Transmembrane Dynamics of Lipids (Wiley Series in Protein and Peptide Science)

TRANSMEMBRANE DYNAMICS OF LIPIDS WILEY SERIES ON PROTEIN AND PEPTIDE SCIENCE VLADIMIR N. UVERSKY, Series Editor Metal...

Author: Andreas Herrmann

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TRANSMEMBRANE DYNAMICS OF LIPIDS

WILEY SERIES ON PROTEIN AND PEPTIDE SCIENCE VLADIMIR N. UVERSKY, Series Editor Metalloproteomics • Eugene A. Permyakov Instrumental Analysis of Intrinsically Disordered Proteins: Assessing Structure and Conformation • Vladimir Uversky, Sonia Longhi Protein Misfolding Diseases: Current and Emerging Principles and Therapies • Marina Ramirez-Alvarado, Jeffery W. Kelly, Christopher M. Dobson Calcium Binding Proteins • Eugene A. Permyakov and Robert H. Kretsinger Protein Chaperones and Protection from Neurodegenerative Diseases • Stephan Witt Transmembrane Dynamics of Lipids • Philippe Devaux and Andreas Herrmann

INTRODUCTION TO THE WILEY SERIES ON PROTEIN AND PEPTIDE SCIENCE Proteins and peptides are the major functional components of the living cell. They are involved in all aspects of the maintenance of life. Their structural and functional repertoires are endless. They may act alone or in conjunction with other proteins, peptides, nucleic acids, membranes, small molecules, and ions during various stages of life. Dysfunction of proteins and peptides may result in the development of various pathological conditions and diseases. Therefore, the protein/peptide structure–function relationship is a key scientific problem lying at the junction point of modern biochemistry, biophysics, genetics, physiology, molecular and cellular biology, proteomics, and medicine. The Wiley Series on Protein and Peptide Science is designed to supply a complementary perspective from current publications by focusing each volume on a specific protein- or peptide-associated question and endowing it with the broadest possible context and outlook. The volumes in this series should be considered required reading for biochemists, biophysicists, molecular biologists, geneticists, cell biologists, and physiologists, as well as those specialists in drug design and development, proteomics, and molecular medicine with an interest in proteins and peptides. I hope that each reader will find in the volumes within this book series interesting and useful information. First and foremost, I would like to acknowledge the assistance of Anita Lekhwani of John Wiley & Sons, Inc., throughout this project. She has guided me through countless difficulties in the preparation of this book series, and her enthusiasm, input, suggestions, and efforts were indispensable in bringing the Wiley Series on Protein and Peptide Science into existence. I would like to take this opportunity to thank everybody whose contribution in one way or another has helped and supported this project. Finally, special thank you goes to my wife, sons, and mother for their constant support, invaluable assistance, and continuous encouragement. Vladimir N. Uversky September 2008

TRANSMEMBRANE DYNAMICS OF LIPIDS EDITED BY PHILIPPE F. DEVAUX ANDREAS HERRMANN

The Wiley Series in Protein and Peptide Science Series Editor: Vladimir N. Uversky

A JOHN WILEY & SONS, INC. PUBLICATION

The cover shows a transparency used by P.D. in presentations given in the days before PowerPoint was available. The cartoons illustrate the principal of the ascorbate assay to assess the transbilayer motion and distribution of spin-labeled lipids in membranes taking the plasma membrane of red blood cells as an example (see Preface and Chapters 1 and 6). Lower cartoon: Spin-labeled lipids (red) were incorporated into the outer leaflet of the plasma membrane and redistributed between both leaflets. Upper cartoon: Ascorbate was added to the cell suspension reducing spin-labeled lipids selectively on the outer leaflet. By comparing the signal intensity of spin-labeled lipids without and with ascorbate, the transbilayer distribution of labeled lipid analogs can be measured. The transparency has been used many times (see the spread colors on the left). Just by chance, it was rediscovered among an amazing pile of reprints in the office of P.D. Copyright © 2012 by John Wiley & Sons, Inc. All rights reserved Published by John Wiley & Sons, Inc., Hoboken, New Jersey Published simultaneously in Canada No part of this publication may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, recording, scanning, or otherwise, except as permitted under Section 107 or 108 of the 1976 United States Copyright Act, without either the prior written permission of the Publisher, or authorization through payment of the appropriate per-copy fee to the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, (978) 750-8400, fax (978) 750-4470, or on the web at www.copyright.com. Requests to the Publisher for permission should be addressed to the Permissions Department, John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, (201) 748-6011, fax (201) 748-6008, or online at http://www.wiley. com/go/permissions. Limit of Liability/Disclaimer of Warranty: While the publisher and author have used their best efforts in preparing this book, they make no representations or warranties with respect to the accuracy or completeness of the contents of this book and specifically disclaim any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives or written sales materials. The advice and strategies contained herein may not be suitable for your situation. You should consult with a professional where appropriate. Neither the publisher nor author shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. For general information on our other products and services or for technical support, please contact our Customer Care Department within the United States at (800) 762-2974, outside the United States at (317) 572-3993 or fax (317) 572-4002. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic formats. For more information about Wiley products, visit our web site at www.wiley.com. Library of Congress Cataloging-in-Publication Data: Transmembrane dynamic lipids / edited by Philippe Devaux, Andreas Herrmann. â•…â•…â•… p. cm. – (The Wiley series in protein and peptide science ; 9) â•… Includes index. â•…â•… ISBN 978-0-470-38845-7 (hardback) â•‡ 1.â•‡ Membrane lipids.â•… 2.â•‡ Cell membranes.â•… 3.â•‡ Membrane transport.â•… I.â•‡ Devaux, Philippe F.â•… II.â•‡ Herrmann, Andreas. â•… QP752.M45T73 2012 â•… 572'.577–dc23 2011021424 Printed in the United States of America eISBN: 9781118120088 ePub: 9781118120101 oISBN: 9781118120118 MOBI: 9781118120095 10â•… 9â•… 8â•… 7â•… 6â•… 5â•… 4â•… 3â•… 2â•… 1

CONTENTS

INTRODUCTION LIST OF CONTRIBUTORS

xiii xxiii

PART Iâ•… ASSESSING TRANSMEMBRANE MOVEMENT AND ASYMMETRY OF LIPIDS

1

â•‡ 1â•… Methods for the Determination of Lipid Transmembrane Distribution and Movement in Biological Membranes

3

Philippe F. Devaux and Andreas Herrmann

1.1 1.2 1.3 1.4 1.5

Introduction,â•‡ 3 Development of Assays for Distribution and Translocation of Lipids across Membranes,â•‡ 4 Overview on Assays for Measuring Distribution and Translocation of Lipids across Cellular Membranes,â•‡ 7 Main Techniques Used to Determine Transbilayer Distribution of Endogenous Lipids in Cell Membranes,â•‡ 9 Main Techniques Used to Determine Transbilayer Distribution of Lipid Analogs in Cell Membranes,â•‡ 12 Abbreviations,â•‡ 21 References,â•‡ 21

â•‡ 2â•… Detection and Measurement of Unlabeled Lipid Transmembrane Movement

25

Iván López-Montero, Marisela Vélez, and Philippe F. Devaux

2.1 2.2

Introduction,â•‡ 25 Measurement of Transmembrane Flip-Flop of Unlabeled Lipids by Shape Change of GUVs,â•‡ 27 v

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2.3 2.4

Measurement of Transmembrane Flip-Flop of Unlabeled Lipids Using AFM,â•‡ 36 Conclusions,â•‡ 41 Acknowledgments,â•‡ 41 Abbreviations,â•‡ 41 References,â•‡ 42

PART IIâ•… LIPID ASYMMETRY IN CELL MEMBRANES

45

â•‡ 3â•… New Insights in Membrane Lipid Asymmetry in Animal and Plant Cells

47

Alain Zachowski

3.1 3.2 3.3

Lipid Asymmetry in Animal Membranes,â•‡ 47 Creating, Maintaining, or Randomizing the Membrane Phospholipid Distribution: Phospholipid Transporters,â•‡ 49 What about Lipid Asymmetry and Translocation in Plant Cell Membranes?,â•‡ 50 Abbreviations,â•‡ 61 References,â•‡ 61

â•‡ 4â•… Sphingolipid Asymmetry and Transmembrane Translocation in Mammalian Cells

65

Gerrit van Meer, Sylvia Neumann, and Per Haberkant

4.1 4.2 4.3 4.4 4.5 4.6

Introduction,â•‡ 65 Sphingosine, Sphingosine-1-Phosphate, and Ceramide,â•‡ 67 Ceramide,â•‡ 68 Glycosphingolipids,â•‡ 68 Sphingomyelin,â•‡ 70 Future Perspectives,â•‡ 71 Abbreviations,â•‡ 71 References,â•‡ 71

â•‡ 5â•… Transbilayer Movement and Distribution of Cholesterol Peter Müller, Anna Pia Plazzo, and Andreas Herrmann

5.1 5.2 5.3 5.4 5.5

Introduction,â•‡ 75 Physicochemical Features of Cholesterol,â•‡ 76 Methods for Measuring Cholesterol Transbilayer Movement and Distribution,â•‡ 77 Transbilayer Movement of Cholesterol in Model Membranes,â•‡ 81 Transbilayer Movement of Cholesterol in Biological Membranes,â•‡ 82

75

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5.6 5.7 5.8 5.9

Transbilayer Distribution of Cholesterol in Lipid and Biological Membranes,â•‡ 82 Cholesterol Flip-Flop: Fast or Slow?,â•‡ 87 Role of Proteins in the Transport of Cholesterol across Membranes,â•‡ 88 Concluding Remarks,â•‡ 90 Acknowledgment,â•‡ 92 Abbreviations,â•‡ 92 References,â•‡ 93

PART IIIâ•… ENERGY-INDEPENDENT PROTEIN-MEDIATED TRANSMEMBRANE MOVEMENT OF LIPIDS

97

â•‡ 6â•… Phospholipid Flip-Flop in Biogenic Membranes

99

Anant K. Menon and Andreas Herrmann

6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8 6.9 6.10

Introduction,â•‡ 99 Assays for Measuring Transbilayer Distribution of Endogenous Phospholipids,â•‡ 100 Assays for Measuring Transbilayer Distribution and Movement of Phospholipid Analogs,â•‡ 102 Shape Changes of GUVs as a Tool to Measure Flip-Flop,â•‡ 106 Transbilayer Movement of Phospholipids in the ER,â•‡ 108 Transbilayer Movement of Phospholipids in the Bacterial Inner Membrane,â•‡ 110 Mechanism of Rapid Lipid Flip-Flop in Biogenic Membranes,â•‡ 112 Efforts to Identify Phospholipid Flippases,â•‡ 113 Flipping of Isoprenoid-Based Glycolipids,â•‡ 115 Conclusion,â•‡ 115 Abbreviations,â•‡ 116 References,â•‡ 116

â•‡ 7â•… Phospholipid Scramblase: When Phospholipid Asymmetry Goes Away Edouard M. Bevers and Patrick L. Williamson

7.1 7.2 7.3 7.4 7.5 7.6

Introduction,â•‡ 119 Historical Overview,â•‡ 120 Physiological Importance of Lipid Scrambling,â•‡ 122 Characteristics of the Phospholipid Scrambling Process,â•‡ 124 Toward Identification: Proposed Candidate Proteins and Mechanisms,â•‡ 132 Concluding Remarks,â•‡ 139 Abbreviations,â•‡ 139 References,â•‡ 140

119

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PART IVâ•… ENERGY-DEPENDENT LIPID TRANSPORT ACROSS MEMBRANES

147

â•‡ 8â•… Flip or Flop: Mechanism and (Patho)Physiology of P4-ATPase-Catalyzed Lipid Transport

149

Patricia M. Verhulst, Joost C.M. Holthuis, and Thomas G. Pomorski

8.1 8.2 8.3 8.4 8.5 8.6

Introduction,â•‡ 149 P4-ATPases are Prime Candidate Phospholipid Translocases,â•‡ 152 Mechanism of P4-ATPase-Catalyzed Lipid Transport: Role of Accessory Subunits,â•‡ 156 Role of P4-ATPases in Vesicle-Mediated Protein Transport,â•‡ 161 P4-ATPase Dysfunction and Disease,â•‡ 162 Future Challenges,â•‡ 166 Acknowledgments,â•‡ 166 Abbreviations,â•‡ 166 References,â•‡ 167

â•‡ 9â•… Coupling Drs2p to Phospholipid Translocation, Membrane Asymmetry, and Vesicle Budding

171

Xiaoming Zhou, Paramasivam Natarajan, Baby-Periyanayaki Muthusamy, Todd R. Graham, and Ke Liu

9.1 9.2 9.3 9.4 9.5

Introduction,â•‡ 171 P4-ATPases in Budding Yeast,â•‡ 172 Evidence That Drs2p Is a Flippase,â•‡ 175 Drs2p in Protein Transport and Vesicle Budding,â•‡ 183 Concluding Remarks,â•‡ 191 Abbreviations,â•‡ 192 References,â•‡ 193

10â•… Substrate Specificity of the Aminophospholipid Flippase

199

Shelley M. Cook and David L. Daleke

10.1 10.2 10.3 10.4 10.5 10.6 10.7

Introduction,â•‡ 199 Substrate Specificity of the PM Aminophospholipid Flippase,â•‡ 200 Identification and Substrate Specificity of Candidate Aminophospholipid Flippases,â•‡ 205 Is the Lipid Specificity of Candidate Aminophospholipid Flippases Unique?,â•‡ 210 Lipid Specificity of Other PS-Binding Proteins,â•‡ 213 Sequence Elements That Bind to PS,â•‡ 215 Conclusions,â•‡ 216 Acknowledgments,â•‡ 217 Abbreviations,â•‡ 218 References,â•‡ 218

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11â•… The Flippase Delusion?

225

Naomi L. Pollock, Petra H.M. Niesten, and Richard Callaghan

11.1 11.2 11.3 11.4 11.5 11.6

ATP-Binding Cassette (ABC) Transporters and Lipid Flip-Flop,â•‡ 225 ABCA4 and Lipid Translocation: Explaining a Phenotype?,â•‡ 228 MsbA and Lipid Translocation: A Key to Survival,â•‡ 230 Drug and Lipid Movement by ABCB1: Is the Mechanism a Flip-Flop?,â•‡ 237 ABCB4: The Forgotten and Likely Lipid Flippase?,â•‡ 240 Conclusions and Perspectives,â•‡ 244 Abbreviations,â•‡ 244 References,â•‡ 245

PART Vâ•… RELEVANCE OF LIPID TRANSMEMBRANE DISTRIBUTION FOR MEMBRANE PROPERTIES AND PROCESSES

251

12â•… Membrane Lipid Asymmetry and Permeability to Drugs: A Matter of Size

253

Adam Blanchard and Cyril Rauch

12.1 12.2 12.3 12.4 12.5

Introduction,â•‡ 253 The Origin of Lipinski’s Second Rule from the Point of View of the Pharmaceutical Industry,â•‡ 254 Solving Lipinski’s Second Rule,â•‡ 257 Lipinski’s Second Law and Potential Application,â•‡ 264 Conclusion,â•‡ 270 Acknowledgment,â•‡ 272 Abbreviations,â•‡ 272 References,â•‡ 273

13â•… Endocytosis and Lipid Asymmetry Nina Ohlwein, Andreas Herrmann, and Philippe F. Devaux

13.1 13.2 13.3 13.4 13.5 13.6

Introduction,â•‡ 275 Bending a Membrane,â•‡ 276 Shape Changes of GUVs Induced by Lipid Asymmetry,â•‡ 278 How Endocytosis Is Linked to Lipid Asymmetry,â•‡ 280 Role of P4-ATPases in the Formation of Endocytic Invaginations,â•‡ 283 Concluding Remarks,â•‡ 284 Acknowledgments,â•‡ 285 Abbreviations,â•‡ 285 References,â•‡ 285

275

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CONTENTS

PART VIâ•… APOPTOSIS AND DISEASES: CONSEQUENCES OF DISRUPTION TO LIPID TRANSMEMBRANE ASYMMETRY

289

14â•… Membrane Lipid Asymmetry in Aging and Apoptosis

291

Krishnakumar Balasubramanian and Alan J. Schroit

14.1 14.2 14.3 14.4 14.5 14.6 14.7 14.8 14.9

Introduction,â•‡ 291 Phospholipid Transporters,â•‡ 292 Lipid Asymmetry in Erythrocytes,â•‡ 294 Lipid Asymmetry during Apoptosis,â•‡ 297 Ca2+ Homeostasis during Apoptosis,â•‡ 298 Membrane Phospholipid Asymmetry: Static or Dynamic?,â•‡ 299 Regulation of Lipid Asymmetry during Apoptosis,â•‡ 300 Significance,â•‡ 304 Concluding Remarks,â•‡ 306 Abbreviations,â•‡ 306 References,â•‡ 307

15â•… Phosphatidylserine Exposure in Hemoglobinopathies

315

Frans A. Kuypers and Eric Soupene

15.1 15.2 15.3 15.4 15.5 15.6 15.7 15.8

Introduction,â•‡ 315 RBC Phospholipid Organization,â•‡ 316 The RBC Flippase,â•‡ 319 PS Exposure in RBCs,â•‡ 324 PS Exposure in Hemoglobinopathies,â•‡ 328 Consequences of PS Exposure,â•‡ 329 Phospholipid Transbilayer Movement in Hemoglobinopathies,â•‡ 330 Conclusion,â•‡ 332 Abbreviations,â•‡ 333 References,â•‡ 334

16â•… Scott Syndrome: More Than a Hereditary Defect of Plasma Membrane Remodeling Florence Toti and Jean-Marie Freyssinet

16.1 16.2 16.3 16.4 16.5 16.6

Introduction,â•‡ 341 Scott Syndrome Features and Phenotype,â•‡ 342 Cell Biology of Scott Syndrome,â•‡ 343 Candidate Proteins in the Transmembrane Redistribution of PS,â•‡ 345 The Significance of Membrane Vesiculation and of Derived MPs,â•‡ 346 What Can Be Learned from Scott Syndrome?,â•‡ 347

341

xi

CONTENTS

16.7

Conclusion,â•‡ 348 Abbreviations,â•‡ 349 References,â•‡ 350

17â•… ABCA1, Tangier Disease, and Lipid Flopping

353

Ana Zarubica and Giovanna Chimini

17.1

Historical Notes: Tangier Disease (TD) and ATP-Binding Cassette Transporter 1 (ABCA1),â•‡ 353 17.2 The ABCA1 Gene and the Regulation of Its Expression,â•‡ 354 17.3 The ABCA1 Protein and Its Interactions,â•‡ 356 17.4 ABCA1: Mutations and Clinical Signs,â•‡ 358 17.5 Targeted Inactivation and Overexpression of ABCA1 in Animal Models,â•‡ 360 17.6 Liver and Macrophage ABCA1: Lipid Efflux and HDL Formation,â•‡ 362 17.7 ABCA1 and Membrane Function,â•‡ 363 17.8 ABCA1: Lipid Flop and Lipid Efflux,â•‡ 364 17.9 ABCA1 and the Lipid Microenvironment at the Membrane,â•‡ 366 17.10 Conclusions,â•‡ 368 Acknowledgments,â•‡ 369 Abbreviations,â•‡ 369 References,â•‡ 371 INDEX

379

Note: Color versions of many of the black and white figures in this book can be viewed at ftp://ftp.wiley.com/public/sci_tech_med/transmembrane_dynamics. Please read the figure captions to learn which figures are available on the ftp site.

INTRODUCTION

HISTORICAL PERSPECTIVES: WHO DID WHAT AND WHAT’S NEXT? Ole Mouritsen, in his recent monograph entitled “Lipids—As a Matter of Fat,” summarized with humor the views of many biologists concerning lipids, as follows: “Lipids appear to play a fairly non-specific role, being rather dull and anonymous compared to fashionable stuff like the proteins that catalyze all biochemical reactions and the genes that contain the information needed to produce proteins” [1]. The present book, which is addressed to researchers, teachers, and students in cell biology and in biochemistry, has the goal of convincing all scientists that lipids, on the contrary, have sophisticated behaviors and play multiple important roles in living organisms. It is also addressed to physicists fascinated by the various spontaneous self-organization of lipids in water (lipid polymorphism) to warn them that lipids in biological systems are not always at thermal equilibrium, and that phase separations and lateral or transmembrane domains seen in model systems can differ fundamentally from biological situations. Indeed, molecule segregation in biological systems results often from the work of ATPases, like the flippases, or is the result of a molecule sorting by “protein gates” (see the “fence and picket model” of Kusumi and collaborators [2]). Such mechanisms are difficult to mimic in model systems. In any case, all lipids are not equivalent and their chemical heterogeneity, for example, between the two sides of a biomembrane, is the result of a long selection during evolution, which allows lipids to fulfill different functions, from that of a fluid hydrophobic medium for membrane proteins to that of selective messenger molecules and enzyme cofactors. In the latter case, they xiii

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have to find their partners in a cell, hence to move rapidly in a very anisotropic environment. To many biologists, lipids form the third class of molecules of living organism after proteins and nucleic acids. Yet, lipids were probably not the third in the evolution nor are they third in importance, since a cell and even many viruses cannot exist without a membrane. The fact is that lipids form the building blocks of biological membranes. They determine the boundary of all living organisms as well as the compartmentalization of organelles in eukaryotes. Regarded as passive molecules forming only viscous cement that holds membrane proteins, filtering out hydrophilic molecules, the lipid bilayer is in reality a sophisticated structure capable of a remarkable polymorphism in water. The physical characteristic of a lipid bilayer permits not only protein movement but also membrane deformations and, coupled to the cytoskeleton, provides the cell membrane with mechanical properties. Not the least astonishing is the bilayer’s ability to divide in two compartments during cell division without losing molecules in the plasma due to efficient self-sealing capacities. Nonetheless, there are still mysteries concerning lipids, which are matters of research, speculation, and controversy. (1) Biophysicists have succeeded in making stable membranes (liposomes) with only one type of lipids, in suspension in water, for example, with egg phosphatidylcholine (PC), while biological membranes harbor several hundred different lipids. Why are there so many chemically different lipids coexisting in nature? (2) Why is the lipid composition of various membranes of eukaryotes different and sometimes even the two sides of biological membranes different (asymmetrical)? This requires numerous specific enzymes for the synthesis and ultimately for the shuttling to the right destination of newly formed lipids. Is such a multiplicity necessary for a fine-tuning of membrane-bound enzymes or is the variety of lipids used to give specific messages to specific proteins? Is the detailed chemical structure of lipids without real importance and does it reflect only the precursor molecules available? Not only do eukaryotic membranes have many chemically different lipids if one considers chain length, unsaturation, and polar head group, but also the lipids are not homogenously distributed within the various organelles and even between the different sides of one membrane. This lipid heterogeneity, a “complication of Nature,” was transmitted more than a million years in eukaryotic cells and has survived the filter of evolution, suggesting that the lipid composition and distribution within a cell is neither accidental nor inconsequential for the activity of cells. Although cells tolerate certain variability in lipid composition, many human diseases have been associated with the inability of mutated cells to synthesize specific lipids or to recycle particular lipids from the nutriments or to address specific lipids to their correct destination. Alternatively, the excess of certain lipids such as cholesterol or saturated phospholipid chains can be poisonous. In the late 1960s, V. Luzzati, in a pioneer work carried out in France, showed by X-ray crystallography that lipids extracted from biological membranes form, in water, lamellar phases, giving rise spontaneously to large multilamel-

INTRODUCTION

xv

lar (onion-style) liposomes made of a superposition of bilayers [3, 4]. Physicists characterized the bilayers as liquid crystals that could be in a fluid state or in a more viscous, gel state. In the early 1970s, the concept of lipid bilayer emerged as the basic model of biomembranes and was popularized in the famous model of “fluid mosaic membrane” of S.J. Singer and G.L. Nicolson [5]. Although the concept of “mosaicity” implies the presence of heterogeneous lateral domains, and in spite of the work carried out by several physical chemists such as H. McConnell, it was only in 1997 (almost 30 years after the initial work of Luzzati and McConnell) that the importance of lateral domains began to be popular among membranologists and that biological functions associated with lateral domains (or rafts) were highlighted (see the work of K. Simons and E. Ikonen [6]). Indeed, the two monolayers of biomembranes form distinct lipid domains: M. Bretscher in England demonstrated in the early 1970s the asymmetrical transmembrane distribution of phospholipids in the plasma membrane of human erythrocytes [7]. Bretscher used the chemical labeling of the amino groups of phosphatidylserine (PS) or phosphatidylethanolamine (PE) and showed that aminophospholipids are principally in the membrane inner monolayer, while PC and sphingomyelin (SM) are essentially in the outer monolayer of human red cells. Subsequent investigation in the laboratory of L.L.M. van Deenen in The Netherlands based on phospholipases and sphingomylinases assays [8, 9] confirmed Bretscher’s results and demonstrated that the transmembrane asymmetry of red cells is an ubiquitous property of the plasma membrane of eukaryotes. In model systems, on the other hand, no transmembrane lipid segregation was found to form spontaneously. Sonication allows one to achieve a lipid sorting between inner and outer monolayers in small unilamellar vesicles (SUVs), but the latter structures are not physiological because of their small size compared with that of vesicles produced in vivo (∼20-nm diameter for SUVs vs. ∼200â•¯nm for endocytic vesicles). Thus, lipid sorting observed in biomembranes had to be caused by a process that does not exist in liposomes and is not a mere thermodynamic equilibrium. Initially, the segregation of aminophospholipids was believed to be due to the topology of enzymes responsible for lipid synthesis or to lipid–cytoskeleton interactions (J.A.F. Op den Kamp [10]). However, Bretscher had the remarkable intuition to postulate the existence of specific lipid enzymes that he named “phospholipid flippase,” which would be responsible for the establishment of the asymmetrical lipid organization at the expense of ATP hydrolysis. In practice, it was later found necessary to specify the orientation of the postulated lipid carrier and the requirement or absence of requirement for ATP hydrolysis. This explains why the habit is now to differentiate among flippase, floppase, and scramblase (Fig. I.1). A prerequisite for stable lipid segregation between the two monolayers of a membrane is a priori a slow transmembrane diffusion. In 1971, R.D. Kornberg and H.M. McConnell at Stanford University demonstrated for the first time, with spin-labeled lipids, the very slow transmembrane diffusion of

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INTRODUCTION

extracellular or lumenal

ATP

flippase

cytoplasm

ATP

floppase

scramblase

flip-flop

Figure I.1.â•‡ Definition of the various lipid transporters in eukaryotic cell membranes. Note that the scramblase is calcium dependent and that “flippase” is a term that is used sometimes to designate an enzyme that catalyzes lipid flip-flop in both directions (inward or outward), for example, in the endoplasmic reticulum.

phospholipids in sonicated lipid vesicles, where the “flip-flop” between the two monolayers was found to require several hours at 30°C [11]. It is now admitted that the spontaneous transmembrane diffusion of lipids is very slow in liposomes of any size as well as in biological membranes. A few exceptions to this rule were discovered recently. Cholesterol, ceramide, phospatidic acid, diacylglycerol, and free fatty acids or esters have a rapid spontaneous diffusion (τ1/2 less than 1 minute). The absence of real polar head groups in such lipids probably explains this unusual result. It was only in 1984, that is, more than 10 years after Bretscher’s hypothesis, that the existence of a phospholipid flippase was demonstrated in France by M. Seigneuret and P.F. Devaux in the human erythrocyte membrane using spin-labeled analogs of naturally occurring phospholipids [12] and the year after by D.L. Daleke and W.H. Huestis, who provided confirmation using an elegant technique involving nonlabeled lipids [13], while A. Schroit’s group [14] took advantage of fluorescent analogs to prove the existence of an erythrocyte aminophospholipid transporter. The requirement of hydrolyzable Mg2+ATP was demonstrated as being necessary for the rapid transport of aminophospholipids, and the specificity was carefully investigated; however, no proteins were identified initially. In 1989, an ATP-dependent flippase activity in chromaffin granules from bovine adrenal medulla was reported by the Paris laboratory and attributed to the so-called ATPase II [15]. This was the first report of aminophospholipid translocase activity in the inner membranes of the eukaryotic cell. The transport observed was in fact from the lumen to the cytosol of the granules but was classified as a flippase activity. In 1996, P. Williamson and R.A. Schlegel’s groups in the United States showed that this granule flippase was homolog to a yeast ATPase (called Drs2p), and studied a mutant deprived of Drs2 that was unable to flip aminophospholipids [16]. The phospholipid flippase seemed to be discovered. However, in 1999 and 2003, the groups of T. Graham in the United States [17] and G. van Meer and J. Holthuis in The Netherlands [18] showed that Drs2p is in fact localized in the yeast trans-Golgi and not in the plasma mem-

INTRODUCTION

xvii

brane, and that five homologs of this protein exist: two in the plasma membrane (Dnf1p and Dnf2p), two in the trans-Golgi (Dnf3p and Drs2p), and one (Neo1p) in endosomes or cis-Golgi. Furthermore, these P-type ATPases seem to be associated with other proteins playing the role of chaperones (CDC50p) or are necessary for the proper targeting to their final destination of the newly formed proteins [19]. In 2006, P. Natarajan and T. Graham [20] showed a flippase activity with fluorescent lipids in yeast Golgi membranes, which they could attribute to the Drs2p. Interestingly, the triple knockout of the Drs2p homologs in yeast led to viable cells, but they were deprived of endocytic activity [18]. In conclusion, the various P-type ATPases may have different specificities but may also be partially redundant. Thus, after about 20 years of research in different laboratories throughout the world, it became obvious that the ubiquitous eukaryotic flippase was in reality a combination of several proteins, including four ATPases called P4ATPase, actually forming a family of five proteins in yeast. In humans, it was predicted from genomic investigation that 14 P4-ATPases were members of the family and could be involved in lipid transport. The purification of specific P4-ATPases and of Drs2p from chromaffin granules or after expression in various systems (yeast and insect cells) is in progress. However, so far the purification has not been achieved on a large enough scale to allow unambiguous tests of lipid transport in reconstituted lipoproteins. Other membrane proteins were reported to have an ATP-dependent lipid translocation activity and correspond to the so-called floppases (see Fig. I.1) with an ATP-binding cassette (ABC). Suggested originally by C.F. Higgins and M.M. Gottesman in 1992 [21], the laboratories of G. van Meer and of P. Borst in The Netherlands [22] showed in 1996 that the ABC transporter P-glycoprotein, also called MDR1, which is responsible for multidrug resistance and is a serious obstacle in cancer therapy, was able to transport fluorescent phospholipids from the inner monolayer to the outer monolayer of the plasma membrane of eukaryotic cells. The low specificity of the P-glycoprotein suggested that this protein could be involved in the transport of SM and PC toward the outer monolayer of the plasma membrane, hence play an important role in the transmembrane lipid asymmetry of the eukaryotic plasma membrane. Other members of the ABC protein family seemed to be responsible for the specific outward transport of PC in transfected epithelial cells [22]. An important point is that ABC proteins are also found in prokaryotes and could be implicated in lipid translocation in bacteria [23]. Besides ATP-dependent flippases, which were found essentially in the plasma membrane of eukaryotes, other ATP-independent proteins also called flippases were postulated to be in specific organelle membranes (endoplasmic reticulum) and could explain the rapid flip-flop observed by several groups. Their primary function would be to facilitate the transmembrane diffusion of lipids in the membranes specialized in lipid synthesis. Already in 1985, W.R. Bishop and R.M. Bell [24] suggested the existence of ATP-independent flippase, catalyzing the diffusion of PC in the endoplasmic reticulum. Since

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then, several researchers have attempted to isolate the protein(s) responsible (A. Menon in the United States [25] and A. Herrmann and collaborators in Germany [26]). Other researchers have attempted to prove that any transmembrane protein should accelerate the flip-flop of lipids, making it unnecessary to search for specific proteins (B. de Kruijff and A. Killian and collaborators in The Netherlands [27, 28]). In practice, the identification of the ATPindependent flippase of low lipid specificity seems even more difficult than it is for the ATP-dependent selective flippase, precisely because the test of ATP requirement cannot be used to discover the latter transporter. BIOLOGICAL ADVANTAGES OF LIPID ASYMMETRY The complexity involved in the regulation of lipid topology, requiring ATP hydrolysis, raises the question of the biological function(s) of such an elaborate system. Actually, one might rephrase this question differently: The lipid composition of a biological membrane is always a mixture of many different lipids. The actual justification of this fact is not obvious, since a stable lipid bilayer can be achieved in liposomes with a single phospholipid species. So what is the biological advantage of the synthesis of many different lipids? A reasonable hypothesis would be that lipid asymmetry is used to tag the two sides of a membrane and to optimize their functionality, which is obviously different. Indeed, the cell outer environment differs fundamentally from the cytosol. One of the first indications of the physiological importance of lipid asymmetry came from the observation by A. Schroit and collaborators (United States) who showed in 1983 and 1985 that the presence of a very small percentage of PS (∼1% of the total lipid composition) in the outer monolayer of red cells was used in vivo as a signal of cell aging and led in the blood circulation to the elimination of aged cells by macrophages [29]. These conclusions, which came originally from experiments associated with the introduction of exogenous PS in the outer monolayer of red cells, were confirmed later by the detection of natural PS with fluorescent Annexin V by J.F. Tait and D. Gibson in 1994 [30]. The exposure of PS in the outer monolayer of platelets is also associated with the formation of clots that stop bleeding (R. Zwaal and collaborators [31]). Thus, lipid flip-flop concerns directly at least two important physiological problems: (1) blood coagulation, which is triggered in vivo by the exposure of PS, a cofactor required for the conversion of prothrombin into thrombin, and (2) elimination of aged and/or apoptotic cells by macrophages. The lipid randomization, that is, loss of lipid asymmetry that is used in vivo as a signal for cell elimination, can be triggered artificially by penetration of calcium ions in the cytosol of platelets, erythrocytes, or lymphocytes with calcium ionophores, and results in “lipid scrambling,” that is, lipid randomization between the two leaflets. This phenomenon is associated with a so far unknown protein named

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“scramblase.” (Note: Suzuki et al. [40] identified the protein TMEM16F as an essential component for Ca2+-dependent exposure of PS.) A rare but severe disease, called “Scott syndrome,” is characterized by the absence of PS redistribution upon calcium entry and has been investigated by R. Zwaal’s group in The Netherlands [32] and by J.-M. Freyssinet and collaborators [33] in France. Various severe diseases such as cancer and Alzheimer’s disease were also reported to be accompanied by defects in lipid asymmetry [34]. ABCA1, another lipid transporter of the family of ABC-ATPases, was considered to be responsible for Tangier disease, characterized by impaired efflux of cholesterol and phospholipids from peripheral cells onto apolipoproteins such as Apo A-1. Cholesterol accumulation in macrophages and apolipoprotein degradation lead to tissue deposition of cholesterol esters and increase the risk of arteriosclerosis in patients. G. Chimini and collaborators in Marseilles studied this particular defect associated with a lipid transporter [35]. In humans, several mutated ABC proteins reputed to be responsible for lipid transport are believed to cause metabolism disorders such as Stargardt syndrome (a genetic disease of vision), progressive intrahepatic cholestasis, pseudoxanthoma elasticum, adrenoleukodystrophy, or sitosterolemia. In 1999, E. Farge and collaborators, in A. Dautry-Varsat’s laboratory, provided evidence of a biological role played by a lipid transporter during the first step of endocytosis [36]. It was shown that the transport of PS and PE from the outer to the inner monolayer by the ATP-dependent flippase is a stimulation of endocytosis and could be the molecular motor of membrane bending involved in the first step of endocytosis. The explanation proposed was that the excess of lipids in one monolayer triggers membrane invagination, as shown in model systems [37]. The yeast knockout experiments mentioned above [18] confirmed that in the absence of flippase proteins, endocytosis was blocked. There are also reports suggesting that PS is important for fusion; hence, it could be useful in the inner monolayer for exocytosis and not only for the regulation of inner leaflet proteins. There are certainly many other enzymes that require specific lipids at specific positions in a cell. Actually, the difference in head group of the lipids from the two sides of a membrane is not the only difference between inner and outer leaflets lipids. Indeed, there is evidence of difference in unsaturation, which is associated with differences in membrane viscosity, as observed in erythrocytes with spin-labeled lipids [38] and with fluorescent lipids [39]: The inner monolayer is more fluid; the outer is more rigid, hence more resistant. It is very likely that this feature is associated with the activity of proteins. When transmembrane lipid asymmetry was demonstrated in red cells and soon after in the plasma membranes of all animals, it was assumed that this feature was a general property of living organisms. This may be true in animal

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and in plant cells, which are both eukaryotes. But the evidence regarding prokaryotes is limited and often concerns rare lipids. PROSPECTS What kind of progress can be expected in a reasonable time? Clearly, the bottleneck for progress in understanding the mechanism of lipid translocation by membrane proteins in eukaryotes has been the difficulty in assigning, isolating, and overexpressing the protein(s) responsible for this process; studying the properties of proteoliposomes; and crystallizing a flippase. Crystallization will be a necessary step for ultimately understanding the mechanism that allows a hydrophobic transmembrane protein to accumulate against a gradient amphiphilic molecule. There are some reports, at low resolution, on the structure of ABC proteins possibly involved in lipid transport. With P4-type ATPases, the data obtained with Ca2+-ATPase can be used as first-order approximation to stimulate the speculations of researchers, but the difference between a lipid and a calcium ion is so large that the detailed analysis of the mechanism is presumptuous. In any case, the determination of the structure and molecular mechanism of a flippase is a challenge for the coming years. It is therefore an objective that cannot be forsaken. Progress in the molecular biology and purification of the P4-type ATPases will lead to this achievement. Other objectives are as follows: 1. isolation of protein(s) responsible for ATP-independent rapid lipid flipflop in the endoplasmic reticulum; 2. isolation of protein(s) responsible for calcium-induced lipid scrambling (scramblase); 3. deeper understanding of all the consequences of lipid asymmetry, including recognition of the diseases caused specifically by a defect (impairment) in flippase activity. ORGANIZATION OF THIS BOOK As shown in the Table of Contents of this book, each chapter concentrates on one particular aspect of lipid asymmetry in biomembranes. However, we are not yet in a situation to give a complete and rational picture. As a consequence, one of the main difficulties in assembling this book was to choose a rational order for the chapters. Although the various chapters are closely linked to each other, there were no compelling reasons to decide which subjects deserved to be first or second. Hence, some repetition is unavoidable and the order of the chapters is rather arbitrary. Nevertheless, we must apologize for this weakness. On the other hand, each chapter can stand alone and does not necessarily require the reading of other chapters.

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ACKNOWLEDGMENTS We would like to express our gratitude to the contributing authors, in particular for being so flexible and sympathetic. Many of the contributing authors belonged to an international research and training network, “Flippases,” which was funded by the European Union from 2005 to 2008. This network provided an important trigger for assembling the book. We thank Professor Sophie Cribier (Paris) and Professor Daniel Picot (Paris) for their help in speeding up the final steps for assembling the book. Finally, we are very much indebted to Anita Lekhwani, Senior Acquisitions Editor at John Wiley and Sons, and to Catherine Odal, Assistant to the Senior Acquisitions Editor, for their efficient collaboration and support as well as for being flexible and handling unforeseen problems pleasantly. November 2010

Philippe F. Devaux Andreas Herrmann

REFERENCES â•‡ 1â•… O. Mouritsen, Lipid—As a Matter of Fat. The Merging Science of Lipidomics, Springer, Berlin, 2005. â•‡ 2â•… C. Nakada, K. Ritchie, Y. Oba, M. Nakamura, Y. Hotta, R. Iino, R. S. Kasai, K. Yamaguchi, T. Fujiwara, A. Kusumi, Nat. Cell Biol. 2003, 5, 626–632. â•‡ 3â•… R. P. Rand, V. Luzzati, Biophys. J. 1968, 8, 125–137. â•‡ 4â•… V. Luzzati, F. Reiss-Husson, E. Rivas, T. Gulik-Krzywicki, Ann. N.Y. Acad. Sci. 1966, 137, 409–413. â•‡ 5â•… S. J. Singer, G. L. Nicolson, Science 1972, 175, 720–731. â•‡ 6â•… K. Simons, E. Ikonen, Nature 1997, 387, 569–572. â•‡ 7â•… M. S. Bretscher, Science 1973, 181, 622–629. â•‡ 8â•… A. J. Verkleij, R. F. A. Zwaal, B. Roelofsen, P. Comfurius, D. Kastelijn, L. L. M. van Deenen, Biochim. Biophys. Acta 1973, 323, 178–193. â•‡ 9â•… R. F. A. Zwaal, B. Roelofsen, P. Comfurius, L. L. M. van Deenen, Biochim. Biophys. Acta 1975, 406, 83–96. 10â•… J. A. F. Op den Kamp, Biochemistry 1979, 48, 47–71. 11â•… R. D. Kornberg, H. M. McConnell, Biochemistry 1971, 10, 1111–1120. 12â•… M. Seigneuret, P. F. Devaux, Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 3751–3755. 13â•… D. L. Daleke, W. H. Huestis, Biochemistry 1985, 24, 5406–5416. 14â•… J. Connor, A. J. Schroit, Biochemistry 1987, 26, 5099–5105. 15â•… A. Zachowski, J. P. Henry, P. F. Devaux, Nature 1989, 340, 75–76. 16â•… X. J. Tang, M. S. Halleck, R. A. Schlegel, P. Williamson, Science 1996, 272, 1495–1497. 17â•… C.-Y. Chen, M. F. Ingram, P. H. Rosal, T. R. Graham, J. Cell Biol. 1999, 147, 1223–1236. 18â•… T. Pomorski, R. Lombardi, H. Riezman, P. F. Devaux, G. van Meer, J. C. Holthuis, Mol. Biol. Cell 2003, 14, 1240–1254.

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19â•… K. Saito, K. Fujimura-Kamada, N. Furuta, U. Kato, M. Umeda, K. Tanaka, Mol. Biol. Cell 2004, 15, 3418–3432. 20â•… P. Natarajan, T. R. Graham, Methods 2006, 39, 163–168. 21â•… C. F. Higgins, M. M. Gottesman, Trends Biochem. Sci. 1992, 17, 18–21. 22â•… A. van Helvoort, A. J. Smith, H. Sprong, I. Fritzsche, A. H. Schinkel, P. Borst, G. van Meer, Cell 1996, 87, 507–517. 23â•… A. Pohl, P. F. Devaux, A. Herrmann, Biochim. Biophys. Acta 2005, 1733, 29–52. 24â•… W. R. Bishop, R. M. Bell, Cell 1985, 42, 51–60. 25â•… A. Menon, W. E. Watkins, III, S. Hrafnsdóttir, Curr. Biol. 2000, 10, 241–252. 26â•… S. Vehring, L. Pakkiri, A. Schroer, N. Alder-Baerens, A. Herrmann, A. K. Menon, T. Pomorski, Eukaryot. Cell 2007, 6, 1625–1634. 27â•… M. A. Kol, A. I. P. M. de Kroon, J. A. Killian, B. de Kruijff, Biochemistry 2004, 43, 2673–2681. 28â•… M. A. Kol, A. I. P. M. de Kroon, D. T. S. Rijkers, J. A. Killian, B. de Kruijff, Biochemistry 2001, 40, 10500–10506. 29â•… A. J. Schroit, J. W. Madsen, Y. Tanaka, J. Biol. Chem. 1985, 260, 5131–5138. 30â•… J. F. Tait, D. Gibson, J. Lab. Clin. Med. 1994, 123, 741–748. 31â•… E. M. Bevers, P. Comfurius, R. F. A. Zwaal, Biochim. Biophys. Acta 1983, 736, 57–66. 32â•… E. M. Bevers, T. Wiedmer, P. Comfurius, S. J. Shattil, H. J. Weiss, R. F. A. Zwaal, P. J. Sims, Blood 1992, 79, 380–388. 33â•… N. Bettache, P. Gaffet, N. Allegre, L. Maurin, F. Toti, J.-M. Freyssinet, A. Bienvenue, Br. J. Haematol. 1998, 101, 50–58. 34â•… A. Castegna, C. M. Lauderback, H. Mohmmad-Abdul, D. A. Butterfield, Brain Res. 2004, 1004, 193–197. 35â•… Y. Hamon, C. Broccardo, O. Chambenoit, M.-F. Luciani, F. Toti, S. Chaslin, J.-M. Freyssinet, P. F. Devaux, J. Neish, D. Marguet, G. Chimini, Nat. Cell Biol. 2000, 2, 399–406. 36â•… E. Farge, D. M. Ojcius, A. Subtil, A. DautryVarsat, Am. J. Physiol. Cell Physiol. 1999, 45, C725–C733. 37â•… E. Farge, P. Devaux, Biophys. J. 1992, 61, 347–357. 38â•… M. Seigneuret, A. Zachowski, A. Herrmann, P. F. Devaux, Biochemistry 1984, 23, 4271–4275. 39â•… G. Morrot, S. Cribier, P. F. Devaux, D. Geldwerth, J. Davoust, J. F. Bureau, P. Fellmann, P. Herve, B. Frilley, Proc. Natl. Acad. Sci. U.S.A. 1986, 83, 6863–6867. 40â•… Suzuki et al., Nature 2010, 468, 834–838.

LIST OF CONTRIBUTORS

Krishnakumar Balasubramanian, University of Pittsburgh, Pittsburgh, PA 15219; Email: [email protected] Edouard M. Bevers, Department of Biochemistry, Cardiovascular Research Institute Maastricht, Maastricht University, The Netherlands; Email: [email protected] Adam Blanchard, School of Veterinary Medicine and Science, University of Nottingham, Sutton Bonington Campus, College Road, Sutton Bonington, Leicestershire LE12 5RD, UK Richard Callaghan, Nuffield Department of Clinical Laboratory Sciences, John Radcliffe Hospital, University of Oxford, Oxford, UK; Email: [email protected] Giovanna Chimini, Centre d’Immunologie de Marseille-Luminy, INSERMCNRS-Université de La Méditerranée, Parc Scientifique de Luminy, 13288, Marseille, France; Email: [email protected] Shelley M. Cook, Department of Biochemistry and Molecular Biology, Medical Sciences, Bloomington, Indiana University School of Medicine, Bloomington, IN 47405 David L. Daleke, Department of Biochemistry and Molecular Biology, Medical Sciences, Bloomington, Indiana University School of Medicine, Bloomington, IN 47405; Email: [email protected] Philippe F. Devaux, Institut de Biologie Physico-Chimique, 13 rue Pierre et Marie Curie 75005 Paris , France; Email: [email protected] xxiii

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Jean-Marie Freyssinet, U. 770 INSERM, Hôpital de Bicêtre, Le KremlinBicêtre, France; Faculté de Médecine, Université Paris-Sud, Le KremlinBicêtre, France; and Faculté de Médecine, Institut d’Hématologie & Immunologie, Université de Strasbourg, Strasbourg, France; Email: [email protected] Todd R. Graham, Department of Biological Sciences, Vanderbilt University, Nashville, TN 37235; Email: [email protected] Per Haberkant, EMBL, Heidelberg, Germany Andreas Herrmann, Department of Biology, Humboldt-University Berlin, Invalidenstr. 42, D-10115 Berlin, Germany; Email: andreas.herrmann@ rz.hu-berlin.de Joost C.M. Holthuis, Department of Membrane Enzymology, Bijvoet Center and Institute of Biomembranes, Utrecht University, 3584 CH Utrecht, The Netherlands; Email: [email protected] Frans A. Kuypers, Children’s Hospital Oakland Research Institute, 5700 Martin Luther King Way, Oakland, CA 94609; Email: [email protected] Ke Liu, NIH Chemical Genomics Center, Bethesda, MD 20892 Iván López-Montero, Departamento de Química Física I, Universidad Complutense de Madrid, 28040 Madrid, Spain; Email: [email protected] Anant K. Menon, Department of Biochemistry, Weill Cornell Medical College, New York, NY 10065 Peter Müller, Department of Biology, Humboldt-University Berlin, InvaliÂ� denstr. 42, D-10115 Berlin, Germany; Email: [email protected] Baby-Periyanayaki Muthusamy, Department of Biological Sciences,Vanderbilt University, Nashville, TN 37235 Paramasivam Natarajan, Department of Biological Sciences, Vanderbilt University, Nashville, TN 37235 Sylvia Neumann, Department of Cell Biology, The Scripps Research Institute, La Jolla, CA Petra H.M. Niesten, Nuffield Department of Clinical Laboratory Sciences, John Radcliffe Hospital, University of Oxford, Oxford, UK Nina Ohlwein, Department of Biology, Humboldt University Berlin, Invalidenstr. 42, D-10115 Berlin, Germany; Email: [email protected] Anna Pia Plazzo, Department of Biology, Humboldt-University Berlin, Invalidenstr. 42, D-10115 Berlin, Germany Naomi L. Pollock, Nuffield Department of Clinical Laboratory Sciences, John Radcliffe Hospital, University of Oxford, Oxford, UK


xxv

Thomas G. Pomorski, Department of Plant Biology and Biotechnology, Faculty of Life Sciences, University of Copenhagen, DK-1871 Frederiksberg C, Denmark Cyril Rauch, School of Veterinary Medicine and Science, University of Nottingham, Sutton Bonington Campus, College Road, Sutton Bonington, Leicestershire LE12 5RD, UK; Email: [email protected] Alan J. Schroit, Department of Pharmacology, The University of Texas Southwestern Medical Center, Dallas, TX 75390; Email: alan.schroit@ utsouthwestern.edu Eric Soupene, Children’s Hospital Oakland Research Institute, 5700 Martin Luther King Way, Oakland, CA 94609 Florence Toti, U. 770 INSERM, Hôpital de Bicêtre, Le Kremlin-Bicêtre, France; Faculté de Médecine, Université Paris-Sud, Le Kremlin-Bicêtre, France; and Faculté de Médecine, Institut d’Hématologie & Immunologie, Université de Strasbourg, Strasbourg, France Gerrit van Meer, Faculty of Science, Utrecht University, Padualaan 8, 3584 CH Utrecht, The Netherlands; Email: [email protected] Marisela Vélez, Instituto de Catálisis y Petroleoquímica, Consejo Superior de Investigaciones Científicas, 28049 Madrid, Spain; and IMDEA Nanociencia, Facultad de Ciencias, Universidad Autonóma de Madrid, 28049 Madrid, Spain; Patricia M. Verhulst, Department of Membrane Enzymology, Bijvoet Center and Institute of Biomembranes, Utrecht University, 3584 CH Utrecht, The Netherlands Patrick L. Williamson, Department of Biology, Amherst College, Amherst, MA; Email: [email protected] Alain Zachowski, Laboratory of “Physiologie Cellulaire et Moléculaire des Plantes,” Université Pierre et Marie Curie—Paris 6 (UR 5) and Centre National de la Recherche Scientifique (EAC 7180); Email: alain.zachowski@ upmc.fr Ana Zarubica, Centre d’Immunologie de Marseille-Luminy, INSERMCNRS-Université de La Méditerranée, Parc Scientifique de Luminy, 13288, Marseille, France Xiaoming Zhou, Department of Biological Sciences, Vanderbilt University, Nashville, TN 37235

PART I ASSESSING TRANSMEMBRANE MOVEMENT AND ASYMMETRY OF LIPIDS

1 METHODS FOR THE DETERMINATION OF LIPID TRANSMEMBRANE DISTRIBUTION AND MOVEMENT IN BIOLOGICAL MEMBRANES Philippe F. Devaux Institut de Biologie Physico-Chimique, Paris, France

Andreas Herrmann Department of Biology, Humboldt-University Berlin, Berlin, Germany

1.1â•… INTRODUCTION To access the transbilayer distribution and movement of lipids and, perhaps in particular, to convince all readers that lipids really are distributed asymmetrically between the two leaflets of eukaryotic cell plasma membranes and to explain how lipid transporters were discovered in biomembranes, it is necessary to give an overview of the main techniques that were and—in many cases—are still used. Our objective here is in fact limited to give solely an overview as well as an indication of the limits of the techniques that have been used during the last 40 years. In this book, it is not possible to detail the various aspects and weakness of each technique. Details can be found in specialized publications, but also in various chapters of this book, to which

Transmembrane Dynamics of Lipids, First Edition. Edited by Philippe F. Devaux, Andreas Herrmann. © 2012 John Wiley & Sons, Inc. Published 2012 by John Wiley & Sons, Inc.

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the readers will be referred. These various techniques, because they have given not always identical but still very consistent results, have allowed some definitive conclusions to be drawn. We will respond to criticisms against the use of lipid analogs carrying a reporter moiety which “could introduce artifacts.” We are well aware, and will outline below, that quantitative but rarely qualitative differences with respect to the behavior of endogenous lipids can be caused by reporter moieties covalently attached to lipids. However, even assays characterizing the transbilayer organization of endogenous lipids can cause modifications of the membrane no longer comparable to the unperturbed, original situation. Nevertheless, looking back over decades of research in this field, analogs in conjunction with assays that may affect membrane properties have provided milestones in understanding the dynamics of transbilayer distribution of lipids. For example, by using spin-labeled phospholipids, two major discoveries in this field were made. First, early in the 1970s, Kornberg and McConnell were able to give for the first time quantitative data on kinetics of passive transbilayer movement (flip-flop) of phospholipids in a bilayer [1]. Second, in 1984, Seigneuret and Devaux discovered by using short-chain spin-labeled lipids that the inward translocation of aminophospholipids in red blood cell membranes is ATP dependent, pointing to an energy-dependent lipid transporter that may also be typical for the plasma membrane of other mammalian or even all eukaryotic cells [2]. Indeed, subsequently similar conclusions of an ATP-dependent inward translocation of aminophospholipids were obtained from studies with fluorescent [3–5] as well as nonlabeled short-chain exogenous [6, 7] or even radioactive long-chain lipids [8]. Hence, being aware that labeling of lipids by reporter moieties affects their properties, lipid analogs provided significant insights into the transbilayer movement and distribution of lipids. Nevertheless, independent complementary methods, in particular those based on endogenous lipids, or at least long-chain lipids with nonperturbing labels (e.g., radioactive), are not only desirable but even mandatory to reach confident conclusions. However, applications of labeled lipid analogs are typically easier to perform as techniques relying on natural lipids. Hence, starting with lipid analogs to address questions on transbilayer lipid organization might efficiently pave the way to apply or even to develop subsequent techniques based on endogenous lipids. 1.2â•… DEVELOPMENT OF ASSAYS FOR DISTRIBUTION AND TRANSLOCATION OF LIPIDS ACROSS MEMBRANES Discoveries of essential aspects of transbilayer lipid organization, in particular of plasma membranes, have been driven by the development of new methods. Likewise, questions that could not be solved with the available repertoire of methods initiated new methods. Early studies on transbilayer organization of lipids addressed whether phospholipids are asymmetrically distributed across

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the membrane. Those studies investigated essentially the transbilayer distribution of endogenous lipids in the plasma membrane of red blood cells, for example, by phospholipases or by chemical modifications of lipids (see below). They revealed that aminophospholipids phosphatidylserine (PS) and phosphatidylethanolamine (PE) are preferentially localized on the cytoplasmic leaflet of red blood cells, while phosphatidylcholine (PC) and sphingomyelin (SM) are predominantly on the exoplasmic leaflet (see reviews by Op den Kamp [9, 10]). In particular, PS, contributing to about 10% of the total phospholipid content of the plasma membrane, was almost exclusively shielded from the external leaflet. These studies also implicated that the asymmetric phospholipid distribution in the plasma membrane is typical for mammalian cells. Of course, this unique distribution immediately raised the question about the molecular mechanism not only generating but also preserving the asymmetric distribution, for example, in the case of the red blood cell circulating for about 120 days in human blood vessels. So, the question was, can lipid asymmetry be created by spontaneous segregation of lipids between the two leaflets of a membrane? Lateral segregation of lipids in domains, for example, in cholesterol- and sphingolipid-enriched domains, so-called rafts [11–14], is a thermodynamic phase separation. In contrast, transversal segregation of lipids does not exist spontaneously in a pure lipid bilayer. However, several factors may trigger such segregation. Membrane bending can cause a spontaneous segregation determined by the size of the polar head group of the lipids, which are mixed initially. In fact, a topological asymmetry exists due to membrane curvature. If one mixes lipids with a large head group and lipids with a small head group and prepares small unilamellar vesicles by sonification, a segregation of lipids is generated due to the large membrane curvature; the preference of lipids with a large head group is to occupy the external side where the curvature allows more space (see the review by Op den Kamp [10]). Other factors spontaneously generating lipid asymmetry are feasible in biological membranes with an asymmetrical environment on each side of a membrane.Transmembrane potential generally creates an electric field that polarizes each side of a membrane. Membrane proteins are not structurally symmetrically organized across the membrane. ATPases of the plasma membrane, for example, have charged residues usually on the cytoplasmic leaflet where ATP binds. Other proteins bind on the external surface or on the cytoplasmic monolayer where the cytoskeleton is attached. One could speculate that positively charged residues of those proteins may interact preferentially with lipids carrying head groups of opposite charges such as PS, or negatively charged amino acid residues with PS via the divalent calcium ion, eventually giving rise to an asymmetric transbilayer arrangement of these lipids. This is how the asymmetrical organization of lipids was initially explained with PS being trapped on the cytoplasmic leaflet of the red blood cell plasma membrane by its negatively charged head group interacting with the cytoskeleton proteins [9, 10]. However, this concept left many questions unanswered.

6


A new twist in understanding the generation and maintenance of lipid asymmetry in the plasma membrane of red blood cells (and other mammalian cells) was provided by the introduction of spin-labeled and fluorescent lipid analogs. When spin-labeled aminophospholipids were inserted into the exoplasmic leaflet of human red blood cells, surprisingly, they rapidly redistributed to the cytoplasmic leaflet [2, 15]. PS analogs disappeared almost completely from the external leaflet within a few minutes at 37°C, while PE reached a stationary distribution with about 80% of the analog on the inner leaflet within about 40 minutes [2]. In contrast, spin-labeled PC and SM moved only very slowly to the cytoplasmic monolayer and remained essentially on the external side. These studies provided two exciting results: a more technical one and a heuristic one. First, the stationary distribution of spin-labeled phospholipids was very similar to that of endogenous phospholipids already known, demonstrating that lipid analogs could qualitatively mimic their endogenous counterparts. Second, there is an energy-dependent transport of specific phospholipids in the plasma membrane. The directed and fast inward redistribution of aminophospholipids immediately provided an explanation for the generation and maintenance of lipid asymmetry in the plasma membrane of human red blood cells and—as implicated by later studies—typically for mammalian cells. In contrast to the model explaining lipid asymmetry by a specific interaction of lipids with the cytoskeleton, the finding of a directed transport could explain how a cell can rapidly repair or readjust any perturbation of lipid asymmetry caused by, for example, endo- and exocytotic processes. Moreover, as mentioned above, lipid asymmetry has to remain during the lifetime of the cells, which lasts sometimes more than several days. The transverse diffusion of lipids or lipid flip-flop, even slow, should lead finally to an equilibration of the lipid distribution between the two monolayers in the absence of a mechanism of lipid distribution repair, which in practice is carried out by a transporter and a flippase protein, and requires ATP as a source of energy. These and other results [16] obtained with lipid analogs also made clear why assays based on chemical labeling or phospholipase treatment of endogenous lipids in the way they were performed could not recognize the rapid inward motion of aminophospholipids. Comparing the time required to perform the assays with the characteristic time of inward and outward motion of phospholipids, the experimental approaches would not have been able to detect the rapid inward transport of aminophospholipids. The observation of an energy-consuming transport of lipids triggered exciting but very challenging research on the identification and characterization of lipid transporters. Although many important details still have to be unraveled, today, we know that for the rapid inward transport of PS and PE in the plasma membrane of eukaryotic cells, a P-Type ATPase is responsible (see in particular Chapters 8–10). Meanwhile, several members of the ATP-binding cassette (ABC) transporters have been identified also to mediate transport of lipids at the expense of ATP, not only phospholipids but also other lipids, for example,

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sterols (see Chapters 11 and 17). It is important to mention that there are other functions of lipid transport besides generating/preserving lipid asymmetry. Lipids can be secreted to the extracellular space via transport across the membrane (see Chapters 5, 11, and 17). Another function of lipid transporter could be to generate bending of the membrane by creating a difference of the surface area between the two monolayers of the membrane (see Chapters 2, 12, and 13). Finally, essentially based on the use of spin-labeled and fluorescent analogs, the study of transbilayer motion of lipids showed that a fast, rather unspecific rapid transbilayer scrambling of phospholipids in the plasma membrane of mammalian cells can be triggered by activation of a putative scramblase (see Chapter 7). In biogenic membranes, flip-flop of lipids is typically mediated very fast by membrane proteins (flippases) mediating an energyindependent and nonspecific redistribution of lipids (see Chapter 6). 1.3â•… OVERVIEW ON ASSAYS FOR MEASURING DISTRIBUTION AND TRANSLOCATION OF LIPIDS ACROSS CELLULAR MEMBRANES Several prerequisites have to be matched by an assay to generate credible results. Of course, each assay has its limits, and the choice of an assay always depends on the information in which one is interested. That is, while an assay might be useful to detect the transbilayer distribution of endogenous lipids or lipid analogs in a cellular membrane, it may not be useful to measure the kinetics of transbilayer motion of lipids. Several reports and reviews have considered in detail which criteria have to be fulfilled in order to determine the transbilayer distribution and movement of (phospho)lipids [17–22] (see also Chapters 5 and 6). Here, we will only shortly summarize the criteria: (1) The assay has to recognize the lipid of interest on the surface of the membrane in a quantitative manner. (2) The approach must distinguish between lipid species located on one leaflet and those on the opposite leaflet of the membrane. Hence, the recognizing reagent or enzyme must not have access to both sides of the membrane; that is, it must be impermeable. (3) Exchange/ redistribution of lipids between both monolayers should not occur while the assay is performed. That is, the time required to assay a specific lipid quantitatively in a leaflet must be shorter in comparison to the characteristic time of lipid transbilayer movements. (4) The amount of lipid of interest should not change during the assay, for example, due to delivery of new lipids or removal of (already modified by the assay) lipids via endocytosis and exocytosis, respectively. (5) Treatment of lipids during the assay should not modify membranes leading to a perturbation and, hence, an enhanced transbilayer redistribution of lipids during assay. In Figure 1.1, most relevant assays for assessing transbilayer movement and/ or distribution of lipid analogs as well as of endogenous lipids are summarized schematically. We will now briefly describe the various assays, focusing first on

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3. 4.

I.

Figure 1.1.â•‡ Assays for the detection of lipid transbilayer distribution (modified from Pohl et al. [81]). (a) Chemical modification assays for endogenous lipids. Endogenous lipids present on the outer plasma membrane leaflet are typically modified on the level of the head group. Reagents frequently used for modification are trinitrobenzene sulfonic acid (TNBS, specific for PE) and fluorescamine. Assays can also be applied to endogenous lipids synthesized in the presence of radioactive precursors in the cell (see b.2). (b) Enzymatic assays for endogenous lipids. (b.1) Phospholipase A2 (enzyme) treatment converts phospholipids in the outer plasma membrane leaflet to lysolipid and fatty acid. Lipid products are then analyzed by chromatography and can be compared with samples untreated with enzyme. An analogous technique is used for SM, employing sphingomyelinase. (b.2) Enzymatic assays have also been applied to assess transbilayer organization of endogenous lipids that are synthesized in the presence of radioactive precursors (marked with asterisks) in the cell (I) and localize to various cellular membranes, that is, also to the cytoplasmic leaflet of the plasma membrane, due to vesicular or monomeric transport (II), and move to the extracellular leaflet of the plasma membrane, for example, due the presence of transporter proteins (III). Upon appearance on the outer membrane surface, lipids are converted by enzyme treatment (enzyme) (see b.1).

TRANSBILAYER DISTRIBUTION OF ENDOGENOUS LIPIDS

9

Figure 1.1.â•‡ (Caption Continued) (c) Antibody-, peptide-, or protein-binding assay for endogenous lipids. Specific antibodies, peptides (e.g., Ro09-198, binding to PE), or proteins (e.g., Annexin V, binding to PS) with a high affinity for a particular lipid head group bind to endogenous lipids present on the outer plasma membrane leaflet. The amount of bound antibody/peptide/protein is quantified. (d) Albumin-extraction, dithionite, and ascorbate assays for fluorescent and spin-labeled lipid analogs. (d.1) At time tâ•¯=â•¯0, the outer (accessible) leaflet of the membrane is labeled with short-chain spinlabeled or fluorescent analogs. Analogs redistribute to the inner leaflet of the membrane leaflet by passive flip-flop or active transport (tâ•¯>â•¯0). To assess transbilayer distribution, analogs are extracted from the outer leaflet (Ext), for example, phospholipids by albumin or cholesterol by methyl-β-cyclodextrin, followed by separation of cells and media. By comparing the extracted amount of analogs with that of analogs remaining in the membrane, the transbilayer distribution can be estimated. If aliquots of the sample are investigated at different time points after labeling, the transbilayer movement of the analog can be assessed. Alternatively to albumin extraction, fluorescence of lipid analogs on the outer leaflet can be quenched using dithionite, or the spin-label signal can be reduced using ascorbate (Red). (d.2) The short-chain lipid analog precursor integrates into the outer membrane leaflet (1), crosses the plasma membrane (e.g., by passive flip-flop) (2), and distributes to different intracellular membranes (e.g., by monomeric transport) (3). Enzymes of the endoplasmic reticulum or Golgi convert part of the lipid analog precursor to the lipid analog of interest (4), which can distribute back to the cytoplasmic leaflet of the plasma membrane, where it becomes available to outward transport by transporter proteins (5). Upon appearance on the outer leaflet, lipid analog is extracted (Ext) or reduced (Red) (see d.1). Color version on the Wiley web site.

endogenous lipids and subsequently on lipid analogs. Note that assays based on exogenous but nonlabeled lipids are introduced in Chapter 2. 1.4â•… MAIN TECHNIQUES USED TO DETERMINE TRANSBILAYER DISTRIBUTION OF ENDOGENOUS LIPIDS IN CELL MEMBRANES 1.4.1â•… Chemical Labeling The first technique used to investigate the localization of phospholipids in the plasma membrane of a eukaryotic cell, the human red cell membrane, was carried out by Mark Bretscher in England in the early 1970s [23]. It was based on the chemical labeling of aminophospholipids by an NH2 reactive reagent (Fig. 1.1a; see also Chapter 6). Typical NH2 reactive reagents are trinitrobenzene sulfonic acid (TNBS) and fluorescamine. To match the condition that reagents do not permeate to the opposite membrane leaflet, those experiments are usually performed at lower temperatures, between 4 and 10°C. Bretscher demonstrated that PS and PE could react only if the cell membrane was made permeable to those essentially nonpermeable reagents. He was the first to claim and to demonstrate the asymmetrical organization of lipids in the human red cell membrane with the aminophospholipids PS and PE located principally

10


in the inner leaflet of the plasma membrane, while the choline-containing phospholipids PC and SM were principally in the outer monolayer. The bilayer concept had already been suggested a long time before by Gorter and Grendel in 1925 [24], who came to that conclusion by an evaluation of the area covered by the erythrocyte lipids. However, they could not infer the lipid asymmetry because they assumed that all lipids were identical and corresponded to phosphatidylglycerol (PG). 1.4.2â•… Enzymatic Treatment Almost at the same time when Bretscher published his results with chemical labeling of aminophospholipids, the Dutch group of Laurence Van Deenen in Utrecht [25, 26] developed a completely different technique to study the transmembrane distribution of phospholipids in the plasma membrane of eukaryotes, based on lipid degradation by phospholipases and sphingomyelinases (Fig. 1.1b; see also Chapter 6). For example, phospholipase A2 treatment by addition of the enzyme to cells converts phospholipids in the outer plasma membrane leaflet to lysolipids and fatty acids. This technique seems a priori hazardous since there may be slow destruction of the membrane under investigation; analysis of the lipid composition in the time course of the assay proves that the external leaflet was indeed perturbed. Yet the results found by this assay are similar to those obtained by chemical labeling of lipids [23], namely, a preferential location of aminophospholipids in the inner leaflet of the plasma membrane. A similar distribution was also found afterward in other cells of the blood circulation such as platelets and, in fact, in all eukaryotic cells investigated with similar techniques. Small variations could be reported for different animals such as ruminants, which have essentially no PC (but a larger fraction of SM). More data corresponding to different eukaryotic cells including plant cells can be found in Chapter 3 of this book (see Table 3.1). Another, very specific enzymatic treatment of phospholipid head group is the use of a decarboxylase, which transforms PS into PE [27, 28]. These approaches can also be applied to radioactively labeled phospholipids, allowing a more sensitive quantification of reaction products. It is very reasonable that radioactive labeling does not affect the behavior of lipids with respect to endogenous, nonlabeled lipids. Labeling of membranes with radioactively labeled phospholipids with two long fatty acid chains can be achieved by lipid transfer proteins [29–31]. However, labeling of the membrane by this technique is a rather slow process. While the approach should be useful to study the stationary distribution of lipids across the membrane, it might be difficult to resolve fast transbilayer movement of lipids. An interesting variant is the measurement of cell surface exposure of lipids that have been labeled radioactively intracellularly. Kälin et al. incubated red blood cells with [14C]-labeled fatty acids, which—after uptake—were incorporated into lyso-PC via two enzymatic steps [32] (Fig. 1.1b). Subsequently,

TRANSBILAYER DISTRIBUTION OF ENDOGENOUS LIPIDS

11

exposure of [14C]PC on the cell surface was assessed by hydrolysis via phospholipase A2 (see above). Results were in agreement with a complementary assay based on the exchange of surface-exposed PC (including [14C]PC) on PC from added liposomes by a lipid transfer protein [32]. The enzymatic approach has also been applied to lipids other than phospholipids. The transbilayer distribution of cholesterol was assessed using cholesterol oxidase [33–35] (see Chapter 5). Several enzymes and chemical modifications are available to specifically detect the transbilayer organization of glycosphingolipids (see Reference 20; Chapter 4). 1.4.3â•… Protein-Binding Assay for Endogenous Lipids Peptides or proteins that bind noncovalently to phospholipids by recognizing specific parts of their head groups provide noninvasive approaches to assess exposure of endogenous lipids on the membrane surface without modification of lipids (Fig. 1.1c). In particular, fluorescent variants of these molecules allow detection of binding by fluorescence microscopy and—as very often used—by flow cytometry analysis in a rather easy way. The most prominent example is the specific binding of Annexin V to PS, which was first employed by Thiagarajan and Tait [36]. Using Annexin V, they could demonstrate the exposure of endogenous PS on the cell surface of activated platelets (see also Chapter 7). Since then, it has found numerous applications (see Chapters 7, 15, and 16). For example, Annexin V binding is used for the detection of PS exposure as an early event of cell apoptosis [37, 38] (see Chapter 15). While binding of Annexin V to PS requires the presence of Ca2+, the cell adhesion glycoprotein lactadherin recognizes PS on cell surfaces without this cation [39, 40]. Lactadherin, also known as milk fat globuleepidermal growth factor (EGF) 8 (MFG-E8), is secreted by macrophages for mediating engulfment of PS-expressing apoptotic cells. Since the activity of prothrombinase essential for blood coagulation is dependent on binding to PS on the cell surface of thrombocytes, this has been used to assay exposure of PS on the membranes [41, 42]. However, interpretation of protein binding to membranes must always consider affinity to other lipids, for example, in case of Annexin V to other negatively charged lipids. Cinnamycin Ro09-0198 is a cyclic peptide that has been isolated from Streptoverticillium griseoverticillatum. This peptide specifically recognizes the head group of PE [43] and forms an equimolar complex with PE on biological membranes. To use the peptide as a probe for analyzing the surface exposure of PE, the peptide has been fluorescently labeled, preserving its reactivity and specificity [44]. For example, it has been applied to assess the exposure of PE on the cell surface of yeast cells [45]. However, one has to be aware that this peptide is able to induce transbilayer lipid movement, as has been shown for HeLa cells as well as model membranes [46]. To initiate flip-flop by Ro09-0198, the presence of PE is required.

12


1.5â•… MAIN TECHNIQUES USED TO DETERMINE TRANSBILAYER DISTRIBUTION OF LIPID ANALOGS IN CELL MEMBRANES 1.5.1â•… Spin-Labeled and Fluorescent Lipid Analogs 1.5.1.1 Early Studies Using Spin-Labeled Analogs The spin-labeling technique was invented by Harden McConnell, a professor of chemistry in Stanford University, in the early 1970s. (Roger Kornberg, one of his students, went on to win a Nobel Prize in 2006, on a very different subject.) In 1971 Kornberg and McConnell published a paper [1] on the slow spontaneous lipid diffusion from one leaflet to the opposite (flip-flop) in sonicated phospholipids vesicles using a modified PC (dipalmitoyl phosphatidylcholine [DPPC]), which contains in the head group a nitroxide ring (TEMPO) replacing a single CH2. The transbilayer redistribution was measured by selective reduction of spin-labeled lipids on the outer leaflet (see below). The half-time of diffusion measured with this probe was of the order of 6 hours at 30°C [1]. Because there was no value known at the time, this number was very important to obtain. However, one could guess that TEMPO is likely to slow down the transmembrane diffusion of the lipid analog because of the size of the paramagnetic moiety and of the polar character of the probe itself. It was only in 2005 that Liu and Conboy (see below) proved by sum frequency vibrational spectroscopy (SFVS) that the TEMPO–DPPC flip-flop is indeed one order of magnitude slower compared with pure DPPC [47]. The SFVS technique is rather sophisticated but has the advantage of measuring the transmembrane diffusion of nonlabeled lipids (if one admits that deuterated lipids are indeed perfect representatives of natural lipids). The value measured by Liu and Conboy is certainly important to know, but the information obtained by the McConnell laboratory in 1971 (34 years before!) was nevertheless an extremely useful hallmark. 1.5.1.2 Spin-Labeled and Fluorescent Analogs with a Short Fatty Acid Chain The spin-labeled TEMPO–DPPC used by Kornberg and McConnell is not soluble in water because of the long chains and must be added before vesicles are formed, for example, by sonication. To label biological membranes, they have to be fused with those vesicles, or they have to be incorporated with phospholipid transfer proteins. This is a limitation that hampers the use of such probe with natural membranes even with plasma membranes like erythrocyte cell membrane. To overcome this problem, in the early 1980s, fluorescent and spin-labeled lipid analogs, in particular, phospholipid analogs, were developed, which are slightly water soluble due to a short fatty acid chain replacing one of the natural long chains—typically in the sn2 position (Fig. 1.2) [2, 15, 48, 49]. The short chain usually has 5 or 6 or sometimes up to 12 carbon atoms. They actually form in water micelles and monomers, which is an essential property for efficient and rapid labeling of intact/preformed membranes. A mere addition to a suspension or monolayer of cells is sufficient to label the plasma mem-

13

TRANSBILAYER DISTRIBUTION OF LIPID ANALOGS (a)

(b)

H

H

O

P

P O

N

O O

O

O

R-N

NBD

N

N O

(c)

(d)

N

+

N

O O P – O O

+

O O P – O

O

O

O

O

O

O O

O

O

+

–

N -O

O N N O

+

N O N – O

Figure 1.2.â•‡ Structure of short-chain spin-labeled (a,c) and fluorescent (b,d) phospholipid analogs. Principal structure of spin-labeled (a) and fluorescent (b, NBD moiety) analogs. Chemical structures of spin-labeled (c) and NBD-labeled (d) phosphatidylcholine analogs. The label moiety is attached to the sn2 chain. P, phosphate group; H, head group. Color version on the Wiley web site.

brane because the monomers will incorporate spontaneously. This process could be very rapid. For example, spin-labeled analogs insert within a few seconds into the outer layer of the plasma membrane [18, 19]. As a dynamic equilibrium between micelles and monomers exists, insertion of monomers into the membrane leads to a depletion and finally disappearance of micelles and monomers from the medium. Since the amount of analogs used corresponds typically to 1â•›molâ•›% or even less of endogenous lipids, in many cases,

14


almost all analogs incorporate into the membrane. However, this has always to be checked carefully. Of course, analogs are also very useful to assess the transbilayer distribution of organelles and reconstituted systems (see, e.g., Chapters 6, 8, and 9). The reporter moiety should be in principle as small as possible to avoid or minimize steric perturbations. Originally, most studies based on fluorescent analogs used lipids with a short chain of six carbon atoms to which terminally the fluorescent group 7-nitrobenz-2-oxa-1,3-diazol-4-yl (NBD) is attached (C6-NBD analogs). Later on, C5-BODIPY analogs were introduced, which have several advantages such as the BODIPY moiety is integrated into the lipid structure along the fatty acid chain, they are more apolar, and they have better fluorescence properties [50]. It has to be underlined that not only the fluorescent moieties but also the spin-labeled groups are of polar nature that may perturb the normal hydrophobic environment of a lipid chain. As a consequence, a looping of the fatty acid chain to which the reporter moiety (NBD) is attached (usually the short chain) to the polar interface of the membrane is facilitated [51, 52]. The possibility to attach the probe on the polar head group exists, but it raises a new problem: The analog may not be recognized by a membrane protein that is supposed to interact selectively with a lipid. In particular, a lipid has to be recognized and transported by a lipid transporter (flippase), which is usually very specific for one or a few substrates. So in practice, the reporter moieties, in particular, fluorescent groups that are generally rather big, can cause artifacts. In other words, a test of biological activity has to be done before using those probes to verify that the degree of perturbation, which is introduced by the probe, is acceptable. Alternatively, observations obtained by using lipid analogs should be verified by assessing the behavior of endogenous lipids. Indeed, there are multiple examples proving that, in particular, fluorescent probe can cause serious perturbations. Examples of artifacts due to size and polarity of fluorescent probes are given in the articles by Devaux and collaborators [22, 53]. Another important issue of those analogs is their short-chain fatty acids. Early studies have shown that the fatty acid chain may have a significant influence on lipid translocation. In 1986, Middelkoop and collaborators showed that by exposing red cell membranes to exogenous phospholipases A2, phospholipids with at least one unsaturated chain experience a more rapid flip-flop than saturated lipids [54]. Half-times varied from 26.3 to 2.9 hours for 1,2 dipalmitoyl-PC and 1-palmitoyl-2-linenoyl-PC, respectively. Recently, various cholesterol analogs have been studied showing large variations in the potential to mimic endogenous cholesterol [55] (for structure of spin-labeled and fluorescent sterol/cholesterol analogs, see Chapter 5). 1.5.1.3 Assessing Transbilayer Distribution and Movement of Spin-Labeled and Fluorescent Lipid Analogs In principle, there are two ways to assess the transbilayer distribution and movement of lipid analogs, either by chemical reduction of the label moieties on the outer/accessible leaflet of the membrane

TRANSBILAYER DISTRIBUTION OF LIPID ANALOGS

15

by a nonpermeant agent or by extraction of analogs from the outer leaflet (Fig. 1.1d). In any case, treatment of analogs must be leaflet specific. The transbilayer distribution can be estimated from the amount of reduced/extracted analogs, that is, analogs of the outer/accessible leaflet, and the amount of nonmodified analogs, that is, analogs on the inner/nonaccessible leaflet. However, one has always to control the amount of total analogs in the membrane under investigation during the assay procedure, which in the ideal case should not alter (see below). To assess the kinetics of transbilayer distribution, aliquots of the labeled sample will be measured after different times of labeling. 1.5.1.3.1â•… Reduction Assayâ•… Spin labels and fluorescent NBD analogs are typically reduced by addition of ascorbate [1, 22] and dithionite [56–58], respectively (Fig. 1.1d). Both agents are, in principle, nonpermeable to membranes. Addition of the reducing agent to the suspension medium destroys the reporter moiety and, hence, the signal of labeled probes present on the outer monolayer. The remaining signal comes from probes that have flipped to the inner monolayer, which can be determined by electron paramagnetic resonance (EPR) or fluorescence spectroscopy. This allows one to determine the percentage of probes at time t that were exposed to the outer and inner monolayer. However, in case of biological membranes, and in particular at a higher temperature (37°C), reducing agents may cross the membrane reacting also with analogs on the intracellular leaflet. If so, reduction has to be performed at a low temperature, for example, 4°C. In any case, the nonpermeability of agents has to be controlled carefully (for more details, see Chapter 6). By measuring aliquots of the labeled sample at different time points after labeling the membrane, the transbilayer movement of lipid analogs can be assessed provided that the movement is significantly smaller in comparison to the time required to perform the assay, that is, reduction of analogs on the outer leaflet (for critical discussion, see Chapters 5 and 6). In principle, those assays are also applicable to analogs with two long fatty acid chains as long as the reporter moiety is accessible to reducing agent. This depends essentially on the localization of reporter moiety. If the moiety is deeply buried into the hydrophobic phase, access to them by polar reducing agents such as ascorbate or dithionite is strongly impaired. Hence, reduction may take much longer and may even be incomplete. Reporter moieties attached to short-chain fatty acids are known to loop back to the membrane surface because of its (partial) polar character (see above) becoming easily accessible to reducing agents. 1.5.1.3.2â•… Back-Exchange Assayâ•… Fatty acids as well as short-chain phospholipids including labeled analogs can be easily extracted from the outer/ accessible membrane leaflet by albumin, typically bovine serum albumin (Fig. 1.1d), while phospholipids with two long fatty acid chains, as is typical for endogenous ones, are not removed from the membrane by albumin [16, 48, 59, 60]. Similar as described for the reduction assay, the transbilayer distribution of short-chain phospholipids analogs can be measured at a given time t after

16


labeling the membrane by incubation of the labeled sample, for example, of cells or organelles, with albumin for a short time (on the order of 1–2 minutes or even less [see Reference 17]). During this incubation, all analogs from the outer, albumin-accessible leaflet are extracted. Upon subsequent rapid centrifugation of the samples, the amount of analogs in the supernatant corresponding to albumin-extracted analogs and in the sediment containing the sample with the analogs on the inner, albumin nonaccessible leaflet can be measured again via EPR or fluorescence spectroscopy. If the amount of labeled lipid analogs initially incorporated into the outer leaflet (=â•¯100% of analogs) remained constant during the whole procedure, it is sufficient to measure at different time points only one fraction, typically that in the supernatant. However, as often observed for biological samples, this is not the case. Apart from the fact that the analogs can be metabolically converted or degraded into other (lipid) molecules (see below), the reporter moiety could be destroyed as well. In particular, spin labels are rapidly reduced by intracellular redox systems as gluthathione [15, 61]. As analogs modified on the inner leaflet can also redistribute back to the outer leaflet, these lipids with a destroyed reporter moiety may occur also on the outer leaflet and will not be detected by the spectroscopical measurement. However, as shown for spin labels, those signals can be recovered by addition of appropriate reagents [16, 60]. The back-exchange assay can also be applied to assess the kinetics of the transbilayer movement. As in any case, the time resolution depends on the time to carry out the assay procedures. Usually, incubation of a sample with albumin and subsequent centrifugation with standard laboratory equipment takes at least 2 minutes; that is, any transbilayer movement on the order of 2 minutes or even faster cannot be resolved adequately. Figure 1.3 shows the transbilayer movement of spin-labeled phospholipids in red blood cells [62]. Those measurements have been performed with standard laboratory equipment providing a time resolution even sufficient to resolve the rapid inward movement of the spin-labeled PS at 37°C with a half-time of about 5 minutes. However, the inward redistribution of spin-labeled PS is much faster in the plasma membrane of sperm cells or osteoblasts with a half-time of ≤2 minutes [63, 64] (for reviews, see References 21 and 65–69). Here, the assay based on standard equipment is on its limit of time resolution. In 1986, Tilley et al. measured the transbilayer redistribution of long-chain radiolabeled phospholipids inserted in the outer membrane leaflet of intact human erythrocytes with a nonspecific lipid transfer protein [8]. The transbilayer mobility and equilibrium distribution of the radiolabeled phospholipids were assessed by treatment of the cells with phospholipase A2. These experiments confirmed the selective ATP-dependent transport of aminophospholipids toward the inner membrane leaflet. Because probe insertion with a phospholipid exchange protein required at least 30 minutes incubation, and because cells and phospholipases also had to be incubated, no real kinetics could be drawn by this method. Nevertheless, partial kinetics data obtained with long-chain phospholipids using this technique [8] were consistent

17

TRANSBILAYER DISTRIBUTION OF LIPID ANALOGS 100

SL-PS SL-PE

SL analogs inside (%)

80

60

40 SL-PC 20 SL-SM 0 0

3

6

9

12

15

18

Time (hours)

Figure 1.3.â•‡ Kinetics of the redistribution of spin-labeled phospholipid analogs in human red cells at 37°C. Analogs were incorporated in the plasma membrane outer monolayer of those cells at tâ•¯=â•¯0 and their redistribution was followed by the backexchange assay. The final transbilayer distributions derived from the plateaus of the curves are in fact identical to the equilibrium distribution of endogenous phospholipids in those cells (see Chapters 3 and 10). Modified from Reference [62]. Color version on the Wiley web site.

with the more detailed results obtained with spin-labeled and fluorescent phospholipids. Buton et al. succeeded in optimizing the above-described albumin backexchange procedure to be performed in about 30 seconds [17]. Thus, they could resolve much better the fast flip-flop of spin-labeled analogs in organelles such as the endoplasmic reticulum or the Golgi. A much better time resolution of the back-exchange assay can be obtained by employing the stopped-flow technique [18, 19] (see Chapter 6). At this point, it might be interesting to compare spin-labeled and fluorescent analogs with respect to transbilayer distribution and movement. Again, although red blood cells are “simple cells,” experiments on them nicely illustrate the differences between analogs. The short-chain fluorescent C6-NBD-PS analog redistributes much slower from the exoplasmic to the cytoplasmic leaflet in comparison with the short-chain spin-labeled PS analog [53, 62]. Furthermore, the asymmetric distribution of C6-NBD-PS is less pronounced than that observed for spin-labeled PS (see above) and endogenous PS (see Chapter 3). Hence, although still transported via the aminophospholipid translocase, the NBD moiety affects the recognition and/or transport of the analog. Interestingly, while spin-labeled PE is efficiently transported to the cytoplasmic leaflet, C6-NBD-PE is almost not transported, indicating that the fluorescent analog is a very poor substrate for the aminophospholipid translocase

18


[53]. Very likely, the difference between both types of analogs can be explained by the NBD group being more bulky in comparison with the spin-label moiety. Similar differences between spin-labeled and fluorescent PS analogs were observed for the plasma membrane of other mammalian cells, for example, for fibroblasts [49, 70], sperm cells [63, 71], and hepatocytes [72, 73]. Back-exchange assay has also been performed by using phospholipid vesicles as a donor of phospholipid analogs instead of albumin [74–77]. Usually, studies have been done using fluorescent analogs. Exchange of the fluorescent analog under study between acceptor and donor membranes can be followed by Förster resonance energy transfer to a stably anchored fluorescent lipid in the donor vesicles (see also Chapter 6). While albumin is able to extract short-chain phospholipids and fatty acids, it does not extract cholesterol and sterol (analogs). However, methyl-βcyclodextrin (MβCD) can efficiently remove cholesterol from membranes. Indeed, extraction by MβCD has been used to assess the transbilayer dynamics of cholesterol/sterol analogs. As outlined in Chapter 5, several (additional) limitations of MβCD-mediated removal of cholesterol analogs have to be taken into account. For example, those analogs in many cases barely reflect the behavior of endogenous cholesterol. A major issue is that MβCD removes not only cholesterol analogs but also endogenous cholesterol. Hence, in this case, the composition of the membrane is continuously altered. For more detailed information, see Chapter 5. 1.5.1.3.3â•… Consequences of Intracellular Trafficking of Analogs for Assaysâ•… Initially, phospholipid analogs have been applied successfully to characterize transbilayer distribution and movement of lipids in the plasma membrane of human red blood cells. Although one has to take into account hydrolysis of analogs during experiment, the absence of any endo- and exocytotic activity and of intracellular membranes in red blood cells was very favorable for applied assays. Indeed, a drawback in assessing transbilayer distribution of spin-labeled or fluorescence analogs in the plasma membrane of eukaryotic cells is the removal of analogs by endocytosis and by intracellular redistribution of analogs to organelle membranes [49, 64, 70, 78]. Hence, the amount of analogs in the membrane of interest may not be constant during the assay procedure, violating a criterion given above. Figure 1.4 illustrates these disadvantages. In Figure 1.4a, the endocytic uptake of fluorescent C6-NBD-PC into osteoblasts at 37°C is shown [64]. As known for PC, movement from the exoto the cytoplasmic leaflet is slow. However, at the chosen temperature, endocytosis is a significant process, causing the disappearance of a significant amount if not most of the analog from the plasma membrane as visualized by the fluorescent intracellular spots corresponding to endosomal structures. In contrast, the fluorescent PS analog, which is transported rapidly from the exoto the cytoplasmic leaflet by the aminophospholipid translocase activity, disappears from the cytoplasmic leaflet and, thereby, from the plasma membrane to the cytoplasm due to the partial water solubility of analogs. Finally, it inserts

TRANSBILAYER DISTRIBUTION OF LIPID ANALOGS

19

(a)

(b)

Figure 1.4.â•‡ Uptake of fluorescent lipid analogs into osteoblasts. (a) Intracellular uptake of C6-NBD-PC occurs essentially via endocytosis. The yellow dots correspond to vesicles endocytosed; (b) C6-NBD-PS is transported by the aminophospholipid translocase to the ctyoplasmic leaflet of the plasma membrane. Once the probe is exposed to the cytosol, it can redistribute to other intracellular membranes because of its comparatively large solubility in water. See also Reference 64. Labeling was performed by incubation of analogs with cells. Upon incubation, noninserted analogs as well as analogs on the outer leaflet were removed by washing with albumin (see text).

into intracellular membranes, which leads to a bright intracellular staining (Fig. 1.4b). Due to the very rapid uptake of the PS analog by inward transport, the amount of PS taken up by endocytosis is low. In addition, biochemical modifications—apart from hydrolysis—can reduce the amount of lipid analogs in the membrane during the assay procedure (see Section 1.5.2).

20


1.5.2â•… Biosynthetic Labeling Nevertheless, the disadvantage of uptake and intracellular redistribution of (short-chain) analogs when assessing transbilayer organization of lipid analogs in the plasma membrane could be advantageous for biosynthetic labeling of lipids. The idea is that a labeled lipid analog, typically a fluorescent one, is taken up by the cell and, by metabolic processes, converted into another lipid analog (Fig. 1.1d). The trafficking of the latter, in particular, its exposure on the cell surface, is then followed, for example, by back-exchange. Thus, to examine outward transport of short-chain lipid analogs, for example, C6-NBDPC, by the multidrug resistance ABC transporter MDR1 Pgp expressed in the plasma membrane, cells were incubated with C6-NBD-PA. This lipid analog is partially converted into C6-NBD-diacylglycerol at the plasma membrane, which rapidly crosses the plasma membrane and becomes available for intracellular synthesis of not only C6-NBD-PC but also C6-NBD-PE [79–81]. The time-dependent appearance of the fluorescent phospholipids in the extracellular membrane surface was followed by incubating the cells with albumin and measuring the fluorescence of the supernatant after centrifugation. A similar approach has been applied to measure sphingolipid translocation. Short-chain analogs of ceramide, for example, C6-NBD-ceramide, were incorporated into the extracellular leaflet of the plasma membrane. These analogs flipped rapidly across the plasma membrane and—upon intracellular redistribution from the cytoplasmic leaflet—were converted to fluorescent SM and glucosylceramide (GlcCer) by enzymes of the Golgi apparatus. Exposure of these intracellularly synthesized fluorescent lipids on the cell surface was measured by the backexchange assay [20, 79, 82]. Those studies have shown that, for example, MDR1 Pgp is capable of transporting various short-chain GlcCer analogs to the extracellular leaflet of the plasma membrane, while the multidrug resistance transporter MRP1 was rather selectively transporting only C6-NBD-GlcCer [79, 82]. For a detailed description of transbilayer asymmetry and dynamics of glycolipids, see Chapter 4. 1.5.3â•… SFVS To circumvent labeling of lipids with bulky moieties, Liu and Conboy [83] introduced SFVS for measuring transbilayer lipid movement. This coherent nonlinear optical vibrational technique, which has been described in detail [84], takes advantage of the selectivity of infrared (IR) and Raman spectroscopy. Excitation induces dipole oscillations of molecules, for example, of C–H bonds. As molecules behave like an aharmonic oscillator, overtone oscillations are excited, including the second harmonic, which is used here. Since a secondorder nonlinear optical process is forbidden in media of inversion symmetry but not on surfaces with broken symmetry, this technique is surface specific in nature, which—under certain conditions (see below)—makes it also suitable for assessing transbilayer distribution and motion of lipids. Experimentally, the

References

21

excitation is read out by overlapping spatially and temporally a visible and a tunable IR laser on a surface (e.g., membrane surface). As a result, a signal (photon) is generated at the sum of their frequencies, which increases on resonance with a vibrational transition. Conboy and coworkers generated asymmetric lipid bilayers: One of the two monolayers contained phospholipids with a deuterated terminal CH3 group. Hence, the transition dipole moments of the C–H vibrational modes that are oriented antiparallel along the C–C bond are different between the two leaflets and do not cancel out, which can be monitored by SFVS. The difference between the transition dipole and therefore the signal becomes reduced and, finally, disappears upon transbilayer redistribution of phospholipids. Using this approach, the flip-flop of various phospholipids in bilayers of synthetic phospholipids has been measured [47, 83, 85, 86]. For example, a decrease of the fatty acid chain length of PC resulting in a significant increase in the flip-flop rate has been shown. Moreover, it was found that spin labeling can substantially affect the transbilayer dynamics of phospholipids (see above). Although the approach requires deuterated lipids, this modification is small in comparison with tagging of lipids by spin or fluorescent labels, and the behavior of natural lipids should be preserved. However, since the technique requires the assembly of asymmetric bilayers, it is restricted to model membranes and not applicable to native, biological membranes. Nevertheless, peptides (transmembrane domains) [87, 88] and membrane proteins can be reconstituted in asymmetric model membranes to study their influence on lipid transbilayer movements.

ABBREVIATIONS MβCD NBD PC PE PG PS SM

methyl-β-cyclodextrin 7-nitrobenz-2-oxa-1,3-diazol-4-yl phosphatidylcholine phosphatidylethanolamine phosphatidylglycerol phosphatidylserine sphingomyelin

REFERENCES â•‡ 1â•… R. D. Kornberg, H. M. McConnell, Biochemistry 1971, 10, 1111–1120. â•‡ 2â•… M. Seigneuret, P. F. Devaux, Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 3751–3755. â•‡ 3â•… J. Connor, A. J. Schroit, Biochemistry 1987, 26, 5099–5105.

22


â•‡ 4â•… â•‡ 5â•… â•‡ 6â•… â•‡ 7â•… â•‡ 8â•…

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â•‡ 9â•… 10â•… 11â•… 12â•… 13â•… 14â•… 15â•… 16â•… 17â•… 18â•… 19â•… 20â•…

21â•… 22â•… 23â•… 24â•… 25â•… 26â•… 27â•… 28â•… 29â•… 30â•… 31â•… 32â•… 33â•… 34â•…

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2 DETECTION AND MEASUREMENT OF UNLABELED LIPID TRANSMEMBRANE MOVEMENT Iván López-Montero Departamento de Química Física I, Universidad Complutense de Madrid, Madrid, Spain

Marisela Vélez Consejo Superior de Investigaciones Científicas, Instituto de Catálisis y Petroleoquímica, Madrid, Spain IMDEA Nanociencia, Facultad de Ciencias, Universidad Autonóma de Madrid, Madrid, Spain

Philippe F. Devaux Institut de Biologie Physico-Chimique, Paris, France

2.1â•… INTRODUCTION Since the pioneering work of Kornberg and McConnell in 1971 [1], the measurement of transverse diffusion of lipids is associated with the use of lipid analogs. A large variety of labeled lipids has been developed mainly for electron spin resonance and fluorescence spectroscopy (see Chapters 1 and 6). These techniques require the use of lipid probes, which bear a nitroxide or a fluorescent group generally in one of their acyl chains. For the study of the spontaneous flip-flop on model lipid bilayers (as large unilamellar vesicles [LUVs]), the (spin or fluorescent)-labeled lipids are easily incorporated from


25

26

FLIP-FLOP OF UNLABELED LIPIDS

the organic solvent during sample preparation. However, the study of transmembrane diffusion (spontaneous or protein mediated) in cell membranes requires the use of water-soluble probes in order to facilitate their incorporation into the biomembrane from solution. The way to do this is to attach the probe to a modified lipid having a short fatty acid chain. This molecule, which is partly water soluble, spontaneously introduces itself into the membrane carrying the probe along. In addition, the short fatty acid chain enables the selective extraction of the probe from the external leaflet with the backexchange method (see Chapters 1 and 6). Once the probe is extracted, the inward transport of labeled lipid is then determined spectroscopically [2]. Likewise, the outward transmembrane lipid asymmetry can also be easily assessed in situ by chemical reduction of the probes exposed on the outer leaflet. The classical techniques of investigation with labeled lipids [1, 2] have proven to give at least reliable information on comparative values of the flipflop half-times (τ1/2’s) of different phospholipids [3]. However, the main objection to this experimental approximation comes from the possible divergences in the values of τ1/2, which may differ slightly from those corresponding to the endogenous lipid. The presence of the label involves a steric hindrance within the bilayer and could have a strong effect on the lipid–protein interactions involved in active transport carried by flippases. Methods suitable for the quantification of the transmembrane distribution of unlabeled lipids were originally based on exogenous phospholipase attack of the cell surface [4]. The precision of this technique is limited, and in situ lipid degradation can induce artifacts. Another approach consists of using radioactively labeled lipids. However, since these measurements of transmembrane motion are restricted to short-chain lipids and demand long and tedious experiments, studies using these molecular probes are relatively scarce [5]. Very recently, efforts have been made to develop new methodological approaches able to detect and to measure the transbilayer movement of unlabeled and long-chain lipids. First, the transbilayer diffusion of unlabeled molecules has been followed from the shape changes of giant unilamellar vesicles (GUVs) associated with the lipid reorganization [6, 7] and quantified considering the elastic and mechanical properties of lipid bilayers. A second methodology used to measure lipid flip-flop without requiring lipid probes is atomic force microscopy (AFM) [8, 9]. The ability of this technique to image with nanometer resolution enables the monitoring of the time evolution of lipid domains on supported bilayers caused by a lipid exchange between monolayers. Third, sum frequency vibrational spectroscopy has been applied to measure the translocation of unlabeled lipids on lipid bilayers [10, 11]. And very recently, time-resolved, small-angle neutron scattering technique has been applied to vesicle systems to determine the interparticle transfer and the flip-flop of unlabeled phospholipids [12]. In this chapter, we review the potential and the most relevant results obtained with the shape change approach on GUVs (see Section 2.2) and with AFM of supported lipid bilayers (see Section 2.3).

Measurement of Transmembrane Flip-Flop of Unlabeled Lipids

27

2.2â•… MEASUREMENT OF TRANSMEMBRANE FLIP-FLOP OF UNLABELED LIPIDS BY SHAPE CHANGE OF GUVs The shape of vesicles is very sensitive to the relative area between the two monolayers. This is the main principle on which the shape change method performed on GUVs to detect and measure lipid flip-flop is based. We start this section with a brief exposition of the bending theory of vesicle shapes. Some examples of shape changes caused by an asymmetric lipid distribution reported in the literature are then presented. A more detailed description of the shape change approach to measure unlabeled lipid flip-flop follows, and a few experimental results are then described. We conclude with a few remarks on the perspectives of new experiments related to energy-dependent flippases. 2.2.1â•… Bending Theory of Vesicle Shapes Lipid bilayers are elastic objects able to support mechanical deformations as stretching, compression, and bending. The mechanical response of the bilayer against lateral stretching or bending is governed by the material parameters, the stretching modulus ε, and the bending modulus κb, respectively. Stretching a lipid bilayer is energetically more expensive (εâ•¯∼â•¯200â•›mN/m) than bending it, which can be achieved by thermal energy (κbâ•¯∼â•¯10â•›kbT). Thus, in the frame of a morphological theory of lipid vesicles, surface area can be considered as fixed. A symmetrical bilayer would adopt a flat conformation in the absence of edge energy effects. Conversely, asymmetrical bilayers (with a different chemical composition or lipid density between monolayers) can adopt more curved shapes characterized by a spontaneous curvature C0, the preferred curvature of the lipid bilayer at equilibrium. Deviations from the preferred curvature have an energetic cost, expressed by the Hookean expression:

1 2 Eb = κ b dA (C − C0 ) . 2

∫

(2.1)

Equation 2.1 was first introduced by Helfrich [13]. A large variety of vesicular shapes were predicted by minimizing Equation 2.1 under different conditions of osmolarity and surface/volume ratio. However, the early work of Helfrich missed the bilayer nature of membranes and considered the vesicles just as two-dimensional elastic sheets. The second model, derived from Helfrich’s ideas, was implemented some years later [14] with the bilayer architecture of membranes. Indeed, when a closed bilayer is bent, an additional contribution to the pure bending mode (the bending of two monolayers as a fixed area) must be taken into account, that is, the relative stretching and compression of monolayers. This leads to enable changes in the differential area between monolayers given by ΔAâ•¯=â•¯Aoutâ•¯−â•¯Ain, where A represents the surface area of the outer (out) and inner (in) monolayers, respectively. Mathematically, the total energy reads,

28


1 1 ακπ 2 Eb = κ b dA (C − C0 ) + (∆A − ∆A0 )2, 2 2 Ah 2

∫

(2.2)

where ∆A0 = A0out − A0in is the difference of the unstressed monolayer areas and κ is the nonlocal bending modulus. h is the bilayer thickness and α accounts for the relative importance of the two contributions to the total bending energy. α has been estimated to be of the order of one for all phospholipids. Equation 2.2 represents the area difference elasticity (ADE) model [14]. Again, the minimization of Equation 2.2, at given parameters of volume-toarea ratio and preferred differential area, leads to the same diversity of vesicular shapes, but new pear, budded, starfish vesicles or vesicles with holes are now also predicted. Vesicle shape regions of minimal energy can be represented into a diagram, which contains different vesicle classes. An ADE shape diagram is reproduced in Figure 2.1 from Reference 15. Shape changes of vesicles triggered by an asymmetric distribution of lipids constitute an experimental evidence of the bilayer structure included on the ADE model (see next section). This is equivalent to maintain constant the vesicle volume and to modify the preferred area difference along the vertical axis of the shape diagram. Lipid flip-flop represents a very simple mechanism to vary the number difference in the ADE contribution (Eq. 2.2). 2.2.2â•… Asymmetry and Shape Changes of Membranes: Examples Following the shape diagram shown in Figure 2.1, it is evident that one could cross the different shape regions by modifying the spontaneous curvature and the preferred differential area of vesicles. Any asymmetry across the membrane will lead to a change in the effective spontaneous curvature. A large variety of mechanisms for monolayer surface area asymmetry generation can be found in the literature [16–18]. Among all of them, an asymmetric distribution of lipids between leaflets has been extensively explored in cells as well as in model vesicular systems. The influence of the incorporation and redistribution of lipids on erythrocyte morphology was first studied by Daleke and Huestis [19]. Red blood cells show a biconcave (discocyte) shape at physiological conditions of pH and osmolarity. Their shapes are strongly modified when exogenous lipid molecules are added to the plasma membrane. A discocyte-to-echinocyte shape change is induced when phosphatidylserine (PS) and phosphatidylcholine (PC) are incorporated to the external monolayer. In addition, PS-induced echinocytes revert to discocytes and then become invaginate stomatocytes. This second stage is produced by PS translocation into the inner monolayer by the aminophospholipid translocase [20] (Fig. 2.2). An asymmetric amount of lipids between both monolayers can also be easily promoted on pure lipid unilamellar vesicles. More curved shapes (budded vesicles) can be obtained from a prolate giant vesicle by the addition of exogenous lyso-PC on the outer monolayer. Conversely, selective depletion of


29

2.5

2.0 Dpro/pear Lpear pears Cpro/pear 1.5

starfish prolates

∆a0

Cnas/obi

Dpro/nas Aobi 1.0

Csto/obi Dpro/obi Asto

oblates Dsto/obi

0.5 stomatocytes Lsto

0.0 0.0

0.2

0.4

0.6

0.8

1.0

v

Figure 2.1.â•‡ Phase diagram of the ADE model [15]. The horizontal axis is the volumeto-area ratio expressed by the reduced volume v (scaled to that of a sphere). The vertical axis shows the effective differential area expressed by Δa0 (scaled to that of a sphere). Along the vertical axis, vesicles can change their shape by modifying the surface area difference between monolayers. Reprinted from Current Opinion in Colloid & Interface Science, 5, Hans-Günther Döbereiner, Properties of giant vesicles, 256–263, Copyright (2000), with permission from Elsevier.

30


(a)

(b)

(c)

(d)

(e)

(f)

(g)

(h)

(i)

(j)

Figure 2.2.â•‡ Scanning electron micrographs of erythrocytes. Discocytes (f) change their shape to echinocytes (a–e) when lipids are accumulated in the external monolayer. When PS molecules are incorporated to the membrane, they are finally transported to the inner leaflet by the action of a flippase, producing a shape change to stomatocyte configurations (g–j). Reprinted from Biochemistry, 24, D.L. Daleke and W.H. Huestis, Incorporation and translocation of aminophospholipids in human erythrocytes, 54065416, Copyright © 1985, with permission from American Chemical Society.

symmetrically incorporated lyso-PC molecules, from the outer monolayer of vesicles by incubation with bovine serum albumin (BSA), a fatty acid transporter, makes vesicles to adopt invaginated shapes [21]. Finally, the redistribution of endogenous lipids constitutes another example of morphological changes caused by an accumulation of some negatively charged lipids (as phosphatidylglycerol [PG]) into one monolayer in response to transmembrane pH gradients. Again, the outward transbilayer transport of PG in LUVs made of a mixture of PC and PG [22] transforms initially invaginated LUVs to long narrow tubular structures, or spherical structures with one or more budded vesicles. On the other hand, when PG inward transport is induced, inverting the pH gradient, noninvaginated LUVs reverse to invaginated vesicular structures. All the experiments described above inspired the technique of shape changes performed on giant vesicles to quantify the transbilayer diffusion of unlabeled lipids presented in the next section. 2.2.3â•… Flip-Flop Detection by Shape Changes in Giant Vesicles Shape of GUVs can be easily modified by varying experimentally the parameters that describe their shape: the preferred area difference between monolayers (asymmetry) and the vesicular surface area/volume ratio (see Fig. 2.1). GUVs are very sensitive to the relative area between monolayers and the minimum asymmetry needed to induce a shape change is of the order of 0.1% of the total area of the vesicle [23, 24]. GUVs are consequently, in comparison with other membrane models, excellent objects to detect lipid asymmetry. They are visible under optical microscopy, which facilitates their observation and manipulation, and their internal volume can be modified. This is important


31

since the reduced volume must be lower than 1 (which corresponds to a sphere) in order to have a vesicular surface excess for allowing shape changes. The most extended method to fabricate giant vesicles is electroformation [25]. Giant vesicles obtained by this technique are unilamellar in a very high percentage but have spherical shape. Vesicles with lower reduced volumes can be obtained by allowing the external solution to evaporate for some minutes. Due to an osmotic pressure difference between inside and outside, spherical vesicles deflate and adopt a prolate shape. The “shape change” approach to infer the flip-flop rates (τ1/2) of nonlabeled lipids is based on the incorporation of a few molecules to the external membrane leaflet that increase its area and trigger a shape change from a prolate to a budded shape vesicle. If a transbilayer movement of lipids occurs, the prolate shape is recovered due to the lipid redistribution into both lipid monolayers. Monitoring the times it takes for the shape changes to occur allows one to calculate the τ1/2’s according to the theoretical model described in the next section. 2.2.4â•… Theoretical Model For quantitatively estimating the translocation rate of unlabeled molecules in shape changes experiments performed on GUVs, a very simple kinetic model can be used [6]. This model is based on the time evolution of the asymmetry function c, which is defined as the difference of the unlabeled lipid concentration between both monolayers:

c (t ) = cout (t ) − cin (t ) ,

(2.3)

where cout(t) and cin(t) are the unlabeled lipid concentrations (in percentage of total lipids) in the outer and the inner monolayer, respectively. Obviously, both the outer and inner concentrations must take into account simultaneously the superposition of the kinetics of exogenous nonlabeled lipid insertion in the outer monolayer and their diffusion toward the inner monolayer. As said before, the threshold for shape changes in GUVs is given by a lipid asymmetry of the order of 0.1% of the total area of the vesicle. This means that budding transition and shape recovery would occur if câ•¯=â•¯cthrsâ•¯=â•¯0.1%. In a first approximation, the time evolution of the two different monolayers must explicitly introduce the two rate constants for insertion and flip-flop and can be written as a differential equation system:

dcout = f (t ) − Kout cout + Kin cin , dt

(2.4)

dcin = Kout cout − Kin cin , dt

(2.5)

where f(t) accounts for an empirical function of lipid incorporation to the external monolayer. Kout and Kin are the translocation rates from the outer

32


to the inner monolayer and from the inner to the outer monolayer, respectively. The expression for f(t) is independently measured [6, 7] and depends on two main parameters: a characteristic time of incorporation and the final concentration of amphiphilic molecules incorporated in the outer monolayer, which is expressed in percentage of total lipids in the GUV. Both parameters depend on the nature of the injected molecules. Several approximations have been assumed in order to simplify Equations 2.4 and 2.5: • The threshold of asymmetry is rigorously defined by ∆S / S = 0.1 ⇒ cthrs = 0.1(avl ) /(aincl ) where avl is the area occupied by a vesicular lipid molecule and aincl is that of the incorporated molecule. Both molecular areas have been assumed to be the same, and therefore, cthrsâ•¯=â•¯0.1. Actually, the theoretical threshold for shape change established on 0.1% is a larger source of error. • The outer aqueous volume is very large compared with the volume enclosed by GUVs. Therefore, only a small fraction of the total added lipid will be incorporated into the vesicle, and the outer aqueous concentration of exogenous lipids can be considered as a constant. • The fraction of exogenous molecules in the inner monolayer, which spontaneously partitions into the internal volume of vesicle, is negligible because the internal volume of the vesicles is very small in comparison to the external volume. Likewise, the water–membrane exchange of the incorporated molecules to the outer monolayer has been neglected. For a GUV, both leaflets are equivalent (in terms of curvature and tension) and therefore, Koutâ•¯∼â•¯Kinâ•¯=â•¯K. By taking into account the above considerations, Equations 2.4 and 2.5 read

dc(t ) = f (t ) − 2 Kc(t ). dt

(2.6)

An analytical solution for c(t) can be obtained once f(t) is experimentally measured. c(t) results as a function that depends on two main kinetic parameters: the incorporation rate of exogenous molecules to the outer monolayer and the flip-flop rate K. The time behavior of the monolayer area asymmetry (Fig. 2.3) has a nonmonotonic character and starts from zero (there is no asymmetry at tâ•¯=â•¯0). The initial increase of c(t) corresponds to the incorporation of exogenous molecules to the external monolayer, whereas the decrease of c(t) with time results from re-equilibration of nonlabeled molecules between the external and the inner monolayer. The curve intercepts the threshold of shape change câ•¯=â•¯cthrsâ•¯=â•¯0.1% at t1 and t2, at the budding transition and the shape recovery, respectively. The experimental values of t1 and t2 allow one to fit the value of K and hence τ1/2â•¯=â•¯ln2 / K.


33

0.12 t1

t2

c (% of total lipid)

0.10

0.08

0.06

0.04

0.02

0.00

0

2

4

6

8

time (minutes)

Figure 2.3.â•‡ Experimental time evolution of the asymmetry function c for C6-ceramide (see Section 2.5) incorporated to a prolate shape egg-PC GUV. At times before t1, the asymmetry is lower than cthrs (solid horizontal line) and the vesicle shape remains as a prolate. When câ•¯>â•¯cthrs, the created asymmetry is high enough to induce the budding transition, which is reversed at t2 when ceramides are redistributed into both monolayers by spontaneous flip-flop. From t1 and t2 (dashed vertical lines), flip-flop half-times can be deduced.

2.2.5â•… Examples A good example for illustrating the use of the “shape change” methodology is the measure of the τ1/2 of unlabeled ceramides [6]. In those experiments, ceramides with different chain lengths (C6-, C10-, and C16-ceramides) were added externally with a micropipette to GUVs made of egg-PC. Figure 2.4 illustrates the shape change pathways followed by giant vesicles when ceramides are injected. Unlabeled ceramides trigger first a budding transition at t1. After a short period of time (t2), a sudden reopening process takes place (Fig. 2.4f), and the initial shape is recovered. Time t1, of the order of 1 minute, varies from one ceramide to another and is longer for longer acyl chains. Likewise, the reopening of the neck was observed 9–15 minutes after t1. From t1 and t2, the τ1/2 for ceramides was calculated to be of the order of 1 minute at 20°C (1.2, 1.6, and 2.4 minutes for C6-, C10-, and C16-ceramides, respectively). Shape change experiments can also be performed at different temperatures ranging from the melting temperature (Tm) of lipids, which conform the giant vesicle (in case Tm was lower than the melting temperature of water, 0°C would obviously be the lower limit temperature) to the highest temperature at which

34


(a)

(b)

(c)

(d)

(e)

(f)

(g)

(h)

(i)

(j)

(k)

(l)

Figure 2.4.â•‡ Shape changes induced by C6-ceramide externally added to an egg-PC giant vesicle. A budding transition is first formed when molecules are accumulated into the outer monolayer (a–e). The prolate shape is recovered when ceramides are redistributed into both monolayers by spontaneous flip-flop (f–l). Scale bar is 10â•›µm. From Reference 6.

shape changes are observable (in case that Kâ•¯>>â•¯incorporation rate, no shape change will be observed on vesicles). For ceramides, the spontaneous transbilayer half-times shifted to values smaller than 1 minute at 37°C (18 and 57 seconds for C6- and C16-ceramides, respectively). It is worth noting that the incorporation of lipids having a very slow transbilayer movement, as lyso-PC [7] or C6-PC, produces a stable budding transition (vesiculation) in GUVs made of PC, as already shown in the previous experiments commented in Section 2.2.2. In a second example [7] illustrating the potentiality of the shape change approximation to detect flip-flop of unlabeled lipids, the prolate shape produced by those lipids was reversed on GUVs prepared with membrane protein extracts from the endoplasmic reticulum (ER) of yeast (see Chapter 6). In eukaryotes, the cytoplasmic leaflet of the ER membrane is the major site of phospholipid biosynthesis. To ensure stable membrane growth, energyindependent flippases mediate rapid, bidirectional, and rather unspecific phospholipid flip-flop with half-times of minutes or less. When egg-lyso-PC or C6-PC was externally added to the reconstituted vesicles, a budding transition


35

occurred within the first 5 minutes, and subsequently, the vesicle regained the prolate shape within 5–10 minutes. Therefore, the reconstituted ER flippase activity triggers the rapid flip-flop that diminishes the surface area difference between the two leaflets (see Chapter 6). 2.2.6â•… Perspectives In comparison with other methods presented in this book, the main advantages of the “shape change” approach are the measurement of flip-flop of unlabeled molecules and long double acyl chain lipids. However, the low solubility in water of the latter molecules arises as the main shortcoming of the method. First, organic solvents are required to accelerate slow diffusion in water of long-chain lipids and moreover to facilitate their incorporation into vesicles. One could argue about possible artifacts associated to the use of organic solvents, but control experiments indicate that the same amount of organic solvent in the absence of the nonlabeled molecule does not trigger any budding transition. Second, a slow rate of incorporation relatively to flipflop disables the triggering of the budding transition. If all incorporated molecules are rapidly redistributed into both monolayers, the accumulation in the outer leaflet needed to induce changes is not reached. Finally, this method is limited to fluid giant vesicles. Indeed, giant vesicles in a liquid-ordered phase can be made of sphingomyelin (SM)/cholesterol (Chol) mixtures. The addition of molecules into the external leaflet triggers the budding transition, but fission of the budded vesicle is produced. This phenomenon prevents the shape recovery and therefore the lipid flip-flop quantification. However, the present approach should also allow investigation of energydependent flippases by reconstituting them into giant vesicles [26]. Although flippases are very likely reconstituted to a similar extent in both opposing directions, unidirectional lipid transport can be ensured by allowing ATP to access only one membrane leaflet. Characterizing the activity of energydependent flippases would not require the addition of lipids. Lipid species to be transported can already be incorporated during GUV formation. Because of the unidirectional fashion of oriented lipid transport, area differences would be created leading to a shape change. This is different from the visualization of energy-independent flippase activity in GUVs where shape changes have to be triggered first, for example, by supplementing the external leaflet with additional lipids. This type of experiment, which requires neither labeled lipids nor short-chain lipids, is illustrated in Figure 2.5 [27]. In this experiment, GUVs were fabricated from erythrocyte membrane extracts lacking the cytoskeleton proteins (spectrin and actin). The addition of magnesium-adenosine triphosphate (Mg-ATP) in the vicinity of a giant prolate vesicle containing solely the endogenous lipids of a red cell membrane triggered the budding transition by the active outward transport of endogenous PS and phosphatidylethanolamine (PE) by the aminophospholipid translocase [20]. This kind of experiments will be important for the identification of the lipid specificity recognition by lipid flippases [28].

36


(a)

(b)

(c)

(d)

(e)

(f)

(g)

(h)

(i)

Figure 2.5.â•‡ Giant vesicle made from erythrocyte membrane extract. The activation of aminophospholipid translocase by external addition of Mg-ATP (1â•›mM) induces a budding transition from a prolate vesicle. PS and PE are transported outward and accumulated into the outer monolayer. Time after Mg-ATP injection: (a) tâ•¯=â•¯0 seconds, (b) tâ•¯=â•¯45 seconds, (c) tâ•¯=â•¯46 seconds, (d) tâ•¯=â•¯47 seconds, (e) tâ•¯=â•¯48 seconds, (f) tâ•¯=â•¯52 seconds, (g) tâ•¯=â•¯1 minute, (h) tâ•¯=â•¯1 minute and 1 second, and (i) tâ•¯=â•¯1 minute and 2 seconds. Scale bar represents 10â•›µm. Temperature: 20°C. From Reference 27.

2.3â•… MEASUREMENT OF TRANSMEMBRANE FLIP-FLOP OF UNLABELED LIPIDS USING AFM AFM is a surface characterization technique that provides topographic information of surfaces [29, 30]. Developed in the mid-1980s within the physics community, the technique has spread among the biologist/biophysicist given its unique capacity to characterize surfaces with nanoscopic resolution while

Transmembrane Flip-Flop of Unlabeled Lipids Using AFM

37

8 7 6

Z (Å)

5 4 3 2 1 0 0

100 200 300 400 500 600 700 X (nm)

Figure 2.6.â•‡ Regions with different lipid-packing density in a supported lipid bilayer made of an Escherichia coli lipid extract (left panel). The height profile (indicated with a solid line on the AFM image) shows the height difference (0.5â•›nm) between both regions.

maintaining the surface immersed in solution. Supported biological membranes have been widely characterized structurally using this technique [31, 32]. The height resolution detects lipid domains that differ in as little as 0.5â•›nm, allowing then to unambiguously identify regions with different lipid aggregation state or packing density (Fig. 2.6). This, combined with its lateral resolution, serves to identify nanometer-range lipid-ordered domains [33]. This size range fills the gap between the micrometer scale accessible through optical microscopy and the high resolution obtained by electron microscopy. The direct topographic information it provides cannot be accessed easily through other techniques available. Since the image contrast is provided by height differences in the membranes, the AFM has the additional advantage that it does not require the use of external labels to provide domain information. Another quality of the AFM microscope that makes it convenient for characterizing biological samples is its ability to image surfaces immersed in a liquid environment. It allows time-resolved monitoring of a surface undergoing modifications. The surface remodeling local effect of lipid-degrading enzyme activities [9, 34, 35] has been detected within the minute time resolution accessible with the AFM. The existing limitations to imaging at higher speeds are due to the mechanical stability of the microscope and not to limitations in the basic operating principles. Great progress has been made recently in developing fast scanning probe equipment [36] that promise an exciting future for this technique. The sample preparation requirement of supporting the membrane onto a solid surface does however impose certain limitations to the type of questions

38


that can be addressed with the AFM in the study of membranes. Anchorage to a solid support requires that the lipid heads of the bottom monolayer interact directly with the solid surface. This interaction however does not completely immobilize the lipids. It is well documented that, although restricted, the lipids are still free to diffuse laterally on supported lipid membranes on different substrates [37]. The water monolayer trapped between the polar head groups and the solid surface is enough to allow them to diffuse. Although it is not obvious if this membrane configuration required by the AFM supports lipid flip-flop of individual lipids from the upper layer to the lower layer of vice versa, there have been a few cases in which phospholipids exchange between the monolayers has been reported [8]. There are basically two circumstances in which this lipid reorganization is clearly observed: (1) when asymmetric supported bilayers are formed initially [8] and (2) when the lipid asymmetry is induced by an enzyme activity affecting only the monolayer exposed to the solution [9]. The nanometric structural information of unlabeled membranes provided by AFM complements the dynamic and spectroscopic information provided by the use of fluorescent techniques, making their combined use of particular interest for understanding at the molecular level the effect of lipid composition on structure and rearrangement of membranes. A good illustration of the contribution of AFM to the study of flip-flop is provided by the study of Lin et al. [8]. Asymmetric supported membranes of two lipids with different transition temperatures, 1,2-dilauroyl-sn-glycero3-phosphocholine (DLPC) and 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC) lipids, were formed. The membranes were prepared by varying the thermal history of the vesicles or the substrate–vesicle suspension temperature differential during deposition. Depending on the detailed temperature history of the sample preparation, three types of membranes differing in the domain height distribution and in stability were observed. Whereas two of the samples were very stable, one of the configurations was observed to undergo domain height redistribution within a few hours. A combination of fluorescent techniques, fluorescence quenching and fluorescence recovery after photobleaching (FRAP), was able to associate the domains to an asymmetric lipid composition in the two membrane leaflets. Symmetric, symmetric/asymmetric, and asymmetric domains of DLPC/DSPC were identified (Fig. 2.7). The reorganization of the domains was interpreted as being due to lipid reorganization induced by lipid flip-flop between the two membranes. The possibility provided by AFM to image with nanometer resolution the time evolution of the domains allowed addressing the question of whether this lipid exchange occurred within the domains or if it occurred preferentially on their perimeter. The evolution of the domain perimeter was quantified, and the values obtained were compatible with the estimates obtained by assuming that the flip-flop occurs at the perimeter and not with the estimates considering exchange along the whole domain area. A rate constant of 76â•¯±â•¯17 per hour was estimated.

Transmembrane Flip-Flop of Unlabeled Lipids Using AFM

(a)

(b)

39

(c)

Figure 2.7.â•‡ Supported lipid bilayers made of DLPC and DSPC and prepared with different methods, which lead to three different configurations: symmetric (a), symmetric/ asymmetric (b), and asymmetric domains (c). The height profiles are denoted with a black dotted line on the AFM image. Reprinted from Biophysical Journal, 90, WanChen Lin, Craig D. Blanchette, Timothy V. Ratto, and Marjorie L. Longo, Lipid asymmetry in DLPC/DSPC-supported lipid bilayers: A combined AFM and fluorescence microscopy study, 228–237, Copyright (2006), with permission from Elsevier.

Another example of the use of AFM in combination with fluorescence techniques to observe the molecular details of membrane modifications induced by lipid flip-flop is given by the work by Lopez-Montero et al. [9]. Bilayers formed by egg-PCâ•›:â•›egg-PEâ•›:â•›SMâ•›:â•›Chol (1:1:1:1â•›mol) present liquidordered domains rich in SM. Exposure of the external monolayer to sphingomyelinase activity induces an in situ asymmetrical SM conversion to ceramide. The smaller area occupied by the ceramide head compared with the SM reduces the area of the external leaflet relative to the inner one, and this mismatch generates a tension. In GUVs, this tension is enough to collapse the vesicles even at low percentage of SM (≈â•¯5%â•›mol). When supported bilayers of the same composition and under the same treatment were analyzed with AFM, nanometer-sized holes and the appearance of higher domains were observed (Fig. 2.8). The results were interpreted as indicating that the pores formed due the lipid area mismatch allowed the redistribution of ceramide across the two leaflets with the subsequent formation of ceramide-rich domains slightly protruding from the surface.

40


(a)

(b)

(c)

(d) 1.4 1.2

Z (Å)

1 0.8 0.6 0.4 0.2 0 0

0.2

0.4 0.6 X (mm)

0.8

1

Figure 2.8.â•‡ Direct observation of nanoscale defects produced by SM to ceramide conversion. Time after injection of sphingomyelinase: (a) 0 minute, (b) 60 minutes, and (c) 65 minutes. The scanned region in (a) and (b) is the same as that delimited by a dashed square in (c). Dimensions of the dashed square are 5â•¯×â•¯5â•›µm. The nanometer-sized holes that appear upon enzymatic treatment are pointed out by arrowheads (b). The ceramide-rich domains indicated by white arrows are also observables in (c). The profile from the black bar on AFM images indicating the relative heights of different kinds of domains is shown in (d). From Reference 9.

These two examples illustrate that the time-resolved topographic information provided by AFM is an invaluable tool to complement the micrometer spectroscopic and dynamic information provided by fluorescence. Observing lipid rearrangements at the nanoscale occurring in the minute timescale contributes to understand the molecular mechanism underlying lipid scrambling and its implications for membrane structure.

Abbreviations

41

2.4â•… CONCLUSIONS Spontaneous lipid flip-flop on biological membranes is a slow process that can take from hours to days. The translocations most relevant to signal transduction and other important cell metabolic functions take place however much faster, more within the minute timescales (Chol, fatty acids, ceramides, or PG). The two ways of detecting unlabeled membrane asymmetry referred to in this chapter, shape changes in GUVs and lipid rearrangement detected by AFM, can provide information within this faster scale that can prove useful to understand the mechanism and regulation of unlabeled lipid translocation in artificial membranes. However, the above-mentioned lipids represent a little percentage of the large variety of lipids present in cells. Therefore, both methodological approaches shown here would be extended to the study of lipid transbilayer movement mediated by energy-dependent flippases. The experiments carried out on GUVs shown in Section 2.2 constitute stimulating evidence that questions referred to unlabeled lipid specificity recognition by proteins for instance, can be approached with both methodologies. The development of such techniques (among others that do not require marked lipids) can give a useful information, which can be, in many cases, complementary to that obtained for labeled lipids during the last four decades.

ACKNOWLEDGMENTS I.L.-M. thanks CAM and MICINN for financial support from NANOBIO-M and from Juan de la Cierva program 2007, respectively. This work was supported by MEC under grants BFU2005-0487-C02-01 and C02-02, S-0505/MAT-0283, and BIO200804478-C03-02.

ABBREVIATIONS ADE AFM BSA C10- C16- C6- Chol DLPC DSPC ER FRAP

area difference elasticity atomic force microscopy bovine serum albumin C10-acylC16-acylC6-acylcholesterol 1,2-dilauroyl-sn-glycero-3-phosphocholine 1,2-distearoyl-sn-glycero-3-phosphocholine endoplasmic reticulum fluorescence recovery after photobleaching

42

GUV kb LUV lyso-PC Mg-ATP PC PE PG PS SM T τ1/2


giant unilamellar vesicle Boltzmann constant large unilamellar vesicle lyso-phosphatidylcholine magnesium-adenosine triphosphate phosphatidylcholine phosphatidylethanolamine phosphatidylglycerol phosphatidylserine sphingomyelin temperature flip-flop half-time

REFERENCES â•‡ 1â•…R. D. Kornberg, H. M. McConnell, Biochemistry 1971, 10, 1111–1120. â•‡ 2â•… P. Fellmann, A. Zachowski, P. F. Devaux, Methods Mol. Biol. 1994, 27, 161–175. â•‡ 3â•… A. Pohl, I. López-Montero, F. Rouviere, F. Giusti, P. F. Devaux, Mol. Membr. Biol. 2009, 26, 194–204. â•‡ 4â•… A. J. Verkleij, R. F. Zwaal, B. Roelofsen, P. Comfurius, D. Kastelijn, L. L. van Deenen, Biochim. Biophys. Acta 1973, 323, 178–193. â•‡ 5â•… C. J. Kirby, G. Colin, Biochem. J. 1977, 168, 575–577. â•‡ 6â•… I. López-Montero, N. Rodriguez, S. Cribier, A. Pohl, M. Vélez, P. F. Devaux, J. Biol. Chem. 2005, 280, 25811–25819. â•‡ 7â•… A. Papadopulos, S. Vehring, I. López-Montero, L. Kutschenko, M. Stöckl, P. F. Devaux, M. Kozlov, T. Pomorski, A. Herrmann, J. Biol. Chem. 2007, 282, 15559–15568. â•‡ 8â•… W. C. Lin, C. D. Blanchette, T. V. Ratto, M. L. Longo, Biophys. J. 2006, 90, 228–237. â•‡ 9â•… I. López-Montero, M. Vélez, P. F. Devaux, Biochim. Biophys. Acta 2007, 1768, 553–561. 10â•… T. C. Anglin, J. Liu, J. C. Conboy, Biophys. J. 2007, 92, L01–L03. 11â•… T. C. Anglin, J. C. Conboy, Biophys. J. 2008, 95, 186–193. 12â•… M. Nakano, M. Fukuda, T. Kudo, N. Matsuzaki, T. Azuma, K. Sekine, H. Endo, T. Handa, J. Phys. Chem. B 2009, 113, 6745–6748. 13â•… W. Helfrich, Z. Naturforsch. 1973, 28c, 693–703. 14â•… L. Miao, U. Seifert, M. Wortis, H. G. Döbereiner, Phys. Rev. E 1994, 49, 5389–5407. 15â•… H. G. Dobereiner, Curr. Opin. Colloid Interface Sci. 2000, 5, 256–263. 16â•…R. Lipowsky, Europhys. Lett. 1995, 30, 197–202. 17â•… J. O. Rädler, I. Koltover, T. Salditt, C. R. Safinya, Science 1997, 275, 810–814.

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P. G. Petrov, J. Lee, H. G. Döbereiner, Europhys. Lett. 1999, 48, 435. D. L. Daleke, W. H. Huestis, Biochemistry 1985, 24, 5406–5416. M. Seigneuret, P. F. Devaux, Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 3751–3755. E. Farge, P. F. Devaux, Biophys. J. 1992, 61, 347–357. B. L. Mui, H. G. Döbereiner, T. D. Madden, P. R. Cullis, Biophys. J. 1995, 69, 930–941. 23â•… K. Berndl, J. Käs, R. Lipowsky, E. Sackmann, U. Seifert, Europhys. Lett. 1990, 13, 659–664. 24â•… E. Farge, P. F. Devaux, J. Phys. Chem. 1993, 97, 2958–2961. 25â•… M. L. Angelova, D. S. Dimitrov, Faraday Discuss. Chem. Soc. 1986, 81, 303–311. 26â•… P. F. Devaux, Cell. Mol. Biol. Lett. 2002, 7, 227–229. 27â•… P. F. Devaux, I. López-Montero, S. Bryde, Chem. Phys. Lipids 2006, 141, 119–132. 28â•… T. Pomorski, A. K. Menon, Cell. Mol. Life Sci. 2006, 63, 2908–2921. 29â•…R. J. Colton, D. R. Baselt, Y. F. Dufrêne, J. B. Green, G. U. Lee, Curr. Opin. Chem. Biol. 1997, 1, 370–377. 30â•… D. Muller, Biochemistry 2008, 47, 7986–7998. 31â•… P. L. Frederix, P. B. Brosshart, A. Engel, Biophys. J. 2009, 96, 329–338. 32â•… E. I. Goksu, J. M. Vanegas, C. D. Blanchette, W. C. Lin, M. L. Longo, Biochim. Biophys. Acta 2009, 1788, 254–266. 33â•… W. C. Lin, T. V. Ratto, M. L. Longo, Methods Mol. Biol. 2007, 400, 503–513. 34â•… M. C. Giocondi, B. Seantier, P. Dosset, P. E. Milhiet, C. Le Grimellec, Pflugers Arch. 2008, 456, 179–188. 35â•… L. Ira, J. Johnston, Biochim. Biophys. Acta 2008, 1778, 185–197. 36â•… T. Ando, T. Uchihashi, N. Kodera, D. Yamamoto, A. Miyagi, M. Taniguchi, H. Yamashita, Pflugers Arch. 2008, 456, 211–225. 37â•… Y. H. Chan, S. G. Boxer, Curr. Opin. Chem. Biol. 2007, 11, 581–587. 18â•… 19â•… 20â•… 21â•… 22â•…

PART II LIPID ASYMMETRY IN CELL MEMBRANES

3 NEW INSIGHTS IN MEMBRANE LIPID ASYMMETRY IN ANIMAL AND PLANT CELLS Alain Zachowski Laboratory of “Physiologie Cellulaire et Moléculaire des Plantes,” Université Pierre et Marie Curie—Paris 6 (UR 5) and Centre National de la Recherche Scientifique (EAC 7180), Paris, France

3.1â•… LIPID ASYMMETRY IN ANIMAL MEMBRANES In the early 1970s, the fact that phospholipids could not be randomly distributed across a biological membrane was established. The human erythrocyte membrane served as a model of plasma membrane, and the methods used were either chemical modification or enzymatic hydrolysis of phospholipids exposed at the cell surface [1–3]. The asymmetrical distribution was striking: ca. 70% of the phosphatidylcholine and more than 85% of the sphingomyelin were localized in the outer membrane leaflet; conversely, more than 80% of the phosphatidylethanolamine and almost all of the phosphatidylserine were found in the cytoplasmic leaflet. This distribution holds for almost all normal erythrocytes, except for ruminant ones that are deficient in phosphatidylcholine and very rich in sphingomyelin, and where phosphatidylethanolamine is almost exclusively found in the inner leaflet [4]. This distribution can be perturbed in pathological (sickle cells, thalassemic cells) or senescent red blood cells, as described in other chapters. When the same methods were used with


47

48

ARE PLANT MEMBRANE LIPIDS ASYMMETRICALLY DISTRIBUTED?

other types of cells, the interpretation of the results was more complex: if erythrocyte membrane is rather impermeant to the chemicals used and resistant to phospholipase-induced lysis and to perturbations induced by phospholipase products, this is not always the case for plasma membranes from nucleated cells. In addition, plasma membrane has to be isolated in order to have access to its total lipid composition. Cell lysis, which is thus the primary step in the isolation protocol, can induce a partial redistribution of the lipids within the bilayer [5]. Finally, if any transmembrane movement of lipids is faster than the reaction time required for labeling or hydrolysis, the description of asymmetry can be erroneous (see Chapters 1 and 6). However, all the data can be interpreted in the same way, namely an enrichment of the outer leaflet in choline-containing phospholipids and an accumulation of aminophospholipids in the inner leaflet. A crucial question was to know whether phosphatidylserine is totally excluded from the outer cell membrane leaflet, or whether it can be present in very small amounts on the cell surface. In other words, were the methods used and the detection of the modified lipid sensitive enough to reveal the presence of a small fraction of a low abundant phospholipid? This was not the case with chemical labeling of the amino group or formation of lysoderivatives by phospholipases A2. A more sensitive method is the prothrombinase assay that can reveal the presence of few percents of phosphatidylserine in a bilayer [6]. However, all these methods work on a cell population and cannot distinguish variations at the single cell level. A very interesting new technical approach to the question of phosphatidylserine asymmetry has been based on the use of fluorescent Annexin V. This protein specifically binds to the lipid in the presence of calcium [7], and if some phosphatidylserine is present on the plasma membrane outer leaflet, living cells become fluorescent (for reviews, see References 8 and 9). Thus, one has access to the situation existing at a single cell level. This noninvasive labeling is now a well-recognized assay and largely confirms that, in most of the “normal, resting” cells, phosphatidylserine is absent from the plasma membrane external layer. On the contrary, it has been shown that apoptotic cells are labeled, indicating that some phosphatidylserine has migrated into the plasma membrane outer layer. Even if some peptides, such as duramycin, can bind phosphatidylethanolamine [10], they have been seldom utilized to label cell surface and to check for the presence of this phospholipid. Determination of lipid distribution in organelle membranes has been much less studied than in plasma membranes. A reason is certainly that one has to isolate these membranes as closed, impermeable vesicles before assaying lipid distribution by either chemical labeling or enzymatic assays (see Chapter 6). On the other hand, the presence of some flippases in given organelles might indicate whether or not a phospholipid asymmetry exists in the membrane. For instance, the presence of an aminophospholipid translocase in the transGolgi network [11] might indicate that phosphatidylserine is asymmetrically distributed in this membrane. Similarly, the presence of a scramblase (see

Phospholipid Transporters

49

Chapter 7) would explain a rather symmetrical distribution of phospholipids in organelles such as exosomes [12]. 3.2â•… CREATING, MAINTAINING, OR RANDOMIZING THE MEMBRANE PHOSPHOLIPID DISTRIBUTION: PHOSPHOLIPID TRANSPORTERS Over the past two decades, numbers of reviews have dealt with the various proteins responsible for lipid translocation within the bilayer, and some of these “flippases” are the subject of other chapters. For some recent reviews, see References 13–17. So, I will just make a brief summary of what has been described in animal or yeast cells. The situation in plant cells will be exposed later in this review. 3.2.1â•… Aminophospholipid Translocase In the 1980s, an ATP-dependent, inward translocation of aminophospholipids in the plasma membrane of erythrocytes and some nucleated cells has been largely documented [1, 3, 18]. Because of the phospholipid specificity, this transporter was named “aminophospholipid translocase.” The transport requires Mg2+ besides ATP, is inhibited by vanadate and calcium, and is sensitive to protein-modifying reagents. The identification of the active peptide, suspected to be identical to the bovine chromaffin granule ATPase II [19], has been established by Tang et al. [20], who showed that this ATPase is able to complement a yeast mutant deficient in the aminophospholipid transport. This was the first member of a large P-type ATPase family, called P4-ATPases, that is present in animals, plants, or yeasts [21] (see Chapters 8 and 9). The P4ATPases maintain aminophospholipid asymmetry and are also involved in membrane trafficking in the endocytic and secretory pathways [22, 23]. A recent question is whether the ATPases can support by themselves the lipid transport or they need a β-subunit to fulfill this function [24]. Peptide homologs to the protein Cdc50p found in yeast [25] would be excellent candidates. 3.2.2â•… ATP-Binding Cassette (ABC) Transporters These transporters would function in an opposite direction to the aminophospholipid translocase. The lipids would move toward the outer membrane leaflet. Few examples of transport of endogenous, long-chain phospholipids have been reported. For instance, MDR3 (ABCB4) is a specific transporter of phosphatidylcholine [26] in bile canalicular membranes [27]. As to MDR1 (ABCB1), it has been reported to catalyze the outward transport of shortchain and 7-nitrobenz-2-oxa-1,3-diazol-4-yl (NBD)-labeled phospholipids and sphingolipids [26]. A question is whether the lipids are recognized per se, or

50


whether they are just xenobiotic “drugs” that are expelled from the cell for detoxification (see Chapters 11 and 17). 3.2.3â•… Scramblase In the presence of calcium, such a protein reorganizes membrane phospholipids and partially or totally quenches the asymmetry. Two putative scramblases have been proposed [28, 29], but that they really work as scramblases is still discussed [30] (see Chapter 7). 3.2.4â•… Flippases of Biogenic Membranes In the endoplasmic reticulum membrane, a bidirectional movement of phosphatidylcholine [31] and other phospholipids [32] takes place (see Chapter 6). It is protein dependent [33] and, until now, has never been described in plasma membranes. The role of this transporter would be to equilibrate newly synthesized phospholipids, which are released on one membrane face, between the two membrane leaflets.

3.3â•… WHAT ABOUT LIPID ASYMMETRY AND TRANSLOCATION IN PLANT CELL MEMBRANES? Looking at lipid asymmetry is a little bit more challenging with plant cells than with animal cells. In fact, plant cells are surrounded by a cell wall. It is composed of polysaccharides (e.g., cellulose), some phenolic compounds (e.g., lignin), and proteins. As plant cells are living in a hypotonic environment, they are turgescent because of water influx. The cell wall prevents the cells from indefinitely swelling, until bursting. It is also an exoskeleton that gives a plant its shape. Due to its selective permeability for relatively low-molecular-weight compounds, the determination of the plasma membrane asymmetry based on phospholipase attack cannot be performed. On the other hand, the cell wall is not a totally hydrophilic structure, so fluorescent or spin-labeled lipid probes can neither be used according to the well-established protocols. Anyway, it is possible to get rid of the cell wall by enzymatic digestion and obtain so-called protoplasts, which resembles animal cells in accessibility from the outer medium. However, protoplasts are very fragile and stressed structures that have to be manipulated very cautiously. 3.3.1â•… Lipid Asymmetry in the Plasma Membrane (Plasmalemma) Until now, no extensive determination of phospholipid distribution in the plasma membrane of protoplasts has been carried out, to the exception of one study on mung bean (Vigna radiata) protoplasts [34]. The protoplasts were not

WHAT ABOUT LIPID ASYMMETRY AND TRANSLOCATION?

51

labeled by fluorescent Annexin V, showing that all the phosphatidylserine was sequestered in the inner membrane leaflet. This is thus in accordance with the general trend found in animal cell plasma membranes. However, it has to be noted that phosphatidylserine is much less abundant in plant cells than in animal cells [35, 36] and that its asymmetric distribution represents a weak potential perturbation in the mass equilibrium that must exist between the two monolayers. Using phospholipase A2 and right-side-out plasma membrane vesicles isolated from the same protoplasts, it has been found that phosphatidylcholine, phosphatidylethanolamine, and phosphatidic acid were almost symmetrically distributed across the bilayer. That this represents the genuine distribution in intact membranes or a partially perturbed state due to membrane rupture that partially randomizes phospholipids [5] is still an open question. Finally, no sphingomyelin asymmetrical distribution can be described, as this lipid is absent from plants. The asymmetric distribution of phosphatidylserine has also been described in plasma membrane from other plants (tobacco, apple), always on the basis of the quasi-absence of Annexin V binding [37, 38]. Interestingly, this asymmetry is severely perturbed during progression of apoptosis, and phosphatidylserine becomes exposed on the outer leaflet of the plasma membrane [38–40]. 3.3.2â•… Lipid Asymmetry in Intracellular Membranes Organization of lipids in organelle membranes has been studied by enzymatic assays, using various phospholipases, and by chemical labeling of the exposed groups. However, interpretation of data in terms of asymmetric distribution may be difficult due to the inaccessibility of the different lipid species to the enzymes or the chemical. A good example is what was found in mitochondria from castor bean (Ricinus communis) endosperm [41]. Approximately, 45% and 60% of the phosphatidylcholine was inaccessible to phospholipase A2 in outer or inner mitochondrial membranes, respectively. These numbers dropped to 25% and 12% for phosphatidylethanolamine and 25% and zero for phosphatidylinositol (and eventually phosphatidylserine, if present). As location of these subpopulations within the bilayer is unknown, the true distribution of each lipid within the bilayer cannot be rigorously established. Anyway, on the basis of the accessible fraction of each phospholipid class, some asymmetry appeared. For instance, all the phosphatidylinositol of the inner membrane is localized in the inner leaflet, while it is twice more abundant in the outer leaflet than in the inner leaflet of the outer membrane. Phosphatidylethanolamine is always preferentially located in the outer leaflet of the outer membrane (44 vs. 32) or of the inner one (70 vs. 18). As to phosphatidylcholine, it is almost symmetrically distributed. The situation is simpler in glyoxysomal membrane, as the inaccessible fraction is very low [41]. The outer leaflet is the preferential location of phosphatidylethanolamine (67 vs. 16 in the inner leaflet), while phosphatidylcholine

52


and phosphatidylinositolâ•¯+â•¯phosphatidylserine are mainly or almost exclusively, respectively, found in the inner leaflet. Two other organelles, specific of plant cells, have also been studied for the lipid distribution in their membranes. The first one is the vacuole. The vacuole is an organelle whose volume represents approximately 90% of the cell volume. Its main role is to maintain a well-balanced water equilibrium in the cell (it acts as a water reservoir) and to keep the turgescent state. The vacuole is surrounded by a membrane (also known as tonoplast). In sycamore (Acer pseudoplatanus) cells [42], this membrane is made of phospholipids (45%), glycolipids (40%) including sterylglucosides, and neutral lipids (15%). Among the phospholipids, phosphatidylcholine accounts for 30%, phosphatidylethanolamine for 47%, and phosphatidylinositol for 15%. When the membrane was treated by chemical reagents (trinitrobenzene sulfonic acid [TNBS] or fluorescamine) or by phospholipases (either A2, C, or D), the relatively large proportion of inaccessible molecules made it difficult to interpret data in terms of phospholipid asymmetry [43]. The inaccessible molecules represented 10– 20% of phosphatidylcholine and 16–34% of phosphatidylethanolamine. Sequential degradation by phospholipase A2 and phospholipase C reduced these numbers to 10%, or less, allowing one to draw a better scheme of monolayer composition. Phosphatidylcholine is equally distributed across the bilayer, while phosphatidylethanolamine is slightly more present (by 20%) in the outer leaflet. Another membrane lipid, namely phosphatidylinositol, was a poor substrate of phospholipase A2, even in the presence of detergent, most probably because palmitic acid (16:0) represented more than three-quarters of fatty acids. The other organelle is the chloroplast. It is the intracellular compartment where photosynthesis takes place. The photosynthetic complexes are located in the thylakoid membrane, which is arranged in a complicated system of membranous cisternae and lamellae. The thylakoid membrane can account for up to 90% of the membranes found in a green leaf. The chloroplast is surrounded by two bilayers, the envelopes, which isolate the chloroplast interior from the cytoplasm. The outer envelope is freely permeable to small molecules, while the inner envelope behaves as a permeability barrier. Plastid membranes are characterized by the presence, in large amounts, of polar lipids with a galactose-containing head group, named galactolipids (Fig. 3.1). Besides, some phospholipids are present, but in low quantities [44]. MonogalactosylÂ� diacylglycerol (MGDG) and digalactosyldiacylglycerol (DGDG), in a ratio close to 2:1, represent more than 80% of the thylakoid membrane lipids. Phosphatidylglycerol is the major phospholipid of this membrane. The composition of the inner envelope membrane resembles that of the thylakoid one. The outer envelope membrane is rich in DGDG and phosphatidylcholine, MGDG being less present than in the other two membranes. It has to be noted that phosphatidylethanolamine is completely absent from these bilayers. There is a marked asymmetry of phospholipids in the outer envelope: phosphatidylcholine is almost completely (>80%) located in the outer (cytoplasmic) leaflet,

53

WHAT ABOUT LIPID ASYMMETRY AND TRANSLOCATION? OH

OH OH

OH O H2C

CH

O

O

R1

R2

OH OH

O

O

CH2OH

CH2

H2C

MGDG

CH

O

O

R1

R2

O

CH2

CH2SO3–

SQDG

OH OH

OH O H2C O R1

CH

CH2

OH

O CH2

O

O

OH CH2OH

O R2

OH

DGDG

Figure 3.1.â•‡ Structure of the major glycolipids found in plastidial membranes. GalactoÂ� lipids contain one or two galactose molecules in sn3 position of a glycerol backbone. The monogalactosyldiacylglycerol (MGDG) is a 1,2-diacyl-3-O-(β-D-galactopyranosyl)sn-glycerol. The digalactosyldiacylglycerol (DGDG) is a 1,2-diacyl-3-O-(α-Dgalactopyranosyl-(1→6)-O-β-D-galactopyranosyl)-sn-glycerol. Galactolipids contain a high amount of polyunsaturated fatty acids. Very often, linolenic acid (18:3) is present at both the sn1 and sn2 positions of the glycerol backbone. In some plants, a palmitolenic acid (16:3) is present at sn2 position of MGDG (and not at sn1 position). The proportions of C18/C18 and C18/C16 MGDG vary among plants [71]. The much abundant sulfolipid is the sulfoquinovosyldiacylglycerol (SQDG), which is a 1′,2′-diacyl-3′O-(6-deoxy-6-sulfo-α-D-glucopyranosyl)-sn-glycerol. Some of it is dipalmitoylated (16:0/16:0), but the major part contains both 16:0 and 18:3 fatty acids. Interestingly, the 16:0 fatty acid can be linked to sn1 or sn2 position of the glycerol backbone.

where phosphatidylglycerol is totally absent [45]. The distribution of the galactolipids in the envelopes was never extensively determined. As to thylakoids, there is a clear asymmetry of all the lipids. That theses structures were isolated from spinach (Spinacia oleracea) or from oat (Avena sativa), phosphatidylglycerol was found more present on the outer leaflet (60–70%) on the basis of susceptibility to exogenous phospholipase A2 [46, 47]. The transmembrane distribution of galactolipids was also determined by lipase attack of thylakoids purified from leaves of lettuce, pea, barley, oat, spinach, or black nightshade [48, 49]. MGDG was always found enriched in the outer leaflet (53–65%) while

54


DGDG resides predominantly in the inner leaflet (78–90%). A reliable distribution of sulfoquinovosyldiacylglycerol could not be reached, as a significant part (30%) was inaccessible to the acyl hydrolase used for the assay. However, as only 10% of this lipid can be hydrolyzed in the outer leaflet, and even if the nonaccessible fraction is also located in this leaflet, it will be more present in the inner leaflet. The transmembrane galactolipid asymmetry already exists in prothylakoids [49], that is, before incorporation of the chlorophyll–protein complexes. So, the asymmetry has to be established very soon in membrane life. In fact, it appears that it is the lipid synthesis in the inner envelope and the lipid transfer into the thylakoid membrane that is at the origin of the asymmetry [50, 51]. Once the asymmetry is created, it is easy to conceive that galactolipids keep their distribution, as their transverse diffusion rate is certainly very low, due to the bulky, fully hydrated head group. 3.3.3â•… Aminophospholipid Translocase(s) in Plant Cells The phosphatidylserine asymmetry described in plant cell plasma membranes (see above) makes it probable that an aminophospholipid translocase is active in this bilayer. Indeed, such an activity was found in Arabidopsis thaliana, and called aminophospholipid ATPase (ALA)1 [52]. No direct measure of lipid translocation was made on intact cells or protoplasts. But two assays showed that ALA1 was capable to transport phosphatidylserine within a membrane. The first one was to compare NBD-labeled phospholipid internalization in yeasts, either wild-type, or drs2, or ALA1-expressing drs2. Results showed that phosphatidylserine translocation was reestablished in the latter strain, similar to what was described with bovine chromaffin granule ATPase II [20]. The second one was to measure ATP-dependent translocation of NBDphospholipids in vesicles made from microsomal yeast membranes and soybean lipids. Wild-type reconstituted vesicles showed an ATP-dependent inward movement of NBD-phosphatidylserine. This movement was much reduced in vesicles prepared from drs2 yeasts, and restored in vesicles from ALA1-expressing drs2 yeasts. The same type of results was obtained for NBDphosphatidylethanolamine, although lower in amplitude. As to movement of NBD-phosphatidylcholine, it was identical in the three preparations and insensitive to ATP, as expected. Thus, the ALA1 protein possesses all the characteristics of an aminophospholipid translocase present in plant cells. Another function of flippases would be to create an asymmetry in the lipid content of each monolayer by selectively transporting a given phospholipid from one monolayer to the other one. This excess in population can generate a local membrane curvature that, if it is amplified, will lead to vesicle formation and budding [53, 54] (see Chapter 2). Such a role has been experimentally confirmed in yeasts that lack plasma membrane aminophospholipid translocases (here, Dnf1p and Dnf2p) and also present a defect in endocytosis [23]. In addition, aminophospholipid translocase is not limited to plasma membrane, but can also be found in Golgi membranes [55, 56]. Here again, lack of


55

activity (Drs2p defective mutant cells) blocks the formation of a subclass of post-Golgi secretory vesicles [57], a phenomenon that can be explained by the importance of lipid transport for vesicle budding. A very similar case has been described in plants. Another isoform (ALA3) of P4-ATPase found in A.â•›thaliana is required for secretory vesicle formation [24] and is localized to the Golgi membrane. To complement yeast mutants defective in Drs2p, Dnf1p, and Dnf2p, expression of ALA3 alone is not sufficient. The presence of a protein from the Cdc50p family (here called ALIS1) is also required. Under these conditions, internalization of NBD-labeled phospholipids is reestablished, especially for NBD-phosphatidylethanolamine, rather than for NBDphosphatidylserine. Surprisingly, the co-expressing yeasts are also capable to internalize NBD-phosphatidylcholine at a higher rate than the defective mutant ones. This is not the case when Drs2p is expressed, showing some differences between the apparent orthologs in plants and yeasts. More recently, the requirement of a second subunit associated to ALA peptide to ensure the correct transport function has also been demonstrated for the ALA2 isoform [58]. In Arabidopsis, there are 12 putative aminophospholipid translocases, ALA1 to ALA12 (Table 3.1). They are all around 130â•›kDa in mass (1107–1247 amino acids), and it is predicted that they contain 10 transmembrane helices. Differences appear in the calculated isoelectric point of these isoforms. They can be distributed in three groups: proteins with a pI below 6 (ALA1, 2, 9, 10, and 12), those with a pI comprised between 6 and 7 (ALA5, 8, and 11), and those with a pI above 7 (ALA3, 4, 6, and 7). Comparing the sequences, it appears that there is at least 30% of identity between two isoforms, showing fairly high sequence conservation within the family (Table 3.2). It has to be pointed out that the identity can reach values as high as ca. 90% (ALA4/ ALA5, ALA6/ALA7, ALA10/ALA11). This is more clearly illustrated by the phylogenic tree (Fig. 3.2) showing that eight isoforms are distributed as four pairs, and that ALA1 and ALA2 are the more distant peptides. Putative aminophospholipid translocases have also been found in other plants whose entire genome is known. Twelve flippases could exist in rice (Oryza sativa), five in grape (Vitis vinifera), seven in a moss (Physcomitrella patens), and two in an algae (Chlamydomonas reinhardtii). It is striking that a high percentage of identity (>25%) exists among these different sequences. In fact, this observation does not concern solely the plants; it holds through all reigns. Comparing a random isoform from various animals, plants, and yeast shows that identity is very often comprised between 30% and 40% (Table 3.3). One can wonder why this enzyme is so conserved, at least in sequence. A reason for maintaining phospholipid asymmetry in animal cell plasma membrane can be found by considering what would happen in its absence (see Chapters 7 and 14–16). The presence of phosphatidylserine on the outer membrane leaflet makes the cell surface procoagulant, which can be deleterious for health. Moreover, the presence of external phosphatidylserine is a signal for cell clearance by monocytes/macrophages. In plants, these responses do

56


TABLE 3.1.â•‡ Properties of the 12 Putative Aminophospholipid Translocase Found in Arabidopsis thaliana Genome Ordered locus name Swiss-Prot accession Number of amino acids Mass (Da) pI

ALA1

ALA2

ALA3

ALA4

ALA5

At5g04930 P98204 1158 130,329 5.92

At5g44240 P98205 1107 124,836 5.97

At1g59820 Q9XIE6 1213 137,753 7.84

At1g17500 Q9LNQ4 1216 138,189 7.53

At1g72700 Q9SGG3 1228 139,342 6.89

The mass and pI are calculated from the amino acid sequence of each peptide (without any potential posttranslational modifications).

TABLE 3.2.â•‡ Percentages of Identity between the 12 Isoforms of Putative Aminophospholipid Translocase Found in Arabidopsis thaliana Genome ALA1

ALA2

ALA3

ALA4

ALA5

ALA6

ALA7

31.3

34.2 31.8

35.0 31.4 44.0

35.6 31.1 43.3 87.7

34.6 30.9 42.2 76.6 75.9

35.7 30.7 42.6 77.3 75.8 88.2

0.993

1 0.998 0.99

sp_P98204_ALA1 sp_Q9XIE6_ALA3 0.992 sp_Q9SLK6_ALA6 sp_Q9LVK9_ALA7 0.985 sp_Q9LNQ4_ALA4 sp_Q9SGG3_ALA5 sp_Q9LK90_ALA8 sp_Q9LI83_ALA10 1 sp_Q9SAF5_ALA11 1 sp_P57792_ALA12 0.997 sp_Q9SX33_ALA9 sp_P98205_ALA2

0.5

Figure 3.2.â•‡ Phylogeny of the 12 potential aminophospholipid translocases from Arabidopsis thaliana. Tree was obtained at http://www.phylogeny.fr [72].

57


ALA6

ALA7

ALA8

ALA9

ALA10

ALA11

ALA12

At1g54280 Q9SLK6 1244 140,682 7.10

At3g13900 Q9LVK9 1247 140,928 7.15

At3g27870 Q9LK90 1189 135,309 6.21

At1g68710 Q9SX33 1200 136,045 5.92

At3g25610 Q9LI83 1202 136,279 5.69

At1g13210 Q9SAF5 1203 136,584 6.54

At1g26130 P57792 1184 133,793 5.91

ALA8

ALA9

ALA10

ALA11

ALA12

36.0 31.9 43.4 55.0 54.9 53.6 53.0

35.2 31.2 44.1 55.8 55.3 54.4 55.7 63.3

34.6 31.0 43.6 56.4 55.3 53.7 54.9 62.8 74.1

33.4 31.5 44.3 55.9 55.3 53.5 54.7 61.2 73.4 87.0

35.7 31.0 42.7 53.6 53.9 53.2 52.8 63.1 80.9 71.6 70.6

ALA1 ALA2 ALA3 ALA4 ALA5 ALA6 ALA7 ALA8 ALA9 ALA10 ALA11 ALA12

TABLE 3.3.â•‡ Percentages of Identity between Some (Putative) Aminophospholipid Translocase from Various Eukaryotic Organisms: Arabidopsis thaliana (ALA1, Swiss-Prot Number P98204), Man (ATP8B1, Swiss-Prot Number O43520), Ox (Chromaffin Granule ATPase II, ATP8A1, Swiss-Prot Number Q29449), Saccharomyces cerevisiae (DRS2, Swiss-Prot Number P39524), Chlamydomonas reinhardtii (ALA2, Swiss-Prot Number A8IVJ6), Rice (Oryza sativa, Swiss-Prot Number Q10LU3), and Grape (Vitis vinifera, Swiss-Prot Number A5BQL2) Arabidopsis

Man

Ox

Yeast

Chlamydomonas

Rice

Grape

35.1

35.4 39.8

32.5 35.3 39.7

31.9 35.4 36.1 33.2

52.0 32.2 33.1 32.6 29.1

35.4 37.0 37.8 34.0 41.1 34.2

Arabidopsis Man Ox Yeast Chlamydomonas Rice Grape

58


not exist, and eventually apoptotic cells have to be kept (to limit pathogen infection, for instance [59]). One of the roles of lipid translocation through the bilayer is to allow vesicle formation from organelles [24], a role that seems common to yeast, animal cells, and plant cells. In accordance with this role, a mutation in ALA3 gene of A.â•›thaliana provokes severe alterations in the polar growth of the cells, leading to reduced primary root growth, longer root hair, and aberrant trichome expansion [60]. Surprisingly, reduced expression of ALA1 in A.â•›thaliana affects the behavior of the plant exposed to low temperature: modified, antisense plants are smaller than wild-type plants when exposed at 8–12°C. No differences in phenotype are seen at normal temperature. It has to be recalled that the drs2 yeast mutant, defective in a protein of the same family, was also temperature sensitive [61]. How could a defect in aminophospholipid transport affect plant growth at low temperature? During adaptation to cold, plants (and in a broader sense, poikilotherms) modify lipid composition of their membranes, both for lipid class and for fatty acid compositions [62]. As fatty acid desaturases and (major part of) lipid synthases are located in the endoplasmic reticulum, lipid trafficking toward the other organelle membranes has to take place in order to make all the adaptive adjustments. So, one can conceive that quenching an aminophospholipid translocase involved in vesicle formation could hamper the correct response to the new conditions. Another possibility relies on the absence of asymmetry of phosphatidylserine (but it represents only few percents of the membrane lipids) and phosphatidylethanolamine in such a mutant. As the fatty acid composition varies with the phospholipid class, the distribution of these fatty acids in each monolayer will be different in the presence or the absence of flippase. It could be possible that favorable lipidic environment of some proteins is not matched at low temperature in the mutants. It is interesting to check whether levels of transcripts of the various ALAs are sensitive to environmental temperature. Mining Genevestigator databases (http://www.genevestigator.com), one can see that cold treatment is not followed by major changes in the transcript levels of the 11 ALAs tested (Fig. 3.3). In fact, the low level of responsiveness can be found in a great variety of situations: abiotic stresses (drought, osmotic stress, oxidative stress, wounding, light quality and intensity) and presence of chemicals (including nutrients) or hormones. The same is true with biotic stresses (attack by pathogens, for instance). Nonetheless, if one gene has to be considered as the most affected, it would be ALA10 whose expression is sometimes increased by a 2–2.5 factor. When the expression is checked against plant organs, again there are only few cases where there is a significant change. Pollen appears to be an organ where transcripts are abundant, at least for some of the ALAs. When genes are clustered as a function of their expression in the different plant organs (Fig. 3.4), it is clear that the tree obtained is different from the phylogenetic tree, indicating that two very close isoforms (e.g., ALA9/Q9SX33 and ALA12/P57792) are not subjected to the same regulations.

59

WHAT ABOUT LIPID ASYMMETRY AND TRANSLOCATION? –2.0

–1.5

–1.0

–0.5 0.0

1.0

1.5

2.0

2.5

# Arrays (exp/ctrl) 3/3 8/4 6/2 6/6 6/6 6/6 6/6

ALA12 ALA7 ALA3 ALA5 ALA1 ALA4 ALA6 ALA2 ALA11 ALA9 ALA8 ALA10

Arabidopsis thaliana (treatment) Stress: cold_2 Stress: cold_3 Stress: cold_4 Stress: cold_green_early Stress: cold_green_late Stress: cold_roots_early Stress: cold_roots_late

0.5

At1g26130 At3g13900 At1g59820 At1g72700 At5g04930 At1g17500 At1g54280 At1g44240 At5g13210 At1g68710 At3g27870 At3g25610

–2.5

Figure 3.3.â•‡ Microarray analysis of ALA expression in Arabidopsis thaliana enduring a cold stress, as obtained from Genevestigator [73]. The bar at the top is giving the color code for the ratio of induction/repression of gene transcription. Color version on the Wiley web site.

Thus, in A.â•›thaliana, the genes coding for the various aminophospholipid translocase behave (almost) as housekeeping genes, a strong indication that this activity could be essential for cell life. Further studies with plants mutated in each of these genes should help in understanding the role of asymmetry. 3.3.4â•… Other Flippases in Plant Cells: ABC Transporters ABC proteins are extremely abundant in plants [63, 64]. For instance, more than 100 members of this family are presumed to exist in A.â•›thaliana. No in vitro study of lipid translocation by one of these proteins has been performed, so one can just hypothesize that there will certainly be some flippases in the family. Examples are the peroxysomal PAX1/CTS/PED3 protein that could transport fatty acids or acyl-CoAs [65–67] and the plastidial TGD1, 2, 3 multicomponent transporter proposed to mediate the transmembrane movement of polar lipids [68, 69]. 3.3.5â•… Other Flippases in Plant Cells: Endoplasmic Reticulum Flippase Very recently, strong experimental data were presented in favor of the existence of a flippase in endoplasmic reticulum membrane [70]. In proteoliposomes reconstituted from solubilized endoplasmic reticulum from spinach leaves, there is an ATP-independent translocation of phosphatidylcholine and phosphatidylethanolamine. This translocation is sensitive to trypsine and chemical reagents. Based on differential inhibition of translocation of each phospholipid, it is highly probable that more than one flippase is present in

ARE PLANT MEMBRANE LIPIDS ASYMMETRICALLY DISTRIBUTED? Q9SGG3 Q9LNQ4 P98205 Q9XIE6 P98204 Q9SX33 Q9LK90 Q9LI83 Q9SAF5 P57792 Q9LVK9 Q9SLK6

60

callus cell suspension seedling cotyledons hypocotyl radicle imbibed seed inflorescence flower carpel ovary stigma petal sepal stamen pollen pedicel silique seed embryo endosperm stem node shoot apex cauline leaf rosette juvenile leaf adult leaf petiole senescent leaf hypocotyl xylem cork roots lateral root root hair zone root tip elongation zone endodermis endodermis+cortex epid. atrichoblasts lateral root cap stele

References

61

Figure 3.4.â•‡ Clustering of the 12 genes coding for aminophospholipid translocase in Arabidopsis thaliana as a function of their expression in the different organs. The darker the color, the higher is the expression level. This graph was obtained from Genevestigator. Color version on the Wiley web site.

the original membrane. As in other organisms, no molecular identification of the corresponding peptides has been done. In conclusion, plants could represent a good model to shed more light on phospholipid asymmetry and the proteins responsible for it. A genetic (mutant) approach is rather easy with a model such as Arabidopsis, as many knockout and knockdown mutants are readily available, and as stable transformation of the plant is well documented. However, the multiplicity of the isoforms, and a possible redundancy, will make the physiological studies quite tedious. ABBREVIATIONS ABC ALA DGDG MGDG NBD TNBS

ATP-binding cassette aminophospholipid ATPase digalactosyldiacylglycerol monogalactosyldiacylglycerol 7-nitrobenz-2-oxa-1,3-diazol-4-yl trinitrobenzene sulfonic acid

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10â•…

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4 SPHINGOLIPID ASYMMETRY AND TRANSMEMBRANE TRANSLOCATION IN MAMMALIAN CELLS Gerrit van Meer Faculty of Science, Utrecht University, Utrecht, The Netherlands

Sylvia Neumann Department of Cell Biology, Scripps Research Institute, La Jolla, CA

Per Haberkant EMBL, Heidelberg, Germany

4.1 INTRODUCTION Sphingolipids are a typical feature of eukaryotic cells, and indeed, they have been found to fulfill a number of intra- and intercellular functions that are specific for eukaryotes. Membrane sphingolipids are organized in specialized membrane domains [1] that are involved in the sorting of membrane proteins and lipids along the cellular vesicular transport pathways. In addition, the domains have been invoked in various types of signaling events, like the formation of the T-cell receptor complex [2] and the formation of cell–cell signaling domains [3]. On the other hand, individual sphingolipids act as lipid second messengers, the clearest examples being sphingosine-1-phosphate [4] and ceramide [5]. Sphingolipids act at discrete locations, and they are synthesized and degraded at defined locations. These are not always on the same side of the membrane, which necessitates transmembrane transport. The sites of transmembrane translocation, the molecular mechanism, and its possible regulation are the topic of the present chapter (Fig. 4.1). Transmembrane Dynamics of Lipids, First Edition. Edited by Philippe F. Devaux, Andreas Herrmann. © 2012 John Wiley & Sons, Inc. Published 2012 by John Wiley & Sons, Inc.

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Cer

Cer

?

SM

SM

DHSph1P

DHSph

DHSph

Lysosome

DHCer

DHCer Cer

Cer

Sph

SphH+

Sph

Sph

Chol

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Cer

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Sph

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NPC1

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Endosome

Lysosome

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GlcCer GlcCer GlcCer

FAPP2

GlcCer

GlcCer

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GlcCer

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GlcCer

GlcCer

SM

GlcCer SM GM3

ER

Golgi

Cer

PM

Cer

Cer

Cer

Cer

CERT

Figure 4.1.â•‡ Transmembrane translocation of sphingolipids. For a detailed explanation of the various pathways, see the text. SM, sphingomyelin; PM, plasma membrane. Color version on the Wiley web site.

DHSph

3-keto DHSph

palmitoylCoA + Ser

DHSph

Sphingosine, Sphingosine-1-Phosphate, and Ceramide

67

4.2 SPHINGOSINE, SPHINGOSINE-1-PHOSPHATE, AND CERAMIDE The first committed step in sphingolipid synthesis is the condensation of serine and palmitoyl-CoA to 3-ketosphinganine. Its reduction yields sphinganine, which serves as a substrate for various (dihydro)ceramide synthases [6]. Most of the dihydroceramide is subsequently desaturated to ceramide by dihydroceramide desaturase [7]. Actually, most ceramide may be synthesized in the endoplasmic reticulum (ER) directly from sphingosine that is derived from sphingolipid breakdown in the lysosomes, via the salvage pathway [8]. Apart from monomeric transport from the lysosomes to the ER, which is not a problem for the relatively hydrophilic molecule, sphingosine that is generated by ceramidase in the lysosomal lumen must translocate across the lysosomal membrane. It has been reported that sphingosine, which has a pKa of 6.7–7.7 in TX micelles, actually has a pKa of 9.1 in phosphatidylserine-containing membranes [9]. From this, it has been argued that the positively charged sphingosine cannot spontaneously flip out of the lysosome but needs a translocator. Because sphingosine was the first lipid to accumulate in lysosomes upon the induction of a Niemann-Pick type C (NPC) disease phenotype, it has been suggested that NPC1 may be the relevant sphingosine transporter [10]. NPC1 belongs to the heavy metal resistance/nodulation/cell division (RND) permeases, a family of bacterial transporters that are driven by the proton motive force [11]. Instead of transporting a proton (or besides transporting a proton), NPC1 may transport protonated sphingosine toward the cytosolic side. This leaves the interesting possibility that NPC1 countertransports a different substrate in the opposite direction toward the lumen or lumenal leaflet of the lysosome. The fact that defects in NPC2, a soluble cholesterol transporter in the lysosomal lumen, result in the same phenotype as defects in NPC1, which has a cholesterol-binding domain next to a sterol-sensing domain [12], strongly suggests that cholesterol transport out of the lysosome is at least one other function controlled by NPC1 [13]. When added from the outside of the cell, sphingosine must cross the plasma membrane and be transported to the ER to be used in cellular synthesis. Unexpectedly, in yeast, the utilization of exogenous dihydrosphingosine was found to involve phosphorylation and dephosphorylation of the sphingoid base on the lumenal side of the ER [14, 15]. It is unclear whether this has anything to do with translocation of the sphingoid base to the lumenal side of the ER membrane, where the active center of the ceramide synthase is thought to be located [16]: Since all lipids tested have been found to be translocated quickly across the ER membrane [17] by a so far unknown translocator [18, 19] (see Chapter 6), it might be expected that also sphingoid bases freely equilibrate between the two leaflets of the ER membrane. Sphingosine-1-phosphate is not only a metabolic intermediate that can be dephosphorylated by a number of phosphatases, or the final step in sphingolipid degradation by the sphingosine-1-phosphate lyase in the ER membrane

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TRANSMEMBRANE TRANSLOCATION OF SPHINGOLIPIDS

[20]; it is also an important lipid messenger with targets in the cytosol (unidentified) and targets on the outside of cells: the sphingosine-1-phosphate receptors. Because sphingosine-1-phosphate is generated by the cytosolic sphingosine kinases SphK1 and SphK2, it must translocate across the plasma membrane to reach the receptor. It has been reported that this translocation is mediated by the multidrug resistance-related protein MRP1, that is the ATP-binding cassette (ABC) transporter ABCC1 [21], and ABCG2 [22]. Their activity may be regulated by estradiol.

4.3 CERAMIDE After the synthesis of dihydroceramide in the ER, it is desaturated to ceramide or, depending on the cell type, converted to a smaller or larger extent to 4-hydroxysphinganine-containing ceramide (phytoceramide) by C-4hydroxylase, a side activity of a delta 4-desaturase [7, 23, 24]. Dihydroceramide may be produced on the lumenal side of the membrane [16], which also requires that acyl-CoA is translocated. In contrast, the desaturase acts on the cytosolic surface [25]. Translocation of ceramide across the ER is probably fast, because general lipid translocation across this membrane is fast and ceramide has a much smaller polar head group than most other lipids tested. In contrast to regular membrane phospholipids, ceramide spontaneously translocates across model membranes with half-times of less than a minute for natural ceramide [26] and 20 minutes for a fluorescent analog [27]. In contrast to sphingosine, ceramide that is produced in the lysosome is unable to leave the lysosome and mix with the biosynthetic pool of ceramide in the ER: mutations that inactivate the acid ceramidase result in the sphingolipid storage disorder Farber’s disease [28]. In parallel to the observation that mutations in the lumenal cholesterol transport protein NPC2 prevent export of cholesterol from the lysosome, it is most likely the inability of intact ceramide to be transported from the intralumenal vesicles to the limiting membrane of the lysosome that blocks its exit.

4.4 GLYCOSPHINGOLIPIDS In a number of cell types, ceramide is partially converted to galactosylceramide (GalCer) by the ceramide galactosyltransferase on the lumenal aspect of the ER [29], whereas in all cells, ceramide reaches the cytosolic surface of the Golgi where glucosylceramide (GlcCer) is formed. Spin-labeled GlcCer rapidly crossed isolated Golgi membranes in one study [30], and also a GlcCer analog with a short fatty acyl chain, C6-NBD-GlcCer, or with two shortened fatty tails, C8C8-GlcCer, did move across the Golgi membrane [31–33]. This translocation was found to be sensitive to inhibitors of the multidrug trans-

Glycosphingolipids

69

porter MDR1, the ABC transporter ABCB1 [33], which is in line with earlier studies showing that ABCB1 and ABCC1 can translocate these lipid analogs across the plasma membrane [34, 35]. Indirect studies also suggest that natural GlcCer can be translocated across Golgi membranes [36–38], and it has been suggested that the cytosolic glycolipid-binding protein FAPP2, which binds to phosphatidylinositol-4-phosphate on the Golgi, plays a role in this translocation. However, such a translocation of natural GlcCer was not found in a direct experiment. Instead, it was proposed that GlcCer is transported in a retrograde fashion to the ER where it will then translocate spontaneously like all other lipids [33]. In this study, we also found evidence that the pools of natural GlcCer mixed with those of GalCer, and that these lipids can be translocated across a post-Golgi membrane. The latter was based on the observation that newly synthesized GlcCer and GalCer reach the outside of the plasma membrane in the presence of brefeldin A, a drug that inhibited vesicular transport as defined by the complex glycolipid sialyllactosylceramide, GM3 [33]. A special case of GlcCer transport is found in keratinocytes, where GlcCer is stored with other lipids in lamellar granules destined for secretion and extracellular hydrolysis to form the intercellular ceramide lamellae of the skin. This process was found to be interrupted in ichthyosis patients due to mutations in the ABC transporter ABCA12 [39–41], which has therefore been proposed to be a GlcCer translocator or “extruder” [42]. Unfortunately, only a few assays are available to study the transbilayer organization of GlcCer [33, 43], and they have not yet been applied to the keratinocyte system. Interestingly, ceramide regulates ABCA12 expression in the skin via peroxisome proliferatoractivated receptor delta [44]. On the lumenal surface of the Golgi, GlcCer is converted to complex glycosphingolipids. By a sorting event that most likely involves their lateral concentration in microdomains [1], the glycosphingolipids are transported on the inside of carrier vesicles to the plasma membrane where they are exposed on the surface. From there, endocytosis cycles them through the lumen of the endosomes, whereby during each cycle, some glycosphingolipids end up in the lysosomes where they are degraded. A number of studies have reported the presence of complex glycosphingolipids on structures like the cytoskeleton where they might be exposed to the cytosol [45–47]. Unfortunately, these localization studies often involved the use of detergents, which may have induced displacement from their original site of residence [48]. Apart from that, the methodology to study the sidedness of (sphingo)lipids is generally not straightforward and has many potential pitfalls [49, 50]. Whereas under normal conditions complex glycosphingolipids are not found in membranes that are not connected to the exo- and endocytotic pathways like the mitochondria [51, 52], it has been reported that specific glycosphingolipids, for example, GD3, do reach the mitochondria [53–56]. For this, the glycolipid must have translocated across a membrane of the exo/endocytotic system. Where and how this occurs is presently unclear.

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4.5 SPHINGOMYELIN Ceramide destined for the synthesis of sphingomyelin reaches the Golgi not via vesicular transport but via the ceramide transfer protein CERT [57], which, like the glycolipid-binding FAPP2, binds to phosphatidylinositol-4-phosphate on the trans-Golgi membrane. There, ceramide has to cross the Golgi membrane to reach the sphingomyelin synthase SMS1, which is situated with its active center on the lumenal side [58, 59] of the trans-Golgi [33]. It is not fully clear to what extent sphingomyelin is limited to the noncytosolic surface of the Golgi, plasma membrane, and endosomes. Early work indicated that some 10%–20% of the sphingomyelin was situated on the cytosolic surface of the plasma membrane [60, 61], and of chromaffin granules [62]. Unfortunately, essentially only a single method has been applied so far and that is sphingomyelinase C, which is prone to artifacts [63]. Contradictory results have been reported on the effect of brefeldin A on the transport of newly synthesized sphingomyelin to the cell surface, the outside of the plasma membrane. In CHO cells, brefeldin A essentially blocked sphingomyelin transport [64], which implies that (1) sphingomyelin could not translocate across the membrane of the mixed ER–Golgi compartment induced by brefeldin A [65], which is unlikely in view of the fact that GalCer under those conditions does have access to the cytosolic surface and does make it to the cell surface [33]; (2) sphingomyelin cannot cross the aqueous space between ER–Golgi and plasma membrane; and/or (3) sphingomyelin cannot flip across the plasma membrane. Others [66] reported that brefeldin A did not interfere with transport of newly synthesized sphingomyelin to the surface of hepatocytes and claimed that there was a brefeldininsensitive vesicular pathway from the ER–Golgi to the plasma membrane. Although it is possible that sphingomyelin traveled across the aqueous cytosol via a transfer protein (e.g., PITPß [67] ) and was translocated outward across the plasma membrane, it is more likely that in these cells, the sphingomyelin synthase did not redistribute to the ER but remained in a trans-Golgi/ endosome compartment that maintained vesicular traffic to the plasma membrane [68]. During the stress-induced activation of the neutral sphingomyelinase, a large fraction of the plasma membrane sphingomyelin can be degraded. Because the neutral sphingomyelinase has its active center on the cytosolic side [69, 70], either sphingomyelin must be translocated to the cytosolic leaflet of the membrane [71], or sphingomyelin was already present in the cytosolic leaflet and the enzyme itself was activated. Whether this implies the activation of a nonspecific scrambling mechanism [72] or a sphingomyelin-specific process is unclear. Along similar lines, the finding that sphingomyelin can be degraded by a sphingomyelinase that was targeted to the mitochondria has been interpreted to indicate that sphingomyelin reaches this organelle [73, 74], which must involve a translocation step toward the cytosol.

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4.6 FUTURE PERSPECTIVES It is clear that we still have a lack of understanding of a number of aspects of the organization and trafficking of sphingolipids. Notably, the transmembrane distribution and translocation of the basic sphingolipids sphingosine, sphingomyelin, and the monohexosylceramides are enigmatic. This is mostly due to a lack of appropriate technology. It will be a challenge to unravel these issues and to identify the translocators involved. From the fact that these lipids are essential for mammalian survival, it can be predicted that such findings will have their impact on the identification and cure of lipid-linked disorders as has already been shown for the ichthyoses and lipid storage diseases mentioned in the chapter. ABBREVIATIONS C6-NBD- N-6-NBD-aminohexanoylGalCer galactosylceramide GlcCer glucosylceramide REFERENCES â•‡ 1â•…G. van Meer, D. R. Voelker, G. W. Feigenson, Nat. Rev. Mol. Cell Biol. 2008, 9, 112–124. â•‡ 2â•… T. Harder, C. Rentero, T. Zech, K. Gaus, Curr. Opin. Immunol. 2007, 19, 470–475. â•‡ 3â•… A. Regina Todeschini, S. I. Hakomori, Biochim. Biophys. Acta 2008, 1780, 421–433. â•‡ 4â•… S. Pyne, S. C. Lee, J. Long, N. J. Pyne, Cell. Signal. 2009, 21, 14–21. â•‡ 5â•…Y. A. Hannun, L. M. Obeid, Nat. Rev. Mol. Cell Biol. 2008, 9, 139–150. â•‡ 6â•…Y. Pewzner-Jung, S. Ben-Dor, A. H. Futerman, J. Biol. Chem. 2006, 281, 25001–25005. â•‡ 7â•… P. Ternes, S. Franke, U. Zahringer, P. Sperling, E. Heinz, J. Biol. Chem. 2002, 277, 25512–25518. â•‡ 8â•… K. Kitatani, J. Idkowiak-Baldys, Y. A. Hannun, Cell. Signal. 2008, 20, 1010–1018. â•‡ 9â•… F. Lopez-Garcia, V. Micol, J. Villalain, J. C. Gomez-Fernandez, Biochim. Biophys. Acta 1993, 1153, 1–8. 10â•… E. Lloyd-Evans, A. J. Morgan, X. He, D. A. Smith, E. Elliot-Smith, D. J. Sillence, G. C. Churchill, E. H. Schuchman, A. Galione, F. M. Platt, Nat. Med. 2008, 14, 1247–1255. 11â•… T. T. Tseng, K. S. Gratwick, J. Kollman, D. Park, D. H. Nies, A. Goffeau, M. H. Saier, Jr., J. Mol. Microbiol. Biotechnol. 1999, 1, 107–125. 12â•…R. E. Infante, M. L. Wang, A. Radhakrishnan, H. J. Kwon, M. S. Brown, J. L. Goldstein, Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 15287–15292.

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5 TRANSBILAYER MOVEMENT AND DISTRIBUTION OF CHOLESTEROL Peter Müller, Anna Pia Plazzo, and Andreas Herrmann Department of Biology, Humboldt-University Berlin, Berlin, Germany

5.1â•… INTRODUCTION Cholesterol is an important biomolecule of mammalian cells and fulfills various biological functions, for example, membrane biogenesis, steroid hormone and bile salt biosynthesis, and embryonic development. The organism derives cholesterol from dietary sources as well as from de novo synthesis. Synthesis is realized from acetyl coenzyme A in a series of enzymatic steps that are mainly confined to the endoplasmic reticulum [1–3]. Because of the extreme relevance of cholesterol for the organism, its level is highly regulated, and any deviation from its physiological concentration may cause pathological situations. The homeostasis of cholesterol underlies complex processes of trafficking, which include desorption and incorporation from/into membranes, intra- and extracellular transport, and transbilayer movement (see Reference 4). Since cholesterol is poorly soluble in water [5], its transfer across aqueous (intra- and extracellular) compartments is accomplished by transfer proteins and vesicles. Cholesterol constitutes a major component of mammalian plasma membranes. Cell membranes are characterized by a gradient of cholesterol among the different organelles, in which its relative fraction on total lipids increases along the secretory pathway from endoplasmic reticulum to Golgi and to the


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TRANSBILAYER MOVEMENT OF CHOLESTEROL

plasma membrane, resulting in a specific enrichment in the latter. The cholesterol content of a membrane is often given in molar relation to the phospholipid concentration and amounts to about 0.8 in mammalian plasma membranes. Apart from its role in stabilizing the membrane bilayer structure, cholesterol has specific and unique physiological functions associated with membrane organization and dynamics. Examples are its role in forming membrane lipid domains [6, 7], modifying the structure and function of membrane proteins [8, 9], and facilitating the interaction of peripheric proteins with the plasma membrane [10]. 5.2â•… PHYSICOCHEMICAL FEATURES OF CHOLESTEROL Similarly to phospholipids, cholesterol has an amphiphilic structure that supports the formation of a lipid bilayer in aqueous solutions. It inserts into a membrane with its hydroxyl group oriented toward the membrane surface and an orientation of the long axis parallel to the normal of the bilayer [11]. However, in comparison to phospholipids, the proportion between the hydrophilic and hydrophobic part is different, as cholesterol contains a small hydrophilic head group represented by the OH moiety and a large hydrophobic structure consisting of a rigid ring system and a short-branched hydrocarbon chain. Due to these unique features, cholesterol has specific impacts on the structure and dynamics of membranes, as it increases the molecular order of the acyl chains of phospholipids enhancing the lipid packing in the membrane [12, 13]. This condensing effect, which has been shown to be much stronger for saturated phospholipids than for unsaturated lipids [14, 15], modulates many membrane properties and functions such as membrane thickness, fluidity, permeability to polar molecules, and lipid phase transition [16–18]. According to the “umbrella model,” it was suggested that in a bilayer, cholesterol relies on the coverage provided by the larger head groups of phospholipid neighbors [19]. Because of its large hydrophobic portion and rather small polar head group, cholesterol can penetrate deeply into the hydrophobic interior of the membrane. This causes its highly dynamic motion parallel to the membrane normal, whereby the molecule may also protrude into the opposite monolayer [20]. The interaction of the ring system of cholesterol with saturated fatty acyl chains mediated by van der Waals interactions and the formation of hydrogen bonds between the hydroxyl groups of cholesterol and the amide group of sphingolipids and ceramides are supposed to cause, in a mixture of lipids, a preferential association of cholesterol with these phospholipid species. This specific lipid–lipid interaction can trigger a lateral separation of lipids into liquid-ordered and liquid-disordered domains of fluid bilayers, which have been extensively characterized in model membranes [21–27]. A segregation of lipids is assumed to be the molecular basis for the formation of lateral domains

Methods for Measuring Cholesterol Transbilayer Movement

77

in biological membranes, in particular of cholesterol and saturated lipidenriched domains called rafts [6]. Those domains have been implicated to play an important physiological role in protein sorting, lipid trafficking, protein– ligand interaction, and signal transduction across membranes [7, 28–32]. For understanding the complex organization and dynamics of cholesterol in membranes, that is, its (1) distribution among membranes, (2) relative amount within a membrane, (3) lateral organization, and (4) its mobility parallel to the membrane normal in a monolayer, the investigation of the transbilayer distribution and translocation (flip-flop) of cholesterol are of particular relevance. The first studies dealing with the latter topics came out in the mid1970s, and meanwhile, a consistent amount of data achieved by different approaches has been published. 5.3â•… METHODS FOR MEASURING CHOLESTEROL TRANSBILAYER MOVEMENT AND DISTRIBUTION The transbilayer movement of cholesterol across the membrane is one important step of its complex trafficking within the organism. As for phospholipids, the basic mechanisms describing how a cholesterol molecule may transverse the membrane are (1) (passive) diffusion across the lipid bilayer, (2) unspecific diffusion facilitated by the presence of membrane proteins or at the boundary of membrane domains, and (3) active protein-mediated transport. The methods that have been developed to measure transbilayer mobility and distribution of cholesterol typically employ chemically nonmodified cholesterol (endogenous, radioactive) or modified cholesterol/steroid molecules, that is, fluorescent or spin-labeled analogs (Fig. 5.1). The rationale of assays characterizing cholesterol movement is to manipulate the molecule of choice specifically in one membrane leaflet (in general, the extracellular or extravesicular), either by its extraction onto acceptor molecules or by its chemical modification, and to measure the kinetics of redistribution of unaffected molecules from the other leaflet (Fig. 5.2). Thus, experimental kinetics consists optimally of two components, one corresponding to the extraction/modification process and the other to the transbilayer movement. However, for obtaining reliable quantitative data of cholesterol flip-flop, the rate constant of the process of modification kmod or extraction kex has to be at least one order of magnitude larger than that of transbilayer movement ki and ko (see Fig. 5.2); otherwise, one can only obtain estimates of the minimum rate constant of cholesterol flip-flop. This relates to the problem that in case the transbilayer movement of cholesterol is very rapid (in the timescale of seconds or faster), assays providing the appropriate time resolution are still lacking (see below). The modification of cholesterol/steroid molecules in one membrane leaflet and the quantification of cholesterol flip-flop rate is achieved by (1) treatment with cholesterol oxidase (modifying the hydroxyl group) and following the

78


CH3

CH3

CH3

CH3

CH3

OH

CH3

CH3

.ON O

HO

3-doxyl-17b-hydroxy-5a-cholestane (SL-androstane)

cholesterol CH3

CH3

CH3

CH3

CH3

CH3

CH3

CH3

O

CH3

.O

N

.O

N O

HO

25-doxyl-cholesterol

3b--doxyl-5a-cholestane (SL-cholestane)

CH3

+ N

H

CH3

O

N

CH3

N

O

CH3

N

HO

O

N CH3

CH3

O

+ N

CH3

CH3

N

O

-

N O

HO

22-(N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino)-23,24bisnor-5-cholen-3b-ol (22-NBD-cholesterol)

25-(N-[(7-nitrobenz-2-oxa-1,3-diazol-4-yl)-methyl]amino)-27norcholesterol (25-NBD-cholesterol)

CH3 CH3

CH3

CH3

CH3

CH3

CH3

CH3

CH3

CH3

CH3

HO

HO

ergosta-5,7,9(11),22-tetraen-3b-ol (DHE) CH3 CH3

CH3

cholesta-5,7,9,(11)-trien-3b-ol (CTL) CH3

CH3 CH3

HO

1-methyl-10-norcholesta-1,3,5(10)-trien-3-ol (sterophenol)

CH3

CH3

CH3 CH3

HS

5-cholestene-3b-thiol (thiocholesterol)

Figure 5.1.â•‡ Chemical structure of cholesterol analogs used for measuring transbilayer movement and distribution of cholesterol.

increase in concentration of the reaction product; (2) reduction of spin-labeled or fluorescent analogs measuring the decline of intact label by electron spin resonance and fluorescence spectroscopy, respectively; and (3) quenching of fluorescent analogs by Förster resonance energy transfer (FRET) or quenchers, measuring the decrease in fluorescence intensity. For extraction of cholesterol or its analogs from membranes, a variety of different acceptors have been employed such as liposomes, red blood cell (RBC) membranes, lipoproteins,

79

kmod

Cholmod

Y

ko

Choli

Cholo

ki

A

kex

0.0

0.2

0.4

0.6

0.8

1.0

0

20

40 time

60

80

100

Choli

Cholo

Figure 5.2.â•‡ Principles of assaying transbilayer movement of cholesterol. Cholesterol molecules or analogs in the inner (Choli) and in the outer (Cholo) leaflet move to the opposite leaflet with time constants ko and ki, respectively. Molecules on the outer leaflet are specifically manipulated by chemical modification into Cholmod with a time constant kmod or by extraction onto an extracellular acceptor (A) with a time constant kex. X and Y are the cholesterol-modifying reactant before and after reaction with cholesterol, respectively. On the right, a typical time course-dependent decrease of signal intensity is shown consisting of a fast component (representing the modification/extraction of molecules localized on the outer membrane leaflet) and a slow component (representing the modification/extraction of molecules originally localized on the inner membrane leaflet).

inside

outside

Cholo

X

signal intensity

80


bile acids, and cyclodextrins. A combination of steroid modification and extraction has also been employed [33]. Notably, the release (extraction) of cholesterol onto extracellular lipoproteins is an important physiological mechanism by which cells export cholesterol (see below). Transbilayer distribution of cholesterol can be measured provided the assays applied fulfill certain conditions. The transbilayer distribution of cholesterol can be deduced from the translocation kinetics. If the rate constants of modification/extraction are very different from those of translocation process, that is, kmod (or kex)â•¯>>â•¯ki and kmod (or kex)â•¯>>â•¯ko (see Fig. 5.2), the distribution can be estimated from the first plateau-like intensity of the respective signal. This plateau is reached after the initial rapid manipulation of all cholesterol (analogs) in the accessible leaflet. Subsequently, the signal intensity decreases only slowly since the redistribution of cholesterol (analogs) from the nonaccessible to the accessible leaflet is slow. In case kmod (or kex)â•¯>â•¯ki and >ko, values for transbilayer distribution can be obtained by fitting the experimental data to an underlying mathematical model [34]. However, if kmod (or kex)â•¯∼â•¯ki and ∼ko or even kmod (or kex)â•¯

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