a chloride sensor mechanism may be present in CFTR that is capable of responding to changes in the extracellular Cl⫺ concentration by modulating the activity of an unidentified ATP flux pathway. These observations are consistent with the hypothesis that specific molecular interactions of CFTR with one or more other proteins are required to create

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the ATP conductance. The interacting molecules may include a separate ATP channel protein regulated by domains in CFTR. Precedents for this mechanism exist in the demonstrations that P-glycoprotein, another ABC transporter protein, regulates distinct cell-volume-activated Cl⫺ channels (74); the sulfonylurea receptor (SUR), another ABC transporter protein, interacts with a K⫹ channel to confer ATP-sensitive K⫹ channel activity (75,76); and coexpression of CFTR with the epithelial Na⫹ channel (ENaC) modifies ENaC gating kinetics and response to phosphorylation (77). Our results in MDCK cells (68) suggest an important role for the R domain in contributing to manifestation of the ATP channel, since its deletion, while leaving the CFTR Cl⫺ channel pore intact, eliminated the ATP channel. It is interesting to note that phosphorylation of the linker region in P-glycoprotein, which is located in a position analogous to that of the R domain of CFTR, is critically involved in P-glycoprotein regulation of cell-volume-activated Cl⫺ channels (74). Alternatively, the molecular interaction of CFTR with the unknown molecules might create a novel permeation pathway. Finally, the interaction could possibly create an ATP channel within CFTR itself. Further studies will be required to clarify the nature of the ATP permeation pathway. Because our results suggest that CFTR is a regulator of an ATP channel that is distinct from the Cl⫺ channel in CFTR, they may provide an explanation for the inability in some studies to observe CFTR-associated ATP permeabilities (70–72). C. Naⴙ Channels In addition to Cl⫺ impermeability, it has been recognized for many years that airway cells from CF patients are more permeable to Na⫹ (39,46,55,78). Apical membrane-localized amiloride-sensitive Na⫹ channels have a higher open probability (79), whereas the number of channels does not appear to be increased (80). The elevated Na⫹ permeability results in enhanced fluid absorption in airways (54), which may contribute to airway mucus dehydration in CF (39). Clarification of the mechanisms that underlie epithelial Na⫹ channel sensitivity to mutations in CFTR have been facilitated by the molecular identification of the Na⫹ channel genes (81–83). ENaC consists of three homologous subunits (alpha, beta, gamma ENaC) associated in a tetramer with 2:1:1 stoichiometry (84,85). Each subunit consists of two transmembrane helices, a large extracellular domain, and two short cytoplasmic tails (81,86,87). By coexpression of CFTR and the Na⫹ channel subunits in fibroblasts, Stutts et al. (88) were able to reconstitute the CFTR-mediated Na⫹ channel regulation observed in the airways. ENaC expressed alone responded to PKA with increased open probability (Po) and mean open time, whereas ENaC coexpressed with CFTR exhibited decreased Po and mean open time under conditions optimal for PKA-mediated protein phosphorylation. Thus, CFTR regulates ENaC at the level of single-channel gating, by

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switching the response of single-channel Po to cAMP from an increase to a decrease. Thus, CFTR appears to exert a negative modulatory influence on ENaC, a regulation that can be observed in several distinct cell types, including human airway epithelia, mouse fibroblasts, and amphibian oocytes (77,88,89). These findings corroborate the long-standing observation that Na⫹ absorption across CF airway epithelia is increased. In CF airways, the abnormally high rate of basal Na⫹ absorption therefore likely reflects the absence of negative regulation of ENaC by CFTR under basal phosphorylating conditions, and increased PKA activity leads only to further absorption. In contrast, CFTR function in normal airways converts the activation of PKA into a stimulus for inhibition of ENaCmediated Na⫹ absorption and possibly stimulation of CFTR-mediated Cl⫺ secretion. The mechanism of this regulation is still unknown. CFTR might transport a mediator that regulates Na⫹ channel activity, as described above for ORCC, or it might regulate the Na⫹ channel by direct or indirect protein–protein interactions. Alternatively, mutations in CFTR might affect intracellular processing of Na⫹ channel subunits or regulatory proteins. Inhibition of ENaC by CFTR was detected in single-channel current measurements when both proteins were reconstituted in planar lipid bilayers (90,91). The central part of the CFTR molecule comprising NBD1 and the R domain interacted with the alpha subunit of ENaC in a yeast two-hybrid interaction screen (89). However, there have been no confirmations of this result from other laboratories. A fragmented CFTR, consisting of NBD1 and R, exerted the inhibitory effect on ENaC when the channels were expressed in oocytes, whereas the G551D mutation located within NBD1 abolished the inhibitory effect of CFTR activation on ENaC Na⫹ current (89). These results may implicate the alpha subunit of ENaC in mediating an interaction with CFTR. However, further studies are necessary to identify which subunits of ENaC are involved in CFTR-dependent regulation of ENaC and whether additional, unidentified proteins are necessary for this regulation. CFTR inhibition of ENaC may depend on anion conduction of CFTR Cl⫺ channels (92). This was shown in several different ways: (1) the degree of inhibition correlates with the magnitude of the stimulated Cl⫺ conductance; (2) mutants of CFTR that conduct less also inhibit less; (3) poorly conducted anions have reduced inhibitory effect; (4) inhibition of CFTR by DPC reduces the inhibition of ENaC; and (5) the degree of inhibition even depends on the direction of Cl⫺ movement. The inhibition apparently is abolished for inward currents (Cl⫺ moving out of the cell). The mechanism of how Cl⫺ movement through activated CFTR contributes to inhibition of ENaC is not clear at this stage. It has been suggested that cytosolic Cl⫺ activity itself regulates epithelial Na⫹ channels. Increases in intracellular Cl⫺ and Na⫹ inhibit the Na⫹ conductance in salivary gland duct cells, which is characteristic of ENaC (93–95). Heterotrimeric GTP binding proteins are possibly activated by an increase in either intracellular Cl⫺ or Na⫹

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concentration which, in turn, inhibits Na⫹ conductance (95), suggesting a basis for the feedback mechanism of Cl⫺ and Na⫹. Activation of CFTR could also interfere with Na⫹ channel activity by its effect on cellular membrane trafficking (96,97). CFTR has a significant impact on both exo- and endocytosis, which are dependent on the presence of Cl⫺ ions (97). Although the mechanism of CFTR-dependent inhibition of ENaC is only now emerging, the functional consequences of the results presented here are obvious. D.

ROMK Kⴙ Channels

The high affinity sulfonylurea receptor expressed in the pancreatic islet beta cell is encoded by a unique ATP-binding cassette protein, SUR1 (75). The sulfonylureabinding protein does not exhibit any channel activity itself but instead interacts with the pancreatic inward-rectifying K⫹ channel subunit, Kir6.2, to form a K⫹ channel, the KATP channel, sensitive to intracellular ATP levels (76,98). Kir6.2 belongs to a family of inwardly rectifying K⫹ channels (Kir) that are distinguished by allowing more inward than outward K⫹ current and are characterized by a structure that contains two transmembrane segments, a probable pore-forming hairpin loop, and cytoplasmic N- and C-terminal domains, in a tetrameric association (99–101). The K⫹ channel properties conferred by coexpression of Kir6.2 and SUR1 were similar to those that are observed in beta-islet cells and that are known to play a critical role in insulin secretion (76). With the subsequent discovery that the cardiac KATP channel comprises a related ATP-binding cassette protein, SUR2A, and Kir6.2, a molecular paradigm for other KATP was established (102). KATP channels are heteromultimers of sulfonylurea receptors and Kir6.x subunits associated in a 1:1 stoichiometry as tetramers (SUR/Kir6.x)4 (103). Kir6.x forms the pore, whereas SUR (SUR1 and SUR2) regulates their activity (98). Changes in [ATP]in and [ADP]in regulate gating of the channel (76,98). The diversity of KATP channels results from the assembly of SUR and Kir6.x subtypes. Kirl.1a appeared to be an excellent candidate for the kidney inward-rectifying K⫹ channel subunit (104); however, neither the beta cell (SUR1) nor the cardiac myocyte (SUR2A) ABC proteins are expressed in the kidney (75,102). Coexpression with CFTR in oocytes modified the single-channel conductance of expressed Kirl.1a and furthermore conferred both ATP and sulfonylurea sensitivity to channel gating (105). The data are compatible with the notion that CFTR and Kirl.1a physically associate to form a hybrid channel with properties of the distal nephron KATP channel. Expression of CFTR also confers sulfonylurea (glibenclamide) sensitivity on ROMK2 (Kirl.1b) (106), also an apical membrane kidney K⫹ channel, and CFTR mutations in NBD1 reduced the glibenclamide sensitivity of coexpressed Kirl.1b (107). Importantly, CFTR is expressed abundantly in the kidney and is localized along the entire nephron on the apical mem-

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brane (108). Its cellular and subcellular expression overlap with Kirl.1 in the cortical collecting duct (109). Thus, the interaction observed in expression systems between Kirl.1 and CFTR may have important implications for renal K⫹ transport. The idea that Kir subunits may interact with ATP-binding cassette proteins beyond the subfamily of sulfonylurea receptors (e.g., CFTR) provides an intriguing possible explanation for functional diversity in KATP channels. IV. MECHANISMS BY WHICH CFTR REGULATES OTHER CONDUCTANCES The diversity of ion channels that have been identified as being regulated by CFTR is surprising. It raises the possibility that CFTR could regulate several different proteins simultaneously and possibly coordinate their activities. At the very least, this idea implies that CFTR may exist within a multiprotein complex that facilitates such regulation. In turn, the presence of such a complex might regulate CFTR activity, for example by stabilizing the protein at the cell surface or by increasing the efficiency with which kinases and phosphatases control the channel. Recent studies suggest that CFTR indeed interacts with other proteins. The C-terminal tail of CFTR is highly conserved among species, and it is unique to CFTR as it is not present in other ABC transporters. The last three amino acids of CFTR, Thr-Arg-Leu (TRL), are similar to a consensus PDZ-binding motif (110). PDZ domains [originally identified in postsynaptic density 95, discs large, and ZO-1 (111,112)] are ⬃90 amino acid motifs that form a fivestranded antiparallel β-barrel flanked by α-helices with a groove on its surface that binds to the carboxy-terminal tail of proteins (113,114). They are found in a large number of multifunctional proteins, existing as single repeats or, more often, as multivalent protein-interaction modules (115), where they mediate protein–protein interactions at structures including the postsynaptic density in neurons and junctional complexes in epithelia. Usually, a short consensus sequence at the COOH terminus of membrane proteins is critical for the interaction with PDZ domains; one consensus sequence consists of the amino acids (D/E)(T/ S)XV. In addition, PDZ domains can interact with other PDZ domains (116,117) and with internal sequences (118). A PDZ domain-containing protein, the Na⫹ /H⫹ exchanger regulatory factor (NHERF), was previously identified as a cofactor that regulates the activity of the Na⫹ /H⫹ exchanger type 3 isoform (NHE3) (119). NHERF contains two PDZ domains and a C-terminal ERM domain that binds to members of the ERM (ezrinradixin-moesin) family of cytoskeletal proteins (120,121). The carboxy tail of the β2 adrenergic receptor, which terminates in -DSLL, was also discovered to interact with the first PDZ domain of NHERF, and mutation of the terminal L to V abolished the interaction (122). Based on this result

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and a database sequence search, the CFTR C-terminus was recognized as a likely NHERF interactor, and full-length as well as C-terminal peptides of CFTR were demonstrated to bind in vitro to PDZ1 of NHERF (123). In another study, affinity selection of random peptides using each NHERF PDZ domain resulted in a similar prediction, which was also verified by in vitro interactions between PDZ1 of NHERF and C-terminal CFTR fusion proteins (124). A more recent study extended these observations by immunolocalizing NHERF to the apical membranes of epithelial cells that express CFTR there, in addition to confirming the interaction between the two proteins using in vitro binding assays (125). Independently, human NHERF was cloned by yeast two-hybrid screening based on its ability to interact with the C-terminal 70 amino acids of CFTR (Raghuram et al., unpublished observations). The results from the two-hybrid system were confirmed using in vitro and in vivo binding assays and by colocalizing the two proteins in heterologous expression systems. The results demonstrated that the terminal three amino acids of CFTR mediate the interaction with NHERF through interactions with NHERF PDZ domains. The two individual PDZ domains of NHERF each interact with CFTR, but the strongest interaction is mediated by a module that contains both domains. The functional implications of the interaction of NHERF with CFTR are unknown. A naturally occurring truncation of the tail of CFTR (S1455X), which deletes the NHERF-binding motif, has been reported in a CF patient (126). The clinical phenotype is unusual, with subjects exhibiting elevated sweat chloride concentrations but no other manifestations of CF. These findings suggest that the binding of NHERF, or NHERF-related proteins, to the tail of CFTR may be important for some aspects of CFTR function. The interaction of PSD-95 with NMDA receptor subunits or Kv1.4 channels causes the formation of channel clusters (127,128). We attempted to determine whether CFTR channel clustering is mediated by the NHERF interaction, by expressing S1455X CFTR in CHO cells and by using patch clamp electrophysiology. CHO cells express NHERF endogenously (119), and we have observed that expressed wild-type CFTR channels in these cells seem to exist in clusters, as evidenced by a strong propensity to find either no or many CFTR channels in a patch. In addition, we noted apparent cooperative ‘‘wavelike’’ gating behavior of the channels in these patches, as observed by others (129,130). Nevertheless, we observed similar clustering and apparent cooperative gating kinetics of S1455X CFTR in these cells, suggesting that, at least in this cell type, this apparent clustering is not mediated by the NHERF interaction (Raghuram et al., unpublished observations). As mentioned above, NHERF (EBP-50) was identified and cloned on the basis of its ability to associate with ezrin, a protein found in the apical domain of epithelial cells (120,121). EBP-50 is also localized at the apical surface through the interaction with ezrin and is expressed in a variety of epithelial tissues as part of a subapical membrane-anchoring cytoskeletal complex. EBP-50 may

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therefore link CFTR to the cortical actin cytoskeleton to localize and/or stabilize it there. Ezrin may also function as a cAMP-dependent protein kinase anchoring protein (AKAP) (131). Binding of NHERF might therefore position the dominant phosphorylation machinery in close proximity to the phosphorylation sites of CFTR (125).

V.

CONCLUDING REMARKS

The identification of CFTR involvement in protein–protein interactions suggests that its multifunctional regulation of other epithelial transport proteins may be mediated by direct protein interactions, and furthermore that such interactions may play important roles in controlling the trafficking, localization, and regulation of CFTR. Recently, a direct physical interaction between the amino terminus of CFTR and Syntaxin 1A was demonstrated, which inhibited the functional activities of normal and disease-associated forms of CFTR Cl⫺ channels (132,133). Using two-hybrid screening, we detected novel proteins that interact with the C terminus upstream of the PDZ-binding motif and with the R domain (Sugita and Foskett, unpublished observations). It is likely, therefore, that CFTR exists within multiprotein complexes, which has important implications for channel localization, interactions, stability, trafficking, and regulation. Detailed investigations in this exciting new area will lead to novel insights into the role of CFTR in physiology and into how mutations in CFTR cause pathophysiology in CF.

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25 Role of Ca2ⴙ-Independent Lysosomal Phospholipase A2 in Turnover of Lung Surfactant Phospholipids Aron B. Fisher University of Pennsylvania, Philadelphia, Pennsylvania

I. INTRODUCTION Lung surfactant is an extracellular phospholipid and protein-rich material that forms an interfacial barrier between the pulmonary alveolar gas phase and the alveolar epithelial cell lining. The presence of lung surfactant is essential for life. Its primary physiological role is to maintain a low surface tension in the alveolar space of the lung, especially at low lung volumes, thereby stabilizing the alveoli and permitting efficient gas exchange. In the absence of surfactant, as illustrated by the respiratory distress syndrome (RDS) of the neonate, alveolar collapse compromises lung gas exchange, leading to asphyxia. Thus, the factors that determine the presence of adequate extracellular surfactant are of major importance for understanding normal respiratory physiology. The major focus of this chapter is the role of a novel phospholipase A2 in the metabolism of lung surfactant. But first the chapter presents an overview of the cell biology of the lung surfactant system with emphasis on lung surfactant turnover. Recent publications explore these facets in greater depth (1,2). Standard physiology textbooks can be consulted for information concerning the physiological role of lung surfactant.

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LUNG SURFACTANT TURNOVER AND METABOLIC PROCESSING

A.

Composition of Lung Surfactant

Lung surfactant is a secreted product that represents a complex mixture of lipid and protein components. Its composition is relatively well defined but may vary depending in part on the source of material. The most commonly analyzed source is surfactant obtained by lavage of lungs with saline (bronchoalveolar lavage) and purified by density gradient centrifugation. Additional sources are the surfactant secretory organelles (lamellar bodies) that can be isolated from the lungs using techniques of subcellular fractionation, and fetal surfactant, which is isolated from amniotic fluid. Phospholipids constitute the bulk of surfactant, making up approximately 80% of the total weight. The phospholipid pool is heterogeneous and includes phosphatidylcholines (PCs) (approximately 75% of total phospholipid), phosphatidylglycerol (PG) (about 10%), and other phospholipids in trace amounts. The phospholipid components are considered further below since they are the major focus of this chapter. The remainder of surfactant by weight is due to neutral lipid components (predominantly cholesterol) and protein. Three specific proteins consistently co-isolate with the phospholipid portion of surfactant and are termed surfactant proteins A, B, and C. About half of the protein content of surfactant comprises a variety of other protein components, such as albumin and surfactant protein D, which are present in bronchoalveolar lavage fluid and contaminate but do not primarily co-isolate with the surfactant phospholipids. Although phosphatidylcholines comprise the largest component of surfactant phospholipids, this group also is heterogeneous as determined by the acyl substituents in the sn-1 and -2 positions. Dipalmitoylphosphatidylcholine (DPPC), with a saturated fatty acid (palmitate) in both acyl positions, accounts for about two-thirds of the total. The phosphatidylcholines constituting the remaining one-third of the PC pool have at least one unsaturated fatty acid, generally in the sn-2 position. Thus, DPPC accounts for 45–50% of the total surfactant by weight, while 20–25% is accounted for by ‘‘unsaturated’’ PC. Current evidence suggests that the surface tension lowering function of lung surfactant is due to DPPC, but the other lipid components and surfactant proteins (especially SP-B and -C) are necessary to ensure rapid spreading of the phospholipid film and to impart other important biophysical features of lung surfactant function. B. Lung Surfactant Processing Type II alveolar epithelial cells (granular pneumocytes) play a central role in the synthesis, secretion, and removal of lung surfactant. These are cuboidal cells composing one of the two major types of cells lining the alveolus, the distal

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airspace of the lung where lung gas exchange occurs. Type I alveolar epithelial cells (membranous pneumocytes) are the other major cell type of the alveolar epithelial surface. Although type I and type II epithelial cells are present in approximately equal numbers, type II cells occupy only 5% of the alveolar surface while the thin, flat type I cells occupy the remainder. Type II cells are readily distinguished by their cuboidal shape and the presence of secretory granules (called lamellar bodies), which are 1–2 µm diameter membrane-limited organelles containing surfactant constituted as lipid bilayers. Surfactant phospholipids are synthesized in the endoplasmic reticulum of the type II cell and transferred to lamellar bodies for intracellular storage. Estimates are that each type II cell has approximately 500 lamellar bodies in various stages of maturity. Physiological secretion of lung surfactant occurs constitutively and also in response to a variety of stimuli including mechanical stretch, intracellular alkalosis, and several circulating agonists. The latter include activators of both protein kinase A and protein kinase C as well as agents that elevate cytosolic calcium. The precise physiological roles for these agonists and their interactions have not yet been elucidated. Secretion is by classical exocytotic mechanisms into the extracellular aqueous compartment termed the alveolar hypophase. Within the alveolar hypophase, the lamellar arrays ‘‘unwind’’ in a Ca2⫹-dependent process to generate tubular myelin, a material that appears lattice-like when examined by electron microscopy. Tubular myelin serves as the extracellular reservoir for DPPC to form the monolayer at the interface between the alveolar hypophase and alveolar air. Generation of the monolayer within physiological time constraints requires the hydrophobic surfactant proteins (SP-B and -C), but much of the extracellular trafficking of surfactant components remains speculative. With cyclic contraction and expansion of the air/liquid interface associated with the respiratory cycle, there is alternate ‘‘squeeze-out’’ and recruitment of DPPC molecules into the interface. DPPC squeezed out from the monolayer forms small liposomal aggregates that are essentially devoid of protein and can be separated from the newly secreted, protein-enriched particles by centrifugation. These ‘‘spent’’ surfactant phospholipids are removed from the alveolar hypophase through endocytosis by type II epithelial cells, although the characteristics of ‘‘spent’’ material are undefined. Reuptake of phospholipids by type II epithelial cells occurs through two different mechanisms (3). The first is classical receptor-mediated endocytosis mediated by clathrin-coated pits. The ligand is SP-A, which binds with high affinity to DPPC and also to SP-A receptors in the coated pits of the type II cell membrane. The SP-A: DPPC complex then is internalized as a unit. The second pathway for DPPC uptake by type II cells is through internalization of cellular plasma membrane with DPPC bound to the extracellular face. DPPC appears to associate with specialized binding sites on the type II cell membrane, possibly

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representing patches of lamellar body membrane inserted into the plasma membrane during exocytosis. Retrieval of this excess plasma membrane by an actindependent process results in phospholipid internalization. Thus, type II epithelial cells are responsible for both secretion of surfactant and its reuptake, thereby constituting a surfactant cycle as illustrated in Figure 1.

Figure 1 Scheme for surfactant cycle in the granular pneumocyte (type II alveolar epithelial cell). The pathways are indicated for processing of surfactant phospholipid including transport of substrate at the basal membrane, synthesis of phospholipid in the endoplasmic reticulum (ER), transport of phospholipid and its storage in the lamellar bodies, secretion at the apical surface into the alveolar hypophase, reuptake in association with surfactant protein A through coated pits and into early endosomes, transfer to multivesicular bodies (MVB) and then to lamellar bodies, degradation in lysosomes, and reutilization of metabolic products for phospholipid resynthesis. A second pathway for phospholipid reuptake is through actin-dependent membrane retrieval. The precise pathways for intracellular transfer of endocytosed material are not well understood.

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C. Coordination of Surfactant Exo- and Endocytosis Because either insufficient or excess surfactant in the airspaces can lead to deranged respiratory function, the amount of extracellular surfactant must be tightly regulated. While the precise mechanisms are not understood, surfactant secretion and reuptake by the type II cells are coordinated, and a perturbation of one results in compensatory changes in the other. For example, agonists that activate PKC or PKA pathways (phorbol esters, β-adrenergic agonists, ATP) result in increased secretion of lung surfactant and also mediate increased surfactant uptake by the lung epithelium (4–7) as shown in Table 1. A possible mechanism has been described because these agonists also increase the cell membrane expression of receptors for SP-A through their recruitment from intracellular pools (8,9). Increased SP-A receptor density can result in increased SP-A binding and internalization with accompanying increased uptake of DPPC through the coated pit/ clathrin-mediated pathway. Physiologically, surfactant secretion is stimulated by increased lung ventilation such as that associated with vigorous exercise; surfactant clearance under these conditions is also stimulated with a gradual return to normal alveolar surfactant pool size (10). D.

Turnover of Alveolar DPPC

An important characteristic of the alveolar surfactant phospholipid is its relatively rapid turnover compared to most membrane bilayers. Studies with different mammalian species have indicated a biological half-time for lung surfactant PC in the alveolar space of approximately 16 h (11–14). This estimate for PC halftime is based on radiolabeled palmitate or glycerol as the PC marker. However,

Table 1 Uptake and Metabolism of Liposomal Dipalmitoyl PC by Isolated Perfused Rat Lunga

Uptake nmol in 2 h % of instilled Degradation nmol in 2 h % of uptake

No additions

8-Br cAMP

116 17%

249 37%

80 69%

197 79%

Lungs were instilled endotracheally with 1 µmol mixed lipid unilamellar vesicles containing 50% 3H-choline-labeled dipalmitoyl PC. 8-Br cAMP (0.1 mM) was added to the lung perfusate. PC ⫽ phosphatidylcholine. Source: Calculated from Refs. 5, 6, 17, and 18.

a

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the biological half-time for the choline moiety of PC is more than twice as long (42 h), compatible with extensive reutilization of the choline component. Turnover time of surfactant, estimated using precursor-product analysis, and assuming that lamellar bodies are the precursor for alveolar surfactant, indicates a PC turnover time of 4–11 h (11–14). These physiological results indicate degradation of PC and reutilization of components as well as possible recycling of PC to account for the discrepancy between PC turnover time and biological half-time. E.

Fate of Internalized DPPC

Dipalmitoylphosphatidylcholine that has been removed from the alveolar space by epithelial endocytosis is processed further within the type II granular pneumocyte as illustrated in Figure 1. Under normal conditions, a small fraction of the internalized DPPC is routed to lamellar bodies for resecretion (14,15). When demand for surfactant is high, such as in the neonatal period and perhaps in association with lung injury, the fraction of internalized DPPC that is resecreted through recycling is increased. Normally, the major fraction of internalized DPPC is degraded in the lysosomal compartment (16–20). The products released with DPPC degradation enter the general metabolic pool of the cell. Choline, the ratelimiting substrate for DPPC synthesis, is avidly retained by the type II cells through a high affinity membrane transport system (21–24). Degradation of DPPC inside the type II cell occurs through the action of phospholipases and other esterases as illustrated in Figure 2 (16,17). The initial step involves the action of phospholipase A2 (PLA2 ) to generate lysoPC. LysoPC is further degraded through the action of lysophospholipase to generate glycerophosphocholine (GPC) and then by esterases to glycerophosphate and choline. LysoPC also can be reacylated (via acyl transferases) in the endoplasmic reticulum to regenerate PC. This pathway is used to ‘‘remodel’’ PC, e.g., to generate saturated from unsaturated PC, but may also function in a ‘‘futile cycle’’ of DPPC degradation and resynthesis (18). Other phospholipases (A1 , C, and D) may participate in the degradation of internalized DPPC, but their role in lung surfactant turnover appears to be relatively minor. Evidence for the major role of PLA2 in degradation of internalized surfactant phospholipid has been obtained through studies with intact isolated perfused lungs as shown in Tables 1 and 2. Radiolabeled phospholipids instilled into the lungs through the trachea are taken up by the pulmonary epithelium. With radiolabel present in the sn-2 palmitate of DPPC, essentially all non-PC radiolabel was recovered in the free fatty acid fraction, indicating degradation of DPPC via phospholipase A2 activity (25); phospholipase C or D would have generated radiolabeled diacylglyceride directly or via phosphatidic acid. When DPPC labeled in the choline moiety was instilled in the rat lungs, the bulk of recovered radioactivity was in water-soluble metabolites, indicating that the lysoPC generated by

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Figure 2 Pathways for phosphatidylcholine (PC) degradation and resynthesis. LysoPC is generated through action of phospholipase A2 (PLA2) (A); lysoPC can be reacylated by acyl transferase (B) with acyl CoA to regenerate PC; lysoPC also can be degraded to constituent products including free choline or choline phosphate through the sequential action of lysophospholipase, nonspecific esterases, and phosphatases (C); choline (or choline phosphate) is reutilized for PC synthesis through the sequential activities of choline phosphate: CTP cytidyl transferase (D) and CDP choline: diacylglycerol phosphocholine transferase (E).

PLA2 was further degraded (6,17,25). In addition, the molecular species of choline-labeled PC showed the presence of unsaturated PC, compatible with generation of lysoPC from DPPC and its subsequent reacylation with an unsaturated fatty acid. Virtually identical results were obtained with rat granular pneumocytes in primary culture after uptake of DPPC liposomes added externally to the medium (16,26). These results indicate a central role for PLA2 in the intracellular processing of internalized surfactant phospholipids.

III. PHOSPHOLIPASE A2 Phospholipase A2 represents a family of enzymes that function to release fatty acid from the sn-2 position of phosphatidylcholine and other phospholipids.

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468 Table 2 Distribution of Metabolic Products of Dipalmitoyl PC Degradation in Isolated Perfused Rat Lunga Radiolabeled DPPC substrate 3

Metabolic product LysoPC Free fatty acid Aqueous metabolites Di- or triglyceride Unsaturated PC

H-Choline label (%)

sn-2-14C-Palmitate label (%)

2 0 65 0 33

⬍1 95 2 3 0

Lungs were instilled with 1 µmol mixed liposomes containing 50% labeled DPPC as described in Table 1. The percent of recovered metabolites in each fraction is indicated. Source: Ref. 25. a

These enzymes are widely distributed throughout the animal and plant kingdoms and participate in a broad range of physiological functions, including digestion, membrane turnover, cell signaling, and toxicity of venoms (27). The PLA2’s have been classified as either secreted or intracellular enzymes. The former are small proteins (approximately 15 kDa monomeric mass) that function predominantly in the extracellular milieu and require a relatively high (mM) Ca2⫹ concentration for activity. They have been classified into three or more ‘‘types’’ based on sequence homologies and biochemical characteristics (27). The intracellular PLA2’s are a much more diverse category. They comprise cytosolic PLA2 (cPLA2) and a heterogeneous (mainly intracellular) group of Ca2⫹-independent PLA2 (iPLA2). The former is a ubiquitous 85 kDa enzyme that has a relatively modest Ca2⫹ requirement (µM) and appears to be linked to eicosanoid metabolism associated with cell signaling processes. This enzyme is inhibited by fluoromethyl ketones. iPLA2 represents a large group of enzymes that have only recently come under investigation (28). Their role and properties are relatively little understood, although new examples are being described with increasing frequency. These enzymes, which are fully active in the absence of Ca2⫹, range in size (so far) from 25 to 85 kDa. An important example is platelet activating factor (PAF) hydrolase, a serum (extracellular) iPLA2 that degrades PAF, a phospholipid analog with an acetate in the sn-2 position (29,30). Bromoenol lactone (BEL) inhibits several members of this group. The optimum pH for in vitro assay of PLA2 enzymes is in the approximately neutral range (7–8), although an iPLA2 activity that is maximal at pH 4 has been described and is the major topic of the remainder of this chapter. Characteristics of the major classes of PLA2 are shown in Table 3.

Snake venoms, pancreatic secretions, inflammatory exudates Extracellular 13–18 mM; catalytic 8.5 Interface, Ca2⫹ His-Asp Phospholipid None p-Bromophenacyl bromide

Source

Localization

Molecular mass (kDa) Ca2⫹ requirement

pH optimum Activation Catalytic site Substrate specificity LysoPLase activity Inhibitor

87 µM; binding to substrate interface 7.4 Phosphorylation Ser 2-Arachidonyl PC yes Fluoromethyl ketones

Cytosol

Ubiquitous

Cytosolic PLA2 (cPLA2)

7.0 ? Ser 2-Acetyl PC (PAF) ? Serine protease inhibitors

40 None

Extracellular

Serum

PAF hydrolase

4.0 ? Ser PC None MJ33, SP-A, serine protease inhibitors

Lysosomes; secretory organelles 25 None

Lung epithelium, probably widespread

Lysosomal PLA2 (aiPLA2)

PC ⫽ phosphatidylcholine; PAF ⫽ platelet-activating factor; MJ33 ⫽ 1-hexadecyl-3-trifluoroethylglycero-sn-2 phosphomethanol; SP-A ⫽ surfactant protein A.

Secreted PLA2 (sPLA2)

Properties of Major PLA2 Classes

Property

Table 3

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A.

PLA2 in Lung Tissue

The lung is a cellularly heterogeneous organ, containing at least 40 major cell types. So it is hardly surprising that one or more examples of each major type of PLA2 (sPLA2 , cPLA2 , iPLA2) have been identified in lungs (31–36). In order to further evaluate the role of PLA2 in surfactant phospholipid degradation, we utilized various PLA2 inhibitors that are directed toward different PLA2 subtypes. The experimental models were the isolated perfused rat lung with DPPC instilled into the trachea or isolated type II cells in primary culture incubated with DPPC (25,26,37). AACOCF3 , a fluoromethyl ketone inhibitor of cPLA2 , had no effect on the rate of DPPC degradation by perfused lung or lung cells. p-Bromophenacylbromide (BPB), an inhibitor of sPLA2 through interaction with histidine at the active site, resulted in a modest inhibition of DPPC degradation. However, MJ33, a novel inhibitor synthesized by Dr. Mahendra Jain (University of Delaware) as an analog of the phospholipid transition state (Fig. 3), decreased the rate of DPPC degradation by approximately 50% as shown in Table 4. This important new inhibitor was effective at a concentration of only 2–3 mol% in mixed liposomes. These observations provided the basis to determine the specific PLA2 responsible for lung surfactant DPPC degradation. The biochemical characterization of PLA2’s as described above indicates two important basic parameters, namely Ca2⫹ requirement and pH optimum, that can be used to screen activities in a heterogeneous organ such as the lung. Accord-

Figure 3 Chemical formula for the transition state phospholipid analog inhibitor MJ33 (1-hexadecyl-3-trifluoroethylglycero-sn-2-phosphomethanol).

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Table 4 Effect of MJ33 and SP-A on Uptake and Degradation of Dipalmitoyl PC by Isolated Rat Granular Pneumocytesa

Uptake, µg/mg cell protein in 2 h Degradation µg/mg cell protein in 2 h % of uptake

Control

8Br-cAMP

MJ33 ⫹ 8Br-cAMP

SP-A

3.3

4.8

4.6

5.2

2.1 64

3.1 65

1.6 35

2.0 39

PC ⫽ phosphatidylcholine; SP-A ⫽ surfactant protein A. a Isolated granular pneumocytes (type II alveolar epithelial cells) after 24 h in primary culture were incubated with 1.2 µM lipid (unilamellar liposomes) containing 50% 3H-methyl choline-labeled dipalmitoyl PC. 8Br-cAMP was added at 0.1 mM, MJ33 at 3 mol%, and SP-A at 10 µg/mL. Source: In part from Ref. 45.

ingly, we assayed the effect of MJ33 on lung homogenate PLA2 activity in the presence or absence of Ca2⫹ (10 mM) and at pH 4.0 or 8.5 (25,35). Lung homogenate demonstrated considerable activity at both pH values in the presence of Ca2⫹ , although activity in the absence of Ca2⫹ was present only at pH 4. The presence of MJ33 at 3 mol% phospholipid resulted in marked inhibition of the Ca2⫹-independent PLA2 activity at pH 4, but Ca2⫹-dependent activity at pH 8.5 was unaffected (Fig. 4). BPB, on the other hand, markedly inhibited PLA2 activity of lung homogenate when measured at pH 8.5 (plus Ca2⫹) but had no effect at pH 4. The reducing agent, dithiothreitol (DTT), also inhibited activity at pH 8.5, compatible with the known importance of disulfide bonds in sPLA, but had no effect on the pH 4 activity. Thus, MJ33, when applied to lung homogenates, appears to specifically inhibit Ca2⫹-independent, acidic pH optimum PLA2 . We have named this enzymatic activity aiPLA2 . B. Isolation and Characterization of aiPLA2 Using MJ33 as a probe, a protein with the characteristic activity of aiPLA2 was purified to near homogeneity from rat and bovine lungs by anion exchange and hydrophobic interaction chromatography (35,36). Subsequent isolations of the protein were facilitated by development of a polyclonal antipeptide antibody based on the identified amino acid sequence (36,38). Amino acid sequencing, Nterminal for the bovine enzyme and internal after tryptic digest for the rat enzyme, identified a total of 49 amino acids for the two species. Of these, 48 showed identity to the deduced amino acid sequence for a cDNA of previously unknown function isolated from a human myeloblast cell line (35,36). Expression of this

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Figure 4 Phospholipase A2 (PLA2) activity of rat lung homogenate. Assay was carried out at pH 4 in the presence of EGTA or at pH 8.5 with 5 mM Ca2⫹. Additions included MJ33 (3 mol%), p-bromophenacylbromide (BPB, 20 µM), and dithiothreitol (DTT, 100 µM). Activity at pH 4 was inhibited by MJ33, whereas BPB and DTT had no effect. The inverse was observed for activity at pH 8.5. Source: Unpublished data.

human cDNA with a wheat germ in vitro expression system, in Xenopus oocytes, or in a human cell line (NIH3T3 cells) resulted in de novo appearance of aiPLA2 activity (35,39). We subsequently identified cDNA clones from rat, mouse, and bovine sources. Both the deduced amino acid and cDNA sequences are very well conserved with ⬎90% identity for all species. A dramatic recent finding is that the protein generated from the cDNA also demonstrates glutathione peroxidase activity and thus represents a bifunctional enzyme (39,40). A review of the literature has now demonstrated previous isolation of the protein from bovine (ciliary body of the eye) and rat (olfactory epithelium) sources, although these preparations were tested for glutathione peroxidase but not for PLA2 activity (41,42). The tissue source for isolation of aiPLA2 protein has been bovine and rat lung. Based on amino acid homology, the protein has been isolated from several other tissues, and the cDNA by Northern blot is widely distributed (38). However, Northern blot for DNA, Western blot for protein, and activity assay all indicated that the greatest expression of the protein is in the lung, especially the cells of the distal bronchioles (Clara cells) and alveolar epithelium (type II cells) (38).

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C. Properties of aiPLA2 The properties of the enzyme have been studied with partially purified protein isolated from bovine and rat lungs as well as enzyme expressed in vitro (wheat germ expression system) or through genetic engineering in E. coli (35,36,38). Unfortunately, enzyme generated by E. coli has been of relatively low specific activity, and the search continues for an improved expression system to generate sufficient quantities of active enzyme. The apparent molecular mass of the isolated protein on gel electrophoresis is 26–29 kDa under reducing conditions, compared to a monomer molecular mass of 25 kDa determined from the amino acid sequence deduced from the cDNA. Thus, the protein demonstrates little posttranslational modification. PLA2 activity is maximal at pH 4, with essentially no activity above pH 6. Activity is unaltered by the presence of Ca2⫹ or addition of EGTA. Phosphatidylcholine is a preferred substrate with an apparent Vmax for the partially purified bovine enzyme of 70 nmol/(min ⋅ mg protein) and apparent Km of 350 µM. Phosphatidylethanolamine is degraded at approximately half the rate observed for PC, while the enzyme is considerably less active with phosphatidylglycerol (PG), phosphatidylinositol (PI), and phosphatidylserine (PS) (Table 5). The enzyme shows no apparent acyl group specificity; PC with palmitate, oleate, or arachidonate in the sn-2 position of PC are equally effective substrates. Substitution of an alkyl bond for the usual acyl bond in the sn-1 position, i.e., an alkyl acyl PC substrate, reduced activity by approximately 50%. The enzyme has no phospholipase A1 or lysophospholipase activities. The enzyme was screened for the activity of various inhibitors. As described above, MJ33 is an effective inhibitor and was used as a probe for isolation of the enzyme. There was no effect of pBPB (an inhibitor of sPLA2 ), fluoromethyl ketones (inhibitors of cPLA2 ), or bromoenol lactone (an inhibitor of some iPLA2s), and activity was unaffected by the presence of a disulfide-reducing agent

Table 5 Substrate Specificity for aiPLA2 Substrate (a) PC PE PG PI PS

Relative activity

Substrate (b)

Relative activity

100 64 26 12 4

Dipalmitoyl PC sn-2 Arachidonyl PC Alkyl acyl PC LysoPC PAF

100 98 42 0 0

Activity was measured with enzyme isolated from bovine lungs and assayed with (a) fluorescent lipid substrates or (b) radiolabeled substrates in unilamellar liposomes. PC ⫽ phosphatidylcholine; PE ⫽ phosphatidylethanolamine; PG ⫽ phosphatidylglycerol; PI ⫽ phosphatidylinositol; PS ⫽ phosphatidylserine; PAF ⫽ platelet activating factor. Source: Ref. 36.

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(either DTT or mercaptoethanol). The enzyme was markedly inhibited by a variety of ‘‘serine protease’’ inhibitors. All secreted PLA2 enzymes have virtually identical architecture at the active site with a His-Asp pair required for catalysis. By contrast, the Ca2⫹-independent PLA2 enzymes characterized to date depend on serine for catalytic activity (30,43). The catalytic serine appears to be part of a ‘‘lipase’’ motif consisting of GXSXG. aiPLA2 activity is inhibited by ‘‘serine protease’’ inhibitors, and a lipase motif (GDSWG) at positions 30–34 of the deduced amino acid sequence is conserved for all species (human, rat, mouse, and bovine) (35,36,38). However, this sequence has not yet been identified definitively as the locus of activity.

D.

Physiological Regulation of aiPLA2 Activity

Because the enzyme requires low pH for activity, the intracellular localization of activity should be limited to specific subcellular compartments. Subcellular fractionation of lungs has indicated the presence of aiPLA2 in lamellar bodies, lysosomes, and cytosol; activity was not detected associated with the plasma membrane, mitochondrial, or microsomal fractions (36). The internal milieu of lysosomes and lamellar bodies is acidic for both organelles and therefore could support enzymatic activity in those compartments (44). However, the physiological significance of lamellar body activity is not known and is possibly regulated by changing pH during organogenesis and associated with the exocytotic process. The presence of aiPLA2 in the cytosolic fraction could be an artifact of the isolation procedure or could be the locus of peroxidase activity. Interestingly, aiPLA2 activity by in vitro assay is inhibited by surfactant protein A (SP-A) (45). Assay of aiPLA2 activity in lamellar bodies isolated from rat lung demonstrated 70% inhibition when 10 mg/mL SP-A was added to the cuvette (Fig. 5); no inhibition of activity was observed with PLA2 assayed at pH 8.5. Further evidence for a physiological role of SP-A was obtained by an experiment using a reducing agent, 2-mercaptoethanol, which in vitro had no effect on aiPLA2 activity but abolished the inhibition of activity by added SP-A. Treatment of isolated lamellar bodies with 2-mercaptoethanol resulted in a doubling of aiPLA2 activity (45). This result suggests that lamellar body aiPLA2 in its native form is inhibited by endogenous SP-A. Finally, the addition of SP-A to intact granular pneumocytes in primary culture resulted in a concentration-dependent decrease in the rate of metabolism of DPPC internalized by the cells (Table 4). The effective concentration of SP-A is within the physiological range, as contained in lamellar bodies, for example, suggesting that SP-A could function as a physiological regulator of aiPLA2 activity.

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Figure 5 Effect of surfactant protein A (SP-A) concentration on PLA2 activity of isolated rat lung lamellar bodies, the secretory organelle, assayed at pH 4 and pH 8.5. Addition of SP-A resulted in a concentration-dependent inhibition of acidic, Ca2⫹-independent PLA2 (aiPLA2) activity but had no effect on PLA2 activity measured at pH 8.5. (Reproduced from Ref. 45 with permission of the American Physiological Society.)

IV. SUMMARY This chapter has summarized studies carried out over a relatively extended period to investigate the mechanism for turnover of lung surfactant phospholipid. ‘‘Spent’’ phospholipid is removed from the alveolar space by endocytosis by lung type II epithelial cells. We have presented evidence that a newly described enzyme called acidic, Ca2⫹-independent phospholipase A2 (aiPLA2), is responsible for degradation of DPPC following its internalization. aiPLA2 is a 25 kDa protein that shows specificity for phosphatidylcholine, is maximally active at pH 4, and has no requirement for Ca2⫹. The catalytic site appears to be a conserved serine residue, and activity may be physiologically regulated by SP-A, the major surfactant-associated protein. aiPLA2 is greatly enriched in lysosomes and lamellar bodies of lung type II granular pneumocytes. Of particular note, the enzyme appears to be bifunctional and in addition demonstrates glutathione peroxidase

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activity. The relative importance of these two activities in various cell types remains a potentially fertile area for future investigation.

ACKNOWLEDGMENTS Original research described in this chapter was supported by grant HL 19737 from the National Institutes of Health, Bethesda, MD. I thank Chandra Dodia who generated a large fraction of the original data, Dr. Abu Al-Mehdi for assistance with illustrations, and my colleagues who have been part of our lung surfactant team, especially Drs. Avinash Chander, Sandra Bates, Henry Shuman, and Sheldon Feinstein. The advice and support of Dr. Mahendra Jain has been invaluable. I thank Elaine Primerano for typing the manuscript.

REFERENCES 1. Robertsen B, van Golde LMG. Pulmonary Surfactant. New York: Elsevier, 1992. 2. Rooney SA. Lung Surfactant: Cellular and Molecular Processing. Austin, TX: RG Landes, 1998. 3. Muller WJ, Zen K, Fisher AB, Shuman H. Pathways for the uptake of fluorescently labeled liposomes by alveolar type II cells in culture. Am J Physiol (Lung Cell Mol Physiol 13) 1995; 269:L11–L19. 4. Fisher AB, Dodia C, Chander A. Beta-adrenergic mediators increase pulmonary retention of instilled phospholipids. J Appl Physiol 1985; 59:743–748. 5. Fisher AB, Dodia C, Chander A. Secretagogues for lung surfactant increase lung uptake of alveolar phospholipids. Am J Physiol (Lung Cell Mol Physiol 1) 1989; 257:L248–L252. 6. Fisher AB, Dodia C, Chander A. Alveolar uptake of lipid and protein components of surfactant. Am J Physiol (Lung Cell Mol Physiol 5) 1991; 261:L334–L340. 7. Chinoy MR, Dodia C, Fisher AB. Increased surfactant internalization by rat type II cells cultured on microporous membranes. Am J Physiol (Lung Cell Mol Physiol 8) 1993; 264:L300–L307. 8. Chen Q, Bates SR, Fisher AB. Secretagogues increase the expression of SP-A receptors on lung type II cells. J Biol Chem 1996; 271:25277–25283. 9. Chen Q, Fisher AB, Strayer DS, Bates S. Mechanism for secretagogue-induced surfactant protein A binding. Am J Physiol (Lung Cell Mol Physiol 19) 1998; 275: L38–L46. 10. Nicholas TE, Power JH, Barr HA. Surfactant homeostasis in the rat lung during swimming exercise. J Appl Physiol 1982; 53:1521–1528. 11. Wright JR, Clements JA. Metabolism and turnover of lung surfactant. Am Rev Respir Dis 1987; 135:426–444.

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12. Wright JR. Clearance and recycling of pulmonary surfactant. Am J Physiol 1990; 259:L1–L12. 13. Wright JR, Dobbs LG. Regulation of pulmonary surfactant secretion and clearance. Annu Rev Physiol 1991; 53:395–414. 14. Fisher AB. Lung surfactant: clearance and cellular processing. In: Rooney SA, ed. Lung Surfactant: Cellular and Molecular Processing. Austin: RG Landes, 1998:165– 189. 15. Jobe A, Rider ED. Catabolism and recycling of surfactant. In: Robertson B, van Golde LMG, Batenburg JJ, eds. Pulmonary Surfactant. New York: Elsevier, 1992: 313–337. 16. Chander A, Reicherter J, Fisher AB. Degradation of dipalmitoyl phosphatidylcholine by isolated rat granular pneumocytes and reutilization for surfactant synthesis. J Clin Invest 1987; 79:1133–1138. 17. Fisher AB, Dodia C, Chander A. Degradation and reutilization of alveolar phosphatidylcholine by rat lungs. J Appl Physiol 1987; 62:2295–2299. 18. Fisher AB, Dodia C. Regulation of surfactant metabolism: degradation of internalized alveolar phosphatidylcholine. Prog Respir Res 1994; 27:74–83. 19. Rider ED, Ikegami M, Jobe AH. Intrapulmonary catabolism of surfactant-saturated phosphatidylcholine in rabbits. J Appl Physiol 1990; 69:1856–1862. 20. Rider ED, Pinkerton KE, Jobe AH. Characterization of rabbit lung lysosomes and their role in surfactant dipalmitoylphosphatidylcholine catabolism. J Biol Chem 1991; 266:22522–22528. 21. Fisher AB, Chander A, Dodia C, Reicherter J, Kleinzeller A. Choline transport by lung epithelium. Am J Respir Cell Mol Biol 1989; 1:455–462. 22. Fisher AB, Dodia C, Chander A, Kleinzeller A. Transport of choline by plasma membrane vesicles from lung-derived epithelial cells. Am J Physiol (Cell Physiol 32) 1992; 263:C1250–C1257. 23. Kleinzeller A, Dodia C, Chander A, Fisher AB. The Na⫹-dependent and Na⫹-independent systems of choline transport by plasma membrane vesicles of the A549 lung cell line. Am J Physiol (Cell Physiol 36) 1994; 267:C1279–C1287. 24. Oelberg DG, Fang X. Conductive choline transport by alveolar epithelial plasma membrane vesicles. Mol Genet Metab 1998; 65:220–228. 25. Fisher AB, Dodia C, Chander A, Jain MA. A competitive inhibitor of phospholipase A2 decreases surfactant phosphatidylcholine degradation by the rat lung. Biochem J 1992; 288:407–411. 26. Fisher AB, Dodia C. Role of phospholipase A2 enzymes in degradation of dipalmitoylphosphatidylcholine by granular pneumocytes. J Lipid Res 1996; 37:1057– 1064. 27. Dennis EA. Diversity of group types, regulation, and function of phospholipase A2. J Biol Chem 1994; 269:13057–13060. 28. Ackermann EJ, Dennis EA. Mammalian calcium-independent phospholipase A2. Biochim Biophys Acta 1995; 1259:125–136. 29. Tjoelker LW, Wilder C, Eberhardt C, Stafforini DM, Dietsch G, Schimpg B, Hooper S, Trong HL, Cousens LS, Zimmerman GA, Yamada Y, McIntyre TM, Prescott SM, Gray PW. Anti-inflammatory properties of a platelet activating factor acetylhydrolase. Nature 1995; 374:549–553.

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30. Tew DG, Southan C, Rice SOJ, Lawrence MP, Li H, Boyd HF, Noores K, Gloger IS, Macphee CH. Purification, properties, sequencing, and cloning of a lipoproteinassociated, serine-dependent phospholipase involved in the oxidative modification of low-density lipoproteins. Arterioscler Thromb Vasc Biol 1996; 16:591–599. 31. Ohta M, Hasegawa H, Ohno K. Calcium independent phospholipase A2 activity in rat lung supernatant. Biochim Biophys Acta 1972; 280:552–558. 32. Heath MF, Jacobson W. Phospholipases A1 and A2 in lamellar inclusion bodies of the alveolar epithelium of rabbit lung. Biochim Biophys Acta 1976; 441:443–452. 33. Lindahl M, von Schenck HM, Tagesson C. Isolation and characterization of phospholipase A2 from rat lung with affinity chromatography and two dimensional gel electrophoresis. Biochim Biophys Acta 1989; 1005:282–288. 34. Neagos GR, Feyssa A, Peters-Bolden M. Phospholipase A2 in alveolar type 2 epithelial cells: biochemical and immunologic characterization. Am J Physiol 1993; 264:L261–L268. 35. Kim TS, Sundaresh CS, Feinstein SI, Dodia C, Skach WR, Jain M, Nagase T, Seki N, Ishikawa K, Nomura N, Fisher AB. Identification of a human cDNA clone for lysosomal-type Ca⫹⫹-independent phospholipase A2 and properties of the expressed protein. J Biol Chem 1997; 272:2542–2550. 36. Akiba S, Dodia C, Chen X, Fisher AB. Characterization of acidic Ca2⫹-independent phospholipase A2 of bovine lung. Comp Biochem Physiol 1998; 120:393–404. 37. Fisher AB, Dodia C. Role of acidic Ca⫹⫹-independent phospholipase A2 in synthesis of lung dipalmitoyl phosphatidylcholine. Am J Physiol (Lung Cell Mol Physiol 16) 1997; 272:L238–L243. 38. Kim TS, Dodia C, Chen X, Hennigan BB, Jain M, Feinstein SI, Fisher AB. Cloning and expression of rat lung acidic Ca2⫹-independent PLA2 and its organ distribution. Am J Physiol (Lung Cell Mol Physiol 18) 1998; L750–L761. 39. Kang SW, Baines IC, Rhee SG. Characterization of a mammalian peroxiredoxin that contains one conserved cysteine. J Biol Chem 1998; 273:6303–6311. 40. Fisher AB, Dodia C, Manevich Y, Chen JW, Feinstein SI. Phospholipid hydroperoxides are substrates for non-selenium glutathione peroxidase. J Biol Chem 1999; 274:21326–21334. 41. Shichi H, Demar JC. Non-selenium glutathione peroxidase without glutathione Stransferase activity from bovine ciliary body. Exp Eye Res 1990; 50:513–520. 42. Peshenko IV, Novoselov VI, Evdokimov VA, Nikolaev YV, Shuvaeva TM, Lipkin VM, Fesenko EE. Novel 28-kDa secretory protein from rat olfactory epithelium. FEBS Lett 1996; 381:12–14. 43. Balboa MA, Balsinde J, Jones SS, Dennis EA. Identity between the Ca2⫹-independent phospholipase A2 enzymes from P388D1 macrophages and Chinese hamster ovary cells. J Biol Chem 1997; 272:8576–8580. 44. Chander A, Johnson RG, Reicherter J, Fisher AB. Lung lamellar bodies maintain an acidic internal pH. J Biol Chem 1986; 261:6126–6131. 45. Fisher AB, Dodia C, Chander A. Inhibition of lung calcium-independent phospholipase A2 by surfactant protein A. Am J Physiol (Lung Cell Mol Physiol 11) 1994; 267:L335–L341.

26 Role of Sodium Pump in Disease Zhimin (Tim) Cao* and Roland Valdes, Jr. University of Louisville, Louisville, Kentucky

I. INTRODUCTION Sodium-potassium-adenosine triphosphatase (Na,K-ATPase; E.C.3.6.1.37), referred to as the sodium pump, is a plasma membrane-associated protein expressed in most eukaryotic cells. It belongs to a P-type ATPase family that includes Na,KATPase, H,K-ATPase, and Ca-ATPase in mammals and H-ATPase in yeast (1). Members in the P-type ATPase family share both homology in amino acid sequence and mechanism of catalytic reaction in which a covalent phosphate intermediate forms. Na,K-ATPase is currently the only known receptor for the cardiac glycosides. Digoxin is a cardiac glycoside that is used primarily in the treatment of congestive heart failure. Much evidence now also suggests that endogenous ligands structurally similar to digoxin exist and act as regulator(s) of sodium pump activity in heart and other tissues. Identification and characterization of the endogenous naturally occurring ligands specific to Na,K-ATPase may lead to discovery of new hormone-like endocrine systems, a disturbance of which may be involved in the etiology of diseases depending on sodium pump function. Therefore, understanding the sodium pump, its endogenous ligands, and their relationship to disease is of central importance. In this chapter, we review progress to date in understanding Na,K-ATPase relative to its structure and function, expression and regulation, tissue distribution, and interaction with cardiac glycosides and discuss diseases linked to aberrant Na,K-ATPase function.

* Current address: Clinical Chemistry and Hematology, Wadsworth Center, New York State Department of Health, Albany, New York.

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II.

STRUCTURE, FUNCTION, AND SUBCELLULAR DISTRIBUTION OF Na,K-ATPase

A.

Structure and Function

Na,K-ATPase, an integral membrane protein, functions as an Na⫹ /K⫹ exchanger pump (sodium pump) by coupling ATP hydrolysis to the export of three intracellular Na⫹ ions and the import of two extracellular K⫹ ions (2,3). Normal Na,KATPase activity ensures maintenance of the concentration gradients of Na⫹ and K⫹ across the cell membrane, which is essential for many cellular biological functions including but not limited to 1. Generation and maintenance of the cell resting membrane potential, which is required for excitation and neuron transmitter release in nervous and muscular systems. 2. The membrane transport of metabolites and nutrients such as glucose and amino acids and ions such as protons, calcium, chloride, and phosphate. 3. Cell volume regulation and osmotic balance (i.e., inhibition of pump activity causes cell volume expansion as observed in human neuroblastoma cells) (4). 4. Fluid absorption in the lung (5). 5. The coated pit formation during receptor-mediated endocytosis (6). 6. The secretion process of fibroblast growth factor (FGF-2) (7). Figure 1 is a schematic diagram to describe the process of Na,K-ATPase in maintaining relative intra- and extracellular concentrations of Na⫹ and K⫹ ions (8,9). The Na,K-ATPase is composed of an alpha (112 kDa) subunit and a beta (55 kDa) subunit. The α-subunit has transmembrane (TM) domains varying in number from six to 10 (10,11) depending upon interpretations that were based on hydropathy profile (12), proteolytic results (13), and predicted α-subunit topology (14), where tentative models representing membrane-spanning segments of the α-subunit have been depicted. The α-subunit contains all the binding sites for Na,K-ATPase’s substrates and ligands. The binding site for ATP is located in a region of the cytoplasmic loop between the residues Lys354 and Lys774 (15). Various sites of phosphorylation have been identified. In one report, upon hydrolysis of ATP, phosphorylation occurred at the highly conserved residue Asp371 (16). However, phosphorylation at Asp369 has also been reported, and it is shown to be a site for pyrophosphate (Pi) interaction associated with ouabain binding (17). Both ATP and Pi can be substrates for the phosphorylation reaction. Phosphorylation by ATP requires the presence of Mg2⫹ , ATP, and Na⫹ ions, and it is sensitive to ouabain and other related cardenolides (18). Phosphorylation by Pi, while it requires Mg2⫹ , is stabi-

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Figure 1 (a) A schematic diagram of the plasma membrane-spanning Na,K-ATPase transporter complex indicating the positions of the alpha, beta, and putative gamma subunits. (b) One of the folding models for the Na,K-ATPase alpha transmembrane segments. This model has 10 transmembrane segments; others with eight segments have also been proposed. Letters refer to the one-letter amino acid code, and numbers represent the topological location of the particular amino acids. (From Refs. 13 and 223.)

lized by ouabain. The phosphorylation and dephosphorylation of the α-subunit accompany a conformational change of the α-subunit via a cycle of reactions by which Na⫹ and K⫹ ions traverse the membrane (19,20). Other phosphorylation events have been reported for the sodium pump. For instance, phosphorylation may occur during a regulatory process in which residue Ser943 is phosphorylated

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by protein kinase C and cAMP protein kinase A, resulting in the inhibition of Na,K-ATPase activity (21). The binding sites of the α-subunit for the cations Na⫹ and K⫹ are located at residues Glu327, Glu779, Asp804, and Asp808 on transmembrane segment 4 (TM4), TM5, and TM6, respectively (22,23). In addition, residues Asp804 and Asp808 are also involved in the enzyme catalytic reaction (24). The β-subunit is also an integral membrane protein with single hydrophobic transmembrane and intra- and extracellular domains. All the glycosylation sites and the sites for forming disulfide bonds are located on the extracellular domain (25,26). Identified functions of the β-subunit include stabilization and intracellular targeting of the α-subunit and involvement in the K⫹-dependent reactions (27,28). Expression of the β-subunit and assembly with the α-subunit are required for the α-subunit to obtain a correct conformation and to form an active Na,KATPase holoenzyme. The indispensable role of the β-subunit in providing a normal functioning sodium pump was evidenced by altered catalytic properties of the α-subunit ATPase when it was not associated with the β-subunit (29,30). In addition, the γ-subunit has been reported to be associated with the α/ β-subunit complex. The γ-subunit (10 kDa) of Na,K-ATPase is a hydrophobic trypsin-sensitive type I single-pass transmembrane protein. It has been found only in kidney tubules and may play a role in stabilizing the E1 conformations(s) of the enzyme as described by Sen and Tobin (19) and Therien et al. (31).

B. Isoforms of Na,K-ATPase To date at least three α-subunit and four β-subunit isoforms of the Na,K-ATPase have been identified, and they are encoded by separate genes (32,33). The homologous nucleotide sequence between these and other P-type ATPases suggests a common origin of ancestry (34). The epitopes responsible for binding the cardiac glycoside ligands are highly conserved between species, leading to a very stable and specific binding site (35). However, subtle differences do exist in the isoforms as to their sensitivity to cardiac glycosides, levels of transport activity, and even affinity for Na⫹. These, along with differential tissue responses to glycosides, implicate a structure base for differences in their functions. The expression and tissue-specific distribution of these various isoforms are important for several reasons: (1) an isoform can be predominantly and selectively expressed in one tissue and not another (36); (2) affinity of the α3-subunit to extracellular K⫹ and intracellular Na⫹ is different from those of the α1- and α2-subunit isoforms (37); and (3) response of the isoforms to regulators or affectors of Na,K-TAPase activity varies (e.g., insulin induces the subcellular translocation of the α2-subunit but not α1-subunit); to mention only a few (38). Tissue-specific distribution of the isoforms identified at both the mRNA and protein

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Table 1 Tissue Distribution of Na,K-ATPase Isoforms Tissue source Adipose Brain

Eye Heart (ventricle)

Kidney

Lung Skeletal muscle Thyroid Uterus

Subunit

Ref

α1, α2a,b β2a,b α1, α2, α3c,d α3a,d α1, α2, α3a,c,d α1, α2, α3, β1c,d α1, α2, α3c,d α1, α3 neonatea,b α1, α2 adulta,b α1, α3 ageda,b α1c,d α1b,d α1, α2, α3b,e α1, α2, α3, T-αb,f α1c,d α1, α2c,d α1, α2, β1, β2b,c α1, α2c,d α1c,d

39 40 41 42 43 44 41 45 45 45 41 46 47 48 41 41 49 41 41

Superscripts are used to indicate methods and tissue sources: a Isoform-specific expression of protein identified by Western blot analysis. b Rat tissue source. c Isoform-specific expression of mRNA identified by Northern blot analysis. d Human tissue source. e mRNA estimated by RT-PCR. f mRNA estimated by competitive RT-PCR. Source: Modified from Ref. 223.

levels of various species is summarized in Table 1. However, controversy still exists, partially due to the application of different detection techniques with different levels of sensitivity and specificity. A recent finding suggests identification of a truncated α-subunit in rat kidney (48), but its biological function remains a mystery. Various expression levels of the α-subunit isoforms associated with different stages of development have also been reported. Lucchesi and Sweadner (45) showed that the α1-subunit is expressed in all stages of development in rat ventricular muscle. However, the α3-subunit is present at birth through days 14–21

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and is then replaced by an α2-subunit in the adult rat. The α3 again predominates in aged rats. The physiological significance of the change in subunit isoform expression is unknown. The β1-subunit isoform, like the α1-subunit isoform, exists in a wide range of tissues from a number of vertebrate species. However, the β2-subunit isoform is mainly expressed in the brain (40,50) and was also found in ocular ciliary epithelia (51) and in the fast twitch muscles of the hindlimb (49). The β2-subunit isoform is not only a component of the Na,K-ATPase; it has also been found to be a mediator for neuron–astrocyte adhesion (52), suggesting that β2-subunitcontaining Na,K-ATPase may play a role in the neuron–astrocyte interaction. C. Interaction of the α- and β-Subunits of Na,K-ATPase The amino acid residues in the α-subunit that interact with the β-subunit were found by using the yeast two-hybrid system to be a four amino acid sequence of SYGQ located within the loop between TM7 and TM8 (26). However, a truncated isoform of the α1-subunit (α1T), which retains only the first 554 amino acid residues and terminates before the TM7, still possesses the ability to complex with the β-subunit and form a functional pump activity (53,54). This indicates that additional site(s) in the α-subunit may exist for interaction with the β-subunit. The sites responsible for the α–α interaction locate within a cytoplasm region of the α-subunit (G554–P785), which is different from that for interaction with the β-subunit (55). The β-subunit interacts with the α-subunit at the proximal stem of the β-subunit ectodomain (26). D.

Subcellular Distribution of Na,K-ATPase

Upon completion of synthesis, the α-subunit associates with the β-subunit in the lumen of the endoplasmic reticulum (ER) and the α/β-subunit complex is targeted to the cytoplasmic membrane, functioning as an Na⫹ /K⫹-pump (56). In polarized epithelial cells Na,K-ATPase distributes to a restricted cell surface locale, e.g., to the apical of retinal pigmented epithelium (57) or the basolateral of kidney (58) where the enzyme (ion transporter) plays a role in epithelial secretory and absorptive processes. Initial studies on the mechanism of the Na,K-ATPase’s specific subcellular distribution using the MDCK cell line (clone J cells) revealed that Na,K-ATPase was randomly targeted to the cytoplasmic membrane (59), where its interaction with cytoskeletal elements occurs (60), resulting in stabilization of the complex. It was postulated that the extent of the enzyme stabilities at the various surface areas of the cells varied and therefore the half-life differed (59). However, further studies using MDCK cells but not the clone J cells demonstrated that the polarized distribution of the Na,K-ATPase on the cell surface results from the existence of a sorting signal located within the NH2-terminal 519

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amino acids of the α-subunit (61), and the interaction with cytoskeletal components appears not to be the only mechanism (62). The sorting signal in the αsubunit dominates the signal in the β-subunit (61). The controversy between Hammerton et al. (59) and others was perhaps due to unique sorting behaviors of the clone J cells caused by the absence of glucosphingolipid polarity from the plasma membrane (63). A partial or complete reverse polarity of the enzyme distribution at the cell surface has been observed in polycystic kidney disease and ischemic condition, implying that specific subcellular distribution of the enzyme is of importance in maintaining normal organ functions (64,65).

III. REGULATION OF SODIUM PUMP ACTIVITY Regulation of Na,K-ATPase is a complicated topic because of its molecular isomeric nature and the multiple functions of the molecule, i.e., the existence of different isoforms, developmental variation in expression levels and individual isoforms, tissue-specific distribution, and regulation by a number of hormones and endogenous cardiac glycoside-like compounds. Reviews by Ewart and Klip (66) and Lingrel et al. (12) cover broad yet substantial details of this aspect. More recent findings supplement those excellent reviews. The α1-subunit is constitutively expressed in a wide variety of tissues. The promoter region of the gene has been located between ⫺77 and ⫹17 of the transcription initiation site containing a binding site for ATF1-CREB and a GC box where the Sp1/Sp3 binds, which are essential for the promoter activity (67). The α3-subunit expression is the most tissue-specific, predominantly expressed in the brain. A regulatory element conferring its brain-specific expression was found within 210 bp upstream of the transcription initiation site (68). Within this region were also found both negative and positive regulatory cis elements. These facts and the finding of α3-subunits in cultured neonatal rat cardiocytes indicate that expression of α3-subunits may be developmentally regulated (69). The regulatory element of the β1 subunit was located within a 21 base pair region (⫺650 to ⫺630) in the β1 gene promoter. An interaction of mineralocorticoid receptor and glucocorticoid receptor with the β1 promoter results in downregulation of transcription (70). A cis-transcription element of the β2-subunit contains a binding site for Sp1 at position ⫺120 of the transcription initiation site, and a binding of Sp1 to the glia regulatory element results in a positive regulatory effect of astrocytes (71). Various endogenous and exogenous modulators have been shown to affect Na,K-ATPase activity, and a summary is provided below. Aldosterone (72), angiotensin II (73,74), arginine vasopressin (75), cAMP (76), calnaktin (77), dopamine (78), dynorphin A (79), endobains (80),

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estrogens (81), glucocorticoids (82), keratinocyte growth factor (KGF) (83), mineralocorticoids (84), potassium (depletion) (85), serotonin (86), SPAI-1 (endogenous peptide) (87), thyroid hormone (T3) (88,89), antisickling/anaesthetic substances/ionophoric antibiotics (90), cyclosporin A (91), desipramine (92,93), ethanol (94), hyperoxia (95), halothane (5), long-chain acyl coenzyme A (96), propionic acid (97), thymosin alpha-1 (98), 12 (R) HETE (unsaturated fatty acid) (99). Be aware that some of these are still not conclusive and remain controversial. A.

Cardiac Glycosides and Na,K-ATPase

Currently Na,K-ATPase is the only known cellular membrane protein that specifically binds cardiac glycosides. Therefore, Na,K-ATPase has been considered a receptor for those ligands. The ligand–receptor relationship has been a biochemical basis for applying digitalis in the treatment of congestive heart failure and arrhythmias. This section focuses on cardiac glycosides of this nature, their mechanism of action, and some potential problems in therapeutic drug monitoring. Readers are also referred to the reviews by MacGregor and Walker (20) and Jortani and Valdes (100). Cardiac glycosides are extracts from the plant genera Digitalis, Strophanthus, and Acocanthera. Digoxin and digitoxin are products of the foxglove plant Digitalis, and ouabain is a product of the East African ouabaio tree or seeds of the plant Strophanthus gratus (101,102). These extracted substances, because of their cardiac inotropic effect, have been applied in medical use for more than 1000 years (103). Clinically, digoxin is most widely utilized because it can be administered orally, is readily absorbed by the gastrointestinal tract, and has a clinically effective half-life. Ouabain is the most widely used for experimental purposes. Structurally, these cardenolides have a steroid nucleus with one or more sugars at position C-3 and a five-membered lactone ring at position C-17 (Fig. 2). A major difference in structure between digoxin and ouabain lies in the sugar moiety and -OH groups, which may contribute to their differences in affinity and biological responses upon binding to the Na,K-ATPase. Noel et al. (104) demonstrated that both the lactone ring at C-17 and the side chain at C-3 are critical components for an interaction of the ligand digoxin or ouabain with the receptor Na,K-ATPase. Two hairpin loops (TM1–TM2 and TM5–TM6) and a transmembrane segment (TM7) have been found to be the major determinants of the ouabain binding site and therefore of ouabain-mediated inhibition (105–107). Part of the binding site determinant (TM5–TM6) is adjacent to the cation transport sites located in TM5 (23), providing a structural base to the antagonistic effect of cations (potassium in particular) on ouabain binding and a possible mechanism for ouabain or

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Figure 2 Structures of the cardiac glycosides (a) ouabain and (b) digoxin. Three digitoxose sugars (in digoxin) and one rhamnose sugar (in ouabain) are attached at the C-3 position of the steroid backbone. (From Ref. 223.)

other cardiac glycoside to inhibit the Na,K-ATPase, as proposed by Palasis et al. (107). In addition, the residues Glu111, Asp122, Phe786, Leu793, and Phe863 have been identified as determinants of ouabain sensitivity (107,108). For further description, see the review by Lingrel et al. (109), who also proposed two models for ouabain binding and inhibition of pump activity. The human heart contains all three α-subunit isoforms of Na,K-ATPase; on the other hand, the pig heart contains only α1. This may explain the reduced response to cardiac glycosides by pig heart recently reported (110). The cardiac myocyte’s excitation–contraction coupling is initiated after the action potential

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is transmitted to the myofibrils, followed by Na⫹ channel opening, depolarization of the sarcolemmal membrane in response to increased intracellular sodium, and Ca2⫹ channel opening. The increased intracellular free Ca2⫹ as a result of Ca2⫹ influx from extracellular space and release from the intracellular compartments binds to troponin C, consequently releases the inhibitory effect of troponin I, relocates tropomyosin T, and allows myosin to interact with actin. This chain of events results in myocardial contraction. In the presence of cardiac glycosides, Na,K-ATPase activity is inhibited, which results in a transient increase in intracellular sodium concentration and Ca2⫹ concentration by activating Na⫹ –Ca2⫹ exchange (111). As a result, the cardiac contractile force is enhanced (inotropic effect) (1). Somberg et al. (112) determined that a 25% reduction in pump activity was associated with a 20% increase in contractile strength. In clinical practice, digoxin toxicity is a very commonly encountered adverse drug reaction. Symptoms of digoxin toxicity can be manifested in the gastrointestinal system (e.g., anorexia, nausea, vomiting), the central nervous system (e.g., headache, confusion, alteration of color perception), and the cardiovascular system (e.g., dysrhythmia, atrial, or ventricular fibrillation). Many factors contribute to the occurrence of digoxin intoxication including inappropriate dosing, interaction with coadministered drug(s) (113), and various physical and pathological conditions that lower the threshold of the tolerance to digoxin. In addition, decreased Na,K-ATPase activity has been reported with aging (114). This indicated that while serum digoxin levels may be within the therapeutic range (0.8– 2 ng/mL), digitalis toxicity may occur in the elderly due in part to inhibition of an already less active sodium pump. Due to the existence of severe adverse effects of digoxin, narrow therapeutic index, and numerous factors affecting the elimination of digoxin, therapeutic drug monitoring for digoxin is a critical process to ensure a therapeutic effect and avoid an adverse reaction. However, a misleading decision may result from preanalytical and analytical errors (115). The following error-causing factors and conditions need to be considered during therapeutic drug monitoring for digoxin: existence of biologically active and inactive digoxin metabolites with variable reactivity with Na,K-ATPase and cross-reactivity of digoxin immunoassays (116,117); cross-reactivity of antibodies against digoxin with endogenous steroidlike hormones and digoxin-like immunoreactive factors (118); inconsistent results from assays using different immunoassay methods (117); interference by the presence of Digibind (digoxin antidote) (119); and interference by ingestion of exogenous cardenolide-like compounds that cross with digoxin antibodies (120). In addition, certain biological conditions also need to be considered, such as the difference in affinity of various Na,K-ATPase isoforms for digoxin binding (104) and alteration of isoform expression in response to aging, hormonal levels, and some pathological conditions (36). Among those conditions, the key issues are to improve specificity and to measure the biologically active form of digoxin.

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A pharmacodynamic and pharmacogenetic issue worth considering is whether the expression levels of a recently identified digoxin-binding protein (121) should be evaluated, especially for those patients who are elderly. B. Endogenous Digoxin-Like Immunoreactive Factors Several classes of compounds with digitalis- or ouabain-like activity have been identified in mammals (122). Here we limit our discussion mostly to those in the digoxin-like immunoreactive family because they more greatly influence the measurement of and drug treatment with digoxin. The therapeutic advantage of using immunoassays to measure digoxin has also led to the identification of digoxin-like immunoreactive factors (DLIFs) or endogenous digitalis-like factors (EDLFs). Gruber et al. (123) first described the presence of a digoxin-like immunoreactive substance in blood of volume-expanded dogs. Subsequently, Valdes et al. reported the presence of similar compounds in humans as factors that were originally defined as giving rise to apparent digoxin values where no digoxin was present. The initial findings included the fact that elevated apparent digoxin levels remained after discontinuation of digitalis therapy (124) and detectable digoxin levels were obtained in the subjects not treated with digoxin (125,126). The population of subjects showing this phenomenon soon included premature infants (125), newborns (127,128), healthy adults (129), pregnant women (130), and patients with renal impairment (131) or liver dysfunction (132,133). These observations indicated the existence of endogenous substances cross-reacting with the antibodies against digoxin used in the immunoassays. Most studies now indicate that a family of compounds designated as digoxin-like immunoreactive factors (DLIFs) are endogenous mammalian molecules (MW 780 Da) with substantial structural similarity to digoxin (118). Evidence from many laboratories also suggests that these compounds interact at the digitalis binding site on the α-subunit of Na,K-ATPase (105–108). However, DLIF was estimated to have about 1000-fold less relative molar immunoreactivity than digoxin using specific antibodies directed at the lactone portion of digoxin (134). Most antibodies to digoxin are sensitive to the lactone ring moiety, and deglycosylation (removal of the digitoxose sugars) does not affect their reaction to the antibodies. However, some antibodies are sensitive to deglycosylation (117). Other putative mammalian endogenous compounds with and some without immunological cross-reactivity with antibodies against digoxin demonstrate the capability to inhibit Na,K-ATPase. These include DLIF (118), ouabain-like-factors (OLFs) (135), hypothalamic inhibitory factor (HIF) (136,137), a labile digoxin-like factor (DLF) (138), mammalian bufodienolide(s) (139), and some newly described species of digoxin-like immunoreactive substances (140,141). In focusing on DLIF, again because of its effects on therapeutic drug measure-

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ments, recent evidence strongly suggests that DLIFs belong to a family of structurally related species with common molecular origin. For example, both the intact DLIF, its genin derivative (no sugars), its mono- and bis-DLIF components, and even, most interestingly, its reduced lactone ring hydrogenated component (dihydro-DLIF) have been identified (134,142). Establishing criteria for defining DLIFs is important. Presently, DLIFs have the following defining properties (118,134,142): They cross-react 1000-fold less with antidigoxin antibodies specific to the lactone ring on digoxin; have maximal absorbance at 216 nm; sequentially deglycosylate under acid conditions to their bis-, mono-, and genin-DLIF components; are more sensitive to acid deglycosylation than is digoxin; and have dihydrodigoxin counterparts (dihydro-DLIFs) with maximal absorbance at 196 nm. And all these individual DLIF species separate chromatographically in a physical manner similar to but distinct from that of their plant-derived counterparts of digoxin. It is important to understand that immunoreactivity does not in itself imply functional sodium pump inhibitory activity or vice versa. Thus, terminology is very important, and the criteria being used to describe these compounds need to be clearly established in published articles (126,143). Determination of the origin of DLIFs and other sodium pump inhibitory compounds in the body is critical for establishing the concept that they are truly endogenous compounds. Available data indicate that the endogenous compoundproducing organs are likely the adrenal gland for DLIF (118) and OLF (137,144,145) and the hypothalamus for HIF (136). Since biophysical analysis suggests that human OLF and bovine HIF are identical but distinct from plant origin ouabain (146), OLF/HIF are likely of endogenous origin rather than arising from intake or contamination from the environment. Studies on tissue distribution of DLIFs in human and rat by Shaikh et al. (118) found that the adrenal cortex contained DLIF concentrations over seven times higher than the adrenal medulla and any other organs tested, including serum, brain, heart, lungs, liver, and kidney. DLIFs isolated from serum shared similar physical properties with DLIFs isolated from the tissues. A fourfold gradient difference in the DLIF concentrations between the serum samples from lumbar vein and infrarenal inferior vena cava of dogs was also observed (118). These results indicate that the adrenal cortex is likely the principal source of DLIFs. Controversy still remains, and more definitive studies are needed (147). The metabolism of DLIFs is still unknown. For an anabolic pathway, existing evidence suggests that dihydro-DLIF may be an immediate precursor for DLIF biosynthesis, and this transformation is mediated by an NADPH-dependent cytochrome P-450 system in adrenal tissues (142). For a catabolic pathway, the deglycosylation process of DLIF at the C-3 position may be steps in its metabolism (134). In contrast, a study on the biosynthetic pathway of digoxin-immunoreactive substance using rat adrenal cortex cells indicated that hydroxycholesterol and pregnenolone are the intermediates, and a process of side chain cleavage

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could be the first step in the synthesis of digoxin-immunoreactive substance from hydroxycholesterol (148). These latter data still have to be confirmed as applied to the molecule DLIF specifically. Early during the discovery of DLIFs, initial data suggested (129) that these compounds were tightly bound to a protein in serum that was not albumin and had an apparent molecular weight between 22 and 36 kDa (149). Recently, isolation and identification of a cardiac glycoside-binding protein (CGBG) from the globulin fraction of bovine serum (121) seemed to confirm those initial findings. The CGBG protein has been found to be a homodimer with a molecular mass of 26 kDa for each subunit, and it also binds ouabain and bufadienolide proscillaridin A but not the common steroids even at high concentrations. In addition, intracellular CGBG-protein with a molecular mass of about 90 kDa was found in the bovine kidney using antibodies against the serum form of CGBG-protein. The cytosolic CGBG was found to have two negative cooperative binding sites for ouabain. These findings strongly link the origin and target organs of DLIFs or other endogenous mammalian cardiac glycosides and support the hypothesis that these in fact may be regulated and transported hormones.

IV. CLINICAL AND PHYSIOLOGICAL CONDITIONS LINKED TO Na,K-ATPase ACTIVITY A number of diseases have been linked to altered sodium pump activity. Here we review only some of the major ones including hypertension and cardiovascular disease, diabetes, neonatal abnormality, and neurological disorders. We also review other clinical and physiological conditions where putative endogenous inhibitors of the pump have been reported to have increased in the blood. By way of definition, here we use endogenous cardenolide-like inhibitors in a general sense to include compounds such as DLIF, EDLF, OLF, and HIF and not limited to any one of these types of compounds since there is still some controversy over their definition. A.

Hypertension and Cardiovascular Disease

Essential hypertension accounts for more than 90% of all hypertension cases and is defined as an elevation of systemic arterial blood pressure with no definable cause. Pathophysiology of hypertension could be simply described as an imbalance between the cardiac output (or blood volume) and peripheral vascular tension activity. The etiology of essential hypertension is unknown; however, an inherited predisposition has been suggested, and affected individuals may be especially sensitive to dietary salt (150). The Na,K-ATPase has been proposed to play a role in hypertension etiology because it is considered crucial to maintaining

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sodium–potassium–water homeostasis. Because Na,K-ATPase affects smooth muscle reactivity and myocardial contractility, it is believed to be involved in systemic vascular hypertension (151). Evidence suggest links between hypertension and involvement of Na,KATPase as mediated by endogenous digoxin- or ouabain-like factors. Early experiments aimed at understanding the relationship between renal hypertension and Na,K-ATPase were conducted using rat one-kidney, one-clip and reduced renal mass saline models of hypertension (152,153). The results showed a significant decrease in Na,K-ATPase activity in microsomal preparations from both the left and right ventricles of hypertensive rats. Upon restoration of the blood pressure by removal of the clips, Na,K-ATPase activity returned to normal. In addition, upon suppression of the vascular Na,K-ATPase with ouabain, the vascular contractile activity and the contractile responses to vasoactive agents increased (154). In the Dahl salt-sensitive/JR rat model, a mutation A1079T(Q276L) was identified in Na,K-ATPase α1-subunit cDNA (155). The mutation alters the hydropathy profile of a region in proximity to TM3 and causes a decrease in the net 86Rb⫹ influx (indicating K⫹ influx). A correction of the mutation in transgenic Dahl S rats exhibited less salt-sensitive hypertension, less hypertensive renal disease, and longer life span than the native mutation (Q276L)-containing transgenic Dahl S controls (151). These findings indicate that Na,K-ATPase and a change of its activity may play a role in the etiology and maintenance of the hypertension, and the α1-subunit gene may be linked to salt-sensitive hypertension. Are endogenous compounds with cardenolide-like activity involved in the pathogenesis of hypertension? Three molecular species having such activity were chromatographically isolated from peritoneal dialysis fluid from patients with chronic renal failure, one having a retention time identical to ouabain and one with a retention time identical to digoxin (156). A higher amount of OLF was also found in the tissues (hypothalamus/hypophysis) and plasma of Milan hypertensive strain rats than that of normotensive rats (157). Yuan et al. (158) showed that administration of ouabain at low doses for long periods of time was associated with development of hypertension in normotensive rats as well as in rats having varying degrees of reduced renal mass (RRM). These data indicate an association of hypertension and multiple endogenous ATPase inhibitors (139). Do compounds like DLIF or OLF facilitate development of hypertension via their effect on Na,K-ATPase activity? Most results of studies support the idea that a circulating sodium pump inhibitor contributes to the pathogenesis of volume-dependent hypertension, but this is still inconclusive. Krep et al. (159) showed that both ouabain and an endogenous sodium pump inhibitor could induce vascular smooth muscle (VSM) contraction, and administration of Digibind (an Fab fragment antidote to digoxin poisoning that binds digoxin and other cardenolides avidly) was able to rapidly reverse the contraction. It has also been reported that Digibind reversed rat model hypertension induced by ACTH (160)

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or DOCA-salt (a volume-dependent model) (161). Substantial evidence also indicates the effects of digoxin- or ouabain-like factors on vasoreactivity: 1. The plasma of primary hypertensive patients has an elevated Na,KATPase inhibitory activity (153), natriuretic activity (162), and digoxin immunoactivity (117). 2. Spontaneous hypertensive rhesus monkeys have elevated digoxin-like activity (163). 3. DLIF isolated from the plasma of volume expanded dogs constricts third-order arterioles, making them more responsive to noradrenaline (164). 4. Urine preparations with natriuretic activity also cause dose-dependent contractions of isolated anococcygeus muscle of the rat (resembles smooth muscle of blood vessels) (165) and raise blood pressure (166). 5. Peripheral vasoconstriction can be induced by an infusion with ouabain (167,168) or digoxin (169). 6. Blood pressure of hypertensive animal models can be reduced by administration of antidigoxin antibodies (170). 7. Digoxin infusion at high dose can induce constriction of coronary arteries (171). 8. OLF from human serum causes vasoconstriction (172). The evidence supports the hypothesis that endogenous digoxin- or ouabin-like compounds are involved in the etiology and development of the hypertensive state via their effect on the Na,K-ATPase. However, unsupportive data also exist; e.g., spontaneously hypertensive rats (SHR) that had been actively immunized against cardiac glycosides did not show an improvement in blood pressure, in spite of their elevated levels of endogenous cardiac glycosides (173). B. Diabetes Mellitus-Related Hypertension Diabetes mellitus is an endocrinological disease with characteristics of metabolic disorders and long-term complications. It is of interest to understand if and how Na,K-ATPase and endogenous cardenolides play a role in the pathogenesis of diabetes mellitus-associated hypertension. Findings that Na,K-ATPase activity is also regulated by insulin have established a biochemical basis to this aspect. Insulin stimulates cellular glucose uptake through a glucose transporter and also increases cellular K⫹ intake by stimulating Na,K-ATPase activity (174). In addition, proinsulin C-peptide was also found to have an effect similar to that of insulin on Na,K-ATPase, with the functional domain located on the C-terminal of the C-peptide (175,176). Clinical studies recently demonstrated a decrease in Na,K-ATPase activity of erythrocyte membrane from NIDDM patients (177) and an increase in serum

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of pump inhibitors from patients with NIDDM and patients with essential hypertension and hyperinsulinemia (178). However, it is uncertain if the elevated level of compounds in serum is a result of, or associated with (1) sodium retention/ volume expansion, or (2) diabetic nephropathy, or (3) hyperinsulinemia. The key issue is whether abnormal levels of these endogenous sodium pump inhibitors could result in pathophysiological consequences, i.e., in diabetic patients, if compounds like DLIF contribute to the development of hypertension and/or abnormality in insulin sensitivity or insulin secretion. Readers are referred to reviews by Clerico and Giampietro (179) and Martinka et al. (180) for further evidence in support of the theory that DLIF (and other related inhibitors) is involved in diabetes mellitus-associated hypertension. A review by Weidmann and Ferrari (181) describes possible mechanisms of Na⫹ retention in both type I and type II diabetic patients that may significantly contribute to the development of diabetic hypertension. Based on the current understanding, a relationship between Na,K-ATPase activity, its inhibitors, insulin, and hyperglycemia can be described as Na,KATPase activity being inhibited by both hyperglycemia and pump inhibitors but upregulated by insulin. The evidence includes findings that (1) protein-free plasma from insulin-dependent diabetic patients decreased purified Na,K-ATPase activity (182); (2) insulin at physiological concentrations activated the sodium pump and reversed the inhibition of the pump activity by hyperglycemia (183); and (3) in experimental hyperinsulinemic/hypertensive rats, vascular Na,KATPase pump activity increased its sensitivity to insulin (184). Based on these findings, it has been postulated that a change in the response of vascular Na,KATPase activity to insulin may be followed by changes in vascular responsiveness associated with hypertension. How does insulin regulate Na,K-ATPase? Regulation of Na,K-ATPase by insulin could occur by various mechanisms. For example, 1. Gene transcription of the α2-subunit (α1 transcript is not affected by insulin) (185). 2. Translocation of Na,K-ATPase from the endoplasmic reticulum (ER) to the cytoplasmic membrane, which again, is α2-isoform-specific (38). This finding could be very significant because the α2-isoform is found predominantly in rat skeletal muscle, which accounts for a major portion of body mass (36). 3. Phosphorylation/dephosphorylation. The dephosphorylation of the αsubunit is catalyzed by an insulin-dependent serine/threonine protein phosphatase-1 (PP-1), and the process is mediated by phosphatidylinositol-3 kinase-generated signals resulting in activation of Na,K-ATPase (186).

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C. Neonatal Abnormalities A connection among neonate disorders, Na,K-ATPase, and endogenous sodium pump modulators has been shown in mainly two areas: neonate electrolyte imbalance and neonatal myocardium hypertrophy. Available data that suggest such connections are summarized as follows: 1. Na,K-ATPase isoforms switch from α3 to α2 during the neonatal developmental stage observed in rat heart (45). 2. DLIF is significantly increased in the plasma of newborns but usually returns to normal within several days after birth (127,187). 3. Increased levels of digoxin-like immunoreactivity have been observed in perinates with various pathological conditions including growth retardation, renal abnormality, hydrocephalus, aneuploidy, and nonimmune hydrops (188). 4. Ouabain at subtoxic concentration was shown not only to inhibit Na,KATPase but also to cause neonatal rat myocardium hypertrophy (189,190). This was found to be a result of a downregulation of the α3-subunit by ouabain via signal pathways that involve Ras or/and p42/44 mitogen-activated protein kinases (MAPKs) (190). These observations suggest that Na,K-ATPase and molecules like DLIF or OLF are involved in pathogenesis of these disorders, but again the relationships remain to be defined. D.

Neurological Disorders

The importance of Na,K-ATPase in maintaining normal neurological physiology is indicated by study of simple life forms. For instance, mutant worms (Caenorhabditis elegans) containing an amino acid change (Leu to Phe) in a highly conserved region in the α-subunit Na,K-ATPase showed abnormal feeding behavior (191). Human neurological abnormalities associated with altered Na,KATPase function have been described in bipolar illness, Alzheimer’s disease, McArdle disease (192), and an animal model mimicking neurological disorder of patients with propionic acidemia or ketotic hyperglycinemia (97). Since very little is known about the latter two disorders, in the following we focus on bipolar illness and Alzheimer’s disease. Bipolar illness (manic depression) is a disorder characterized by severe mood swings from manic status, with characteristics of episodes of irritability, excessive energy, and distractibility, to depression such as mental and motor slowdown to the point of stupor, or vice versa. Bipolar illness patients also exhibit altered ion distribution and transport. Since one of the Na,K-ATPase biological

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functions in neuronal systems is to maintain neuronal cell membrane potential, El-Mallakh et al. (193) proposed that in the biphasic phenomenon a mild or moderate reduction in Na,K-ATPase activity could lead to mania by increasing membrane excitability and neurotransmitter release. A greater degree of pump inhibition, and consequently depolarization block, could result in depression by decreasing neurotransmitter release (194). In patients with manic states, sodium retention and intracellular calcium concentration are higher than normal. Therefore, therapeutic modalities such as lithium or calcium channel blockers are theorized to restore sodium and calcium homeostasis by affecting sodium–calcium exchange (193). Interestingly, symptoms mimicking those observed in bipolar patients occur with digitalis toxicity. These can include confusion, disorientation, drowsiness, lethargy, agitation, and hallucinations (195). All these findings suggest that the Na,K-ATPase may be involved in the pathophysiology of bipolar illness. Results from experiments using samples from humans or using animal models further support the hypothesis that Na,K-ATPase associates with bipolar illness. The Na,K-ATPase in erythrocytes from lithium-free patients with bipolar manic depression showed a significantly lower activity than normal, and the decreased activity can be normalized by lithium therapy (196,197). However, this change in Na,K-ATPase activity was not seen in patients with neurotic depression, or schizoaffective disorder, or schizophrenia. These results indicate that Na,K-ATPase activity directly associates with bipolar illness and that a decrease of its activity in bipolar affective disorder can be trait-dependent. Naylor et al. (198) reported that the reduced Na,K-ATPase activity in manic depressive psychosis was not caused by a change in Na,K-ATPase molecule concentration in cell membrane. Evidence from our laboratory, using a digoxin-specific immunoassay, suggests that DLIF is elevated in acutely psychiatrically ill bipolar patients compared to recovered bipolar patients (unpublished). Increased endogenous ouabain-like compounds that suppress the Na,K-ATPase have been proposed as a mechanism in this disorder (199). An animal study in which rats were administered ouabain at sublethal doses intracerebroventricularly demonstrated a significantly increased motor activity, indicating that an inhibition of Na,K-ATPase in the central nervous system may play a pathophysiological role in bipolar illness (200). A new finding by Mynett-Johnson et al. (201) that a Na,K-ATPase αsubunit gene (ATP1A3) may be responsible for bipolar disorder further solidifies the hypothesis of El-Mallakh (200). Alzheimer’s disease (AD) is a neurodegenerative disorder with characteristics of decreased choline acetyltransferase activity, amyloid beta-peptide deposits, senile plaques, and loss of neuronal cells (202). The predominant clinical symptom is dementia with unknown etiology. AD is also associated with lower cerebral blood flow and decreased use of oxygen and glucose, especially in areas exhibiting neuropathological changes (203). In a normal brain, approximately

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50% of the energy expenditure at rest is used in maintaining Na,K-ATPase activity. Is abnormality of the Na,K-ATPase involved in Alzheimer’s disease etiology? Currently, there is no answer to this. However, there are some observations that may provide some indications for future studies. Studies using brain tissue from AD patients as opposed to age-matched controls showed a decrease in ouabain binding (203), especially in the neocortex, which is predominantly affected in AD patients. Correspondingly, a decrease in ouabain-inhibitable Na,K-ATPase activity in the brain subcortical but not cortical areas of patients with AD was noted (204). Interestingly, altered levels of Na,K-ATPase isoform expression were observed in the brain tissue of Alzheimer’s disease patients, i.e., an elevation in the α1-subunit expression and suppression in the α3-subunit expression (205). These findings may not be completely consistent with those of others (203,204), but they may be related to an increased reactive gliosis as a result of increase in the α1-subunit expression and to a prestage of plaque formation as reflected by a decline in α3-subunit expression. Clinically, a reduced risk of developing AD in postmenopausal women was shown by using estrogen replacement therapy. This is consistent with the observation that pretreatment with 17-beta-estradiol or estriol largely prevented the oxidative effect of Na,K-ATPase activity in an animal model of Alzheimer’s disease with impaired Na,K-ATPase at the cortical synaptosomes (206). E.

Other Clinical Conditions

The therapeutic application of digoxin in noncardiac clinical situations suggests that endogenous counterparts may play a role in other diseases. For example, in urology, digoxin has been therapeutically used in the treatment of urethrocavernous fistula after a penile trauma (207) and recurrent priapism states (208). Gupta et al. (208) found that digoxin was capable of altering human penile erectile function in part by inhibiting corporeal smooth muscle sodium pump activity, which promotes contraction and impedes nitric oxide induced relaxation. Another example is in oncology. The anticancer drug cis-diamminedichloroplatinum(II) (CDDP) has been used in treatment of both non-small-cell lung cancer (NSCLC) and small-cell lung cancer (SCLC). Resistance of lung cancer cells to the anticancer drug CDDP seems to correlate with an intracellular accumulation of CDDP. A study by Bando et al. (209) showed a correlation between decreased intracellular CDDP accumulation and reduced Na,K-ATPase activity in both NSCLC and SCLC cell lines after treatment with ouabain. Therefore, an inhibitor of Na,KATPase may have potential application to enhance anticancer treatment. In fact, administration of ouabain during pre- or postirradiation treatment of cancer showed a synergistic effect on reducing the survival of tumor cells, with less effect on normal cells, indicating the feasibility of application of ATPase inhibitor(s) in radiotherapy (210).

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V.

ENDOGENOUS SODIUM PUMP INHIBITORS IN OTHER PATHOLOGICAL OR PHYSIOLOGICAL CONDITIONS

A rise in the levels of factors such as DLIF or OLF has been noted in various pathological or physiological conditions. These include infarction of the myocardium (211), congestive heart failure (212), reversible cardiac dysfunction induced by physical exhaustion (noted in human subjects by electrocardiographic evidence (213), a secondary hypertension due to hypothyroidism (214), acute and chronic renal disease (131,215), fluid retention due to hepatic failure (132,133), pregnancy-induced hypertension (PIH) (preeclampsia) (126), and normal pregnancy. For example, in normal pregnancy and PIH the elevated DLIF values resolve rapidly upon delivery of the fetus (130,216–218). In addition, healthy children (5–16 years of age) who had detectable levels of these endogenous factors also showed significant hyponatremia, hypernatriuresis, and an elevated systolic blood pressure ( p ⬍ 0.01) compared with children without, for example, DLIF detected (219). What physiological or pathological role do these compounds play? De Angelis and Haupert (220) demonstrated in studies both in vitro and in vivo that release of an endogenous Na,K-ATPase inhibitor from Wistar rat midbrain and adrenal tissues was significantly increased by hypoxia. An elevated DLIF level was also observed in the serum of patients with advanced chronic respiratory failure (221). This observation indicates a hormonal response to hypoxia or ischemia resulting in reducing Na,K-ATPase activity and ATP consumption (222). Because these are cellular energy-conserving responses, these inhibitors of sodium pump activity may play a protective role in certain pathological and physiological conditions.

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200. El-Mallakh RS, Harrison LT, Li R, Changaris DG, Levy RS. An animal model for mania: preliminary results. Prog Neuro-Psychopharmacol Biol Psychiat 1995; 19: 955–962. 201. Mynett-Johnson L, Murphy V, McCormack J, Shields DC, Claffey E, Manley P, McKeon P. Evidence for an allelic association between bipolar disorder and a Na⫹, K⫹ adenosine triphosphatase alpha subunit gene (ATP1A3). Biol Psychiatry 1998; 44:47–51. 202. Markesbery WR. Alzheimer’s disease: a mini review. J KY Med Assoc 1989; 87: 333–335. 203. Harik SI, Mitchell MJ, Kalaria RN. Ouabain binding in the human brain. Arch Neurol 1989; 46:951–954. 204. Liguri G, Taddei N, Latorruca S, Nediani C, Sorbi S. Changes in Na⫹,K⫹-ATPase, Ca2⫹-ATPase and some soluble enzymes related to energy metabolism in brains of patients with Alzheimer’s disease. Neurosci Lett 1990; 112:338–342. 205. Chauhan NB, Lee JM, Siegel GJ. Na,K-ATPase mRNA levels and plaque load in Alzheimer’s disease. J Mol Neurosci 1997; 9:151–166. 206. Keller JN, Germeyer A, Begley JG, Mattson MP. 17Beta-estradiol attenuates oxidative impairment of synaptic Na⫹ /K⫹-ATPase activity, glucose transport, and glutamate transport induced by amyloid beta-peptide and iron. J Neurosci Res 1997; 50:522–530. 207. Seftel AD, Matthews LA, Herbener TE, Spirnak JP. Corpus cavernosum-spongiosum fistula after blunt pelvic trauma: successful resolution with digoxin. J Urol 1996; 156:1769. 208. Gupta S, Salimpour P, Saenz de Tejada I, Daley J, Gholami S, Daller M, Krane RJ, Traish AM, Goldstein I. A possible mechanism for alteration of human erectile function by digoxin: inhibition of corpus cavernosum sodium/potassium adenosine triphosphatase activity. J Urol 1998; 159:1529–1536. 209. Bando T, Fujimura M, Kasahara K, Matsuda T. Significance of Na⫹, K(⫹)-ATPase on intracellular accumulation of cis-diamminedichloroplatinum(II) in human nonsmall-cell but not in small-cell lung cancer cell lines. Anticancer Res 1998; 18(2A): 1085–1089. 210. Verheye-Dua F, Bohm L. Na⫹, K⫹-ATPase inhibitor, ouabain accentuates irradiation damage in human tumor cell lines. Radiat Oncol Invest 1998; 6:109–119. 211. Kohn R, Lichardus B, Rusnak M, Fridrich V, Zelenay J, Mizera S, Sumbal J, Margitfalvi P, Hricak V, Riecansky I. Personal experience with determination of endogenous, digoxin-like substances in patients with myocardial infarct and other cardiopathies. Bratisl Lek Listy (Slovak, abstract in English) 1995; 96:82–87. 212. Gottlieb SS,Rogowski AC, Weinberg M, Krichten CM, Hamilton BP, Hamlyn JM. Elevated concentrations of endogenous ouabain in patients with congestive heart failure. Circulation 1992; 86:420–425. 213. Valdes R Jr, Hagberg J, Vaughan TE, Lau BWC, Seals DR, Ehsani AA. Endogenous digoxin-like immunoreactivity in blood is increased during prolonged strenuous exercise. Life Sci 1988; 42:103–110. 214. Argento NB, Hamilton BP, Valente WA, Hamlyn JM. Increased circulating levels of a ouabain-like compound in hypothyroid hypertension. Hypertension 1991; 18: 425.

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215. Izumo H, Izumo S, Deluise M, Flier JS. Erythrocyte Na,K pump in uremia: acute correction of a transport defect by hemodialysis. J Clin Invest 1984; 74:581–588. 216. Delva P, Capra C, Degan M, Minuz P, Covi G, Milan L, Steele A, Lechi A. High plasma levels of ouabain-like factor in normal pregnancy and in pre-eclampsia. Eur J Clin Invest 1989; 19:95–100. 217. Gusdon JP Jr, Buckalew VM Jr, Hennessey JF. A digoxin-like immunoreactive substance in preeclampsia. Am J Obstet Gynecol 1984; 150:83–85. 218. Valdes R Jr, Graves SW, Knight AB, Craig HR. Endogenous digoxin immunoactivity is elevated in hypertensive pregnancy. Prog Clin Biol Res 1988; 192:229–232. 219. Garbagnati E. Serum digoxin-like immunoreactive factor in children and its relation to sodium metabolism. Acta Paediatr 1998; 87:500–504. 220. De Angelis C, Haupert GT Jr. Hypoxia triggers release of an endogenous inhibitor of Na(⫹)-K(⫹)-ATPase from midbrain and adrenal. Am J Physiol 1998; 274(1 pt 2):F182–F188. 221. Varsano S, Shilo L, Bruderman I, Dolev S, Shenkman L. Endogenous digoxin-like immunoreactive factor is elevated in advanced chronic respiratory failure. Chest 1992; 101:146–149. 222. Stanimirovic DB, Ball R, Durkin JP. Glutamate uptake and Na,K-ATPase activity in rat astrocyte cultures exposed to ischemia. Acta Neurochir (suppl) 1997; 70: 1–3. 223. Rose AM, Valdes R Jr. Understanding the sodium pump and its relevance to disease. Clin Chem 1994; 40:1674–1685.

Index

Acid phosphatases, Apo-CPO, 314–317, 316f conserved active site, 317–324 haloperoxidases, 313–317 HPP superfamily, 317–319 LPP family, 319–320 reaction scheme, 322–324, 323f Actin cytoskeleton, Akt1 insulin activation, 292f glucose transporter translocation, 289–290 GLUT4 translocation, 290–293 PI 3-kinase, 291f Acute respiratory distress syndrome (ARDS), 233 Acylglycoxylceramides, skin, 80 α-Dihydroxy bile salts, CMC, 260, 260f localization, 265 ADP, binding sites, 273 African clawed frog, 2 Africanized honeybees, 45

Akt, glucose transporter translocation, 287–289 glucose transport stimulation, 296f Alamethicin, hydrophobicity, 11–12 lipid-bound peptides, 14t origin and properties, 3t structure, 8, 9f, 32 Alamethicin channels (barrel stave model), 12–15, 13f, 30–32, 31f Alkylating agents, skin lipids, 82 Allipticine, 112 Alveolar dipalmitoylphosphatidylcholine (DPPC), turnover, 465–466 Alveolar type II cells, LPS, 238–240 Alzheimer’s disease (AD), amyloid β-peptide, 335–336 familial, 353–354 Na, K-ATPase, 495–497 oxidative stress, 335–336 Amphipathic helical peptides, 1–18 antibacterial activity, 10 513

514 [Amphipathic helical peptides] anticancer agents, 18 hydrophobicity, 11–12 membrane-permeabilization models, 12–17 membrane-permeabilizing peptides, 2–7 prospective applications, 17–18 secondary effects, 17–18 structural motifs, 7–12, 9f study models, 7 Amyloid β-peptide, Alzheimer’s disease, 335–336 EPR, 338, 338f methionine residue, 35, 340, 345 Amyloid β-peptide-associated free radical oxidative stress, neurotoxicity, 337–340, 339f Amyloid β-peptide-induced lipid peroxidation, 340–342 Amyloid β-peptide-induced protein oxidation, 342–345 Anionic vesicles, melittin, 50, 52f melittin-mediated lipid exchange, 56f Antagonistic lipid A analogs, endotoxins, 222–223 Antimicrobial peptides, bacterial membrane, 30 vertebrates, 4 Apicomplexa parasite mitochondria, 184–186 Apis mellifera, 45 Apo-CPO, acid phosphatase activity, 314–317, 316f phosphatase, 322–323, 323f Apolar α-dihydroxy bile salts, 255 Apolar bile salts, erythrocyte membranes, 263–264, 263f, 264f Apoptosis, 370–371 initiation, 370–371 physiological changes, 381 TUNEL visualization, 371

Index Apoptotic membrane structure analysis, 372–382 acridine orange staining, 373 apoptosis, detection, 373–374 induction, 373 cell culture, 372 cell membrane characteristics, 378– 381 cell separation, 372 DNA fragmentation analysis, 373 ESR spectroscopy, 374–375, 375f, 376f malignant vs. nonmalignant cells, 377–378, 377t results, 377–381 spin labeling, 374 TUNEL test, 374 APP proteolytic cleavage, PS1, 361 Arachaea, 127–128 Arachaeal lipids, 128–129 structure, 153, 154f Archefect ⫹ (AF⫹), synthesis, 156 Archefect AF1, synthesis, 156 Archefect AF2, synthesis, 156 Archefects, synthesis, 154–156 Arginine, V-BPO, 317 Ascophyllum nodosum V-BPO, 310, 313–314, 314f amino acid sequences, 313–314, 315f Asian toad, 2 Atovaquone, 186–189, 186f antimitochondrial activity, 187– 188 bc 1 complex, 187–188 mitochondrial membrane potential collapse, 188 parasite death, 188–189 Atovaquone-binding region, atovaquone-resistant mutants, 192 structural changes, 192–196 Atovaquone-mediated mitochondrial membrane potential (∆Ψ m ) collapse, 188–190

Index Atovaquone-proguanil synergy, 189– 192, 190f Atovaquone-resistant mutants, atovaquone-binding region, 192 ATP, binding sites, 273 ATP channels, CFTR, 444–445 AU-1421, 112 Autosomal dominant polycystic kidney disease (ADPKD), clinical aspects, 409 etiology, 409 genetic heterogeneity, 410 PKD1 gene structure, 410–412 Bacterial membrane, antibacterial mechanism, 33–34 cytolytic peptides, 30–33 diastereomers, 34–41 negatively charged, 30 Bactericidal permeability-increasing protein (BPI), 228 β-agonists, mononuclear cell membranes, 427–435 Barrel stave model, 12–15, 13f, 30–32, 31f Basolateral plasma membranes, apolar bile salts, 263, 263f β-defensin hBD-2, origin and properties, 3t, 4 Bee venom, cellular responses, 45–46, 46t components, 45–46 Bee venom toxicity, 45–48 antigenicity and allergenicity, 47–48 melittin-induced lysis, 56–58 melittin phospholipid exchange, 56 phospholipase-activating peptides, 48–49 phospholipid exchange mediation, 49–58 PLA2 pathophysiology, 46–49 Benzoyl peroxide, cellular effects, 84 β-galactosidase, LacZ, 357 Bile acids, cholestasis, 254–257 Bile salts, binding, 257–261

515 [Bile salts] erythrocyte membranes, 261–264, 263f, 264f hepatocyte membranes, 261–264 liposomal damage, 257–261 localizations, 265f mechanisms, 254f membrane diffusion, 259f therapeutic effect, 264–267 Bioenergetic metabolism, liver fluke infection, 207–208 Bipolar illness, Na, K-ATPase, 495– 496 B lymphocytes, 229–230 activation, 229f Brain, oxidative stress markers, 337 Bromobimane, TEL liposomes, 140, 140f Bromoenol lactone (BEL), 468 Brown adipose tissue (BAT), 118– 119 Buforin, 2 Bupivacaine, oxidative phosphorylation, 116f Byssinosis, 233 Caenorhabditis elegans, 355, 361 Ca 2⫹-independent PLA2 (iPLA2), 468 Caldariella acidophila, 146 Caldariomyces fumago, CPO, 308 Calorigenic effect, 271 Campothecin, chemical structure, 371, 371f CAMP-regulated Cl⫺ channel, CFTR, 439–442 Campylobacter jejuni lipid A, aggregate structures, 226 biological activity, 227f Cannalicular liver plasma membranes (cLPM), apolar bile salts, 263, 263f Carbonyl cyanide m-chlorophenylhydrazone (CCCP), 109t, 204 Carboxyfluorescein (CF), CDC, 258, 258f LUV, 261f

516 [Carboxyfluorescein (CF)] TEL liposomes, 141–144, 142f– 144f, 143t Cardiac glycosides, sodium pump, 486– 489 Cardiolipin, liver fluke infection, 206 Cardiovascular disease, Na, K-ATPase, 491–493 Carpet model, 13, 15–17, 30, 31f, 32– 34 Catalytic subunit model, G-6-Pase, 304 Cationic protonophores, 112 Cationic uncouplers, phosphatedependent, 115–116 CD14, LPS binding, 230 CD18, LPS-binding, 231 CD34⫹, 229 CD95 (APO-1Fas) receptor, 370–371 Cecropin A, origin and properties, 3t Cecropin P1, hydrophobicity, 11–12 lipid-bound peptides, 14t origin and properties, 3t structure, 8, 9f Cecropin P carpet (carpet model), 13, 15–17, 30, 31f, 33–34 Cecropin PS, Schiffer-Edmundson wheel projection, 29, 29f Cecropins, origin and properties, 2, 3t, 4 Cell death, 370 Cepharanthine, 114 Ceramides, skin, 78–79 Chenodeoxycholate, erythrocyte membranes, 263, 264f Chenodeoxycholic acid (CDC), 255– 256, 255f, 256f binding, 257–258 LUV, 260f Chloroperoxidase (CPO), 308 Chlorpromazine, mechanisms, 254f Cholate, binding, 257–258 localization, 265 Cholera toxin binding subunit (CBT), 167–179 FRET, 169–173, 171f

Index Cholera toxin (CT), 167–169 action mechanism, 168–169 structure, 168 Cholestasis, 253–267 bile acids, 254–257 bile salt liposomal damage, 257–261 bile salt membrane effect, 261–264 bile salt therapy, 264–267 defined, 253 mechanisms, 254f pathophysiology, 253–254 UDC, 264–267 Cholesterol, binding site, 265 phase transition, 266–267 skin, 78–79, 81 Chromobacterium violaceum lipid A, aggregate structures, 226 biological activity, 226–227, 227f Circular dichroism (CD), amphipathic helical peptides, 7 model diastereomers, 39 PxB, 71–72 Cl⫺ channel, outwardly rectifying CFTR, 443–444 P-glycoprotein, 445 Cl⫺ permeability, cystic fibrosis, 442– 443 Coenzyme Q, 186f Combined conformational flexibility substrate transport model, G-6-Pase, 304–305 Compound 406, 323 structure, 221–222, 221f Compound 506, structure, 221–222, 221f Compound E5531, 323 structure, 221–223, 221f Corallina pilulifera V-BPO, amino acid sequence, 314, 315f Core, enterobacterial LPS, 220–221, 220f Counterions, lipid A, 226 Coupling, loose, oxidative stress, 96– 101 Creatine kinase (CK), 344–345

Index Critical micellar concentration (CMC), α-dihydroxy bile salts, 260, 260f Cromoglycate, cell survival, 430, 431t mononuclear cell membranes, 427– 435, 434, 434f order parameters, 429, 429t Cromoglycin, structural formula, 429f Crotalus adamentus, 56 CTB, membrane structure, 175–178 pH, 173–175, 174f pyreneGM1, 175–179, 176t, 177f C-terminal tail, CFTR, 448 Cumene hydroperoxide, cellular effects, 84 Curvularia inaequalis V-CPO, 308–314 amino acid sequence, 313–314, 315f structure, 310–312, 311f vanadium binding site, 310–311, 312f Cycloguanil, 189–190 Cyclooxygenase-2, 236 Cystic fibrosis, dysfunctional Cl⫺ permeability, 442–443 Cystic fibrosis transmembrane conductance regulator (CFTR), 439–450 ATP channels, 444–445 cAMP-regulated Cl- channel, 439– 442 C-terminal tail, 448 Na⫹ channels, 445–447 NHERF, 448–449 outwardly rectifying Cl⫺ channel, 443–444 ROMK K⫹ channels, 447–448 Cystogenesis, loss of function hypothesis, 418–419 molecular mechanism, 418–420 Cystolic PLA2 (cPLA2), 468 properties, 469t Cytochalasin D, glucose transporter translocation, 289–290, 290t insulin-dependent Akt1 activation, 292, 292f PI 3 kinase, 292–293

517 Cytochrome b genes, malarial parasites, 193 Cytochrome c oxidase, 185–186, 186f ascorbate respiration, 274–277, 275f, 276f 3,5-diiodothyronine, 274–277 nucleotides, 273–274 Cytochrome P450, liver fluke infection, 208–209 Cytolytic peptides, 27–29 action mode, 30–33 origin and properties, 2, 3t sequences, 28t Cytoskeleton, actin (see Actin cytoskeleton) Cytosolic binding proteins (CTBPs), 271

5-DAS, erythrocyte membranes, 262f 16-DAS, erythrocyte membranes, 262f Decouplers, 115–117 Deoxycholate (DC), erythrocyte membranes, 263, 264f 2-Deoxyglucose, 294t Dermaseptin, carpet model, 17 Dermaseptin-S, minimal inhibitory concentration, 37t Schiffer-Edmundson wheel projection, 29, 29f Dexamethasone, liver fluke infection, 210–211 ∆-hemolysin, hydrophobicity, 11–12 origin and properties, 2, 3t Diabetes mellitus, insulin-dependent, 293–294 noninsulin-dependent, GLUT 4 protein, 294–295 Diabetes mellitus-related hypertension, Na, K-ATPase, 493–494 Diacid, synthesis, 155, 155f Diacid dichloroanhydride, synthesis, 155, 156f Diastereomeric peptides, vs. antibacterial peptides, 40

518 Diastereomers, bacterial membrane, 34– 41 Dicumyl peroxide, cellular effects, 84 Diethylstilbestrol (DES), 204 Differential scanning calorimetry (DSC), UDC, 265–266 Digitoxin, sodium pump, 486–487 structure, 487f Digoxin, 499 sodium pump, 486–489 structure, 487f therapeutic drug monitoring, 488– 489 toxicity, 488 Digoxinlike immunoreactive factors (DLIFs), metabolism, 490–491, 498 origin, 490 sodium pump, 489–491 Dihydrofolate reductase (DHFR) inhibitor, 189–190 Dihydroorotate dehydrogenase (DHOD), 184–186 3,5-diiodothyronine, cytochrome c oxidase, 274–277 formation, 272 subunit Va, 274 Dimiristoylphosphatidylcholine, UDC, 266, 266f Dipalmitoylphosphatidylcholine (DPPC), alveolar turnover, 465–466 degradation, 466–467, 467f, 470, 471t, 478f internalized, 466–467, 467f metabolism, 465, 465t PLA2, 466–467, 467f surfactant, 462–463 Diquat, 82 DLIF, diabetes mellitus-related hypertension, 494 Docking proteins, 284 16-doxylstearic acid (16-DSA), spin label, 374–375, 375f–376f, 378, 379f–380f

Index Drug carriers, TEL liposomes, 139–142, 148 Edema, LPS-induced pulmonary, 235– 236 Egg lecithin, TEL liposomes, 135, 136t, 147 Eicosanoid, production, 47 Electrolyte imbalance, Na, K-ATPase, 495 Electron paramagnetic resonance (EPR), amyloid β-peptide, 338, 338f TEL liposomes, 130–131, 146 Electron spin resonance (ESR), defined, 369–370 free radicals, 337–338 Embellisia didymospora, V-CPO, 309– 310 amino acid sequence, 314, 315f ENaC, 444–445 Endocytosis, surfactant, 465, 465t Endogenous digitalis-like factors (EDLFs), sodium pump, 489– 491 Endogenous sodium pump inhibitors, 498 Endothelial cells, 230 Endothelin, 237 Endotoxemia, 228, 233 Endotoxins, 217–224 antagonistic lipid A analogs, 222–223 bronchoconstriction, 236 cell activation model, 232, 233f chemical structure, 220–222 critical aggregate concentration, 224– 226 edema, 235–236 leukocyte pulmonary sequestration, 234–235 lung tissue LPS, 234 mechanisms, 254f membrane aspects, 218–220 molecular conformation, 225–226 phase states, 224–225 physical structure, 224–226 pulmonary hypertension, 236–237

Index [Endotoxins] supramolecular structure, 225–226 surfactant homeostasis, 237–240, 239f target cells, 228–230 transition temperature, 224–225 Energy metabolism, thyroid hormones, 271–272 Enterobacterial wild-type lipopolysaccharide, structure, 220–221, 220f Epithelial cells, type I alveolar, surfactant, 463 type II alveolar, surfactant, 462–464, 464f Erythrocyte membranes, apolar bile salts, 263–264, 263f, 264f bile salts, 261–264 5-DAS, 262f 16-DAS, 262f Escherichia coli, peptide treatment, 38f PxB, 65–68, 65f–68f Escherichia coli lipid A biological activity, 227f structure, 221–222, 221f Ethynyl estradiol, mechanisms, 254f Eukaryotic cell transfection, 153–165 archefect-mediated transfection, 160– 162, 162f archefect penetration, 160, 161f archefects synthesis, 154–159 β-galactosidase cell assay, 157 DNA-archefect complex preparation, 157, 159–160, 159f fluorescent microscopy, 157 liposome preparation, 156 phospholipids, 162–165, 163f–164f Eukaryotic membrane, lipid composition, 5, 6f European honeybee, 45 Exocytosis, surfactant, 465, 465t Familial Alzheimer’s disease (FAD), 353–354 Farmer’s lung, 233

519 Fasciola gigantica, 201 Fasciola hepatica, 201–203 life cycle, 201–202 pathology, 202–203 Fasciolosis, 201 FCCP, 109t Fenoterol, cell survival, 430, 431t mononuclear cell membranes, 434, 434f order parameters, 429t, 430, 433 structural formula, 429f F 0 F 1-ATPase, liver fluke infection, 204–205 First mechanism respiratory control, 274 Fluazinam, 111, 112f Fluidity, 369–370 Fluid system, 99f Fluorescence resonance energy transfer (FRET), CTB, 169–173, 171f Fourier transform infrared (FTIR), amphipathic helical peptides, 7 Free fatty acids (FFA), skin, 78–79 uncoupling, 118 Free radicals, ERP, 337–338 French pressure cell extruded, vs. sonicated liposomes, 132–134 Friedreich ataxia, 411 Giant lamellar bodies, 238 Glucose-6-phosphatase (G-6-Pase), gene, 305–307, 306f GSD, 304–307 liver fluke infection, 209 models, 304–305 new membrane topology, 320–322, 321f vanadium-containing haloperoxidases, 307–313 Glucose transport, maximal stimulation, 296f Glucose transporter 4 (GLUT4), 284 Glucose transporter 7 (GLUT7), 204 Glucose transporter 4 (GLUT 4) protein type 2 diabetes, 294–295

520 Glucose transporter 4 (GLUT4) translocation, actin cytoskeleton, 290– 293 PI 3-kinase, 286–289 SB203580, 294t Glucose transporters, actin cytoskeleton, 289–290 Glucose transporter translocation, insulin signaling pathway, 293 IRSs, 284–285 PI 3-kinase, 285–287 PKB, 287–289 signaling pathway, 284–289 Glutamine synthetase (GS), 344–345 Glutathione (GSH), liver fluke infection, 209–210 Glycerol-3-dehydrogenase, 185–186, 186f Glycerol ethers, 128 Glycerophosphocholine, liver fluke infection, 206 Glycerophosphoethanolamine, liver fluke infection, 206 Glycogen storage disease (GSD), G-6Pase, 304–307 Glycolipids, skin, 80 Glycosides, sodium pump, 486–489 Glycosylceramide A, skin, 80 Gramicidin, 113, 113f Gram-negative bacteria membrane, architecture, 5, 6f Granular pneumocytes, surfactant, 462– 464, 464f Γ-secretase, 362 Halide binding, V-CPOs, 312–313 V-HPOs, 309–310 Haloperoxidases, acid phosphatases, 313–317 defined, 307–308 groups, 308 heme-containing, 308 vanadium-containing, 307–313 Helical peptides, amphipathic (see Amphipathic helical peptides)

Index Heme-containing haloperoxidases, 308 Hemolytic peptides, origin and properties, 2, 3t Hepatocyte membranes, bile salts, 261– 264 Histidine phosphatase/peroxidase (HPP) superfamily, 317–319 conserved domains, 317, 318f Homoviscous adaptation, 219 HOP-1, 355 protein sequences, 359, 360f H⫹ pumps, molecular slip, 117 Human embryonic kidney (HEK) cell line, IRS-4, 284 Hydrazone, detection, 344 Hydroperoxides, skin, 83–84 Hydroxynaphthoquinones, 186–187 Hyperglycemia, 294 Hyperosmotic shock, PxB, 67–69 Hypertension, diabetes mellitus-related, Na, K-ATPase, 493–494 Na, K-ATPase, 491–493 pregnancy-induced, DLIFs, 498 Hypothalamic inhibitory factors (HLFs), sodium pump, 489–490 IgE, melittin, 48 Imminium ions, skin lipids, 82 Indolicidin, origin and properties, 3t Inflammation, defined, 46–47 Insulin, actin cytoskeleton, 292f glucose transporter translocation, 293 PI 3-kinase, 285–287 PKB, 287–289, 288f p38 MAP kinase, 293 125 insulin tetraether lipid liposomes, 142–144 Insulin-dependent diabetes mellitus, 293–294 Insulin receptor substrates (IRS), glucose transporter translocation, 284–285, 296f Insulin receptor substrates (IRS-1), 284–285

Index Insulin receptor substrates (IRS-2), 284–285 Insulin receptor substrates (IRS-3), 284 Insulin receptor substrates (IRS-4), HEK cell line, 284 Insulin receptor substrates (IRS) proteins, glucose transport stimulation, 296f PH domains, 284 PTB domain, 284 Interleukins, 228 Intrinsic uncoupling, 118–122 permeability transition, 120–122 protein uncoupling, 118–120 Inulin efflux, CDC, 258, 258f Ischemia, 96 mitochondria, 97–98 6-Ketocholestanol (kCH), 112, 113f Kidney disease, autosomal dominant polycystic, 409 Kirl.1a, 447–448 LacZ, β-galactosidase, 357 Lapinone, 187 Large unilamellar vesicles (LUVs), bile salt binding, 257–261 CDC, 260f CF, 261f UDC, 260f Latrunculin B, glucose transporter translocation, 289–290, 290t insulin-dependent Akt1 activation, 292, 292f Leukemic blast cells, 269f Leukocytes, pulmonary sequestration, 234–235 Lin-12, 355 Linear polypeptides, structure, 27–29, 34 Linoleic acid, 80 Lipid A, aggregate structures, 226 biological activity, 226–228, 227f counterions, 226 enterobacterial LPS, 220–221, 220f

521 [Lipid A] molecular conformation, 222–223, 223f Rhodobacter capsulatus, structure, 223 structure, 221–222, 221f Lipid liposomes, tetraether (see Tetraether lipid liposomes) Lipid peroxidation, amyloid β-peptide-induced, 340–342 brain, 337 ozone-induced, 85 Lipid peroxidation products, skin, 83– 84 Lipid phosphate phosphohydrolase (LPP) family, acid phosphatases, 319–320 Lipids, stratum corneum, 78 Lipopolysaccharide-binding protein (LBP), 219–220 Lipopolysaccharides (LPS), 5, 217–241 alveolar type II cells, 238–240 bronchoconstriction, 236 endotoxin activity, 218–220 endotoxin bioactivity, 222–224 endotoxin structure, 220–222, 224– 226 endotoxin target cells, 228–230 enterobacterial wild-type, structure, 220–221, 220f hypoxic vasoconstriction, 237 lung tissue distribution, 234 physicochemical parameters, 226– 228 proinflammatory properties, 233–240 pulmonary vasculature, 237 PxB, 64, 73 smooth (S-form), 221 surfactant homeostasis, 237–240, 239f in vivo endotoxins, 233–240 wild-type, 221 Liposomes, bile salts, 257–261 tetraether lipid (see Tetraether lipid liposomes)

522 Liver fluke infection, 201–211 bioenergetic metabolism, 207–208 F 0F 1-ATPase, 204–205 host immune response, 210–211 liver function, 202 microsomes, 208–209 mitochondria uncoupling, 203–204 NEFAs, 205–207 oxidative stress, 209–210 parasite life cycle, 202 pathology, 202–203 phospholipids, 206–207 Loose coupling, oxidative stress, 96–101 Low-density microsomes (LDM), glucose transport stimulation, 296f PI 3-kinase, 287 LPS-binding proteins (LBP), CD14, 230 CD18, 231 mCD14, 231 sCD14, 231 LPS-induced pulmonary edema, 235– 236 Lung, fluid homeostasis, 442–443 PLA2, 470–471 Lung surfactant (see Surfactant) Lymphoblast cells, 369f Lysine, V-BPO, 317 Lysis, melittin-induced, 56–58 LysoPC, 466 Lysophosphatidylethanolamine, liver fluke infection, 206 Lysophospholipid, defined, 47 Lysosomal PLA2 (aiPLA2), isolation, 471–472 physiological regulation, 474, 475f properties, 469t, 473–474 SP-A, 474, 475f substrate specificity, 473f Lytic peptides, origin and properties, 2, 3t MAb anti-breast cancer 6C6, 390–405 antigen biochemical characterization, 395–399, 397f, 399f

Index [MAb anti-breast cancer 6C6] antigen expression, 402–403, 403f antigen localization and orientation, 399–402, 400f–401f cDNA clone isolation, 394–395, 396f cDNA library screening, 390–391 cell lines and media, 390 cell transfection, 392 epitope mapping, 393–394 immunofluorescence analysis, 392 materials and methods, 390–394 membrane protein biotinylation, 393 metabolic labeling and immunoprecipitation, 392–393 mRNA preparation, 394 phage DNA preparation and sequencing, 391 results, 394–403 in vitro transcription and translation, 391–392 Macrophages, endotoxemia, 228 Magainin, antimicrobial, 18 diabetic foot ulcer infections, 18 discovery, 2–4 hydrophobicity, 11–12 structure, 8, 9f, 40 Magainin 2, lipid-bound peptides, 14t origin and properties, 3t Magainin analogs, histamine-releasing activity, 18 Magainin Pharmaceuticals, web site, 18 Magainin pores (toroidal model), 13f, 15–17 Mal-6, 343, 343f Malarial parasite mitochondria, 184– 186 electron transfer chain complexes, 185–186 Malondialdehyde (MDA), skin, 83 Manic depression, Na, K-ATPase, 495– 496 Mast cell degranulation, mononuclear cell membranes, 434, 434f Mast cells, 48

Index Mastoparans, histamine-releasing activity, 18 origin and properties, 2, 3t PLA2, 48–49, 52 McArdle disease, Na, K-ATPase, 495 MCD14, LPS-binding, 231 Melittin, antigenicity and allergenicity, 47–48 diastereomers, 35, 35t hydrophobic segment, 51f hydrophobicity, 11–12 minimal inhibitory concentration, 37t origin and properties, 2, 3t phospholipid exchange, 56 phospholipid exchange mediation, 49–58 phospholipid interface, 49–50 PLA2, 48–49 PLA2 catalysis, 52–56 POPC hydrolysis, 55, 55f primary sequence, 49f Schiffer-Edmundson wheel projection, 29, 29f vesicle binding, 50, 52f zwitterionic phospholipids, 57 Melittin-induced lysis, 56–58 Melittin-mediated vesicle-vesicle contact, 53f Membrane-associated antigens, mAbs, 388–390 Membrane permeabilization models, 12–17 alamethicin channels, 12–15, 13f, 31f carpet model, 13, 15–17, 30, 31f, 32–34 magainin pores, 13f, 15–17 Membrane-permeabilizing peptides, amino acid sequences, 10 classification and biological function, 2–4 target membrane action and properties, 4–7 Membrane ruffling, 289 Membranes, composition, 218–219 Membranous pneumocytes, surfactant, 463

523 Metacercariae, 202 Methionine residue, 35 amyloid β-peptide, 340, 345 Microsomes, liver fluke infection, 208– 209 Mitochondria, apicomplexa, 184–186 ischemia, 97–98 malarial, 184–186 Mitochondrial binding sites, T3, 272 Mitochondrial loose coupling, oxidative stress, 96–101 Mitochondrial membrane potential (∆Ψm) collapse, atovaquone, 188–190, 190f cellular physiology, 188–189 flow cytometry, 188–190 proguanil, 190, 190f rotenone, 188 Mitochondrial respiration, thyroid hormones, 272 Mitochondria uncoupling, liver fluke infection, 203–204 MJ33, chemical formula, 470f DPPC degradation, 470, 471t Model diastereomers, zwitterionic phospholipid membranes, 39–40 Model phospholipid membranes, cyclopeptide interaction, 37–39 Monoclonal antibodies (mAbs), 387– 390 (see also mAb anti-breast cancer 6C6) cancer, 387–388 membrane-associated antigens, 388– 390 Monocytes, endotoxemia, 228 Mononuclear cell membranes, action sites, 434f electron paramagnetic resonance spectroscopy, 428 functional structure, 432f mast cell degranulation, 434, 434f materials and methods, 428 mononuclear cell separation, 428 results, 429–430

524 [Mononuclear cell membranes] spin labeling, 428 stabilization, 434, 434f MRNA, presenilins, 354 MtDNA, malarial parasites, 193 Myelin, surfactant, 238 Myeloperoxidase, 308 Myocardium hypertrophy, Na, K-ATPase, 495 Na, K-ATPase, cardiovascular disease, 491–493 diabetes mellitus-related hypertension, 493–494 hypertension, 491–493 isoforms, 482–484, 483f neonatal abnormalities, 495 neurological disorders, 495–497 plasma membrane, 255, 255f structure and function, 480–482, 481f subcellular distribution, 484–485 α-subunit and β-subunit interaction, 484 Na⫹ channels, CFTR, 445–447 Na⫹ /H⫹ exchanger regulatory factor (NHERF), CFTR, 448–449 Naja nigricola, 56 Necrosis, 370 Neonatal abnormalities, Na, K-ATPase, 495 Neurological disorders, Na, K-ATPase, 495–497 Nitrogen dioxide, 84 skin, 85–86 N,N′-dicyclohexylcarbodiimide (DCCD), 204 Nonesterified fatty acids (NEFAs), liver fluke infection, 205–207 Noninsulin-dependent diabetes mellitus, GLUT 4 protein, 294–295 Nonitol ethers, 128 Notch proteolytic cleavage, PS1, 361 Nuclear magnetic resonance (NMR), amphipathic helical peptides, 7 phospholipids, 206 PxB, 71–72

Index Nucleotides, cytochrome c oxidase, 273–274 Oligomycin, 204 Organic peroxides, toxic effects, 83–84 Organ transplantation, TEL liposomes, 149 OsmY gene promoter, PxB, 64–67 O-specific chain, enterobacterial LPS, 220–221, 220f Ouabain-like-factors (OLFs), pathological conditions, 498 sodium pump, 489–490 Outwardly rectifying Cl⫺ channel (ORCC), CFTR, 443–444 Oxidative phosphorylation, bupivacaine, 116f protonophoric uncouplers, 109–114 uncouplers, 108f Oxidative stress, 95–101 ischemia, 96 liver fluke infection, 209–210 mitochondrial loose coupling, 96– 101 Oxidative stress markers, brain, 337 Oxidizing dusts, skin, 84 Oxidizing gases, skin, 84 Ozone, 84 skin, 84–85 Paenibacillus, 63–64 Paraquat, 82 Parasite mitochondria, 184–186 apicomplexa, 184–186 malarial, 184–186 Pardachirus marmoratus, 32 Pardachirus pavoninus, 32 Pardaxin, carpet model, 17 circular dichroic spectra, 36f diastereomers, 34, 35f, 36f minimal inhibitory concentration, 37f structure, 32 Parvaquone, 187 PbX, septic shock, 71 toxicity, 71

Index PDZ domains, 448 Penile trauma, digoxin, 499 Peptide membrane lysis, 12, 13f Peptides, amphipathic helical (see Amphipathic helical peptides) antimicrobial, bacterial membrane, 30 vertebrates, 4 cytolytic (see Cytolytic peptides) diastereomeric, vs. antibacterial peptides, 40 hemolytic, origin and properties, 2, 3t lytic, origin and properties, 2, 3t membrane-permeabilizing, amino acid sequences, 10 classification and biological function, 2–4 target membrane action and properties, 4–7 phospholipase-activating, bee venom toxicity, 48–49 Permeability transition pore (PTP), 100 Permeability transition (PT), 108, 120– 122 Peroxide binding, V-HPOs, 309 Peroxides, organic, toxic effects, 83–84 skin, 83–84 Peroxyacetyl nitrate, 84 P-glycoprotein, Cl⫺ channel, 445 PH, CTB, 173–175, 174f Phosphatases, acid (see Acid phosphatases) Phosphate-dependent cationic uncouplers, 115–116 Phosphatidylcholine (PC), 5, 6f liver fluke infection, 206 surfactant, 238, 462 Phosphatidylethanolamine (PE), 5, 6f liver fluke infection, 206 Phosphatidylglycerol, periplasmic space, 69 PxB, 72, 73f

525 Phosphatidylinositol 3-kinase (PI 3kinase), glucose transport translocation, 285– 287 insulin, 285–287 Phosphatidylinositol, liver fluke infection, 206 Phosphatidylserine, liver fluke infection, 206 Phosphocholine, liver fluke infection, 206 Phosphoester binding, PxB, 71–73 Phospholipase-activating peptides, bee venom toxicity, 48–49 Phospholipase A2 (PLA2), antigenicity and allergenicity, 47–48 characteristics, 468, 469t classification, 468 DPPC degradation, 466–467, 467f kinetic properties, 52 lung tissue, 470–471 melittin, 48–49 pathophysiology, 46–49 rat lung homogenate, 472f structure, 52 surfactant, 467–475 Phospholipids, eukaryotic cell transfection, 162–165, 163f–164f liver fluke infection, 206–207 melittin, 49–58 NMR spectroscopy, 206 skin, 79 surfactant, 462–463 Phospholipid vesicles, PxB-induced contacts, 71–73 Phosphotyrosine binding (PTB) domain, IRS proteins, 284 PI 3-kinase, cellular localization, 291f glucose transport stimulation, 296f GLUT4 translocation, 286–289 LDM, 287 PKD1 gene, chromosome 16 location, 411f duplicated, 410 polypyrimidine tract, 411

526 PKD2 gene, targeted disruption, 420– 421 PKD1 mutation, mice, 419–420 PLA2 catalysis, kinetic scheme, 54f melittin, 52–56 Plasma membrane, CDC, 255f, 256f Na-K-ATPase, 255, 255f Plasmodium, electron transfer protein complexes, 186f mitochondrial genomes, 184, 185f Plasmodium falciparum, atovaquone, 187 mtDNA, 193 Plasmodium vivax, 187 Plasmodium yoelii, 192–196, 194f Platelet-activating factor (PAF), 228 Platelet-activating factor (PAF) hydrolase, 468 properties, 469f Pleckstrin homology (PH) domains, IRS proteins, 284 P38 MAP kinase glucose transport stimulation, 296f insulin, 293 Pneumocystis, atovaquone, 187 Pneumocytes, granular, surfactant, 462–464, 464f membranous, surfactant, 463 P-NPP, kinetics, 316–317 Point mutagenesis, PxB, 70 Polycystic kidney disease, autosomal dominant, 409 Polycystin-1, endogenous expression, 415–416 membrane association, 416–418, 417f polycystin-2 interaction, 420 structure, 412–414, 413f in vitro expression, 414–415 Polycystin-2, 420–421 polycystin-1 interaction, 420 Polycystins, cystogenesis, 418–420 novel members, 421

Index Polymorphonuclear leukocytes (PMN), 228 Polymyxin B (PxB), 63–74 circular dichroism, 71–72 E. coli, 65–68, 65f–68f hyperosmotic shock, 67–69 lipopolysaccharides, 73 nonmutational adaptation, 70–71 osmY gene promoter, 64–67 periplasmic space, 69–70 phosphatidylglycerol, 72, 73f phosphoester binding, 71–73 phospholipid vesicles, 71–73 point mutagenesis, 70 prototype, 73–74 rapid responses, 67–69 structure, 64f Polymyxin nonapeptide (NP), 64 Polymyxins, 63–74 lipopolysaccharide binding, 64 PLA2, 48–49 PLA2 catalysis, 52 untoward effects, 71 Polypeptides, linear, structure, 27–29, 34 Preeclampsia, DLIFs, 498 Pregnancy, DLIFs, 498 Pregnancy-induced hypertension (PIH), DLIFs, 498 Preoxypropionyl nitrate, 84 Presenilin 1 (PS1), 353–354 Presenilin 2 (PS2), 353–354 Presenilins, cell biology, 354–355 functions, 359–362 homologs, 355 membrane topology, 355–359 mRNA, 354 synthesis, 354–355 Priapism, digoxin, 499 Primary pump, molecular slip, 117, 118f Programmed cell death (PCD) (see Apoptosis) Proguanil, 190–192, 190f Protein carbonyls, 343–344

Index Protein kinase B (PKB), glucose transporter translocation, 287– 289 Protein oxidation, amyloid β-peptide-induced, 342–345 brain, 337 Proteins, surfactant, 462 Protonophoric uncouplers, 109–114 cationic, 112 recouplers, 112–114 weakly acidic, 109–111 Protoporphyrin, mechanisms, 254f PS1, hydrophobic domains, 357, 358f hydrophobicity profile, 355–356, 355f protein sequences, 359, 360f proteolytic cleavage, 361 secondary structure, 356 PS2, protein sequences, 359, 360f Pulmonary edema, surfactant, 237 Pulmonary hypertension, endotoxins, 236–237 Pulmonary sequestration, leukocytes, 234–235 Pulmonary surfactant homeostasis, 237– 240 PyreneGM1, CTB, 175–178, 176t, 177f, 178–179 Quinones, skin lipids, 82 Reactive nitrogen species (RNS), Alzheimer’s disease, 336 Reactive oxygen species (ROS), Alzheimer’s disease, 336 liver fluke infection, 209–210 Recombinant apoenzyme (rCPO), 314– 317 Reproterol, cell survival, 430, 431t mast cell degranulation, 434, 434f mononuclear cell membranes, 434, 434f order parameters, 429t, 430 structural formula, 429f Respiratory control, mechanism, 274

527 Rhodamine archefect (Rh-AF), 156, 161f Rhodobacter capsulatus lipid A, aggregate structures, 226 biological activity, 226–227, 227f chemical structure, 223 Rhodobacter sphareoides lipid A, structure, 221–224, 221f Rhodocyclus gelatinosus lipid A, aggregate structures, 226 biological activity, 227f Rhodopseudomonas viridis lipid A, aggregate structures, 226 biological activity, 227f Rhodospirillum fulvum lipid A, aggregate structures, 226 Rigid system, 99f ROMK K⫹ channels, CFTR, 447–448 Rotenone, ∆Ψ m , 188 Rough mutant (R-LPS) lipopolysaccharide, 221 Salbutamol, cell survival, 430, 431t mast cell degranulation, 434, 434f mononuclear cell membranes, 434, 434f order parameters, 429t, 430, 433–434 structural formula, 429f SB203580, 293, 294t GLUT4 translocation, 294t SCD14, LPS-binding, 231 Sebum, 80–81 Second mechanism respiratory control, 274 Secreted PLA2 (sPLA2), properties, 469t Sel-12, 355, 361 protein sequences, 359, 360f Septic shock, 233 PbX, 71 SF6847, 109–110, 109t intramolecular rotation, 111 Shuttle mechanism, uncouplers, 110, 110f Shwartman reaction, 71

528 Skin, barrier function, 78–79 cornified epidermal layers, 78 morphology, 78f Skin lipids, 79–81 environmental oxidants, 81–86 epidermal, 79–80 sebaceous gland, 80–81 Slip inducers, 115–117 Smooth muscle cells, 230 Smooth (S-form) lipopolysaccharide (LPS), 221 Sodium-potassium-adenosine triphosphatase (see Sodium pump) Sodium pump, 479–498 cardiac glycosides, 486–489 endogenous digoxinlike immunoreactive factors, 489–491 regulation, 485–491 Sodium pump inhibitors, endogenous, 498 Sonicated liposomes, vs. French pressure cell extruded, 132–134 SP-A, 240 SP-D, 240 SPE-4, 361 Sphingomyelin (SM), 5, 6f liver fluke infection, 206 surfactant, 238 Squalene, skin, 79–81 Staphylococcus aureus, 2 Sterol-regulatory element-binding protein (SREBP), 361–362 Sterols, skin, 79–80 Stratum corneum, lipids, 78–79 Substrate transport model, G-6-Pase, 304 Subunit IV, 273 Subunit Va, 3,5-diiodothyronine, 274 Subunit VIaH, 273 Succinate dehydrogenase, 185–186, 186f SUR1, 447 Surfactant, composition, 462 defined, 461

Index [Surfactant] exocytosis/endocytosis coordination, 465, 465t PLA2, 467–475 processing, 462–463 Surfactant homeostasis, 237–240, 239f Surfactant protein A (SP-A), lysosomal PLA2 (aiPLA2), 474, 475f Synaptosomal membrane proteins, EPR, 343, 343f T3, mitochondrial binding sites, 272 TApar, diastereomers, 34–35, 35t minimal inhibitory concentration, 37t Tert-butyl-hydroperoxide, cellular effects, 84 Tetracycline, minimal inhibitory concentration, 37t Tetraether, synthesis, 154–155, 155f Tetraether lipids (TELs), dimensions, 128 Tetraether lipid (TEL) liposomes, 127– 149 alcohols, 137, 138t bile salts, 137, 138t bromobimane, 140, 140f 6-carboxyfluorescein, 142–144, 143t carboxyfluorescein fluorescence, 141– 142, 142f– 144f cell interaction, 138–139 cell membrane interaction, 131 detergents, 137, 138t drug carriers, 139–142, 148 egg lecithin, 135, 136t, 147, 148 electron paramagnetic resonance measurement, 130–131, 146 French pressure cell extruded vs. sonicated liposomes, 132–134 125 insulin, 142–144, 145f laser light scattering, 133–134, 133f liposome preparation, 129–130 membrane order parameters, 135, 135t, 147 methods and materials, 129–131 mixed liposomes, 135, 136t, 147

Index [Tetraether lipid (TEL) liposomes] protons, 137 results, 132–146 size, 130, 132, 132t, 146 skin penetration, 131, 145–146 special applications, 145 stability, 131, 136–137, 136t–137t, 147 sterilization, 137–138 temperature, 137 toxicity, 139 in vivo intestinal absorption, 144– 145 zeta potential, 130, 134, 134f, 147 Tetraether macrocycle, 128 Theileria, mitochondrial genomes, 184, 185f parvaquone, 187 Thermoplasma acidophilum, 127–128 main tetraether phospholipid, 128, 129f Thermoplasma volcanium, 127–128 Thiobarbituric acid reactive substances (TBARS), Alzheimer’s disease, 336 skin, 83 Thromboxane, 236 Thyroid hormones, energy metabolism, 271–272 UCPs, 272–273 uncoupled mitochondrial respiration, 272 T lymphocytes, 229–230 activation, 229f Toll-like receptors (TLR2), 232 Toroidal model, 13, 15–17, 33 Toxoplasma, atovaquone, 187 Toxoplasma gondii, 192–196, 194f Trichoderma virides, 2, 32 Triclabendazole, 201 Tru-S-C 7, 115, 115f Type 1 diabetes, 293–294 Type 2 diabetes, GLUT 4 protein, 294– 295 Type I alveolar epithelial cells, surfactant, 463

529 Type II alveolar epithelial cells, surfactant, 462–464, 464f Type 1 uncoupling protein (UCP1), 119–120 Tyrosine kinase, glucose transport, 284 Ubiquinol-cytochrome c oxidoreductase (bc 1) complex, 185–186 atovaquone, 187–188 Ubiquinol (QH2), 185–186, 186f Ultraviolet radiation, skin, 82–83 Uncoupled mitochondrial respiration, thyroid hormones, 272 Uncouplers, atypical, 115–118 classification, 114t defined, 107 oxidative phosphorylation, 108f protonophoric, 109–114 cationic, 112 recouplers, 112–114 weakly acidic, 109–111 shuttle mechanism, 110, 110f Uncoupling, intrinsic, 118–122 permeability transition, 120–122 protein uncoupling, 118–120 mitochondria, liver fluke infection, 203–204 Uncoupling proteins (UCPs), thyroid hormones, 272–273 Uncoupling protein (UCP-1), 273 Uncoupling protein (UCP-2), 273 Uncoupling protein (UCP-3), 273 Urethrocaval fistula, digoxin, 499 Uridine diphosphoglucose (UDPG), liver fluke infection, 210 Ursodeoxycholate (UDC), binding, 257–258, 265 dimiristoylphosphatidylcholine, 266, 266f DSC, 265–266 liver disease, 256–257 localization, 267f LUV, 260f

Index

530 [Ursodeoxycholate (UDC)] phase transition, 266–267 therapeutic effect, 264–267 Vanadium-containing bromoperoxidases (V-BPO), arginine, 317 Ascophyllum nodosum, 310, 313– 314, 315f Corallina pilulifera, 314, 315f discovery, 308 lysine, 317 Vanadium-containing chloroperoxidases (V-CPO), catalysis, 312–313, 313f Curvularia inaequalis, 310–314, 312f, 315f discovery, 308–309 Embellisia didymospora, 309–310, 314, 315f Vanadium-containing haloperoxidases (V-HPO), 307–313 catalysis, 312–313 cofactors, 309–310 discovery, 307–309 enzyme kinetics, 309–310 function, 307–309

Vascular cells, 230 Vertebrates, antimicrobial peptides, 4 Vesicle-vesicle contact, melittin-mediated, 53f Vibrio cholerae, 167 Von Gierke disease, G-6-Pase, 304– 307 W88, 170, 172f, 175 Weakly acidic protonophores, 109– 111 Wild-type lipopolysaccharide (LPS), 221 Wormhole model, 13, 15–17, 33 Wound healing, amphipathic helical peptides, 18 Wumen protein, 319 Yellow jackets, 45 Zwitterionic phospholipid membranes, model diastereomers, 39–40 Zwitterionic phospholipids, melittin, 57 Zwitterionic vesicles, melittin, 50, 52f