Advances in Protein Chemistry, Volume 45: Lipoproteins, Apolipoproteins, and Lipases

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ADVANCES IN PROTEIN CHEMISTRY Volume 45

Lipoproteins, Apolipoproteins, and Lipases

This Page Intentionally Left Blank

ADVANCES IN PROTEIN CHEMISTRY EDITED BY C. 6. ANFINSEN

JOHN T. EDSALL

Department of Biology The Johns Hopkins University Baltimore, Maryland

Department of Biochemistry and Molecular Biology Harvard University Cambridge, Massachusetts

FREDERIC M. RICHARDS

DAVID S. EISENBERG

Department of Molecular Biophysics and Biochemistry Yale University New Haven, Connecticut

Department of Chemistry and Biochemistry University of California, Los Angeles Los Angeles, California

VOLUME 45

Lipoproteins, Apol ipoproteins, and Lipases EDITED BY VERNE N. SCHUMAKER University of California Los Angeles, California

ACADEMIC PRESS, INC. A Division of Harcourt Brace & Company San Diego New York Boston London Sydney Tokyo Toronto

This book is printed on acid-free paper. @

Copyright 0 1994 by ACADEMIC PRESS,INC. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher.

Academic Press, Inc. 525 B Street, Suite 1900, San Diego, California 92101-4495

United Kingdom Edition published by

Academic Press Limited 24-28 Oval Road, London NW 1 7DX

International Standard Serial Number: 0065-3233 International Standard Book Number: 0-12-034245-6 PRINTED IN THE UNITED STATES OF AMERICA 9495

9 6 9 7 9 8 9 9

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1

CONTENTS CONTRIBUTORS PREFACE

.

.

ix xi

Structure and Function of Lipases

ZYCMUNTS. DEREWENDA

Introduction . Lipases: Crystallographic Database . Molecular Basis of Liquid Degradation by Lipases Evolutionary Relationships . Lipases with Unknown Three-Dimensional Structures . VI. Conclusions . References .

I. 11. 111. IV. V.

.

1 3 10 29 38 46 46

Structure and Catalytic Mechanism of Secretory PhospholipasesA2

DAVID L. Scorr AND PAULB. SICLER I. Introduction

.

11. Primary Structure and Classification 111. Secondary and Tertiary Structure

.

.

IV. Chemistry of Catalysis of Secretory Phospholipases A2 V. Interfacial Catalysis . References .

.

53 54 58 66 75 80

Lipid-Binding Proteins:A Family of Fatty Acid and Retinoid Transport Proteins

LEONARD BANASZAK, NATHANWINTER, ZHAOHUIXu, DAVIDA. BERNLOHR, SANDRA COWAN, AND T. ALWYNJONES

I. Introduction . 11. General Structural Features

. V

90 92

vi

CONTENTS

111. Conformational Similarity among Intracellular

IV. V. VI. VII. VIII.

IX. X. XI. XII.

Lipid-Binding Proteins . Intracellular Lipid-Binding Protein Cavity . Apo- versus Holo-iLBP Structure . Gap between PD and PE Binding Affinity and Specificity Based on Biochemical Studies . Binding Affinity and Specificity Based on . Crystallographic Studies Members of iLBP Family with Known Crystal Structure . Members of eLBP Family with Known Crystal Structure . Factors Involved in Lipid Binding in iLBPs . Concluding Remarks . References .

99 110 114 116 119 122 126 137 141 145 148

Structure of Serum Albumin

DANIEL C. CARTER AND JOSEPH X. Ho

I. Introduction

.

11. Albumin Structure 111. Nature of Ligand Binding

.

IV. Evolution of Albumin Structure V. Summary and Future Directions References .

153 155 176 190 194 196

. .

Apolipoprotein B and Low-Density Lipoprotein Structure: Implications for Biosynthesis of Triglyceride-Rich Lipoproteins VERNEN.SCHUMAKER, MARTINL. PHILLIPS, AND JON E. C H A ~ R T O N

I. Introduction

.

11. Apolipoprotein B Structure . 111. Low-Density Lipoprotein Structure

.

IV. Structural Studies of Apolipoprotein B on Low-Density Lipoprotein Surfaces . V. Lipoprotein Assembly . VI. Summary . References . * :

205 207 2 13 226 240 243 244

vii

CONTENTS

Apolipoprotein E: Structure-Function Relationships KARL H. WEISCRABER

Introduction . Apolipoprotein E . Function . Impact of Structure on Function . Three-Dimensional Structure of Apolipoprotein E 22-kDa Fragments . VI. Lipid Binding . VII. Future Directions . References . I. 11. 111. IV. V.

249 25 1 260 277 285 290 294 295

The Amphipathic a Helix: A MultifunctionalStructural Motif in Plasma Apolipoproteins

P. SECREST,DAVID W. GARBER, CHRISTIE G. BROUILLEITE, AND G. M. ANANTHARAMAIAH STEPHEN C. HARVEY, 303 I. Plasma Lipoproteins and Apolipoproteins . 11. The Amphipathic a Helix . 309 111. Conclusions . 363 References . 363 JERE

Lipophorin:The Structure of an Insect Lipoprotein and Its Role in Lipid Transport in Insects

Jost L. SOULACES AND MICHAELA. WEI. Introduction . 11. Lipid and Apolipoprotein Composition of Lipophorins . 111. Size, Molecular Weight, Heterogeneity, and Shape of Lipophorins . IV. Organization of Lipids and Proteins in Lipophorins V. Metabolism . VI. Metabolic Implications of Lipophorin Structure . VII. Concluding Remarks and Future Directions . References .

37 1 372

.

384 388 393 405 408 409

AUTHOR INDEX

417

SUBJECT INDEX

45 1


CONTRIBUTORS Numbers in parentheses indicate the pages m which the authors’ conhibutions begin.

G. M. ANANTHARAMAIAH (303), Departments of Medicine and Biochemistry, Atherosclerosis Research Unit, University of Alabama at Birmingham Medical Center, Birmingham, Alabama 35294 LEONARD BANASZAK (89), Department of Biochemistry, University of Minnesota, Minneapolis, Minnesota 55455 DAVIDA. BERNLOHR (89), Department of Biochemistry and Institute of Human Genetics, University of Minnesota, St. Paul, Minnesota 55455 CHRISTIE G. BROUILLETTE (303),Southern Research Institute, Birmingham, Alabama 35205 DANIELC. CARTER (153), Space Science Laboratory, Biophysics Branch, National Aeronautics and Space Administration, Marshall Space Flight Center, Huntsville, Alabama 358 12 JON E. CHATTERTON (205), Department of Chemistry and Biochemistry, and the Molecular Biology Institute, University of California, Los Angeles, California 90024 SANDRA COWAN (89), Department of Molecular Biology, Biomedical Center, Uppsala University, S-75 124 Uppsala, Sweden ZYCMUNTS. DEREWENDA (l), MRC of Canada Group in Protein Structure and Function, Department of Biochemistry, University of Alberta, Edmonton, Alberta, Canada T6G 2H7 DAVID W. GARBER (303), Departments of Medicine and Biochemistry, Atherosclerosis Research Unit, University of Alabama at Birmingham Medical Center, Birmingham, Alabama 35294 STEPHENC. HARVEY (303), Department of Biochemistry, University of Alabama at Birmingham Medical Center, Birmingham, Alabama 35294 JOSEPH X. Ho (153), Space Science Laboratory, Biophysics Branch, National Aeronautics and Space Administration, Marshall Space Flight Center, Huntsville, Alabama 358 12 T. ALWYN JONES (89), Department of Molecular Biology, Biomedical Center, Uppsala University, S-75 124 Uppsala, Sweden MARTINL. PHILLIPS (205), Department of Chemistry and Biochemistry, and the Molecular Biology Institute, University of California, Los Angeles, California 90024 VERNEN. SCHUMAKER (205), Department of Chemistry and Biochemistry, and the Molecular Biology Institute, University of California, L o s Angeles, Los Angeles, California 90024 ix

CONTRIBUTORS

X

DAVIDL. Scorn (53), Department of Molecular Biophysics and Biochemistry, Howard Hughes Medical Institute, Yale University, New Haven, Connecticut 065 10 JEREP.SECREST (303),Departments of Medicine and Biochemistry, Atherosclerosis Research Unit, University of Alabama at Birmingham Medical Center, Birmingham, Alabama 35294 PAULB. SICLER (53), Department of Molecular Biophysics and Biochemistry, Howard Hughes Medical Institute, Yale University, New Haven, Connecticut 065 10 Josk L. SOULACES (371), Department of Biochemistry and Center for Insect Science, Biological Sciences West, University of Arizona, Tucson, Arizona 8572 1

KARL H. WEISGRABER (249), Gladstone Institute of Cardiovascular Disease, San Francisco, California 94 14 1 MICHAELA. WELLS(371), Department of Biochemistry and Center for Insect Science, Biological Sciences West, University of Arizona, Tucson, Arizona 8572 1 NATHAN WINTER(89), Department of Biochemistry, University of Minnesota, Minneapolis, Minnesota 55455 ZHAOHUI Xu (89), Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, Connecticut 06510

PREFACE

This volume contains eight beautifully written, well-illustrated chapters that present both new and reviewed information fundamental to a clear understanding of lipid catabolism and transport at the molecular level. Three-dimensional structures of important serum lipoproteins, apolipoproteins, and lipases, utilizing X-ray data when available, are emphasized, and an attempt is made to relate structures to functions. Among this rich fare, Zygmunt Derewenda has contributed the opening chapter on the triglyceride lipases. Lipid metabolism begins with a meal, and after a preliminary digestion in the stomach, the gastric contents are sprayed into the intestine where dietary lipids combine with the biliary secretion to form an emulsion providing a large lipid-aqueous interface where lipases may act. Triglyceride hydrolysis is accomplished principally by pancreatic lipase, which, together with colipase, binds to the interface where it has access to the lipid. Derewenda describes the three-dimensional structure of this enzyme as well as the structures of several fungal lipases; both the common features and the structural differences provide insight into the functional mechanisms of all of these enzymes. The chapter also describes the currently available structural database and provides fascinating insights into the mechanism of interfacial activation and ester bond hydrolysis by this group of lipases. In the second chapter, phospholipases are reviewed by David Scott and Paul Sigler. In the gut, phospholipids are hydrolyzed by a variety of lipases, but principally by pancreatic phospholipase AS. Over sixty members of this group of enzymes have been studied, and features of particular relevance to catalysis are emphasized by Scott and Sigler who develop from high resolution crystallographic analyses a detailed examination of secondary and tertiary structures. A reaction pathway is proposed, including substrate binding and the formation of a complex possessing highly specific interactions between the substrate, calcium, and active site residues and the formation and collapse of the tetrahedral intermediate. The chapter concludes with a detailed discussion of catalysis at an interface, “hopping” or “scooting,” and the interfacial adsorption surface. This insightful contribution provides a paradigm for the mechanism of phospholipase activity. The extensive, multigenic family of fatty acid and retinoid binding proteins is clearly described by Leonard Banaszak and co-workers in their elegant third chapter. The products of hydrolysis of the intestinal lipids, including fatty acids, cholesterol, monoglycerides, and lysophosxi

xii

PREFACE

pholipids, have very low solubilities and are absorbed by biliary micelles in the gut. These micelles diffuse through the glycocalyx,which stabilizes an unstirred water layer at the surface of the enterocyte. The enterocytes are a single layer of cells which line the villi of the intestinal cavity and absorb all products of digestion. Transfer of fatty acids from the micelle to the membrane occurs principally by passive diffusion and direct absorption. Once inside the enterocyte, the fatty acids are bound to intracelMar fatty acid binding proteins. These proteins are found in all cell types and abundantly within the enterocyte and hepatocyte, facilitating the solubilization and transport of the lipid substrates of metabolism. This third chapter focuses on structural analyses and comparisons between members of this multigenic family, primarily emphasizing the intracellular lipid binding proteins. It discusses the structural motif, general characteristics of the binding cavity, ligand entry, and the portal hypothesis and provides a detailed comparison of intra- and extracellular lipid binding proteins with known crystal structures. The chapter concludes with a discussion of the results of site-directed mutagenesis studies, the thermodynamics of lipid binding, and considerations of protein stability and folding. The fourth chapter provides the first review of the high resolution X-ray structure of serum albumin by the scientists who performed these very important studies, Daniel Carter and Joseph Ho. In contrast to the enterocyte which receives lipids directly from the gut, the hepatocyte, the characteristic cell of the liver, receives lipids from a variety of sources. Within the liver new fatty acids may be synthesized from dietary carbohydrates, particularly when present in excess. In addition, the liver removes most of the chylomicron and VLDL remnants and LDL from the blood, which are then hydrolyzed in the lysosome providing an abundant source of fatty acids. Dietary short-chain fatty acids absorbed by the enterocytes diffuse to the intestinal capillaries where they can be bound to serum albumin. Long-chain free fatty acids released from adipose tissue and those from the stomach released by gastric lipase are also transported to the liver bound to serum albumin. Because it is cheap, readily available, binds a wide range of ligands, and plays a central role in the transport of fatty acids and many pharmaceutical agents, serum albumin has been one of the most extensively studied of all proteins. But crystals of diffraction quality have been difficult to obtain. Recently, the high resolution structure has been determined by Carter and his colleagues, and this beautifully written chapter correlates the very extensive and significant studies of the past with the current three-dimensional structure of serum albumin. Domain structure and the arrangement of the disulfides, the surface charge distribution, and the conformational flexibility of the

PREFACE

...

xu1

albumin molecule are described in detail. The nature of ligand binding, including small organics, long-chain fatty acids, and metals, to multiple sites on the albumin molecule is clearly depicted. Comparative studies and the evolution of albumin structure are described next, and the chapter concludes with perceptive comments on future directions being taken to explore the structure and function of this fascinating protein. Apolipoprotein B, which plays an essential role in lipoprotein biosynthesis, is reviewed in the fifth chapter by Verne Schumaker, Martin Phillips, and Jon Chatterton. The intestines and liver are the two organs which synthesize and secrete lipoproteins. Chylomicrons and very low density lipoproteins (VLDL), the characteristic lipoproteins secreted by intestine and liver, respectively, function to transport nonpolar lipids, principally triglyceride, from these organs to the other tissues of the body. Cholesteryl esters are transported by the low density lipoproteins (LDL), which are derived from the VLDL during their metabolism, and also by high density lipoproteins (HDL). Elevated plasma concentrations of LDL are directly associated with risk for atherosclerosis, while HDL apparently exert a protective effect. Lipoprotein biosynthesis occurs in the lumen of the endoplasmic reticulum, and from there the newly synthesized lipoproteins proceed through the regular secretory pathway including the Golgi, where carbohydrate processing occurs and where they are packed into secretory vesicles. The secretory vesicles migrate to and fuse with the plasma membrane, discharging their contents into the intestinal lymph or hepatic blood. Apolipoprotein B (apoB) is the protein essential for biosynthesis and secretion of the triglyceride-rich chylomicrons and VLDL. Both the molecular biology and chemistry of this critical protein are reviewed in the chapter by Schumaker, Phillips, and Chatterton, who then proceed to develop a detailed model for the LDL, relating size to physical properties and composition. The distribution of apoB on the surface of the low density lipoproteins appears to be that of a belt, engirding the LDL about the equator. The beltlike structure also seems to be a critical feature of lipoprotein assembly for in studies involving expression of C-terminally truncated fragments the size of the apoB apparently determines the circumference of the triglyceride-filled core of the primary lipoprotein particle. Lipoprotein assembly is cotranslational, thus apoB and nonpolar lipids dissolved in the membrane are packaged together to form the nascent lipoprotein, while the still growing apoB polypeptide is translocated through the bilayer of the rough endoplasmic reticulum. The intestinal lymph containing the secreted chylomicrons flows into the thoracic duct where it ascends from the gut to join the great veins of the head and neck. The VLDL are secreted by the liver into the space of

XV i

PREFACE

Disse, plasma-filled cavities separated from the flowing blood by a fenestrated membrane through which the VLDL diffuse. Both the chylomicrons and VLDL contain the single molecule of apolipoprotein B essential for their formation; apolipoprotein B is often referred to as an “integral” apolipoprotein since it does not transfer to other lipoproteins but remains associated with the same particle throughout its life span. Chylomicrons and VLDL also acquire additional “peripheral” apolipoproteins, especially apolipoproteins A-I and A-IV, during biosynthesis and later, after secretion, by transfer from the high density lipoprotein fraction which contains an abundant supply of the peripheral apolipoproteins A-I, C-I, C-11, C-111, and E. The triglyceride-rich chylomicrons and VLDL are metabolized by lipoprotein lipase, an enzyme attached to the glycocalyx lining the lumenal surface of cells forming the capillaries which supply tissues throughout the body with nutrients and oxygen. Apolipoprotein C-I1 is an essential cofactor in this hydrolysis. Lipoprotein lipase belongs to the multigene family which includes pancreatic lipase and hepatic lipase, enzymes to which it is closely related, as discussed in the first chapter. Free fatty acid is abundantly released through hydrolysis by lipoprotein lipase and serves to feed the adjacent tissues passively diffusing across cellular membranes and being transported through aqueous spaces both by intracellular binding proteins and extracellular albumin. During this hydrolytic process excess lipoprotein surface molecules, including phospholipids, cholesterol, and the peripheral apolipoproteins, are transferred to the HDL particles. The original chylomicron or VLDL becomes a much smaller “remnant” particle during this hydrolysis by lipoprotein lipase, and acquires a substantial quantity of apolipoprotein E (apoE). This protein possesses a site complementary to and recognized by the B/E receptor and also by the lipoprotein remnant receptor, the two receptors in mammalian liver which remove the lipoprotein remnants from the circulation. The function of apoE in lipoprotein metabolism is reviewed in the sixth chapter by Karl Weisgraber. The three-dimensional structure of a 22-kDa fragment of human apoE (34.2 kDa) has been solved by X-ray crystallography; the relation of this structure to the role of apoE in lipoprotein metabolism is discussed in detail, together with a critical and extensive examination of the chemistry and biology of this apolipoprotein which plays such a central role in lipoprotein metabolism. Apolipoprotein E has three major isoforms in the human population which affect lipoprotein metabolism differently, resulting in different levels of the plasma lipoproteins. The impact of structure on function and how plasma lipid concentrations are affected by the different apoE isoforms are the themes of this important chapter.

PREFACE

xv

The seventh chapter by Jere Segrest and colleagues reviews the amphipathic helix, the dominant structural motif of the peripheral apolipoproteins found abundantly in the high density lipoprotein fraction. High density lipoproteins play multiple roles in lipid metabolism, and plasma levels are inversely correlated with risk for atherosclerosis. The excess surface lipids generated during lipolysis of the triglyceride-rich chylomicrons and VLDL enter the HDL density class where they are converted to cholesteryl esters via the lecithin cholesterol acyl transferase (LCAT) reaction. HDL also function in the removal of cholesterol from peripheral tissues, catalyzing the conversion of this cholesterol to cholesteryl ester via the LCAT reaction, and then in the transfer of the cholesteryl esters to VLDL remnants for their return to the liver. Many peripheral apolipoproteins are found within the HDL class, although apolipoproteins A-I and A-11, which play structural as well as functional roles, comprise over 90% of the HDL proteins. Abundant quantities of apoC-I, apoC-11, apoC-111, apoE, and some apoA-IV are present in the human HDL fraction where they are available for transfer to nascent chylomicrons and VLDL to serve essential metabolic roles as cofactors, regulators, and ligands for lipoprotein receptors. The dominant structural motif of the peripheral apolipoproteins is the amphipathic helix which is responsible for the reversible association of these proteins with lipids, as well as for many biological functions mediated by these apolipoproteins. In their well-illustrated chapter, Segrest and his colleagues review the different classes of amphipathic helices, using a combination of powerful computer programs to develop a comparison database and to analyze these structures. Then the chapter proceeds to review their evolutionary origins, their physical-chemical properties, X-ray structure determination and conformational analysis, and a variety of structure-function studies, including the activation of lipoprotein lipase, receptor recognition, LCAT activation, and antiviral and anti-inflammatory activities. A fascinating and fundamentally very different way of organizing a lipid transport system is provided by insect lipoproteins which have been carefully studied to provide an interesting and rewarding contrast to the mammalian lipid transport system. In insects, a single lipoprotein, lipophorin, acts as a reusable shuttle to transfer diglycerides between the fat bodies, where they are stored as triglycerides, and the peripheral tissues. In their delightful review of this wondrous system, Jose Soulages and Michael Wells describe the composition and the organization of lipids and proteins in the lipophorins and the structure of apolipophorin-111 which has been determined by X-ray crystallography to 2.5 A. Lipophorin is reloaded with lipid by another remarkable protein, the lipid trans-

xvi

PREFACE

fer particle, which can effect a net transfer of diglycerides from the fat bodies to the depleted lipophorin particles. The major progress which has been achieved in describing the lipid transport system in insects provides results of significance for understanding lipid transport in general, an illustration of comparative biochemistry at its best. VERNE N. SCHUMAKER

STRUCTURE AND FUNCTION OF LIPASES By ZYQMUNT 5. DEREWENDA MRC of Canada Group in Protein Structure and Functlon, Dopartment of Blochemlatry, University of Alberta, Edmonton, Alberta, Canada T6G 2H7

I. Introduction

.................................................................................................... ................................................................

11. Lipases: Crystallographic Database

Ill.

IV.

V.

VI.

A. Fungal Lipases ......................................................................................... B. Human Pancreatic Lipase ........................................................................ Molecular Basis of Lipid Degradation by Lipases .......................................... A. Hydrolysis of the Ester Bond .................................................................. B. Molecular Basis of Substrate Specificity .................................................. C. Molecular Mechanism of Interfacial Activation ...................................... Evolutionary Relationships ............................................................................. A. Structure of the P-eSer-a Motif ............................................................... B. Tertiary Fold of alp Hydrolases .................................................. C. Lipases and Proteinases: Possible Relationships ..................................... Lipases with Unknown Three-Dimensional Structures ................................. A. Prokaryotic Enzymes ............................................................................... B. Other Fungal Lipases .............................................................................. C. Human Lipases Involved in LipidlLipoprotein Metabolism .................. Conclusions ..................................................................................................... References .......................................................................................................

1 3 6 9 10 10 18 20 29 30 33 37 38 39 39 40 46 46

I. INTRODUCTION Lipases (acylglycerol acylhydrolases, EC 3.1.1.3) constitute one of the structurally less well-characterized families of hydrolases. They hydrolyze the ester bonds in tri-, di-, and monoacylglycerols, although some will degrade a fairly broad range of compounds containing an ester linkage. A characteristic property common to all lipases is their behavior in a heterogeneous medium in which the enzyme functions almost exclusively at the oil-water interface. Adhered to the interface, the enzyme displays a much higher activity than in the aqueous phase (where in many cases the activity is immeasurably low), and accordingly the activity against water-soluble substrates is negligible. This phenomenon is known as interfacial activation and has also been observed among phospholipases. Lipases are ubiquitous enzymes playing a pivotal role in all aspects of fat metabolism. In humans and other vertebrates a variety of lipases control the digestion, absorption, and reconstitution of fat, as well as lipoprotein metabolism (Desnuelle, 1986). In plants, for example, liADVANCES IN PROTEIN CHEMISTRY. Vol. 45

1

Copyright Q1994 by Academic Press. Inc. All rights of reproduction in any form reserved.

2

ZYCMUNT S. DEREWENDA

pases are abundant in energy reserve tissues (Hassanien and Mukherjee, 1986). Microorganisms are also known to produce a wide spectrum of extracellular lipid-degrading enzymes (Lie et al., 199 1). The recent interest in lipases is stimulated to some extent by their industrial potential (Harwood, 1989). Many of these enzymes withstand exhaustive drying and continue to be active in organic solvents, in which, in the absence of water, they catalyze several potentially useful reactions, including transesterification, regioselective acylation of glycols and menthol, and even synthesis of peptides (e.g., Margolin and Klibanov, 1987; Lagrand et al., 1988; West and Wong, 1986; Valivety et al., 1992). Until recently relatively little was known about the molecular basis of lipid hydrolysis. The first amino acid sequence of a triacylglycerol lipase was given by De Caro et al. (1981). As more sequence data became available, it was noted that lipases and esterases share a short consensus sequence, G-X-S-X-G (Boel et al., 1988; Datta et al., 1988; Antonian, 1988; Brenner, 1988). The role of the invariant serine at the center of this sequence was debated (Maraganore and Heinrikson, 1986). Some authors speculated about a “lipid recognition site,” others compared this pentapeptide to the sequence around the nucleophilic serine in serine proteinases. Conflicting hypotheses were also put forward with respect to the mechanism of interfacial activation. Desnuelle et al. (1960) were the first to suggest that a conformational change in the enzyme could be responsible for the enhancement of activity at the oil-water interface. There were also other hypotheses. For example, Wells (1974) suggested that the apparent activation of lipases is due to the orientation of the scissile ester bond on the surface of micelles; Brockerhoff (1968), on the other hand, pointed to the possibility of differences in solvation of the ester bond in solution versus a lipid phase, whereas Brockman et al. (1973) postulated that a steep substrate concentration gradient at the interface may provide an explanation. Clearly, high-resolution three-dimensional structures were needed to address these problems. The first crystallographic study of a lipase is credited to Hata et al. (1979), who investigated the structure of a fungal enzyme (GcL)’ from Geotrichum candzdum, although at a very low resolu-

’

Abbreviations: GcL, Geotrichum candidurn lipase; hPL, human pancreatic lipase; RmL, R h i m w o r michei lipase; hHL, human hepatic lipase; hLPL, human lipoprotein lipase; hLAL, human lysosomal acid lipase; hGL, human gastric lipase; BAL, bile salt-activated lipase; HSL, hormone-sensitive lipase; CLP, colipase; AChE, Torpedo californica acetylcholinesterase: cDNA, complementary deoxyribonucleic acid; VLDL, very low-density lipoprotein: IDL, intermediate-density lipoprotein; HDL, high-density lipoprotein; apoC11, apolipoprotein C-11.

LIPASE STRUCTURE/FUNCTION

3

tion (581). It was not until 1990, however, that two independent groups reported high-resolution refined crystal structures of two unrelated lipases: one (RmL) purified from a fungus, Rhzromucor miehei (Brady et al., 1990), and the other (hPL) from human pancreas (Winkler et al., 1990). Both enzymes were shown to contain triads of Ser, His, and Asp, reminiscent of those found in the subtilisin and chymotrypsin families of serine proteinases (Blow et al., 1969). Their catalytic function was soon confirmed by site-directed mutagenesis experiments on several enzymes (e.g., Lowe, 1992; Yamaguchi et al., 1992). In contrast to the proteinases, however, these putative catalytic centers were not exposed to solvent, but were instead buried under surface loops, immediately suggesting that significant conformational changes must precede the catalytic event. The crystal structure of a fungal lipase, that of GcL, has also been elucidated at high resolution (Schrag et al., 1991). In addition to these native structures, crystallographic analyses of two complexes of RmL with covalently bound phosphorus-based inhibitors revealed the conformational changes in the enzyme associated with interfacial activation (Brzozowski et al., 1991; U. Derewenda et al., 1992). The human pancreatic enzyme has been successfully studied in a complex with its cofactor colipase (van Tilbeurgh et al., 1992), and extensive conformational changes in the enzyme were subsequently observed when the crystallization was carried out in the presence of mixed phospholipid/ bile salt micelles (van Tilbeurgh et al., 1993). The present-review assesses at some length current progress in crystallographic studies of neutral lipases and the impact these studies have had on our understanding of the structure-function relationships in lipolytic enzymes in general [short reviews have been given by Smith et al. (1992), Lawson et al. (1992), and Derewenda and Sharp (1993)l. The main objective of this paper is to familiarize the reader with the currently available structural database generated by crystallographic analyses. The structural basis of the enzymatic hydrolysis of the ester bond and activation of lipases at an oil/water interface will be described. Possible relationships between lipases and other serine hydrolases will be discussed in the context of structural comparisons. Finally, it will be shown that even the relatively limited structural database available to date may have a considerable impact on further research into the properties of those lipases whose structures are not known at the present time. 11. LIPASES: CRYSTALLOGRAPHIC DATABASE

Table I includes the crystallographic details of all X-ray studies on lipases completed through June, 1993. All native structures were deter-

TABLE I Structural Database of Neutral L i m e s and Related Enzymes

Resolution Structure Native lipases Rhimmucor muhei lipase

*

Humuoh lanupnosa lipase"

Pauillium camembed lipase

Geotrkhum candidum lipase

Crystal data P212121,one molecule in the AU"; a = 71.6, b = 75.0, c = 55.0 A P212121,one molecule in the AU; a = 103.2, b = 52.0, c = 47.7 A P I , one molecule in the AU; a = 45.6, b = 47. I , c = 33.5 A; a = 79.5", B = 112.1", y = 70.3" P21, one molecule in the AU; a = 59.4, b = 84.0, c = 56.0 A;

Lipid-degrading esterase Ftuarium solani cutinase

1.9

R factor

(%I 12.9

Ref. Brady et al. (1990)

1.8

-17.0

Author's laboratory

2.1

-17.0

Author's laboratory

2.2

19.5

Schrag et al. (1991)

P21, two molecules in the AU; a = 47.8, b = 112.8, c = 91.0 A; /3 = 99.3"

2.3

21.9

Winkler et al. (1990)

P2 I , one molecule in the AU; a = 35.12, b = 67.3, c = 37.05

1.6

15.4

Martinez et al. (1992)

p Human pancreatic lipase

(4

= 100.1"

A; 0 = F)J.!,"

u1

Complexes of lipases with inhibitors or cofactors Rhizomwor miehei with nC2221, one molecule in the hexyl phosphonate A U ; a = 48.3, b = 93.9, ethyl ester c = 122.1 A C222 I , one molecule in the Rhiwmwor miehei with A U ; a = 48.6, b = 93.9, diethyl p-nitrophenyl phosphate c = 122.1 A hPL with procolipase Ps221, one complex in the AU; a = b = 80, c = 251 8, P42212, one complex in the AU; hPL with procolipase in a = b = 133.4, c = 92.6 8, the presence of mixed rnicelles in the active conformations Other enzymes exhibiting homology andlor structural similarities Haloalkane dehalogenase P21212,one molecule in the AU; a = 94.8, b = 72.8, c = 41.4 8, Dienelactone hydrolase F'212121. one molecule in the AU; a = 48.9, b = 71.5, c = 78.2 8, Acetylcholinesterase P3,2 1, one molecule in the A U ; a = b = llO,c= 1358, Wheat serine P41212,one molecule in the AU; a = b = 98.9, c = 209.5 A carboxypeptidase a

AU. asymmetric unit.

3.0

18.5

Brzozowski et al. (1991)

2.6

16.2

U . Derewenda et al. (1992)

3.0

23.0

van Tilbeurgh et al. (1992)

3.0

18.6

van Tilbeurgh et al. (1993)

2.4

17.9

Franken et al. (1991)

1.8

15.0

Pathak and Ollis (1990)

2.8

18.6

Sussman etal. (1991)

2.2

16.7

Liao et al. (1992)

6

ZYGMUNT S. DEREWENDA

mined at high resolution and have been subject to extensive crystallographic refinement, but it is easy to see that this database is fairly limited. Four of the enzymes studied so far are extracellular fungal proteins, whereas human pancreatic lipase is the only known representative of enzymes from higher eukaryotes. It is noteworthy, however, that there are several reports of successful crystallizations of other lipases (e.g., Cleansby et al., 1992; Kordel et al., 1991; Kim et d.,1992); this offers hope that the database will soon expand. The review of the available structural information begins with a description of the molecular architecture of the fungal enzymes. A . Fungal Lipases I . Three-Dimensional Structure of a Lipase from a Filamentous Fungus (Rhizomucor miehei)

This extracellular enzyme is a relatively small (269 amino acids) single polypeptide chain and one-domain protein. The gene coding for it has been cloned and a cDNA-derived amino acid sequence is available (Boel et al., 1988). RmL crystallizes readily from high concentrations of phosphate buffers and yields orthorhombic crystals (see Table I) of high quality, diffracting to beyond 1.9 A. A detailed description of the RmL structure appears elsewhere (Z. S. Derewenda et al., 1992); an overview is presented here. All but the first five residues of RmL have been positioned in the 1.9-A electron density map. The molecule adopts an uncommon fold that consists of a sequential, predominantly parallel, singly wound ninestranded fl sheet, with all connecting fragments showing the classic right-handed twist, and six main (Y helices (Fig. 1). One face of the sheet is adjacent to a single N-terminal helix and the other face makes numerous interactions with most of the loops, turns, and interstrand connections. This results in unique asymmetry of the molecule. Three disulfide bridges stabilize the fold and link the following pairs of cysteines: 29-268, 40-43, and 235-244. Four cis peptide bonds are present in RmL and they all precede prolines. A number of hydrophilic internal cavities containing ordered solvent molecules were found. This is an interesting observation, particularly in the light of the GcL structure (see below), in which Schrag et al. (1991) also note the presence of a number of internal water molecules. Whether these internal hydrophilic cavities play some role in the stability of lipases in the hydrophobic environment, or perhaps ensure the availability of water for hydrolysis, remains to be seen.


7

FIG. 1. The molecular structure of RmL: the /3 strands are shown as lightly shaded arrows, helices are depicted as darker cylinders, and the three residues that constitute the catalytic triad are shown in full using the ball-and-stick convention and are labeled. The lid covering the active site is seen at the top.

The catalytic site in RmL was identified originally from the location of the known lipasejesterase consensus sequence G-X-S-X-G (Brenner, 1988)containing the nucleophilic serine (Ser-144). This amino acid was found to be involved in a hydrogen-bonded constellation also including His-257 and Asp-203. Overall this hydrogen-bonding network is very reminiscent of the catalytic triad of serine proteinases. However, in contrast to proteinases, the triad is concealed under a short helix, the “lid,” and is therefore inaccessible to solvent. 2. Geotrichum candidum Lipase

Lipases from the fungus G . candidum have been extensively studied over a number of years. The first report of a crystallographic study is credited to Hata et al. (1979), but their low-resolution structure (5.0 A) was too inaccurate to reveal the correct molecular architecture of the enzyme. The resolution was subsequently extended to 2.8 A (Sugihara et al., 1991), but the structure fell short of the quality required for a detailed description at an atomic level. A crystallographically refined structure was finally reported by Schrag et al. (1991)at 2.2 8, resolution.

a


GcL contains 544 amino acids in a single chain folded into one domain, making it one of the largest structural domains observed to date in a protein. Like RmL, GcL is an alp structure with a central, predominantly parallel p sheet. There are 11 strands in the central sheet, 3 more in a small additional sheet, and 17 a helices (Fig. 2). T h e catalytic Ser-217, a part of the G-X-S-X-G pentapeptide, is located at a tight turn between the C terminus of a /3 strand and an N terminus of an a helix, exactly as observed in RmL. The hydroxyl of Ser-217 is hydrogen bonded to the imidazole of His-463, which in turn donates a hydrogen bond to Glu-354. Thus, GcL constitutes the first known example of a serine hydrolase in which the acid residue of the triad is a glutamate and not an aspartate. The putative active site is covered by two nearly parallel a helices (residues 66-76 and 294-310) coming from different parts of the polypeptide chain. Schrag et al. (1991) note that both of these helices can be easily displaced following relatively minor adjustments in the mainchain conformation.

FIG. 2. The molecular structure of GcL; other details as in Fig. 1 . The two lids obscuring entry to the active site are the two parallel dark helices in the foreground.

LIPASE STRUCTURElFUNCTlON

9

B . Human Pancreatic Lapase Pancreatic lipase is one of the mammalian key digestive enzymes. It completes the dietary triacylglycerol breakdown initiated by preduodenal lipases, including lingual and gastric enzymes, (see below). The enzyme is inhibited in the intestine by bile salts, but the activity is restored in the presence of colipase (CLP), a relatively short (95 residues) heatstable polypeptide secreted by the pancreas (Semeriva and Desnuelle, 1979; Borgstrom and Erlanson-Albertsson, 1984). The structural details of the interaction of colipase with lipase are described in Section III,C. Single crystals of hPL were grown in the presence of LiCl and poctylglucoside, and the structure was determined at 2.2 8, resolution (Winkler et al., 1990). In contrast to fungal enzymes, hPL is clearly divided into two domains. The larger N-terminal domain is made up of the first 335 amino acids and exhibits an alp architecture, including a predominantly parallel central p sheet (Fig. 3). In general terms, the folding of this unit shows significant similarities to folding of the fungal enzymes (see also Section IV,B). The additional C-terminal domain is a p sandwich formed by two sheets, each made up of four antiparallel strands. Most biochemical studies relating to triacylglycerol lipases have been performed using the pancreatic enzyme, leading to a wealth of accumulated experimental data. Prior to the elucidation of the crystal struc-

FIG.3. The molecular structure of hPL; other details as in Fig. 1 . The main lid is seen on top of the catalytic domain, which is to the right.

10


ture of hPL it was generally assumed (Chapus et al., 1988)that there are two distinct sites in the protein: a catalytic center involving a histidine, and a “lipid recognition” site with Ser- 153,* which was subsequently shown to be at the center of the consensus sequence G-X-S-X-G. T h e X-ray structure identified Ser-153 as a part of a catalytic triad that also includes His-264 and Asp- 177. T h e hydrogen-bonding network is structurally analogous to that found in all the catalytic triads. As in fungal lipases, the catalytic center is not accessible to solvent. However, instead of a simple helical ‘‘lid’’found in RmL, or two ‘‘lids’’ seen in GcL, hPL appeared to have several loops that collectively obscure the entrance to the active site. It was subsequently observed that the displacement of two of these leads to the enzyme assuming an active conformation (van Tilbeurgh et al., 1993). One of these loops is a surface loop between residues 237 and 261, both of which are cysteines linked by a disulfide bridge. T h e second fragment is a shorter loop spanning residues 78 and 84. BY LIPASES 111. MOLECULARBASISOF LIPIDDEGRADATION In general terms, the crystallographic results show that lipases contain several distinct sites, each responsible for a specific function. T h e hydrolysis of the ester bond is accomplished by the catalytic triad, responsible for nucleophilic attack on the carbonyl carbon of the scissile ester bond, assisted by the oxyanion hole, which stabilizes the tetrahedral intermediates. T h e fatty acid recognition pocket defines the specificity of the leaving acid. There is also one o r more interface activation sites, responsible for the conformational change in the enzyme. In this section the discussion is on the available structural data relevant to the function of all these sites.

A. Hydrolysis of the Ester Bond

1 . Stereochemistry of the Catalytic Triad in Lipases

The discovery of catalytic triads in lipases and in related esterases, such as acetylcholinesterase (AChE) (Sussman et al., 1991) and cutinase (Martinez el al., 1992), revived interest in this otherwise well-known constellation of amino acids (like GcL, the AChE triad includes a glutamate). It should also be remembered that there are other functionally ‘The sequence numbering for hPL used in this paper is sequential, and therefore differs from that originally used by Winkler ct al. (1990), who number the amino acid inserted at position 31 (with respect to the porcine enzyme) as 30a.


11

analogous triads, with a thiol group acting as a nucleophile; these are found in thiol proteinases such as papain (Kamphuis et al., 1984) and actinidin (Baker, 1980), and in dienelactone hydrolase (Pathak and Ollis, 1990; Pathak et al., 1991). It might be argued that a catalytic triad made up of a Ser/Cys, a histidine, and a carboxylic acid is nature’s favorite hydrolytic tool. The discovery of the triad in lipases prompted Blow (1990) to search the Protein Data Bank to see if there are any fortuitous triads in known protein structures. He found two such triads, in the Taka a-amylase (Matsuura et al., 1984) and immunoglobulin Kol (Marquart et al., 1980), but neither showed the required stereochemistry. In fact, highresolution studies of two homologous a-amylases, including the Taka enzyme (Boel et al., 1990), ruled out any similarity of the environment of His-108 with a catalytic triad. It can therefore be concluded from the currently available structural database that a hydrogen-bonding network analogous to the chymotrypsin triad must result in enhanced nucleophilic activity of Ser (or Cys), and can therefore be found only in a catalytic center; fortuitous triads are probably eliminated by evolution because they create unnecessarily reactive sites. Many excellent reviews have already been written on the subject of the catalytic centers of serine and thiol proteinases (e.g., Kraut, 1977; Baker and Drenth, 1987; Warshel et al., 1989). In this paper the focus is specifically on the structure of the catalytic triad in lipases, with emphasis on the differences from and similarities to the catalytic centers of proteinases. The atomic coordinates for the G. candidurn lipase were not available when this review was written, and the analysis of the stereochemistry of the active centers is therefore restricted to lipases from R. miehei and the human pancreas. In RmL the nucleophilic triad consists of Ser-144, Asp-203, and His257, whereas in hPL the analogous amino acids are ser-153, Asp-177, and His-264. All hydrogen bonds identified in this system are analogous to those observed in serine proteinases: Ne2 of the histidine is bonded to the serine hydroxyl, and N61 is H-bonded to the aspartic acid (Fig. 4). Further details of the H-bonding stereochemistry are given in Table 11. The following discussion will first address the interaction of the nucleophilic serine with the histidine. In the X-ray structures of most serine proteinases, the hydrogen bond between the His Ne2 and Ser Oy is fairly long and therefore appears to be weak [for a discussion of this, see Read and James (1988)l. Early on, these observations led Matthews et al. (1977) to question even the existence of this bond. Nuclear magnetic resonance (NMR) studies (Smith et al., 1989) carried out in solu-

12


A\ vAL205

I

?

TYR 260

ASP176

!

FIG. 4. The stereochemistry of hydrogen bonding in the the catalytic triads in (A) RmL and (B) hPL; the crosses denote water molecules and the numbering in B is taken from the original set of hPL coordinates, in which amino acids beyond residue 30 are numbered one less than in this text.

tion indicate, however, that the protonation of the catalytic histidine may be responsible for the disturbance of the hydrogen-bonding network in crystals. It is interesting to note in this context that in RmL the stereochemistry of the side chains of His-257 and Ser-144 favors the formation of a strong hydrogen bond-whatever the donor-acceptor relationship. A careful analysis of both alternatives (Table 11) suggests, however, that the protonated imidazole forms particularly favorable inter-

TABLE I1

Hydrogen Bonding al Active Sites of RmL and hPL Bond

Lipase

Ser/His

RmL (high pH) RmL (low pH) hPL (high pH) hPL (low pH)

His/Asp

a

RmL (061 syn) hPL (061 syn) RmL (061 anti) hPL (061 anti) RmL (062 syn) hPL (062 syn) RmL (062 anti) hPL (062 anti)

Angle on the hydrogen atom.

d--...a (A)

H-.-a (A)

Ser-153-H . , . . . . . . . . His-264N~2

His-264N~2-H . . . . . . . . . . Ser-1530y

2.88 2.88 2.84 2.84

1.93 2.06 1.94 1.99

161.2 142.6 152.4 110.6

His-257N61-H . . . . . . . . Asp203061 Free Tyr-26007)-H . . . . . . . . . Asp-20306 1 His-264N61-H . . . . . . . . Asp-17706 1 Free Free Val-205N-H . ... . . . . . . Asp-203062 Thr-205N-H . . . . . . . . . . Asp-177062 Water-H.. . . . . . . .. . . . . Asp-177062

2.78 2.66 2.46 2.97 3.14 2.79

1.80 1.77 1.51 2.04 2.21

176.4

Donor (d)

Acceptor (a)

His-257N~2-H . . . . . . . . . . Ser-1440y

Ser-144-H . . . . . . . . . . His-257N~2

?

d-...fi..a

-

153.3 164.0

-

158.3 157.1 ?

(")a

14


actions. This is also consistent with the observed acidic pH in the crystallization medium (pH 3.9-4.0). In hPL the geometry of the interaction between Ser-153 and His-264 is similarly indicative of a strong hydrogen bond. The angle on the hydrogen atom is approximately 150°, regardless of whether the imidazole is protonated. Again, the pH of the crystals (pH 5.5) favors protonation. The second vital interaction in any triad is that of the histidine imidazole with a carboxylic acid. Generally, a hydrogen-bonded Asp-His pair is found in the catalytic centers of a variety of enzymes, including Zn-containing enzymes, serine proteinases, esterases, and several other hydrolases and oxidoreductases. The two H-bonded side chains assist in a nuclephilic attack during hydrolysis either directly by activating a serine hydroxyl, a cysteine thiol, or a water molecule (as is the case, for example, in phospholipase A*), or indirectly through a metal ion that polarizes the carbonyl attacked by a nucleophile (as in carboxypeptidase A). In RmL, 0 6 1 of Asp-203 accepts a syn-type H-bond from N61 of His-257 and an anti-type bond from 0 7 of Tyr-260 (Fig. 5A). On the other hand, 0 6 2 is within a small distance of both N61 of His-257 (3.02 A) and the main-chain N of Val-205 (2.97 A). However, in the former case the angle on the hydrogen atom (062 * * * H-Nc2) is much too acute (-80') for a strong H-bond. It is therefore the interaction with Val-205 that is relevant with respect to the stabilization of the side-chain conformation of Asp-203. This stereochemistry is very close to that found in chymotrypsin (Fig. 5B), where 0 6 1 of Asp-102 also accepts two H-bonds: a syn type from N E of~ His-57 and an anti from the hydroxyl of Ser-114. The overall effect is such that in lipases and in chymotrypsin the carboxyl group of the catalytic Asp is oriented in the same way with respect to the plane of the imidazole of the catalytic histidine. In subtilisin, a representative of the second family of serine proteinases, the relative dispositions of Asp and His are in some contrast to those described above (Fig. 5C). The plane of the carboxyl group of the catalytic Asp is now rotated with respect to the imidazole in a direction opposite to the one observed in RmL and trypsin; 0 6 1 (which accepts an H-bond from N61 of the catalytic His) is now stabilized by an additional H-bond donated by the main-chain amide of Thr-33. In hPL the H-bond between Asp-177(061) and His-264(N61) is unique in that it is of an anti type (Fig. 5D). Moreover, the syn pair of electrons on 0 6 1 is free. The stability of the Asp-177 side chain is achieved by a a three-centered H-bond involving the anti pair of electrons on 062, the amide hydrogen of Thr-204, and a buried water molecule. Again, the syn pair of electrons on 0 6 2 is free.

-


?gl Ser214

15

O\H Tyr260

\

\

\\.anti

anti

N61

!

N€2

\,anti

sy;.!,j

\N His57

B

A

Thr33 I

WATER

C

D

FIG.5 . A representation of the relative dispositions of the side chains of the catalytic aspartates and histidines in (A) RmL, (B) trypsin, (C) subtilisin, and (D) hPL. (A-C) the view is parallel to the plane of the imidazole, looking from where the catalytic serine would be, and along the H bond of His(NSl)-Asp(062). The histidine nitrogens and the carboxyl oxygens of Asp are identified in A and are the same in B and C. (D) The view is rotated -90' with respect to the preceding view so that the interaction of Asp and His in hPL can be better visualized.

In the past, considerable significance was attached to the syncarboxylate interaction. Gandour ( 1981)suggested that the syn electron pair may be more basic than the anti pair by a factor 103-104.This seemed to explain the enhanced basicity of imidazole. However, studies of databases (Allen and Kirby, 1991)and model compounds (Zimmerman et al., 1991) indicate that the difference in the relative basicities between syn- and anti-carboxylates is marginal (0.4-0.6 units). The hPL triad and the Asp-His couple in the active center of phospholipase A2 appear to support this view. Thus, the preference toward syn-type bonds observed in most serine proteinases may be due to packing effects in the active centers rather than to stereoelectronic effects. In spite of the obvious similarities, there are also some subtle differences in the stereochemistry of the environments of the Asp-His

16


couples in lipases and chymotrypsin-like proteinases. In the latter, the residue that stabilizes Asp-102 by donating an H-bond, Ser-214, is much closer to the molecular surface than the structurally analogous and completely buried Tyr-260 in RmL. According to a proposal put forward by Meyer et al. (1988), catalytic triads could function by shuttling a proton from the reactive histidine to the solvent through Asp102, Ser-214, and a network of flip-flop H-bonds. No such networks can be seen in lipases. In addition, site-directed mutagenesis studies carried out on rat anionic trypsin (McGrath et al., 1992) clearly indicate that the Ser214-Asp102H-bond is not essential for the function of the triad. Th e substitution of Ser-214 with Ala even increases k,,, from 490 to 900 min-' and has a less pronounced effect on K , (a change from 192 to 260 p M ) (McGrath et al., 1992). Another difference between lipases and chymotrypsin-like proteinases is in the second H-bond to 0 6 2 of the catalytic aspartate. In chymotrypsin the main-chain conformation of His-57 allows its amide group to provide the stabilizing H-bond, in contrast to lipases in which amides of Val-205 (RmL) and Thr-205 (hPL) serve a similar purpose. As was already mentioned, in both GcL and AChE triads an aspartate is replaced by a glutamate. It is interesting to note in this context that in hPL a glutamate can effectively replace Asp-177, with an accompanying sevenfold reduction in K , (Lowe, 1992). This result prompted Lowe (1992) to express reservations about the assignment of Asp-177 as the catalytic acid. Usually, an X-ray structure provides an unambiguous answer, but in the case of hPL there exists a possibility that the assignment may not be simple. Schrag el al. (1992) showed that Asp-206 (located at the end of strand 7) can be rotated into a position suitable for an interaction with His-264. Such a triad would have a syn-type interaction, more in concert with other serine hydrolases. Further site-directed mutagenesis studies should resolve this controversy. In conclusion, structural data indicate considerable variability in the environments of the catalytic triads. T h e only features that appear to be invariant and sufficient for the creation of a nucleophilic site are the strong hydrogen bonds between the hydroxyl (or the thiol), the imidazole of the histidine, and a carboxyl group of an aspartic or a glutamic acid side chain.

2 . The Oxyanion Hole The purpose of an oxyanion hole is to stabilize the tetrahedral intermediates that occur during the acylation and deacylation steps (Kraut, 1977). In chymotrypsin the H-bond donors that make up the oxyanion hole are the peptide amides of Ser-195 and Gly-193 (Henderson, 1970); in the subtilisins the amide of Ser-221 and another from the side chain

17


of Asn-155 serve the same purpose (Robertus et al., 1972). The situation in thiol proteinases is much less clear because of the limited structural database [for a review of the structure and function of the oxyanion hole in proteinases, see Menard and Storer (1992)l. As opposed to the catalytic triad, which is made up of side chains that can now be mutated at will, the structure-function relationships in the oxyanion hole are not equally susceptible to experimental verification. Only in subtilisin the involvement of the side-chain amide of Asn-155 allows for quantitative assessment of the role of the oxyanion H bonds in the stabilization of the tetrahedral intermediates. The replacement of Asn-155 with an isosteric Leu reduces the A,,, by 200- to 300-fold, but leaves the K , essentially unaffected (Bryan et al., 1986). This observation is fully consistent with Asn-155 not contributing to substrate binding, but playing a key role in the stabilization of the intermediate. In lipases the existing database regarding the oxyanion holes is still limited. In RmL two amide groups (residues 145 and 146) were originally proposed as likely candidates for this function (Brady et al., 1990). However, structural analyses of the two RmL-inhibitor complexes (Brzozowski et al., 1991; U. Derewenda et al., 1992) revealed that the oxyanion hole is likely to be fully formed only after the conformational change associated with interfacial activation, and that it is made up of both the amide and the side-chain hydroxyl of Ser-82 (Fig. 6). A hy-

e e hE2 2 5 7

hE2 2 5 7

FIG. 6. A stereo view of the structure of the oxyanion hole in RmL, as inferred from the structure of the complex with n-hexyl phosphonate ethyl ester. The dotted lines represent the two hydrogen bonds to the tetrahedral intermediate.

18


droxyl group in this position is invariant in the entire family of lipases from filamentous fungi, although in the enzyme from Rhizopus delemar a threonine replaces the serine. Yamaguchi et al. (1992) mutated Ser-83 (homologous to Ser-82 in RmL) in a Penicillium camemberti lipase to a Gly and found the mutant enzyme to be inactive. A preliminary study of the R . delemar enzyme with an Ala in the position of Thr-82 gave the same result (R. Jorger and M. Haas, USDA, personal communication). In hPL Winkler et al. (1990) originally suggested that the amides of Leu-154 (structurally equivalent to 145 in RmL) and Phe-78 may make u p the oxyanion hole. van Tilbeurgh et al. (1993) succeeded in the structural characterization of a complex of hPL with bovine procolipase (see Section III,C for the description of this cofactor and its function) crystallized in the presence of mixed phospholipid/bile salt micelles. In this complex hPL assumes the active conformation, and it is clear that after the accompanying conformational change the amides of Leu- 154 and Phe-78 can indeed serve as electrophiles for the oxyanion. In GcL Schrag et al. (1991) postulate that Ala-218 (again structurally equivalent to 145 in RmL) and Ala-132 play the same role in the stabilization of the intermediates. However, assuming that conformational rearrangements will also occur in this lipase during interfacial activation, the confirmation of this proposal will have to await a structural description of an enzyme-inhibitor complex. B . Molecular Basis of Substrate Specijicity Lipases may show specificity for the type or position of the fatty acid in the tri-, di-, or monoacylglycerol substrate and they may also exhibit stereospecificity toward a wide variety of substrates in organic solvents. I n general, however, specificity is low and in many cases the hydrolytic potential extends even to phospholipids and other organic compounds with an ester bond (e.g., Deckelbaum et al., 1992). This is a logical consequence of the primary function of most lipases the hydrolytic degradation of acylglycerols in general. Other esterases and thioesterases, most notably those involved in the terminal stages of fatty acid biosynthesis (e.g., Ferri and Meighen, 1991; Voelker et al., 1992), are much more specific with respect to the type of fatty acid. In proteins, molecular recognition and substrate specificity is often achieved by a network of specific hydrogen bonds to polar groups of the substrate or ligand. Fatty acids have no such groups and their interaction with the protein is restricted to much weaker, hydrophobic effects. Among lipases with known X-ray structures, substrate specificity is generally low. GcL is probably the most specific enzyme, favoring unsaturated fatty acids with a cis double bond at the 9-position (oleic,


19

linoleic, or linolenic). This specificity varies from strain to strain, probably due to different isoenzymes present in the crude extracts (Baillargeon, 1990; Baillargeon and McCarthy, 1991; Charlton and Macrae, 1992). However, no X-ray structures GcL complexed with inhibitors or substrate analogs have been reported and hence the molecular basis of this specificity is not known. On the other hand, the specificities of hPL and RmL are very low. RmL strongly favors positions 1 and 3 (rather independent of the type of the fatty acid) in triacylglycerols. To date it is the only lipase that has been crystallized in a complex with an inhibitor containing a short aliphatic chain (Brzozowskiet al., 1991). However, the low resolution of the study does not allow for a detailed description of the molecular basis of substrate specificity. Also, no structure-based analysis of stereospecificity in lipases is possible; results of purely chemical studies (Kazlauskas et al., 1991, and references therein; Xie et al., 1990) fall outside the scope of this review. The inhibitor used by Brzozowski et al. (1991), n-hexyl phosphonate ethyl ester, has a hydrocarbon moiety (n-hexyl) that occupies the place of the leaving fatty acid; the general location of this group is clearly visible. It is noteworthy that, on the dislocation of the “lid” from the active center, the n-hexyl is bound in a long groove lined with several hydrophobic side chains (Fig. 7). It is quite clear from this structure that

FIG. 7. A stereo view of the n-hexyl moiety bound in the fatty acid-binding pocket in

RmL.

20


even a much longer chain can be easily accommodated. It is not known, however, if the entire length of the fatty acid has to be buried in the complex to allow for hydrolysis; more likely, only the first 12 or 14 carbons are immobilized to allow for cleavage of the scissile bond.

C . Molecular Mechanism of Interfacial Activation

Since the pioneering studies of the phenomenon by Sarda and Desnuelle (1958), many authors have hypothesized about the molecular basis of interfacial activation (see Section I). T h e present structural evidence supports the original proposal put forward by Desnuelle et al. (1960), who postulated a conformational change in the enzyme, fixing itself at the interface. All crystal structures of lipases clearly show that a change is necessary to expose the catalytic centers, which in the native enzymes are buried under various surface loops, or lids. 1 . Interfacial Activation Site: The Lids The lids constitute the interface recognition and activation sites. Their topology with respect to the active sites in the three known structures of lipases is shown in Fig. 8. In RmL the analysis of the structural features of the lid is simplified by the availability of structures of both native and complexed molecules; this allows for clear identification of the mobile fragments. T h e lid is created by a long surface loop made up by residues 80- 109. This loop (Leszczynski and fragment defies a classical definition of an Rose, 1986) in that it exhibits well-defined secondary structure in its central helical fragment. Residues 82-96 (which include a short helix) directly obscure the entrance to the active site in the native enzyme. It is notable that between Arg-80 and Val-95 this fragment is not involved in any hydrogen bonds with any other parts of the molecule. Thus, the lid interacts with the main body of the protein only through hydrophobic interactions. In hPL there is a long surface loop between the disulfide-bridged cysteines 238 and 262 (Winkler et al., 1990). Positioned directly above the catalytic triad, this loop contains at its apex a short helix (residues 249 to 255), similar to the RmL lid, which is connected to the main body of the protein by two elongated stretches of the polypeptide chain. One of the central amino acids in this helix is a tryptophan (Trp-253), again reminiscent of the RmL structure. The second, smaller lid is made u p of the loop that connects strand p 5 of the central sheet with the helix a2 [secondary structure nomenclature follows that of Winkler et al. (1990)l

21


B. hPL

C. RmL FIG. 8. A representation of the general locations of the putative “lids” in (A) GcL, (B) hPL, and (C) RmL. The arrows indicate the directions of the helices; the loop covering the site in hPL is shown as an arc. The site of the oxyanion hole is at the top of the serine.

22


and includes residues 77 to 86. In the closed, inactive conformation, this loop interacts exclusively with the residues of the main lid. In GcL the active site is covered by two nearly parallel helices, residues 66-76 and 294-3 10, and the outside surfaces of these helices and of the bordering region are very hydrophobic (Schrag et al., 1991). This is in concert with the observations relating to the environment of the lid in RmL. 2 . Anatomy of Conformational Change A detailed analysis of the stereochemistry of the conformational changes associated with interfacial activation is possible for both RmL (Brzozowski el al., 1991; U. Derewenda et al., 1992) and hPL (van Tilbeurgh et al., 1993), although the low resolution of the latter study imposes some limitation on the accuracy of the observations. In the complexes of RmL with inhibitors, the central, helical part of the lid (residues 85-92) is transported 8 8, across the molecular surface (Fig. 9). During this change the helix actually rolls back from the active site, rotating by 167" about its own axis. This dramatic shift is a rigid body movement: when the two helices, in the native and inhibited enzymes, are superimposed, the root-mean-square deviation for all mainchain atoms is 0.31 A, well within the accuracy limits imposed by the experimental methods used. The conformational change is therefore brought about by specific rotations about conformational dihedral angles within two hinge regions (Fig. 10). This is not uncommon, and similar hinge regions have been identified in other proteins. What is unusual about RmL is the extent of change, which significantly alters the secondary structure in the hinge regions. At the N-terminal end of the lid, Ser-83 and Ser-84 constitute one hinge. In spite of a change of 60" in the 4 angle, Ser-83 remains within the YR region of the Ramachandran plot [the conformational space regions are denoted using the notation introduced by Efimov (1986)l. On the other hand, Ser-84 changes its secondary main-chain conformation from 6 to p. T h e latter transition is accomplished by a dramatic change of the angle. At the C-terminal end of the lid the hinge includes four consecutive amino acids: Asp-91, Leu-92, Thr-93, and Phe-94, all of which change their secondary structures (6 to a,p to a,/3 to 6, and p to YR, respectively). Among the side chains of the amino acids that make up the lid, only Trp-88 undergoes a significant change during the activation process (the xI and x 2 angles are 179" and 79", respectively, in the native form

+

A

B

FIG. 9. A representation of the conformational and surface changes in RmL during interfacial activation; the shaded amino acids are hydrophilic, others are hydrophobic. The cylinder represents the lid, rolling in the direction shown by the arrows, across the molecular surface. The active center is labeled.

24


180

-

-l80!.

. . . . .

.

.

.

.

.

.

'

'

'

.

'

residue number FIG. 10. Changes in the main-chain conformational angles in RrnL observed in the lid hinge regions on activation.

and -74" and - 125" in the inhibited species). However, this residue is involved in the intermolecular contacts in the crystal, and it is therefore quite probable that the conformations of some side chains in this region are affected by crystal packing. An interesting question is how each of the two conformations is stabilized, and what triggers the change. Let us first analyze the hydrogenbonding patterns in the two structures. In the native (inactive) molecule the lid contains a number of hydrogen bonds, all internal. The hydroxyl of Ser-82 is hydrogen bonded to the amide of Ser-84, and from this point the main-chain atoms form the usual 1 t 5 a-helical bonds with Leu-92 in the 3,o-helix conformation. T h e side chains of Asn-87 and Asp-91 are exposed to solvent. T h e C-terminal fragment (residues 93-96) is stabilized by the interactions of the main-chain atoms of Val-95: N to N61 of His-108 and 0 to N of Lys- 109. The activation brings about a significant change in this network. The hydroxyl of Ser-82 now interacts with 0 6 1 of Asp-91, while Asn-87,

LIPASE STRUCTURElFUNCTION

25

which becomes buried beneath the lid, forms several new H-bonds: 0 6 1 binds the amide nitrogens of Thr-93 and Phe-94, while N62 forms a bond with the carbonyl of Tyr-60. The hydroxyl group of Ser-84 is Hbonded to 0 6 1 of Asp-61. The helical central part of the lid retains its structural integrity and the carbonyl oxygen of Ile-89 forms an additional H-bond with the hydroxyl of Thr-93. The a helix is actually extended by one residue (Leu-92), while Thr-93 assumes the 310 conformation to become the terminating residue of the helix. The imidazole of His-108 changes its H-bonding partner from the amide nitrogen of Val-95 to the carbonyl oxygen of Ala-90, with N.92 involved in place of N61. Thus, the active enzyme actually gains three additional direct H-bonds between the lid and the main domain of the protein molecule. T h e water-mediated interactions may also be of considerable importance. In the activated form the lid occupies a new position on the surface of the molecule, some 8 8, away from the original location. This deep surface depression extends 10 8, into the molecule and is filled in the native enzyme by 18 water molecules, half of which are directly hydrogen bonded to the polar protein groups. During the conformational change ail but three of these solvent molecules are expelled. In the lipase-inhibitor complex these three molecules become buried and mediate the polar contacts formed primarily by Asn-87. The moving lid causes a change in the nature of the exposed surface of the active lipase molecule (Table 111). There are 12 distinctly hydrophobic amino acids (Ile-85, Trp-88, Ile-89, Leu-92, Phe-94, Val-205, Leu-208, Phe-2 13, Val-254, Leu-255, Leu-258, and Leu-267) that become more exposed on activation. The newly exposed area created by these residues amounts to 732 A2,or 7% of the total surface of the molecule. At the same time we observe a significant loss of predominantly polar surface. The polar residues of the lid (Ser-84, Asn-87, Asp-91, and Thr-93) account for 329 8,' of the lost polar surface, whereas the tetrapeptide 58-6 1 (Tyr-AspThr-Asn), which interacts with the lid in its new position, loses 124 of the surface exposed to solvent in the active enzyme. Prior to the detailed crystallographic studies of hPL a considerable amount of biochemical data relating to the activation mechanism of hPL had been accumulated. hPL is the only lipase among the lipases with known X-ray structures for which systematic structure-function relationship studies have been initiated using site-directed mutagenesis (Lowe, 1992). Substitutions were made at positions Ser-153, His-264, and Asp-177, and measurements of catalytic activity and interfacial binding were carried out. These studies showed that the catalytic site is

w2

26

ZYCMUNT S. DEREWENDA TABLE I11 Changes in Exposed Surface of RmL Molecule on Activation

Exposed surface (Az) Surface Hydrophobic zipper Lid

Main domain

Hydrophilic surface Lid

Main domain

Amino acid

Native

Active

Change (A)

Ile-85 Trp-88 lle-89 Ile-92 Phe-94

8 36 23 35 22

127 136 96 113 83

119 I00 73 78 61

Val-205 Leu-208 Phe-2 I3 Val-254 Leu-255 Leu-258 Leu-267

0 35 92 49 119 21 90

25 86 I39 97 129 58 113

25 51 47 48

Ser-84 Asn-87 Asp-9 1 Thr-93

61 91 111 103

38 2 34 53

-33 -87 - 77 -50

Tyr-58 Asp-59 Thr-60 Am-6 1

161 124 98 71

141 100 60 29

-20 - 24 - 38 -42

10

37 23

divorced from the site responsible for interfacial binding, an observation in concert with all the conclusions inferred from the structural data relating to the fungal enzymes. There was also convincing spectroscopic evidence supporting the hypothesis that conformational changes must accompany the activation of hPL. This evidence came from fluorescence and near-ultraviolet circular dichroism studies of hPL during its inactivation by a covalent specific inhibitor, tetrahydrolipstatin, in the presence of bile salts (Luthi-Peng and Winkler, 1992). It is, however, the role of colipase, a cofactor unique to the pancreatic enzyme, that is of particular significance. CLP was originally believed to function by anchoring the lipase molecule to the bile salt-covered surface of the lipid micelle (Canioni et d., 1977; Verger et al., 1977), thereby playing a key role in the interfacial activation. The enhancement of activity of hPL in the presence of CLP is approximately 10fold. Abousalham et al. (1992) used limited proteolytic degradation of pancreatic lipases by chymotrypsin as a tool to obtain information re-


27

garding the function of individual domains and the location of the colipase-binding site. Using horse pancreatic lipase they observed cleavage of only one peptide bond, i.e., between Leu-410 and Thr-411. This bond is located approximately in the middle of the C-terminal domain. The cleavage generates two fragments, a 45-kDa catalytic fragment (with an additional -80 amino acids from the C-terminal fragment) and a small 2-kDa fragment. As expected, the 45-kDa fragment retained its catalytic activity toward emulsified tributyrin in the presence of a noninhibitory concentration of bile salts, but could not be reactivated by colipase once the bile salt concentration was raised. A similar experiment using dog and human pancreatic enzymes resulted in complete degradation of the catalytic domains, but yielded intact C-terminal fragments following the cleavage of the Phe-335-Ala-336 bond. These C-terminal fragments were shown to cross-link with colipase. Abousalham et al. (1992) postulated that the two-domain organization of mammalian pancreatic lipases may reflect their cofactor binding requirements. A vivid confirmation of these hypotheses came from the crystal structure of the complex of the human lipase with bovine procolipase determined at 3.0 8, resolution by van Tilbeurgh et al. (1992). This is not only the first structure showing the stereochemistry of cofactor binding by a lipase, but also the first structural I characterization of colipase (Fig. 11). In these crystals procolipase appears as a flattened molecule made up of three regions shaped like fingers and stabilized by disulfide bridges. Many amino acids adopt an extended conformation, but no regular @-sheetstructure is observed. Two short loops at the tips of the fingers are disordered. This molecule binds exclusively to the C-terminal domain of lipase, although there are few direct interactions. This is consistent with the observation that in the absence of an oilwater interface the binding constant of the colipase-lipase complex is low (Donner et al., 1976), but it raised the possibility that further conformational changes (in lipase, which in the lipase-procolipase complex is not affected) are necessary to achieve full expression of enzymatic activity when the two proteins bind to the interface. Indeed, van Tilbeurgh et al. (1993) demonstrated that in the presence of a mixed micelled containing phospholipids and bile salts a conformational change can be induced in the lipase-procolipase complex, and that this change involves the expected two lids of the lipase, i.e., fragments 238262 and 77-86. The changes are spectacular both in magnitude and character and, in contrast to RmL, involve complicated reorganization of the secondary structure of the lids. In the active conformation the main lid assumes a helix-turn-helix structure, with the new helices

28


FIG. 1 1 . The lipase-procolipase complex: details are as in Figs. 1-3; the procolipase molecule is the more darkly shaded molecule, and is bound to the noncatalytic C-terminal domain of hPL.

formed by segments 242-247 and 252-260, which in the inactive molecule form the extended arms. The turn between these newly formed helices coincides with the part of the polypeptide chain that forms the apical helix in the inactive lipase. The maximal main-chain movement during this structural transition is 29 A and occurs at Ile-249. The main consequence of this structural reorganization is the exposure of the active site; however, there are other, equally important effects. The loop that contains residues 78-86, having lost its interactions with the lid, undergoes structural reorganization, which results in the formation of the oxyanion hole. Finally, the lid in its new position makes specific contacts with the molecule of colipase, presumably stabilizing the complex in the active state. The new interactions include the formation of three hydrogen bonds (van Tilbeurgh et al., 1993). Thus some of the original suggestions concerning the role of the lids in the interactions with lipase cofactors (Derewenda and Cambillau, 1991) have been confirmed at least in the case of hPL. Taken together, the surfaces around the newly reorganized active site and the adjacent surfaces of the colipase molecule create a substantial hydrophobic area in a fashion analogous to that seen in RmL. It is possible that the creation of such hydrophobic patches will be found to occur in other lipolytic enzymes that undergo an activation process.

LIPASE STRUCTURElFUNCTION

29

The crystal structures of lipases described to date are static descriptions of conformational states, possibly stabilized in some cases by crystal packing interactions. What is lacking is the dynamic component? What initiates the conformational transition? Is there a true lipidrecognition site that triggers the change, and does a simple two-state model come sufficiently close to the actual phenomenon? Or, does the interface simply stabilize one of the conformations existing in solution in an equilibrium? And just how stable is the inactive conformation? It appears that crystallography is able to provide at least partial answers to these questions. Two very interesting new results indicate that the so-called native, or inactive, molecule may be far less stable than has been inferred from the original crystal structures, and that the physicochemical properties of the solution (e.g., ionic strength, pH) may significantly destabilize the lid. T h e first result is a preliminary report by Grochulski et al. (1992), who succeeded in crystallizing the Candidu rugosa (formerly cylindracea) lipase (an enzyme homologous to GcL) in the open conformation, although without any inhibitor or substrate analog. It is not clear yet if the crystal packing forces are responsible for the stabilization of the active conformation in this new crystal structure. On the other hand, the high-resolution (1.85 A) structure of a lipase homologous to RmL and purified from the fungus Humicola lanugznosa (currently in the final stages of refinement in the author’s laboratory) shows the lid to be partly disordered. Both of these structures were studied in crystals obtained from solutions of organic solvents, i.e., from low-ionic-strength media. Thus the present evidence points to a fairly labile equilibrium, which in the immediate proximity of the water-lipid interface may favor the open conformation.

IV. EVOLUTIONARY RELATIONSHIPS Our knowledge of protein evolution is limited. This is not surprising, given the fact that to date only about 400-500 protein structures (excluding homologous molecules and other derivative studies) have been determined. Every time a representative of a new family of proteins is characterized at the molecular level it is tempting to speculate about possible evolutionary relationships with other groups and families of proteins. Such comparisons frequently help in the understanding of the structure-function relationships in the newly determined protein. Lipases are no exception, and in this section I will explore some of the more interesting similarities and differences both within this new superfamily and with other proteins.

30

Z Y G M U N T S. DEREWENDA

A.

Structure of the P-ESer-a Motif

The earliest observation that implied evolutionary links between all lipases was that of the consensus pentapeptide G-X-S-X-G, subsequently shown to contain the nucleophilic serine. The apparent similarity of this sequence to that found around the active serine in the chymotrypsin and subtilisin families of serine proteinases prompted a number of authors to infer an evolutionary relationship between the three families. Further evidence in support of such a link came from secondary structure prediction studies indicating that the nucleophilic serine in a lipase is most likely within a P turn, structurally reminiscent of proteinases (Reddy et al., 1986). In fact, one of the commonly used phrases found in introductions to many papers dealing directly or indirectly with lipases refers to “the consensus G-X-S-X-G pentapeptide found in the active site of all serine proteinases and esterases.” We now know that the implication that homology and/or structural similarities exist between the enzymes belonging to these diverse groups is incorrect. T h e matter has been dealt with in the literature (Derewenda and Derewenda, 1991; Liao el al., 1992), but it seems appropriate to review some of the conclusions. Table IV shows the secondary structures of the loops containing the nucleophile in serine proteinases (including wheat carboxypeptidase) and lipases. In trypsin and its homologues, apart from the two invariant glycines of the so-called consensus pentapeptide, G-X-S-X-G, there is a third invariant glycine in position 4. Glycines 1 and 4 both adapt secondary conformations outside the allowed regions of the Ramachandran plot. On the other hand, glycines 1 and 5 are both involved in the oxyanion hole and this may result in additional steric restraints on the side chains in these positions. In subtilisin, however, residue 5 adopts a classical a-helical conformation, and it is an alanine instead of glycine. Glycine 1 occupies a disallowed region of the Ramachandran plot, but different from that seen in trypsin. In contrast to these proteinases, the lipases and wheat carboxypeptidase exhibit a very different secondary structure, with both glycines 1 and 5 within the allowed Ramachandran region, and the nucleophile adopting a strained, &-typeconformation. Glycines appear to be invariant not because of the restraints imposed by local secondary structure, but by packing considerations (see below and Fig. 12). Thus, it must be concluded that there is no structural similarity between the pentapeptides containing the active serine in the two families of serine proteinases and that in lipases. The next interesting question is whether the consensus sequence shared by lipases and esterases corresponds to a common conserved structural motif. In all three known structures of lipases (RmL, hPL,

TABLE IV Main-Chain Conformational Angles of Active-Site “Consensus” Pentapeptlde C-X-S-X-G(A)in Serine Proteinaces and LipaseP

Chymotrypsin Residue

G

x S

X G(A) a

99 -81 -48 93 -74

Subtilisin

RmL

Wheat carboxypeptidase

hPLb

9

R‘

4

9

R

4

$

R

4

9

R

4

9

R

-17 -19 140 -26 175

ne

152 -74 -59 -85 -76

169 -10 -26 -22 -46

ne

-172 - 105 62 -57 -56

146 144 -121 -39 -35

fi

-132 -104 57 -63 -66

130 141 -126 -33 -10

p p

-169 -128 39 -64 -69

179 149 -101 -15

p p

O

a

a /3

ne

p

a a a

a

E

a a

E

a a

&

a

T h e values for chymotrypsin, subtilisin, and wheat carboxypeptidase are extracted from the Protein Data Bank and were reported by Liao et al.

(1992).

T h e values given for hPL are the mean values from the two polypeptide chains.

‘ R denotes the region of the conformational space of the Ramachandran plot occupied by the given residue; ne, nonallowed for nonglycine residues.

32


FIG. 12. The packing of amino acids within the p-eSer-a motif in RmL.

and GcL) the consensus pentapeptide G-X-S-X-G containing the active serine forms a very sharp turn, connecting one of the central strands of the p sheet (usually strand 4 or 5) with a buried a helix. This turn can be described either as the y turn (Matthews, 1972)or as a derivative of a type 11' /3 hairpin (Sibanda et al., 1989). Its uniqueness warrants some discussion. In a general case the second residue of the classic type 11' turn (the position occupied in the lipase turn by the nucleophilic serine) adopts the so-called 8 conformation within the Ramachandran space [the nomenclature is again that of Efimov (1986), extended by Sibanda et al., (1989)l. This secondary structure of the nucleophile is preserved among all the structurally characterized members of the lipase-esterase superfamily (see Section IV,B). Although unusual, it has been observed several times in other protein structures (for a discussion, see Derewenda and Derewenda, 1991). In RmL the interatomic distances between CP of Ser-144 and other atoms of the turn are very short (N of Leu-145, 2.85 A; C of His-143, 2.77 A; 0 of His-143, 2.81 A). Similar


33

distances are seen in hPL. In addition to these short contacts between nonhydrogen atoms, there is a particularly unfavorable interaction of the amide hydrogen that follows the E residue with the p carbon of the latter. An ultra-high-resolution (- 1.O A) study will be necessary to resolve the stereochemical questions associated with this unique structure. The close proximity of the p strand and of the helix enforced by the sharpness of the turn containing the G-X-S-X-G pentapeptide leads to a number of close contacts between amino acids from these two secondary structure elements (Fig. 12). It was this repetitive pattern of small andlor hydrophobic residues observed in amino acid sequences adjacent to the consensus pentapeptide that originally prompted Derewenda and Derewenda (1991) to postulate that, in all lipaseslesterases in which the G-X-S-X-G motif contains the nucleophile, the structure of this pentapeptide will be similar to that of RmL. So far all experimental data support this conclusion (see Section IV,B and Table V). There are a number of possible reasons why this particular motif was conserved during evolution. First, the tight turn brings a rigid nucleophile into the center of the active site, with plenty of free space to introduce other amino acids assisting in catalysis and to create room for the incoming substrate. Additionally, the nucleophile is in a perfect position to benefit from the macrodipole of the helix. It has been noted that the buried character of this unusual a helix may enhance the electrostatic effect of the macrodipole. Putting these hypotheses to a test by site-directed mutagenesis, or by other means, will be challenging.

B . Tertiary Fold of alp Hydrolases It is well established that the same three-dimensional scaffolding in proteins often carries constellations of amino acids with diverse enzymatic functions. A classic example is the large family of alp, or “TIM,” barrel enzymes (Farber and Petsko, 1990; Lesk et al., 1989). It appears that lipases are no exception: to date five other hydrolases with similar overall tertiary folds have been identified. They are AChE from Torpedo californica (Sussman et al., 1991); dienelactone hydrolase, a thiol hydrolase, from Pseudomonus sp. B13 (Pathak and Ollis, 1990; Pathak et al., 1991); haloalkane dehalogenase, with a hitherto unknown catalytic mechanism, from Xanthobacter autotrophicus (Franken et al., 1991); wheat serine carboxypeptidase I1 (Liao et al., 1992); and a cutinase from Fusarium solani (Martinez et al., 1992). Table I gives some selected physical and crystallographic data for these proteins. They all share a similar overall topology, described by Ollis et al. (1992) as the alp hydrolase

TABLE V Selecttd Lipases and Related Hydrolytic Enzymes Containing G-X-S-X-G Consensus Motif" Enzyme Known 3D structures Human pancreatic lipase Rhiwmtuor miehei lipase Humuola lanuginosa lipaseb Penuillium camemberti lipaseb Geotrichum candidum lipase Torpedo c a l f m i c a acetylcholinesterase Pseudonumcrcsp. dienelactone hydrolase Fusarium solani cutinase Unknown 3D structures Human lipoprotein lipase Human hepatic lipase Human gastric lipase Bile salt-stimulated lipase Hormone-sensitive lipase MwaxeNO TA144 lipase (Lips) Mwaulla TA144 lipase Staphylococcus aureus lipase Pseudumanas fragi lipase

Sequence N K R E K T K T

V V V L V V V L

H A V V M T G l

V V F V I I L A

I T T V F F V G

G G G G G G G G

H H H H E E Y Y

S S S S S S C S

L L L L A A L Q

G G G G G G G G

A G G A A G G G

H A A A M A A A

A T L V S S L A

A A A A V V A L

N H Q N R N R K R

V V L I I T L V V

H H H T C H G H N

L L Y L L V A L L

L I V F A G I V I

G G G G G G G G G

Y Y H E D N W H H

S S S S S S S S S

L L A A A M M M Q

G G G G G G G G G

A A T G G G G G A

H H T A N A G Q L

A V I S L I G T T

A S G V C S A I A

The boldface type denotes residues whose side chains pack closely in the interior of the motif; the active nucleophile is in italic type. The structural studies of both H . lanuginosa lipase and P . camemberti lipase are in the final stages in the author's laboratory and the cr-ESer-p motif in these molecules is clearly resolved. a


35

fold (Fig. 13), reminiscent of that found among lipases, and all contain the above described 0-ESer-a motif. In each molecule there is a central p sheet, with identical topology of the central eight strands +1, +2, -lx, +2x, and (+1x)3 (as defined by Richardson, 1981). Wheat carboxypeptidase is slightly different in that it has a hairpin loop inserted between strands 7 and 8. Cutinase, on the other hand, is a truncated molecule in which the first strand is missing altogether, the second is not in a regular p conformation, and the last helix-turn-strand fragment is also removed. Among the lipases GcL is the one most similar to the consensus fold, and its topology is identical to that of AChE. In this context RmL can be seen as a related subgroup in which a deletion of the third strand eliminated the crossover connection and simplified the fold (alternatively it could be argued that the alp topology originated with an insertion of the third strand into the RmL sheet!). Strand 9 of RmL constitutes an additional insertion. Human pancreatic lipase deviates from the consensus fold by having the order of the first strands reversed, while an extra hairpin loop is inserted in the N-terminal end. In all cases the p-ESer-a motif is found to be made up of strand 5 (strand 3 in cutinase) and the following helix. The nucleophiie is commonly Ser, although a cysteine is found in dienelactone hydrolase, and even more surprisingly, an aspartate in haloalkane dehalogenase. In the latter case, however, the catalytic mechanism has not been elucidated and the function of the aspartate is inferred somewhat speculatively from its location. The packing rules imposed on the side chains of the p-ESer-a motif are largely obeyed, although in wheat carboxypeptidase the second Gly of the consensus pentapeptide is replaced by an Ala, whereas in haloalkane dehalogenase the first Gly becomes a Val. Although the structural similarities between the nucleophile turns of different hydrolases have been conclusively demonstrated, their evolutionary history is much more difficult to establish. Is it possible that all of these enzymes have evolved from one distant ancestor, o r has the hydrolase fold with its p-ESer-a motif been reinvented in the course of evolution several times? Alternatively, was this motif introduced once, and then built into a variety of hydrolases by means of exon shuffling? Our present database is much too insufficient to address these complicated issues. However, in the enzymes of the human lipase gene family (pancreatic, lipoprotein, and hepatic lipases; see below), the entire p-ESer-a motif is exclusively coded for by a single exon. This is in contrast to a much simpler organization of the RmL gene, wherein only one intron is present (Boel et al., 1988), but still provides some circumstantial support for the exon shuffling theory.

B

D

F


37

T h e p-eSer-a motif is likely to be present in other enzymes with esterase or thioesterase activity, although their structure may be different from that of the alp hydrolases. Some of these proteins have already been crystallized for X-ray studies [e.g., Vibrio harveyi acyltransferase (Swenson et al., 1992)], and their crystal structures should throw some more light on the structural and evolutionary relationships within the lipaselesterase superfamily. T h e general location of the acid member of the triad in the alp hydrolase fold is also conserved. The catalytic Asp (or Glu) occurs at the carboxyl end of the seventh strand. The only significant deviation from this pattern is in the pancreatic lipase in which the catalytic Asp occupies a position at the end of strand six. It has already been pointed out that Asp-206 (at the carboxyl end of strand seven) can also be rotated into a position favorable for a hydrogen bond to the catalytic His-264, after a rotation around the x1 angle (Schrag et al., 1992). Thus there exists a possibility that hPL constitutes an evolutionary intermediate in the pathway of migration of the catalytic acid to a new position within the fold (Schrag et al., 1992). Th e third member of the catalytic triad, a histidine, is located on a loop that follows the last strand of the consensus sheet. T h e exceptions to this rule are RmL, wherein this C-terminal loop follows the extra strand inserted in position 9, and cutinase, wherein the catalytic His188 is located on the same C-terminal loop as the catalytic Asp-175, owing to the absence of the last p strand. Ollis et al. (1992) postulate that the similarities between the enzymes that make up this new family of hydrolases are sufficient to infer divergent evolution of these enzymes from a common ancestral gene. It will be interesting to see if this view prevails. C . Lipases and Proteinases: Possible Relationships

We are now returning to the recurring issue of possible evolutionary links between lipases and serine proteinases. We already know that detailed comparisons ruled out any homology between the two families of

FIG. 13. The alp hydrolase fold and it variations. (A) cutinase, (B) dienelactone hydrolase and haloalkane dehalogenase, (C) wheat carboxypeptidase, (D) RmL, (E) hPL, (F) GcL and AChE; the three catalytic residues, always in the order Ser, Asp/Glu, and His, appear as dark dots. The folds are aligned in such a way as to show the structural homologies within the hydrolytic domains, somewhat divorced from the N-terminal part of the sheet. The hPL is a two-domain protein, and the location of the additional Cterminal domain is indicated.

38


serine proteinases on the one hand and lipases on the other, and that the common consensus sequence (G-X-S-X-C) is fortuitous. However, we now address a somewhat different question: is it possible that there are other serine proteinases, possibly more ancient and even more abundant in nature, which may be related to the esterases and lipases of the alp hydrolase family? First, the structure of wheat serine carboxypeptidase I1 described by Liao et al. (1992) serves as good evidence of possible relationships. Serine carboxypeptidases are found in virtually every higher organism (Breddam, 1986), and some are of considerable importance, such as the proteinase involved in the regulation of blood pressure in humans (Odya and Erdos, 1981). Second, it is interesting to note that a new family of serine proteinases, the prolyl peptidase family, has been identified (Rawlings et al., 1991). T h e following enzymes have been included in this group: prolyl oligopeptidase (Rennex et al., 199l), rat liver acylaminoacylpeptidase (Kobayashi et al., 1989), human protein 3p21 (Naylor et al., 1989), rat liver dipeptidylpeptidase IV (Ogata et al., 1989), and yeast dipeptidylpeptidase B (Roberts et al., 1989). They all share two important features that make them different from the subtilisin and chymotrypsin families, but resemble in some ways lipases and esterases. First, these proteinases contain a G-X-S-X-G pentapeptide that appears to occur within a context indicating the presence of the p-eSer-a motif (as described for RmL and other lipases). Second, the order in which the members of the putative triad occur along the polypeptide chain appears to be the same as in lipases (i.e., Ser . . . Asp . . . His) and is therefore in contrast to other serine proteinases. T h e essential catalytic serine and histidine have been identified in prolyl oligopeptidase (Rennex et nl., 1991; Stone et al., 1991) as Ser-554 (part of the consensus pentapeptide) and His680. Both are conserved throughout the family. In addition, there are two conserved aspartates, Asp-529 and Asp-642. T h e latter position of a catalytic Asp would result in a sequence order on the three triad residues (Ser . . . Asp . . . His) identical to that found in lipases. Any reliable assessment of structural similarities must, of course, await an elucidation of at least one crystal structure of any of the members of this newly described family of proteinases. WITH UNKNOWN THREE-DIMENSIONAL STRUCTURES V. LIPASES A broad variety of lipases isolated and purified to a varying degree from many organisms have been investigated to date. I t is beyond the scope of this review to discuss the wealth of biochemical data generated

LIPASE STRUCTUREIFUNCTION

39

by these studies. However, I would like to end with a discussion of some key lipases whose structures have not yet been determined, in the hope that such discussion in the context of the available crystallographic data may be of interest. A . Prokaryotic Enzymes

Owing to their industrial potential, prokaryotic lipases have been extensively studied. The first amino acid sequence of a bacterial enzyme elucidated by gene cloning was, to our knowledge, that of Staphylococcus hyicus (Gotz el al., 1985). Other known cDNA-derived sequences include those of Staphylococcus aureus (Lee and landolo, 1986), Pseudomonm fragi (Kugimiya et al., 1986; Aoyama et al., 1988), lip2 and lip3 gene products of Moraxella TA144 (Feller et al., 1991a,b),Pseudomom cepacia (Jorgensen et al., 1991), Alcaligenes denitrijicans (Odera et al., 1986), Pseudomonas sp. LS107d2 (Johnson et al., 1992), Bacillus subtilis 168 (Dartois el al., 1992), and several others. None of the reported sequences has detectable homology to any of the structurally characterized lipases. So far the only common feature shared by these and other lipases appears to be the presence of the consensus pentapeptide G-X-S-X-G [the only exception being the B. subtilis lipase reported by Dartois et al. (1992), in which the homologous sequence fragment is A-H-S-M-GI. This may indicate at least some localized structural similarities generated by the presence of the p-ESer-a motif either within a central p sheet or in a different structural context. On the other hand, some examples of significant homology have been detected within the prokaryotic enzymes, e.g., S . a u r m and S . hyicus share 46% of their amino acid sequence. Lipases from different species of Pseudomonm have been crystallized successfully: Pseudomom glumue (Cleansby et al., 1992), Pseudomonm sp. strain ATCC 21808 (Kordel et al., 1991), Pseudomom fluorescens (Larson et al., 1991), Pseudomom putidu (Sarma et al., 1991), and Pseudomom cepacia (Kim et al., 1992); their three-dimensional structures should soon be known. B. Other Fungal Lipases RmL is not the only studied representative of lipases purified from the family of filamentous fungi. The other three enzymes cloned and investigated thoroughly for their catalytic potential are the R. delemur lipase (Haas et al., 1991), the enzyme from P. camemberti (Yamaguchi and Mase, 1991; Yamaguchi et al., 1991), and a lipase from H . lunuginosa (Boel et al., 1991). All three show homology to RmL. However, the

40


P . camemberti lipase is unique in that it shows no activity toward triacylglycerols. Instead, it degrades mono- and diacylglycerols, with highest activities observed on 1- and 3-monopalmitoyl-sn-glycerol and l-monostearoyl-rac-glycerol (Yamaguchi and Mase, 1991). Both the H . lanuginosa lipase and the P. camemberti enzymes are in the final stages of high-resolution refinement in the author’s laboratory. A detailed comparison of these molecules will not only help in understanding the structural basis of substrate specificity, but may also provide valuable guidelines for a design of novel enzymes with potential industrial applications.

C. Human Lipases Involved in LipidlLipoprotein Metabolism

I. Human Lipase Gene Family The human pancreatic lipase is homologous to two other lipases playing pivotal roles in lipoprotein metabolism: lipoprotein lipase and hepatic lipase. Together these three proteins constitute the human lipase gene family. Lipoprotein and hepatic lipases are important enzymes involved in the metabolism of chylomicrons and various fractions of lipoproteins. Both have been the subject of attention, as evidenced by numerous reviews (e.g., Garfinkel and Schotz, 1987; Wang et al., 1992). This interest stems from the fact that abnormal lipoprotein metabolism has been linked to various disorders, including hyperchylomicronemia, hypercholesterolemia, hypertriglyceridemia, obesity, diabetes, and premature atherosclerosis. Genetic defects in both HL and LPL are now known to be the cause of at least some familial disorders of lipoprotein metabolism. The activity of human lipoprotein lipase in plasma was first observed by Hahn (1943), who called it the “clearing factor.” Anfinsen et al. (1952) established that this factor is in fact an enzyme. It is present throughout the circulatory system, and is localized on the surface of the capillary endothelium in extrahepatic tissue such as muscle (both skeletal and cardiac) and adipose tissue. Its lipolytic activity is directed primarily against the triacylglycerols in chylomicrons and very low-density lipoproteins (VLDLs), but it requires a cofactor, apoliprotein C-I1 (apoC-11), for full expression of activity [for a review, see Smith and Pownall ( 1984)3. Apolipoprotein C-I1 is a 78-amino-acid-long molecule found on both chylomicrons and VLDLs and also on high-density lipoproteins (HDLs). The tissue-specific action of LPL directs fatty acids into the peripheral tissues for storage and to satisfy the energy requirements, or the need for energy storage.


41

Hepatic lipase is involved in the metabolism of high-density lipoproteins and intermediate density lipoproteins (IDLs), converting the HDL2 fraction to HDL3 and generating LDLs from IDLs. T h e enzyme appears to have broad specificity: it hydrolyzes tri-, di-, and monoacylglycerols, acyl-CoA thioesters, and even phospholipids. hHL is secreted by the liver parenchymal cells and does not require any cofactors for its activity. Neither HL nor LPL has been crystallized, although intensive studies are underway in several laboratories. However, given the homology within the family (Komaromy and Schotz, 1987; Ben-Zeev et al., 1987; Kirchgessner et al., 1989; Datta et al., 1988; Persson et al., 1989), structural arguments can be inferred from this relationship (human HL is 45% identical with hPL, and human LPL shares 36% of its amino acids with hPL). A direct correlation of sequence homology with structural similarity is never easy. Site-directed mutagenesis studies of HL and LPL based on the molecular model of PL indicate that the general three-dimensional structure is the same in all three enzymes. In particular, site-directed mutagenesis studies have shown that all three residues inferred from the X-ray structure of PL to be involved in the catalytic triad are indeed critical for the expression of the enzymatic activity (e.g., Faustinella et al., 1991, 1992; Lowe, 1992). Derewenda and Cambillau (1991) aligned the sequences of all three lipases based on the molecular structure of hPL. Their results show that the regions of most significant homology correspond largely to the secondary structure elements that build the core of the hPL molecule. In addition, secondary structure prediction calculations identified the critical stretches of the central p sheet. Thus, the overall similarity of the molecular architecture of all three members of the lipase gene family seems to be well established. Both PL and LPL require cofactors for full expression of activity (CLP and apoC-11, respectively); no such cofactor is necessary for HL. Derewenda and Cambillau (1991) postulated that, in the human lipase gene family of enzymes, the loops of the N-terminal domain, which exhibit the most pronounced variation in their amino acid sequences, may be responsible for conferring specificity with respect to cofactors. The structure of the lipase-procolipase complex (van Tilbeurgh et al., 1992; see above) does not support this hypothesis. However, in the case of LPL the structural basis of its interaction with apoC-I1 may be quite different. Wong et al. (1991) and Davis et al. (1992) produced hybrid molecules by interchanging the C-terminal domains between the rat hepatic and lipoprotein lipases. Their HL chimera, made u p of the HL N-terminal catalytic domain and the LPL C-terminal fragment, exhibited the salt-resistant catalytic properties characteristic of HL, but was

42


not stimulated by apoC-11. T h e LPL fragment was still recognized by monoclonal antibodies raised against LPL. Thus, the binding of apoCI1 occurs within the catalytic domain of LPL. It is tempting to hypothesize that the surface loop between disulfide-bridged residues 237-26 1 (homologous to loop 238-262 in hPL; see above) might in this case constitute the apoC-I1 binding site. T h e importance of this surface loop for LPL activity has been probed in an elegant study by Faustinella et al. (1992), who replaced it in the lipoprotein lipase with the corresponding lid from the hepatic enzyme and also from the pancreatic lipase. T h e hepatic loop hybrid showed no significant loss of enzyme activity, whereas the pancreatic one resulted in complete loss of activity. Should the lid be the actual cofactor-binding site in LPL, what is the purpose of the C-terminal domain in LPL and HL? Both enzymes bind to the vascular endothelium by interactions with heparan sulfate proteoglycans. Davis et al. ( 1992) show that the heparin-binding activity resides in the C-terminal domain. The same authors also found that an exchange of the C-terminal domain between HL and LPL resulted in an enhanced ratio of phospholipase-to-lipase activity in the chimeras. This observation suggests that in some way the C-terminal domain affects substrate specificity. How can this be possible given the fact the active site is far away from the C terminus? Wong et al. (1991) note several reports that indicate that LPL and HL function as dimers and oligomers, respectively (Osborne et al., 1985; Garfinkel et al., 1983; Jensen and Bensadoun, 1981; Twu et al., 1984). They proposed, therefore, head-to-tail dimerization of lipase molecules in a fashion such that the C-terminal domain would bind and present the substrate to the catalytic center of the second enzyme molecule. The similarity of the C-terminal domain to the fatty acid-binding protein (Sacchettini et al., 1988, 1989) and the presence of two molecules in the asymmetric unit of the hPL crystals (Winkler et al., 1990) are given, somewhat circumstantially, as evidence in support of this theory. A subsequent report by Giller et al. (1992) indicates that the human lipase gene family may be more complex than was originally thought. These authors have isolated human cDNAs, cloning for two pancreatic lipase-related proteins, hPLRPl and hPLRP2, with 68 and 65% identity, respectively, with the pancreatic lipase. hPLRP2 was shown to have lipolytic activity (although only marginally dependent on colipase) whereas hPLRPl was inactive. The two novel proteins probably exist in other species, and some of the sequences reported to be pancreatic lipase may be orthologues of hPLRPl and hPLRP2. Clearly, despite the impetus created by the elucidation of the pancreatic lipase crystal structure, there is much more work ahead before we


43

can formulate a cohesive picture of structure-function relationships in the human lipase gene family. 2. Preduodenal Lipases The digestion of fat is not initiated by the pancreatic enzyme, although the physiological significance of preduodenal lipolysis has long been underestimated. In cases of complete deficiency of pancreatic lipase activity, patients are still able to absorb 70% of dietary fat (Hamosh, 1984). In humans this preduodenal activity is located only in the gastric mucosa and not in the lingual, pharyngeal, or esophageal areas, where similar lipolytic activity is found in other mammals [for a review of gastric and other preduodenal lipases, see Gargouri et al. (1989b)l. The cDNA-derived amino acid sequences of human gastric lipase (Bodrner et al., 1987) and rat lingual lipase (Docherty et al., 1985) show clear homology, with 78% identity. There are 379 and 377 amino acids, respectively, and three of the four putative glycosylation sites in the human enzyme were also found in the rat lingual protein. However, with the exception of the G-X-S-X-G consensus sequence motif, there appears to be no homology with any other lipase family. Gastric lipases were until recently thought to be sulfhydryl enzymes. Both rat and human enzymes have one free SH group, as shown by incubation with 5,5’-dithiobis(2-nitrobenzoicacid) (DTNB) and 4,4’dithiopyridine (4-PDS). In both cases incubation results in the loss of enzymatic activity (Gargouri et al., 1988; Moreau et al., 1988). Similar inactivation of both human and rat gastric lipases was achieved using ajoene, derived from ethanol extracts of garlic (Gargouri et al., 1989a). However, the crystallographic database of lipases strongly suggests that the G-X-S-X-G motif contains a nucleophilic serine. The reevaluation of the enzymatic mechanism of gastric lipases focused on the inactivation of the enzyme by diethyl p-nitrophenyl phosphate (Moreau et al., 1991). I t was found that both gastric and pancreatic enzymes were stoichiometrically inactivated, though the binding to the oil-water interface was not impaired. The essential sulfhydryl group was not modified during the reaction with E600. The essential role of a serine residue is further supported by the inhibition of gastric lipase by tetrahydrolipstatin (Gargouri et al., 1991; Ransac et al., 1991). It is quite probable that the free sulfhydryl group is topologically close to the active site, and its chemical modification may distort the environment of the catalytic triad or the or the oxyanion hole. A similar phenomenon has been observed among subtilisins, wherein a subfamily including proteinase K, thermitase, yeast proteinase B, and yeast KEX-2-encoded protein is sensitive to

44


thiol reagents. In this case crystallographic studies have shown that the reactive cysteine (Cys-68) is located very close to the catalytic histidyl residue. Introduction of Cys into savinase (a Cys-free subtilisin) rendered the latter sensitive to thiol reagents, but had no influence on the kinetic properties (Bech et al., 1992). Thus steric effects are responsible for the observed inhibition in thiol-sensitive subtilisins. Intense attempts to obtain single crystals of gastric lipase are underway (Moreau et al., 1992). 3, Lysosomal Acid Lipase

Lysosomal acid lipase/cholesteryl ester hydrolase catalyzes the deacylation of triacylglyerols and cholesteryl esters in low-density lipoproteins acquired through receptor-mediated uptake, as well as intracellular triacylglycerols. This is a key step in intracellular cholesterol metabolism (Goldstein and Brown, 1977). Low levels of human lysosoma1 acid lipase (hLAL) have been associated with two hereditary disorders: Wolman’s disease (WD) and cholesteryl ester storage disease (CESD) (Schmitz and Assman, 1989). cDNA encoding the human enzyme was cloned (Anderson and Sando, 1991). The deduced amino acid sequence includes 378 residues in a single chain; a 2 l-residue-long signal peptide precedes this sequence. Identifiable homology was detected only with respect to the human gastric lipase and rat lingual lipase (58 and 57% sequence identity, respectively). The inhibition of hLAL by boronic acids and diethyl p-nitrophenyl phosphate (Sando and Rosenbaum, 1985; G. N. Sando and H. L. Brockman, unpublished, cited by Anderson and Sando, 1991) indicates that hLAL is a serine hydrolase. T w o lipase/esterase consensus pentapeptides, G-X-S-X-G, are found, but only one of them appears to be consistent with the packing requirements of the P-eSer-a nucleophilic motif (see above). Susceptibility of the enzyme to sulfhydryl reagents, and the requirement of thiols for the stability of purified hLAL, prompted Anderson and Sando (1991) to propose that a cysteine residue, or rather a Cys/Ser couple, may be involved in an internal transacylation reaction. It must be pointed out, however, that hLAL has all three cysteines of the gastric enzyme (as well as six additional ones), and so the “inhibitory” Cys is also there. The same argument proposed herein with respect to hGL, i.e., that a free cysteine is topologically close to the active site, also holds for hLAL. 4. Bib Salt-ActivatedLipase (Cholesterol Esterase)

Human milk contains a bile salt-activated lipase (BAL). This enzyme helps infants in the digestion of milk fat (for a review, see Olivecrona


45

and Bengtsson, 1984). BAL is stable in the stomach, and its activation in the duodenum by bile salts prevents premature hydrolysis of triacylglycerols. The function of bile salts appears to be complex and involves both substrate binding and a catalytic process (Wang et al., 1988). The cDNA of human BAL was cloned and an amino acid sequence was derived (Baba et al., 1991). The protein is a single polypeptide chain made up of 722 amino acids and an additional signal sequence of 20 amino acids. The C-terminal 176 residues contain 16 repeating units with a basic motif P-V-P-P-T-G-D-S-G-A-P;this accounts for the unusually high proline content, as well as the low isoelectric point (pZ 3.7). Two disulfide bridges (64-80, 246-257) stabilize the structure. The G-X-S-X-G motif is there, and inhibition by diisopropyl fluorophosphate strongly suggests that Ser-194, which is a part of this pentapeptide, is indeed the nucleophile (Baba et al., 1991). The amino acid sequence of human BAL is closely related to those of rat pancreatic lysophospholipase (Han et al., 1987), cholesterol esterase (Kissel et al., 1989), bovine pancreatic cholesterol esterase/lysophospholipase (Kyger et al., 1989), AChE (Schumacher et al., 1986), cholinesterase (Lockridge et al., 1987), and thyroglobulin (Mercken et al., 1985). It was shown, in fact, that bile salt-activated lipase, lysophospholipase, and cholesterol esterase are one and the same protein (Hui and Kissel, 1990). The observed homology prompted Hui and Kissel (1990) to predict that the catalytic triad in BAL includes Glu-78 or Asp-79, Ser-194, and His-435. However, more recent site-directed mutagenesis studies based on sequence alignments using the reported X-ray structures of AChE and GcL pointed to Asp-320 as the catalytic acid (DiPersio and Hui, 1993). BAL appears, therefore, to be yet another member of the alp hydrolase family.

5. Hormone-Sensitive Lipase Hormone-sensitive lipase (HSL) is a key enzyme in fatty acid mobilization and one of the most complex lipolytic enzymes. It is under acute hormonal and neuronal control through reversible CAMP-dependent phosphorylation of a single serine (Belfrage et al., 1984). T h e activation is mediated by catecholamines (Stralfors and Belfrage, 1983), whereas the antilipolytic effect of insulin arises from the prevention of phosphorylation (Stralfors et al., 1984). Although originally thought to occur only in the adipose tissue, HSL has been found in a number of other tissues, notably heart and skeletal muscle, where its function is not always clear. The enzyme shows the same catalytic activity toward both triacylglycerols and cholesterol esters and is thought to be involved significantly in the biosynthesis of steroids in ovaries, adrenal cortex, testes, and placenta (Holm et al., 1988).

46


HSL cDNA from rat adipocytes was cloned and sequenced (Holm et al., 1988). The predicted protein is 757 amino acids in length and over 82.8 kDa in size. There is no homology with any other protein. The only sequence feature shared by this enzyme with other lipases is the G-X-S-X-G pentapeptide, which occurs within a stretch reasonably consistent with the p-eSer-ar motif, Thus, Ser-423, which is a part of this pentapeptide, is the likely nucleophile. Nothing is known about the other members of a catalytic triad, if indeed it exists in this enzyme. The regulatory serine that undergoes phosphorylation by CAMPdependent kinase has been identified as Ser-563 (Holm et al., 1988). Nothing more is known about the molecular structure of this important enzyme. VI.

CONCLUSIONS

A quick look at the list of references shows that work in the past few years has resulted in a major breakthrough in our understanding of lipases. However, with less than a handful of high-resolution crystallographic analyses, the structural studies of these fascinating enzymes are still in their infancy. T o date no systematic investigations using sitedirected mutagenesis along with X-ray crystallography have been reported. Also, only two complexes of a lipase with covalent inhibitors have been analyzed, both involving the same enzyme from R. miehei. On the other hand, enzymes for which extensive site-directed mutagenesis experiments have been carried out (e.g., lipoprotein lipase) continue to resist crystallization attempts. Clearly, a fascinating and important new field of structural biochemistry has been opened and will attract many investigators in the coming years. ACKNOWLEDGMENTS Mr. J. Chlebek and Dr. U. Derewenda are gratefully acknowledged for their help in the preparation of the figures. Drs. C. Kay and D. Brindley, both of the University of Alberta, are thanked for the critical reading of the original manuscript. Dr. Cecilia Holm provided valuable comments on selected sections. The coordinates of hPL were kindly provided by Dr. F. Winkler, those for the a carbons of GcL were provided by Dr. M. Cygler, and those for the hPL-procolipase complex were provided by Dr. van Tilbeurgh. Financial support from the Medical Research Council of Canada (grant to the Group in Protein Structure and Function), the Alberta Heritage Foundation for Medical Research, and NOVONordisk (Copenhagen)is gratefully acknowledged. The author is a Medical Scholar of the Alberta Heritage Foundation.

REFERENCES Abousalham, A., Chaillan, C., Kerfelec, B., Foglizzo, E.. and Chapus, C. (1992).Protein Eng. 5, 105-111. Allen, F. H., and Kirby, A. J. (1991).j.Am. Chem. SOC.113,8829-8831. Anderson, R. A., and Sando, G. N. (1991).J.Biol. Chem. 266,22479-22484.

LIPASE STRUCTUREIFUNCTION

47

Anfinsen, C. B., Boyle, E., and Brown, R. K. (1952).Science 115,583-586. Antonian, E. (1988).Lipids 23, 1101-1 106. Aoyama, S.,Yoshida, N., and Inouye, S. (1988).FEES Lett. 242,36-40. Baba, T., Downs, D., Jackson, K. W., Tang, J., and Wang, C . 4 . (1991).Biochemistty 30,

500-510. Baillargeon, M. W. (1990).Lipids 25,841-848. Baillargeon, M. W.,and McCarthy, S. G. (1991).Lipzds 26,831-836. Baker, E. N. (1980).J.Mol. Biol. 141,441-484. Baker, E. N.,and Drenth, J. (1987).In “Biological Macromoleculesand Assemblies” (F. A. Jurnak and A. McPherson, eds.), Vol. 3,pp. 314-368.Wiley, New York. Bech, I. M., Branner, S., Hastrup, S., and Breddam, K. (1992).FEES Lett. 297, 164-

166. Belfrage, P., Fredrikson, G., Stralfors, P., and Tornqvist, H. (1984).In “Lipases” (B. Borgstrom and H. I. Brockman, eds.), pp. 365-416.Elsevier, Amsterdam. Ben-Zeev, O., Ben-Avram, C. M., Wong, H., Nikazy, J., Shively, J. E., and Schotz, M. C. (1987).Biochim. Biophys. Acta 919, 13-20. Blow, D. M. (1990).Nature (London) 343,694-695. Blow, D. M., Birktoft, J., and Hartley, B. (1969).Nature (London) 221,337-339. Bodmer, M. W., Angal, S., Yarranton, G. T., Harris, T. J. R., Lyons, A., King, D. J., Pieroni, G., Riviere. C., Verger, R., and Lowe, P. A. (1987).Biochim. Biophys. Acta 909,

237-244. Boel, E., Huge-Jensen, B., Christiansen, M., Thim, L., and Fiil, N. P. (1988).Lipzds 23,

701-706. Boel, E., Brady, L., Brzozowski, A. M.. Derewenda, Z., Dodson, G. G., Jensen, V. J., Petersen, S. B., Swift, H., Thim, L., and Woldike, H. F., (1990).Biochemistry 29,

6244-6249. Boel, E., Huge-Jensen, B., Woldike, H., Gormsen, E., Christensen, M., Andreasen, F., and Thim, L. (1991).GBFMonogr. Ser. 16,207-219. Borgstrom, B., and Erlanson-Albertsson, C. (1984).In “Lipases” (B. Borgstrom and H. L. Brockman, eds.), pp. 152-183.Elsevier, Amsterdam. Brady, l.,Brzozowski, A., Derewenda, Z., Dodson, E., Dodson, G., Tolley, S., Turkenburg, J., Christiansen, L., Huge-Jensen, B., Norskov, L., Thim, L., and Menge, U. (1990).Nature (London) 343,767-770. Breddam, K. (1986).CarLtberg Res. Cummun. 51,83-128. Brenner, S. (1988).Nature (London) 334,528-530. Brockerhoff, H. (1968).Biochim. Biophys. Acta 159,296-303. Brockman, H.L., Law, J. H., and Kezdy, F. J. (1973).J.B i d . Chem. 248,965-970. Bryan, P., Pantoliano, M. W., Quill, S. G., Hsiao, H.-Y., and Poulos, T. (1986).Proc. Natl. Acad. Sci. U.S.A. 83, 3743-3745. Brzozowski, A. M., Derewenda, U., Derewenda, Z. S., Dodson, G. G., Lawson, D. M., Turkenburg, J. P.,Bjorkling, F., Huge-Jensen, B., Patkar, S. A., and Thim, L. (1991). Nature (London) 351,491-494. Canioni, P., Julien, R., Rathelot, J., and Sarda, L. (1977).Lipids 12,393-397. Chapus, S., Rovery, M., Sarda, L., and Verger, R. (1988).Biochimie 70, 1223-1234. Charlton, E., and Macrae, A. R. (1992).Biochim. Biophys. A d a 1123,59-64. Cleansby, A., Garman, E., Egmond, M. R., and Batenburg, M. (1992).J. Mol. Biol. 224,

281-282. Dartois, V., Baulard, A., Schanck, K., and Colson, C. (1992).Biochim. Biophys. Acta 1131,

253-260. Datta, S., Luo, C.-C., Li, W.-H., VanTuinen, P., Ledbetter, D. H., Brown, M. A., Chen, S.-H., Liu, S.,and Chan, L. (1988).J.Biol. Chem. 263, 1107-1110.

48


Davis, R. C., Wong, H., Nikazy, J., Wang, K., Han, Q.. and Schotz, M. C. (1992).J. Biof. Chem. 267921499-2 1504. De Caro, J., Boudouard, M.,Bonicel, J., Guidoni, A.. Desnuelle, P., and Rovery, M. (1981).Biochim. Biophys Actu 671, 129-138. Deckelbaum, R. J., Ramakrishnan, R., Eisenberg, S., Olivecrona, T., and BengtssonOlivecrona, G. (1992).Biochistty 31,8544-8551. Derewenda, U., Brzozowski, A. M., Lawson, D. M., and Derewenda, Z. S. (1992).Biochemistry 31, 1532-1541. Derewenda, Z.S., and Cambillau, C. (1991).J.Biol. C h . 266,23112-231 19. Derewenda, Z. S., and Derewenda, U. (1991).Biochem. Cell B i d . 69,842-851. Derewenda, 2.S.,and Sharp, A. M. (1993).Trends Biochem. Sci. 18,20-25. Derewenda, Z.S.,Derewenda, U., and Dodson, G. G . (1992).J.Mol. Bzol. 227,818-839. Desnuelle, P. (1986).In “Molecular and Cellular Basis of Digestion” (P. Desnuelle, ed.), pp. 275-296. Elsevier, Amsterdam. Desnuelle, P., Sarda, L., and Ailhard, G. (1960).Biochim. Biophys. Actu 37,570-571. DiPersio, L. P., and Hui, D. Y. (1993).J.Biof. Chem. 268,300-304. Docherty, A. J. P., Bodmer, M. W.,Angal, S., Verger, R., Riviere. C., Lowe, P. A., Lyons, A., Emtage, J. S., and Harris, T. J. R. (1985).NucleUAcidcRes. 13, 1891-1903. Donner, J., Spink, C., Borgstrom, B., and Sjoholm, I. (1976).Biochemistry 15,5413-5417. Efimov, A. V. (1986).Mof. Bwl. (Moscow) 20,250-260. Farber, G . K., and Petsko, G . A. (1990).Trends Biochem. Sci. 15,228-234. Faustinella, F., Smith, L. C., Semenkovich, C. F., and Chan, L. (1991).J.Biol. Chem. 266,

9481-9485. Faustinella, F., Smith, L. C., and Chan, L. (1992).Biochemistry 31, 7219-7223. Feller, G., Thiry, M., and Gerday, C. (1991a).Biochim. Biophys. Acta 1088,323-324. Feller, G., Thiry, M., and Gerday, C. (1991b).DNA Cell Biof. 10,381-388. Ferri, S.,and Meighen, E. A. (1991).J.Biof. Chem. 266, 12852-12857. Franken, S., Rozeboom, H. J., Kalk, K. H., and Dijkstra, B. W. (1991).EMBOJ. 10,

1297- 1302. Gandour, R. D. (1981).Bioorg. C h . 10, 169-176. Garfinkel, A. S., and Schotz, M. C. (1987).In “Plasma Lipoproteins” (A. M. Gotto, Jr., ed.), pp. 335-357. Elsevier (Biomed. Div.), New York. Garfinkel, A. S., Kempner, E. S., Ben-Zeev, O., Nikazy, J., James, S. J., and Schotz, M. C. (1983).J. Lipd Res. 24,775-780. Gargouri, Y., Moreau, H., Pieroni, G.. and Verger, R. (1988).J.B i d . Chem. 263, 2159-

2162. Gargouri, Y.,Moreau, H., Jain, M., deHaas, G. H., and Verger, R. (1989a).Biochim. B q h y s . Acta 1006,137-139. Gargouri, Y., Moreau, H., and Verger, R. (1989b).Biochim. Biophys. Acta 1006,255-271. Gargouri, Y., Chahinian, H., Moreau, H., Ransac, S., and Verger, R. (1991).Biochim. Biophys. Acta 1085,322-328. Giller, T., Buchwald, P., Blum-Kaelin, D., and Hunziker, W. (1992).J. Biof. Chem. 267,

16509-16516. Goldstein, J. L., and Brown, M. S. (1977).Annu. Rev. Biochem. 46, 897-930. Gotz, F.,Popp, F., Korn, E., and Schleifer, K. H. (1985).Nucleic Acids Res. 13,5895-5906. Grochulski, P., Schrag, J., and Cygler, M. (1992).Proc. Am. Crystullogr. Assoc., 1992 Annu. Meet., p. 122. Haas, M.J., Allen, J., and Berka, T. R. (1991).Gene 109, 107-1 13. Hahn, P. F. (1943).Science 98, 19-20. Hamosh, M. (1984).In “Lipases” (B. Borgstrom and H. L. Brockman, eds.), pp. 49-81. Elsevier, Amsterdam.


49

Han, J. H., Stratowa, C., and Rutter, W. J. (1987). BiochemGtty 26, 1617-1625. Harwood, J. (1989). TrendcBiochem. Sci. 14, 125-126. Hassanien, F. R., and Mukherjee, K. D. (1986).J.Am. Oil C h . SOL.63,893-897. Hata, Y., Matsura, Y., Tanaka, N., Kakudo, M., Sugihara, A., Iwai, M., and Tsujisaka, Y. (1979).]. Biochem. (ToRyo)86,1821-1827. Henderson, R. (1970).J. Mol. Biol. 54,34 1-354. Holm, C., Kirchgessner, T. G., Svenson, K. L., Fredrikson, G., Nilsson, S., Miller, C. G., Shively, J. E.,Heizmann, C., Sparkes, R. S., Mohandas, T., Lusis, L. J., Belfrage, P., and Schotz, M. C. (1988). Science 241,1503-1506. Hui, D. Y., and Kissel. J. A. (1990). FEBSLett. 276, 131-134. Jensen, G. L., and Bensadoun, A. (1981).Anal. Biochem. ,113,246-252. Johnson, L. A., Beacham, 1. R., MacRae, I. C., and Free, M. L. (1992). Appl. Environ. Microbial. 58, 1776-1779. Jorgensen, S . , Skov, K. W., and Diderichsen, B. (1991).J.Bactmiol. 173,559-567. Kamphuis, L. G., Kalk, K. H., Swarte, M. B. A., and Drenth, J. (1984).J. Mol. Biol. 179, 233-256. Kazlauskas, R. J.. Weissfloch, A. N. E., Rappaport, A. T., and Cuccia, I. A. (1991).J. Org. Chem. 56,2656-2665. Kim, K. K., Hwang, K. Y., Jeon, H. S., Kim, S . , Sweet, R. M., Yang, C. H., and Suh, S. W. (1992).J.Mol. Biol. 227, 1258-1262. Kirchgessner, T. G., Chaut, J. C., Heinzmann, C., Etienne, J., Guilhot, S., Svenson, K., Ameis, D., Pilon, C., d’Auriol, L., Andalibi, A., Schotz, M. C., Galibert, F., and Lusis, A. J. (1989).Proc. Natl. Acad. Sci. U.S.A. 86,9647-9651. Kissel, J. A., Fontaine, R. N., Tuck, C. W., Brockman, H. L., and Hui, D. Y. (1989). Biochim. Biophys. Acta 1006,227-236. Kobayashi, K., Lin, LW.,Yeadon, J. E., Klickstein, L. B., and Smith, J. A. (1989).J. Biol. Chem. 264,8892-8899. Komaromy, M. C., and Schotz, M. C. (1987). Proc. Natl. Acad. Sci. U.S.A. 84, 1526-1530. Kordel, M., Hofmann, B., Schomburg, D., and Schmid, R. (1991).J.Bacterial. 173,4836484 1 . Kraut, J. (1977).Annu. Rev. Biochem. 46,331-358. Kugimiya, W., Otani, Y., Hashimoto, Y., and Takagi, Y. (1986). Biochem. Bzophy. Res. Commun. 141, 185-190. Kyger, E. M., Weigand, R. C., and Lange, L. G. (1989). Biochem. Biophys. Res. Commun. 164,1302-1309. Lagrand, G., Triantaphylides, C., and Baratti, J. (1988). Biohchnol. Lett. 10,549-554. Larson, S., Day, J., Greenwood, A., Oliver, J.. Rubingh, D., and McPherson, A. (1991). J . Mol. Biol. 222, 21-22. Lawson, D. M., Brzozowski, A. M., and Dodson, G. G. (1992). Cum. Biol. 2,473-475. Lawson, D. M., Dodson, G. G., Hubbard, R. E.,Huge-Jensen, B., Boel, E., Brzozowski. A. M., and Derewenda, Z. S . (1993). In “Lipases, Their Biochemistry, Structure and Applications” (P. Wooley, ed.). Keminsk Institut, Aarhus Universitet (in press). Lee, C. Y.,and Iandolo, J. J. (1986).J. Battniol. 166,385-391. Lesk, A., Branden, G I . , and Chothia, C. (1989). Proteins: S t r w t . , Funct. Genet. 5, 139148. Leszczynski,J . F., and Rose, G. D. (1986). Science 234,849-853. Liao. D.-I., Breddam, K., Sweet, R. M., Bullock, T., and Remington, S. J. (1992). Biochemislv 31,9796-9812. Lie, E., Persson, A., and Molin, G. (1991).Appl. Murobiol. Biotechnol. 35, 19-20. Lockridge, D.. Bartels, C. F., Vaughen, T. A., Wong, C. K., Norton, S. E., and Johnson, L. L. (1987).J.Biol. Chem. 262, 549-557.

50


Lowe, M. E. (1992).J.Biol. Chem. 267, 17069-17073. Luthi-Peng, Q.,and Winkler, F. K. (1992).Eur.1. Biochem. 205,383-390. Maraganore, J. M., and Heinrikson, R. L. (1986).Trends Biochem. Sci. 11,497-498. Margolin, A. I., and Klibanov, A. M. (1987).J.Am. Chem. SOL.109,3802-3804. Marquart, M., Deisenhofer, J., Huber, R., and Palm, W. (1980).J . Mol. Biol. 141, 369-

391. Martinez, C., De Geus, P., Lauwereys, M., Mathyssens. G., and Cambillau, C. (1992). Nafure (London)356,615-6 18. Matthews, B. W. (1972).Macromolecules 5,818-819. Matthews, D. A., Alden, R. A., Birktoft, J. J., Freer, S. T., and Kraut, j.(1977).J.Bzol. Chem. 252,8875-8883. Matsuura, Y., Kunusoki, M., Harad, W.,and Kakudo, M. (1984).J . Biochem. (Tokyo) 95,

697-702. McGrath. M. E., Vasquez, J. R., Craik, C. S., Yang, A. S., Honig, B., and Fletterick. R. J. (1992).BimhLmistry 31,3059-3064. Menard, R., and Storer, A. C. (1992).Biol. C h . Hoppe-Seyler 373,393-400. Mercken, L., Simons, M. J.. Swillens, S., Massaer, M., and Vassart. G. (1985).Nafure (London)316,647-65 1. Meyer, E., Cole, G . , Radhakrishnan, R.,and Epp, 0. (1988).Acta Crystallogr., Sect. B BU,

26-38. Moreau, H., Gargouri, Y., Pieroni, G., and Verger, R. (1988).FEES Left. 236,383-387. Moreau, H., Moulin, A., Gargouri, Y., Noel, J.-P., and Verger, R. (1991).Biochemistry 30,

1037- 1041. Moreau, H., Abergel, C., Carriere, F., Ferrato. F.. Fontencilla-Camps. J. C., Cambillau, C., and Verger, R. (19921.1.Mol. Biol. 225, 147-153. Naylor, S. I., Marshall, A., Hensel, C., Martinez, P. F., Holley, B., and Sakaguchi, A. Y. (1989).Cenomics 4,355-361. Odera, M., Takeuchi, K., and Toh-e, A. (1986).J.Fennenf. Technol. 64,363-37 1. Odya, C. E., and Erdos, E. G. (1981).In “Methods in Enzymology” (L. Lorand, ed.), Vol. 80, pp. 460-466. Academic Press, New York. Ogata, S., Misumi, Y., and lkehara, Y. (1989).J.Biochem. (Tokyo) 106,548-551. Olivecrona, T., and Bengtsson, G. (1984).In “Lipases” (B. Borgstrom and H. 1. Brockman, eds.), pp. 205-261. Elsevier, Amsterdam. Ollis, D. I., Cheah, E., Cygler, M., Dijkstra, B., Frolow, F., Franken, S. M., Harel., M., Remington, S. J., Silman, I., Schrag, J., Sussman, J., Verschueren, K. H. G., and Goldman, A. (1992).Proleins 5, 197-21 1. Osborne, J. C., Bengtsson-Olivecrona, G . , Lee, N.S., and Olivecrona, T. (1985).Biochemistry 44,5606-56 1 1. Pathak, D., and Ollis, D. (199O).J.Mol. Biol. 214,497-525. Pathak, D.,Ashley, G., and Ollis, D. (1991).Proteins: Sfmf., Funct., Genet. 9,267-279. Persson, B., Bengtsson-Olivecrona, G., Enerback, S., Olivecrona, T., and Jornval, H. (1989).Eur.1. Biochem. 179,39-45. Ransac, S., Gargouri, Y., Moreau, H., and Verger, R. (1991).Eur. J. Biochem. 202,

395-400. bwlings, N. D., Polgar, I., and Barrett, A. J. (1991).Bioehcm. J. 279,907-91 1. Read, R. J., and James, M. N. G. (1988).J.Mol. Biol. 200,523-55 1. Reddy, M. N., Maraganore, J. M., Meredith, S. C., Heinrikson, R. I., and Kezdy, F. J. (1986).J.Biol. Chem. 261,9678-9683. Rennex, D., Hemmings, B. A., Hofsteenge, J., and Stone, S. R. (1991).Biochemisfry 30,

2195-2203.


51

Richardson, J. S. (1981).Adv. Protein Chem. 34, 167-339. Roberts, C. J., Pohling, G., Rothman, J. H., and Stevens, T. H. (1989).J.Cell Biol. 108,

1363-1373. Robertus, J. D., Kraut, J., Alden, R. A., and Birktoft, J. (1972).Biochemistry 11, 4293-

4303. Sacchettini,J. C., Gordon, J. L., and Banaszak, L. J. (1988).J.Biol. C h . 263,5815-5819. Sacchettini, J. C., Gordon, J. L., and Banaszak, L. J. (1989).J Mol. Biol. 408,327-339. Sando, G. N., and Rosenbaum, L. M. (1985).J.B i d . C h . 460,15186-15193. Sarda, L., and Desnuelle, P. (1958).Biochim. Biophys. Acta 30,513-521. Sarma, R., Dauberman, J., Poulose, A. J., van Bielen, J., Power, S., Shew, B., Gray, G., Norton, S., van Kimmenade, A., Jarnagin, A., Ashizawa, E., Boston, M., Requandt, C., Kolattukudy, E., and Bott, R. (1991).GBFMonogr. Ser. 16,71-77. Schmitz, G., and Assman, G. (1989).In “The Metabolic Basis of Inherited Disease” (C. R. Scriver, A. I. Beaudet, W. S. Sly, and D. Valle, eds.), 6th ed., Vol. 2, pp. 1623-1644. McGraw-Hill, New York. Schrag, J. D., Li, Y., Wu, S., and Cygler, M. (1991).Nature (Londcm)351,761-764. Schrag, J. D.,Winkler, F. K., and Cygler, M. (1992).J. Biol. C h . 467,4300-4303. Schumacher, M., Camp, S., Maulet, Y., Newton, M., MacPhee-Quigley, K.,Taylor, S. S., Friedman, T., and Taylor, P. (1986).Nature (London) 319,407-409. Semeriva, M., and Desnuelle, P. (1979).Adv. Enzymol. 48,319-370. Sibanda, B. I., Blundell, T. L., and Thornton, J. M. (1989).J.Mol. Biol. 406, 759-777. Smith, L. C., and Pownall, H. J. (1984).In “Lipases” (B. BorgstrOm and H. 1. Brockman, eds.), pp. 263-305.Elsevier, Amsterdam. Smith, L. C., Faustinella, F., and Chan, 1. (1992).Curr. Opin. S t m t . Biol. 4,490-495. Smith, S. O., Farr-Jones, S., Griffin, R. G., and Bachovchin, W. W. (1989).Science 444,

961-964. Stone, S. R., Rennex, D., Wokstrom, P., Shaw, E., and Hofteenge, J. (1991).Biochem. J.

276,837-840. Stralfors, P., and Belfrage, P. (1983).j. Biol. Chem. 258, 15146-15152. Stralfors, P., Bjorgell, P., and Belfrage, P. (1984).Proc. Natl. Acud. Sci. U.S.A. 81, 3317-

3321. Sugihara, A., Hata, Y., Shimada, Y., Tominaga, Y., and Katsube, Y. (1991).CBF Monogr. Ser. 16,47-54. Sussman, J. L., Harel, M., Frolow, F., Oefner, C., Goldman, A., Toker, L.,and Silman, L. (1991).Science 453,872-879. Swenson, L., Ferri, S. R.,Green, R., Sharp, A. M., Meighen, E. A., and Derewenda. Z. S. (1992).J. Mol. Bwl. 447,572-574. Twu, J.-S., Garfinkel, A. S., and Schotz, M. C. (1984).Biochim. Blophys. Acta 794, 330-

337. Valivety. R. H., Halling, P. J., and Macrae, A. R. (1992).FEES Lett. 301,258-260. van Tilbeurgh, H., Sarda, L., Verger, R., and Cambillau, C. (1992).Nature (London) 359,

159-162. van Tilbeurgh, H., Egloff, M.-P., Martinez, C., Rugani, N., Verger, R., and Cambillau, C. (1993).Nature (London) 362,814-820. Verger, R., Rietsch, J.. and Desnuelle, P. (1977).J.Biol. C h . 452,4319-4325. Voelker, T. A., Woreell, A. C., Anderson, I., Bleibaum, J., Fan, C., Hawkins, D. J.. Radke, S. E., and Davies, H. M. (1992).Science 457,72-74. Wang, C.-S., Heartsuck, J.. and Downs, D. (1988).Biochemktty 47,4834-4840. Wang, C.-S., Heartsuck, J., and McConathy, W. J. (1992).Biochim. Biophys. Acta 1123,

1-17.

52

ZYGMUNT S . DEREWENDA

Warshel, A., Naray-Szabo, G . , Sussman, F., and Hwang, J.-K. (1989). Biochemistry 28, 3629-3637.

Wells, M. A. (1974). Biochemistry 13,2248-2257. West, J. B., and Wong, C.-H. (1986). Tetrahedron Lett. 28, 1629-1632. Winkler, F. K., DArcy. A., and Hunziker. W. (1990). Nature (London) 343,771-774. Wong, H., Davies. R. C., Nikazy, J., Seebart, K. E., and Schotz, M. C . (1991). Proc. Natl. A c ~S .C ~U.S.A. . 88, 1 1290- 1 1294. Xie, Z.-F., Suemune, H., and Sakai, K. (1990). Tetrahedron: Asymmetry 1, 395-402. Yamaguchi, S., and Mase, T. (1991). AH1. Microbzol. Biotechnol. 34, 720-725. Yamaguchi, S., Mase, T., and Tekeuchi, K. (1991). Gene 103,61-67. Yamaguchi, S., Mase, T., and Tekeuchi, K. (1992). Bioscz. Biotechnol. Biochem. 556, 3 15-3 19.

Zimrnerrnan, S. C., Korthals, J. S., and Kramer, K. D. (1991). Tetrahedron 47,2649-2660.

STRUCTURE AND CATALYTIC MECHANISM OF SECRETORY PHOSPHOLIPASESA2 By DAVID L. SCOTT and PAUL B. SIGLER The Department of Molecular Biophyslca and Biochemistry and the Howard Hughes Medical Institute, Yale University, New Haven, Connectlcut 06510

I.

Introduction

.......................................................

11. Primary Structure and Classification .................................. 111. Secondary and Tertiary Structure ....................................

A. Conserved Substructures ........................................ IV. Chemistry of Catalysis of Secretory Phospholipases A2 .................. A. Productive-Mode Substrate Binding .............................. B. Formation and Collapse of Tetrahedral Intermediates .............. C. Comparison with Serine Proteases ................................ V. Interfacial Catalysis ................................................. A. “Hopping” versus “Scooting” ..................................... B. Interfacial Adsorption Surfaces .................................. C. Electrostatic Considerations ...................................... D. Interfacial Adsorption and Catalytic Efficiency ..................... References .........................................................

53 54 58 60 66 67 71 74 75 75 76 77 78 80

I. INTRODUCTION Phospholipases A2 (PLA~s,EC 3.1.1.4) stereospecifically hydrolyze the sn-2 ester bond of 1,2-diacyl-3-sn-phosphoglycerides (van Deenen and de Haas, 1963).Interest in PLA2s arises from their unusual adaptation to perform catalysis at the interface between two bulk phases. As such, they represent a paradigm in our understanding of the interaction between proteins and organized lipids (membranes). In addition, release of arachidonic acid from the sn-2 position of membrane glycerophospholipids plays a key regulatory role in the synthesis of prostaglandins and other mediators of inflammation (Dennis, 1987;Burgoyne and Morgan, 1990;Chock et al., 1991). Two types of PLA2s have been described. Secretory PLA2s (sPLA2s) are robust, small enzymes (14 kDa) that are highly resistant to denaturation (Verheij et al., 1981;Dennis, 1983; Waite, 1987). The most striking kinetic feature of sPLAzs is their preference for substrates organized into micelles, monolayers, vesicles, o r membranes. Activity of sPLA2s on lipid aggregates is often three orders of magnitude greater than that on dispersed (monomeric) substrate. Calcium ion, although weakly bound by sPLAls (Kd > M),is essential for catalysis. Other metal ions either fail to support catalysis or competitively inhibit calciumADVANCES IN PROTEIN CHEMISTRY, Val. 4 1

53

Copyright 01994 by Academic Press, Inc. All rights of reproduction in any form reserved.

54

DAVID L. SCOTT AND PAUL B. SICLER

mediated activity (Pieterson et al., 1974a; Hershberg et al., 1976). One or possibly two calcium ions are required per enzyme molecule for optimal activity (Scott et al., 1990a; Scott and Sigler, 1993). Members of this large family of homologous proteins are readily isolated in milligram quantities from exocrine pancreas, reptile and insect venoms, and mammalian nonpancreatic sources. In contrast, the more complex (1 10 kDa) intracelM a r enzymes have only recently been isolated in amounts sufficient for detailed biochemical and kinetic investigations (Clark et al., 1990, 1991; Kramer et al., 1991). This review will focus on aspects of sPLA2 structure that are of particular relevance to catalysis. We have proposed a formal reaction pathway based largely on high-resolution structure analysis of crystalline complexes of sPLA2s with a transition-state analog (Scott et al., 1990a). This mechanism, which is consistent with the available biochemical and genetic data (Waite, 1987), implies a highly specific interaction between the calcium cofactor, the glycerophospholipid substrate, and the active site residues. Conserved water molecules, associated with either the catalytic histidine or the primary calcium ion, are essential participants in the proposed chemistry. 11. PRIMARY STRUCTURE AND CLASSIFICATION

More than 60 secretory PLA2s have been isolated and sequenced (Dufton and Hider, 1983; Heinrikson, 1991). With a few exceptions, these proteins are highly homologous and contain between 119 and 143 amino acids. Sequence alignment is based on strong three-dimensional structural homology and is simplified by the large proportion of conserved or invariant residues (Fig. 1). Most of the conserved residues either directly participate in catalysis or occupy structurally sensitive positions (Table I). Conserved cysteine residues are particularly abundant (10 or 14 cysteinedsequence) and form a rigid network of disulfide bridges that stabilizes much of the tertiary structure. Secretory PLA2s can be divided into three classes based on sequence criteria (Randolph et al., 1980). The highly homologous class I and class I1 enzymes are distinguished by the location of one of their seven disulfide bridges and by the sequence of the carboxyl terminus. T h e sevenamino acid carboxyl-terminal extension of class I1 enzymes forms a distinctive bannister overhanging the primary calcium-binding site but has no known function. Indeed, from a kinetic viewpoint, class I and class I1 sPLA2s are identical (Heinrikson, 1991). Class 1 enzymes are abundant in Elapidae/Hydrophidae snake venoms and in mammalian exocrine pancreas and have recently been isolated from human spermatazoa (Langlais

Apismelljrcra

3

1

I

9

I

I

m

3

l

0

Q

I I YPGTLWCOHONKS SGPNELORPKHTDACCRTH 3 6 4 0

!MCP

m

45

ss

so

60

DVMS A O E S K H O L T N T A S HTRLSCDC! 7s

80

rug

90

95

4

D KF Y D 100

C L K N S ADTI S S YFVOKMYF N L I DT KCYKL E H P VT 0 1M 110 11s im 123 130 134 C G E R T E O R C L H Y T V D K SK P K V Y O W F D L R K Y

B N

Class I

C h s II

Class

m

FIG. 1. Secretory phospholipase A2 classes: primary and secondary structure. (A) Representative sequences for class 1 [porcine exocrine pancreas (Puijk el ul., 1977), Nuju nuju atra venom (Tsai et al., 1981)], class I1 [human nonpancreatic (Seilhamer etal., 1989)], and class I l l [bee venom (Kuchler etal., 1989)l enzymes.The sequences ofclass 1/11 enzymes are highly homologous and have been aligned using a common numbering system (Renetseder et al., 1985). Asterisks denote residues that are identical to those of the porcine enzyme at that position. W, Residues involved in coordination of the primary calcium ion; 0, the catalytic histidine; 0 ,supporting residues of the catalytic network. (B) Schematic representations of class I, class 11, and class I11 sPLAls based on the known three-dimensional structures. The diagrams are arranged to highlight the marked structural homology and are not necessarily to scale. The a helices are indicated by rectangles, p strands by arrows, disulfide bridges by black bars, and the primary calcium ion by a circle. The amino and carboxyl termini are labeled N and C, respectively.

56


TABLE I Conserved or Invariant Amino Acid Residues among Class 1111 Secretory Phospholipases A20 Sequence number

Residue(s) present

2 3 4 5 8 9 1 I* 18 19 22 25 26 27 28 29 30 31 32 33 35 37 39 40 41 42 43 44 45

L-V(4) Hydrophobic Q-E(WN(4) F M-L-V

48

49 50** 51 52 55 61 62* 63* 64* 65* 66* 68 69 73 75

1

c-c77 Hydrophobic Hydrophobic F-Y

Y G C-C 126 Y-F(1) c-c45 G Nonanionic G G G P-A(2) D A-E-D L-T-S D R-K(2)-A(1) C-C 105 C-C29 H D C-C 133 C-C98 Y A-D(1) c-c91 K-R( 1) F-V(2) L L-v D P-T(4) Y-K Y Y-F(3)-W(2)

Apparent function Contributes to entrance of hydrophobic channel Interfacial adsorption surface Stabilizes amino terminus and adjacent loop (73-78) Supports His-48 in active site Contributes to active site, may stabilize helix (96-99) Adjacent to residue 5 in hydrophobic channel Anchors amino-terminal helix and tip of p wing Contributes to interfacial adsorption surface Contributes to interfacial adsorption surface Contributes to hydrophobic channel Stabilizes Asp-39 and backbone at position 112 Structural (pre-calcium-binding loop) Anchors calcium-binding loop and carboxyl terminus* Stabilizes distal segment calcium-binding loop Anchors calcium-binding loop Calcium-binding loop Contributes to interfacial adsorption surface Calcium-binding loop Calcium-binding loop Calcium-binding loop Structural (forms turn proximal to helix C) Forms cap at proximal base of helix C ?

Contributes to back wall of hydrophobic channel Stabilizes calcium-binding loop (N-27, N-28, and N-38) ?

Braces antiparallel helices (C and E) Anchors calcium-binding loop Catalysis Coordinates primary calcium ion Stabilizes carboxyl terminus Braces antiparallel helices (C and E) Catalytic network-structural support Lies under Tyr-52 Stabilizes helix D*/loop** ? ? Contributes to interfacial adsorption surface Contributes to interfacial adsorption surface H bonds to residue 62: stabilizes loop (62-66) Structural-backbone directional change Binds p r o 3 nonbridging oxygen of sn-3 phosphate Catalytic network-structural support Structural support for distal amino-terminal helix

SECRETORY PHOSPHOLIPASES

A2

57

TABLE I (continued) Sequence number

Residue@) present

77* 83 84 95 96 98 99

c-CI 1 Hydrophobic C-C96

102 I03 I05 I06 I13 126 I33**

I-v-L C-C84 c-c5 1 D A A

c-C44 F-L( 1) Y C-C27 C-C50

Apparent function ?

Stabilizes p wing Underlies/support Tyr-52 Stabilizes /3 wing Catalytic network Contributes to back wall of channel Contributes to back wall of channel

-

Contributes to back wall of channel Structural (hydrogen bonds to 0 - 2 1 2 others)

-

“This analysis was based on 50 sequences chosen from the literature (Heinrikson, 1991). Parentheses indicate the number of sequences that bear a “minority” residue. One or two asterisks indicate residues found only in class I or class I1 species, respectively.

et al., 1992). Class I1 proteins are present in Crotalidae/Viperidae snake venoms and are broadly distributed among a variety of mammalian cell types, including platelets, gastric mucosae, and the vascular endothelium (Johnson et al., 1990; Kramer et al., 1989; Komada et al., 1989; Kurihara et al., 1991). Class I1 I sPLAps include the evolutionarily divergent venom enzymes from the European honeybee Apis mellifera (Kuchler et al., 1989), the Gila monster Heloderma suspectum (Gomez et al., 1989), and the Mexican bearded lizard Heloderma hom’dum horridum Wiegman (Sosa el al., 1986). The bee enzyme is relatively insensitive to the aggregation state of its substrate and hydrolyzes dispersed and aggregated substrates at similar rates (Raykova and Blagoev, 1986; Lin et al., 1988).Although several of the bee venom isoforms are N-glycosylated, their kinetic characteristics are identical to their nonglycosylated counterparts (Marz et al., 1983; Dudler et al., 1992). In addition to the three classes of active sPLAps, a group of sPLAp-like proteins has also been reported (Maragnore et al., 1984). These proteins bear high sequence homology toward class 1/11 enzymes but are catalytically inactive due to one or more amino acid substitutions. The K49 protein family, in which the normally invariant Asp-49 is replaced with a

58


lysine residue, is among the best studied members of this group (van den Bergh et al., 1989; Yoshizumi et al., 1990; Francis et al., 1991; Scott et al., 1992). 111. SECONDARY AND TERTIARY STRUCTURE

High-resolution X-ray crystal structures are available for representatives of all three sPLA2 classes (Fig. 2). T h e three-dimensional structures are complemented by detailed solution studies [infrared, Raman, and nuclear magnetic resonance spectroscopy (Areas et al., 1989; Fisher et al., 1989; Williamson et al., 1989; Kennedy et al., 1990; Roberts, 1991)] and molecular dynamics simulations (Campbell et al., 1988; Sessions et al., 1989; Gros et al., 1990; Demaret and Brunie, 1990). Interpretation of the three-dimensional models is also aided by kinetic results obtained from an extensive library of chemically o r genetically modified enzymes (Waite, 1987). Secretory PLA2 species have been cloned and overexpressed from bovine and porcine exocrine pancreas (Deng el al., 1990; De Geus et al., 1987), human nonpancreatic sources [synovial fluid (Seilhamer et ul., 1989; Kramer et al., 1989; Bomalaski et al., 1991)], and the venom glands of Naja naju nuja (Kelley et al., 1992) and A. mellifera (Dudler et al., 1992). Several kinetic studies are compatible with models in which sPLA2 forms dimers or higher order oligomers at the interface in order to exhibit full enzymatic activity (Cho et al., 1988; Tomasselli et al., 1989; Bell and Biltonen, 1989a,b). Comparison of the refined crystal structures of three sPLA2s that form stable dimers in solution fails to suggest a catalytic advantage to dimerization (Brunie et al., 1985; Achari et al., 1993; Scott et al., 1993a). Indeed, sPLA2 aggregation is not obligatory because the monomeric form of at least one of these enzymes efficiently hydrolyzes substrate when trapped on anionic vesicles (Jain et al., 1991). The overall dimensions of a typical class 1/11 sPLA2 are roughly 22 X 30 X 42 8, with 50% of the structure Q helix and 10% p sheet. Crystalline class I and class I1 enzymes share a homologous “core” of invariant tertiary structure (Renetseder et al., 1985). This core includes three a helices (residues 1 to 12,37 to 54, and 90 to 109) and a distinctive backbone loop (residues 24 to 30). Amino acid side chains arising from the homologous core coordinate the primary calcium ion, define the substrate-binding pocket, and directly mediate the bond-breaking events of catalysis. Not surprisingly, two of the three a helices, as well as the backbone “calcium-binding” loop, are also present in the class 111 bee venom sPLA2 (residues 25-37, 61-74, and 8-12). Throughout this review, functional homology between class 1/11 and class 111 residues will be

SECRETORY PHOSPHOLIPASES A2

59

16

6

79

FIG. 2. The tertiary structure of secretory phospholipases A2: a-Carbon traces of (A) the class I Naja nuja atra venom sPLA2 (White el al., 1990), (B) the class I1 human nonpancreatic sPLAz (Scott etal., 1991),and ( C )the class I l l bee venom enzyme (Scott etal., 1990b). The location of the primary calcium ion is indicated by the large black sphere, the position of the cocrystallized transition-state analog is shown in A and C, and the side chains of His-48 and Asp-99 are shown in B. The face of the enzyme that corresponds to the proposed interfacial adsorption surface is inscribed. The residues in A and B are labeled according to the common numbering system (Renetseder et af., 1985).

60

DAVID L. SCOTT AND PAUL B. SIGLER

indicated by displaying the corresponding sequence numbers separated by a slash mark (e.g., His-48/34).

A . Conserved Substructures 1 . Amino Termini The amino-terminal nitrogen of a class 1/11sPLA2 lies at the center of an extensive network of hydrogen bonds that interconnect the aminoterminal helix, an adjacent segment of the /3 wing (residues 74-84), and the active site Asp-99 (Fig. 3). Disruption of this network markedly impairs adsorption at lipid-water interfaces. For example, transamination of the N-terminal amino group of the bovine pancreatic enzyme reduces catalysis on micelles by two orders of magnitude without affecting the k,,, measured on dispersed substrates. The failure of the transaminated enzyme to exhibit the normal preference for aggregated substrates resembles that of the native proenzyme, which does not bind to the interface (Abita et al., 1972; Pieterson et al., 1974b). In both cases, structural studies indicate that residues 1-4 and 62-73 are partially disordered (Dijkstra et al., 1982, 1984). This disorder is eliminated by tryptic removal of the N-terminal heptapeptide during the proenzyme-toenzyme maturation (Dijkstra et al., 1983). In contrast to its class 1/11counterparts, the amino terminus of the class 111 bee venom sPLA2 does not communicate with the catalytic machinery (Scott et al., 1990b). Instead, the amino-terminal nitrogen stabilizes the conformation/position of the calcium-binding loop by forming a watermediated hydrogen bond with the side chain of His- 11. The active site Asp-64 (the class 111 analog of Asp-99) is, however, connected by a network of water-mediated hydrogen bonds to the backbone nitrogen of Leu-59 (Fig. 3). This nitrogen, like the amino-terminal nitrogen of class 1/11 enzymes, also lies at the base of an a helix and forms several additional stabilizing contacts with adjacent residues. 2 . Amino-Terminal Helix (Residues 1 to 12) In class 1/11 crystal structures, the side chains of residues 2, 5, and 9 contribute to the “right” wall and mouth of the hydrophobic channel occupied by the acyl chains of productively bound substrate (Fig. 4) (White et al., 1990; Thunnissen et al., 1990; Scott et al., 1990a, 1991). Residue 2 lies directly at the opening of the channel and is always either a leucine or a valine, whereas sequence positions 5 and 9 are occupied by an invariant Phe and Ile, respectively. Conservation of Phe-5 is particularly critical because the phenolic ring lies sandwiched at the bottom of the hydrophobic channel between Ile-9 and the catalytic histidine.

SECRETORY

PHOSPHOLIPASES A*

61

d4

B 48

/

\

A

ll73

FIG.3. The catalytic networks of secretory phospholipases AP:comparison with a serine protease. Comparison of the catalytic networks of (A) the class 1 Naju naja atra venom enzyme (White el al., 1990), (B) the class Ill bee venom phospholipase A2 (Scott et al., 1990b),and (C) that of a serine protease Sfreptomycesgrisezcstrypsin (Read and James, 1988). The histidine-aspartate couple is highlighted in all cases. W indicates a water molecule; the sn-2 phosphonate of the transition-state analog is labeled P2. The oxygen atom, which is thought to serve as the attacking nucleophile during general base-mediated attack on the substrate, is crosshatched.

-

class I Naja naja aha

-

Claaa II Humrn nonpancreatic

-

Class 111 Bee venom (Apb mellifera)

FIG.4. The hydrophobic channel of secretory phospholipases AP. A schematic display looking directly “into” the channel showing the residues that comprise the mouth and internal contours; m-1 and sn-2 refer to the protruding termini of the alkyl substituents of the transition-state analog. The ellipses indicate residues whose side chains have been shown to be involved, or are likely to be involved, in interfacial adsorption. The hatched box corresponds to the location of the amino-terminal end of the catalytic network. Top:The class I enzyme from Naja mja atra venom (White el al., 1990, copyrighted by the American Association for the Advancement of Science), m&&: the class I 1 enzyme from human nonpancreatic sources (Scott et al., 1991), and bottom: the class I11 enzyme from the venom of A+ melliftro, the European honey bee (Scott et al., 1990b).

SECRETORY PHOSPHOLIPASES

A2

63

The side chain of residue 4 anchors the amino-terminal helix with respect to the enzyme proper (Fig. 5). Replacement of the highly conserved Gln at this position with Asn, as in the human nonpancreatic sPLA2, may permit small movements of the amino terminus ( CRBP X R B P I I > P2 > ALBP. V. APO- VERSUS HoLo-iLBP STRUCTURE A. General Considerations

Three of the iLBPs have had both their apo and holo structures determined. This is true for ALBP, CRBPII, and IFABP (see references in Table I). Other than the presence or absence of ligand, there seems to be little difference in the conformation of the protein. For example, in ALBP there is a significant shift in the position of Phe-57 on binding of ligand, but there is no overall change in the protein conformation. In

LIPID-BINDING PROTEINS

115

crystalline CRBPII, one ordered water molecule is displaced on binding of ligand. Others that are either disordered or not detected at the experimental resolution are almost certainly displaced. In summary, comparisons of the crystal structures of the apo- and holo-iLBPs show that the conformational differences with and without their respective hydrophobic ligands are minimal, often within the experimental error of the structural study. Ligand binding, therefore, does not apparently induce large conformational changes in the protein. One interesting difference suggests a portal for lipid to entedexit the binding site within the cavity, and will be discussed in more detail below. Because of the internal nature of the binding site and with no other conformational changes evident between apo and holo forms, association of the hydrophobic ligand must be facilitated by dynamic motions of the protein. Preliminary theoretical studies of such dynamic effects have been carried out for serum retinol-binding protein, a member of eLBP family (Aqvist et al., 1986). B . Ligand Entry into Binding Cavity: Portal Hypothesis From the data given above and unlike most enzymes, it should be clear that the iLBPs bind the hydrophobic ligand in a cavity as opposed to a cleft. The cavity that was discussed above is not generally connected to the solvent by any obviously large opening. Instead, as can be seen in Figs. 9 and 10 for most of the iLBPs, the binding site is essentially surrounded by protein atoms. Furthermore, if the location of the binding cavity is studied (Figs. 3 and 8), the fact that the binding cavity is not centered in the iLBP envelope is clear. Rather the cavity is near the surface formed by the helices a1 and aII, and the turns connecting PC to PD and PE to PF. Without any large opening that might serve as the ligand entry point, preferential pathways for binding must be surmised from the crystal structure. The first hypothesis about a portal for ligand entry and exit was based on the packing density of atoms on the surface of the IFABP molecule (Sacchettini et al., 1989). It appeared that in studying the crystal structure, a small opening in the molecular surface was visible in the region of aI,aII, and the turns PC-PD and BE-PF mentioned above. This opening connected to the cavity. This evidence was reinforced by crystallographic studies of fatty acid binding to ALBP. In the crystal structure of the oleate : ALBP complex?the last two carbon atoms of the bound fatty acid protrude from the cavity into the surrounding solvent (Xu et al., 1993).

116

LEONARD BANASZAK E T AL.

The location of the atoms that protrude into the solvent clearly points to this segment of ALBP as a favored ligand entry point. Near the alleged portal in ALBP, P2, and IFABP is a phenylalanine side chain. Crystallographic studies of the apo, the stearate, and the oleate complex of ALBP revealed that one side chain, Phe-57, was located in different positions in the different crystal forms depending on the presence of ligand (Xu et al., 1993).The comparisons are shown in stereo in Fig. 11. Similarly in the P2 study, there are three molecules in the asymmetric unit. T w o have Phe-57 in one conformation; one molecule has it in another conformation that differs primarily by a X I change. T h e stereo drawing in Fig. 11 includes both the Ca trace of the holo form with bound oleate, and the apo form of ALBP. Note that the C a models of the apo and holo forms are essentially superimposable. In the stereo image of the Ca model, the location of the putative portal region is circled. The helix that isjust visible in the enlargement is aII. For orientation relative to the schematic shown earlier, recall that Phe-57 is on the PC-PD turn and Asp-76 belongs to the BE-PF turn. Both are shown in Fig. 11. Phe-57 is present in its two orientations; the one that is labeled belongs to the apo form of ALBP. When viewed in stereo, the enlarged portion of Fig. 11 shows that the side chain is positioned in the space between a11 and the turns PC-PD and PE-PF. It should also be apparent that the reorientation of the side chain of Phe-57 requires changes only in the torsional angle (x,)around Ca-CP.

VI. GAPBETWEEN PD A N D PE In the iLBPs, all of the adjacent /3 strands of the barrel are connected by interstrand hydrogen bonds except for PD and PE, wherein the separation or gap is too large. T h e gap region should be visible in any of the stereo diagrams containing the Ca model. As noted above, in spite of the lack of interstrand hydrogen bonds, the separation between PD and PE does not provide access to the ligand-binding cavity. Hence both the structural and functional bases of the gap are unknown despite the fact that it is present in all of the iLBPs. One hypothesis is that it is the result of a favored folding pathway (Xu et al., 1992). The idea is that an intermediate step in folding leads to a more open /3 sheet. This then closes into a P barrel but cannot give a tight hydrogen-bonding arrangement between PD and BE. Incidentally, the top of the gap is near the purported ligand portal mentioned above. The gap regions found in ALBP, CRBPII, and IFABP are pictured in Fig. 12. The two strands that make u p the sides of the gap have their 4, J, angles appropriate for P strands and have a series of water molecules


117

FIG. 11. A stereodiagram of the Ca trace of apo- and holo-ALBP. The small stereo diagram at the bottom shows the Ca models of both the apo and the holo forms of crystalline ALBP. Every tenth residue is numbered, and because of the close similarity in the conformation of the two proteins much of the diagram appears as one solid line. At the top of the molecule (shown in the enlargement) Phe-57 appears twice. This is due to the fact that this side chain has two different conformations in the structures of the apo and holo forms. With Phe-57 swung to the left, the holo-form is viewed. When located in the rightmost position, Phe-57 is as appears in the apo form, seemingly closing the portal to the ligand-binding cavity.

forming bridges between main-chain atoms in both strands. T h e remainder of the space between these strands is filled with amino acid side chains. If one compares the location of the water molecules in the PD-PE region, only two waters are in the same position in all of the known crystal structures. They are shown by the crosses near the very bottom of Fig. 12. In addition to these two waters, the close conformational similarity between all of the three proteins in this region should also be apparent. Overall, the loop between the p strands D and E is very conserved in both its primary and three-dimensional structure. Note the close conformational positioning of Phe-64 and Phe-70. If viewed in stereo, the two

118

LEONARD BANASZAK ET AL.

FIG. 12. A stereodiagram of the gap region of three iLBPs. The coordinates of all atoms in the residues, including /3D and BE of three crystalline proteins (ALBP, IFABP, and CRBPII), are shown in stereo. The alignments were calculated as described in Fig. 5 and in the text. Waters that are conserved in all three crystal structures are displayed as crosses and three conserved phenylalanines are labeled according to the ALBPlR numbering scheme.

side-chain rings are parallel and relatively close. Along with a Gly-67, they are highly conserved in all of the iLBPs. Note also in Fig. 12 how both IFABP and CRBPII fill the middle portion of the gap with tyrosines, but they use different strands to provide the side chain. Although it is difficult to see because of the three overlayed structures, the molecule without a tyrosine at this position is ALBP. Because ALBP does not fill this portion of the gap with any side chain, it has in the gap a small pore that is large enough to admit a water molecule into the binding cavity. In the ALBP structure this pore is filled with ordered waters. The ends of PD and PE are not so structurally conserved. The turns at the top of the gap region as oriented in Fig. 12 are two of the least


119

conserved structural features in the iLBP family. CRBP and CRBPII both have two amino acid insertions in the turn between strands PE and PF. T h e turn between strands PC and PD contains the amino acids that are proposed to be the portal into the binding cavity. Note in Fig. 12 that the orientation of Phe-57 varies somewhat from protein to protein, although this may be a key residue that changes orientation on ligand binding. In summary, hydrogen bonding occurs between all of the /3 strands of the iLBP barrel except between PD and PE. A gap is formed between these two strands. Two water molecules and two phenylalanines appear to be highly conserved in the turn between PD and BE. Presumably this sequence contributes in some important way to the formation and stability of this unusual form of supersecondary structure. Away from the interconnecting turn at PD-BE, conformational homology is somewhat less striking. When the amino acid sequences of the PD-PE turn are compared for all family members, the consensus sequence -F-X-X-GX-X-FIC- is found. AND SPECIFICITY BASED VII. BINDING AFFINITY ON BIOCHEMICAL STUDIES T h e crystallographic structures frequently alluded to in this review are usually derived from studies of the iLBP with and without bound ligand. This has made it possible to tabulate the more obvious interactions between the hydrophobic ligand and the protein. Before studying the structural data, it is of interest to look at the affinity constants from the point of view of chemical measurements. Because they always involve ligands that have very low solubility in aqueous buffers, binding studies using compounds such as fatty acids or retinoids are not easy to do, so different investigators often obtain different dissociation constants. The data are summarized in Table VII. Note that the dissociation constant for retinoids binding to CRBP or CRBPII are generally smaller than those of fatty acids binding to ALBP, P2, o r IFABP. Retinol binding to an iLBP gives a dissociation constant in the range of 10 to 40 nM and fatty acid binding generally appears in the micromolar range. As can be seen in Table VII, retinaldehyde appears to be bound somewhat weaker. Retinoic acid, on the other hand, does not bind to CRBP or CRBPII but it does have a low affinity for ALBP and P2, relative to that for CRABP. The affinities between the binding proteins and fatty acids appear to be similar and vary from roughly 0.2 to 3 pM (Table VII). As the chain length decreases, binding affinity is also reduced. For example, no laurate ( C I ~binding ) to IFABP is detectable (Lowe et d.,1987). ILBPs that

TABLE VII

Ligand-Binding Propertiesfor U P S Protein

Form

Ligand

P2

Native Native Native Native Y 19-phospho ALBP C 117-TNB" C 117-TNB R126Q R126Q Y128W Y128W R126L. Y128F R126L. Y128F Native

HFABP CRABP

Native Native

CRBP

Native

Oleate Retinoic acid 12-Anthroyloxy oleate cis-Paraniric acid 12-Anthroyloxy oleate 12-Anthroyloxy oleate &-Paranirk acid 12-Anthroyloxy oleate cis-Paraniric acid 12-Anthroyloxy oleate cis-Paraniric acid 12-Anthroyloxy oleate cis-Paraniric acid Oleate all-trans-Retinol all-trans-Retinoicacid Oleic acid all-tram-Retinoic acid 13-cis-Retinoicacid Retinal Retinol 4-H ydroxyretinoic acid 18-Hydroxyretinoic acid all-trans-Retinol 13-cis-Retinol 9-&-Retino1 11-cis-Retinol

ALBP

rc, (app) 3 w 2 w 2 w 1.7 f l No binding 6.5 phi No binding 2 w 25 3 w 15 CLM 0.5 pM N o binding Binds Binds Binds 0.5 7nM 156 nM No binding No binding 17 nM 20 nM 10 nM 40 nM No binding No binding

w

Ref. Matarese and Bernlohr (1988) Baxa et al. (1989) Sha et al. (1993) Sha et al. (1993) Buelt et al. (1992) Sha el al. (1993) Sha et ul. (1993) Sha et al. (1993) Sha et al. (1993) Sha et al. (1993) Sha et al. (1993) Sha et al. (1993) Sha et al. (1993) Uyemura et al. (1984) Uyemura et al. (1984) Uyemura et al. (1984) Peeters et al. (1991) Fiorella and Napoli (1991) Fiorella and Napoli (1991) Fiorella and Napoli (1991) Fiorella and Napoli (1991) Fiorella and Napoli (1991) Fiorella and Napoli (1991) Levin et al. (1988) MacDonald and Ong (1987) MacDonald and Ong (1987) MacDonald and Ong (1987)

Q108R

CRBPII

Native

Q108R

e

Q128R

2

IFABP

Native

MFBl MFB2

Native Native

CAST (ileal)

Native

LFABP

Native

a

all-tram-Retinoic acid all-trans-Retinal Methyl retinoate all-tram-Retinol all-tram-Retinal all-tram-Retinoic acid all-trans-Retinol all-tram-Retinaldehyde 134s-Retinol all-tram-Retinoic acid Palmitate all-h-am-Retinol all-tram-Retinalde hyde all-trans-Retinoic acid Palmitate all-trans-Retinol all-tram-Retinaldeh yde all-tram-Retinoic acid Palmitate Oleate Arachidonate Palmitate Oleate Oleate Oleate Palmitate Chenodeoxycholate Glycocholate Oleate

C 117-TNB refers to Cys- 117 modified with thionitrobenzoate. LFABP possesses a high- and low-affinity binding site.

N o binding Weak binding No Binding Threefold weaker binding Binds weakly Binds weakly 10 nM 10 nM 40 nM

No binding No binding Decreased binding Decreased binding No binding Binding Decreased binding Decreased binding No binding No binding 0.2 phd 3.7 phd 3.6 pM 14 m Binds with lower affinity than MFBl 36 phd Binds Binds Binds 0.2l0.9 phdb

Levin et al. (1988) Lietal. (1991) Levin et ul. (1988) Stump et al. (1991) Stump et al. (1991) Stump et al. (1991) MacDonald and Ong (1987) MacDonald and Ong (1987) MacDonald and Ong (1987) Cheng et al. ( 1991) Cheng et al. (1991) Chengetal. (1991) Cheng et ul. (1991) Cheng et al. (1991) Chenget al. (1991) Chenget al. (1991) Cheng et al. (1991) Chengetal. (1991) Cheng et ul. (1991) Miller and Cistola (1993) Lowe et ul. (1987) Lowe et al. (1987) Smith et al. (1992) Smith et al. (1992) Miller and Cistola (1993) Sacchettini et al. ( 1990) Sacchettini et ul. ( 1990) Miller and Cistola (1993) Miller and Cistola (1993) . .

122


bind fatty acids are capable of binding a wide variety of analogs, in particular, fluorescent fatty acids (Kim and Storch, 1992a,b). Storch and colleagues have examined the binding and kinetic properties of a number of FABPs with a series of fluorescent anthroyloxy fatty acids (Kim and Storch, 1992a,b; Wootan et al., 1993). Interestingly, their results suggest that dissociation of fatty acids from some, but not all, binding proteins occurs by a collisional and not a diffusional process. That is, physical association with membranes is rate limiting for dissociation of a ligand from its binding protein. Moreover, the phospholipid composition of artificial membranes had a pronounced effect on this process. These results are novel and suggest that when considering metabolic role(s) played by the iLBPs, interactions with membranes must be considered. In the case of CRBP and CRBPII, a number of retinoids have been tested (MacDonald and Ong, 1987; Cheng et al., 1991). No binding to CRBP or CRBPII occurred with 94s- or 1 l-cis-retinol. However, binding was detected with 13-cis-retinoland 3-dehydroretinol. One unusual finding was that CRBPII but not CRBP had a detectable affinity for all-transretinaldehyde. Later studies (Li et al., 1991) showed that CRBP could bind retinol or retinal but that it had a 100-fold higher affinity for the alcohol. VIII. BINDING AFFINITYAND SPECIFICITY BASED ON CRYSTALLOGRAPHIC STUDIES The crystallographicstudies of the iLBPs not only provide information on the overall location of the hydrophobic ligand-binding site within the cavity but also the conformation of the ligand and the interactions that contribute to the specificity. Because the coordinates for the known iLBP structures were superimposed for the homology comparisons, the same transformations can be applied to the ligands. A stereodiagram showing the superimposed ligands is found in Fig. 13. Residues that are in contact with the bound lipid for these proteins have been discussed in a previous section on the binding cavity. Recall that the number ranges from 18 to 21. Note the overall curvature in the conformation of both the bound fatty acids and the retinols in Fig. 13. The bend appears to occur because of the cavity shape and the anchoring position of the polar head group. The bending is approximately the same for all of the ligands except fatty acid bound to MFBB, the insect protein (Benning et al., 1992). Specificity for the ligand appears to derive from either ionic or polar interactions with the head group of the amphipathic ligand. The recogni-

123


aha CmPn

aha cRnPl1

FIG. 13. A stereodiagram of the ligands of CRBP, IFABP, MFB2, and P2. These structures were aligned as described in Fig. 5. ALBP and HFABP bind their ligands in the same manner as P2, and CRBPII binds its ligand in the same manner as CRBP. The alignment of the crystal coordinates is based only on protein atoms. The striking similarity in the location of the ligands in the crystals of the holo forms is due to the homology in the iLBP family.

tion motifs are shown schematically in Fig. 14. There appears to be three key residues involved in these interactions. They are positions 106//3H, 126//3J, and 128//3J. In addition, for CRBP and CRBPII, a lysine at position 40/PB contacts the isoprene tail of the bound retinol. Positions 106, 126, and 128 are the positions that provide specific interactions with the polar group of bound lipid. Although a number of holo-iLBP structures are known, there are only three apparent motifs that form the interactions with the polar portion of their ligand. We will refer to these ligand interactions as binding motifs, and we will designate these three motifs as the P2 motif, the IFABP motif, and the CRBP motif. In the P2 motif, the carboxylate of the bound ligand is located in a pocket between Arg-106//3H, Arg-l26//3J, and Tyr-l28//3H, as shown in Fig. 14. These three amino acids all hydrogen bond to the carboxylate group of the bound fatty acid; however, the hydrogen bond to Arg- 106 is usually indirect via an intervening water molecule. ALBP, P2, HFABP, and MFB2 all interact with their bound fatty acid using the P2 motif. It is not surprising that ALBP, P2, and HFABP should all bind in the same

124


128

106

FIG.14. The recognition motifs in the iLBPs. The amino acids are numbered according to their location in crystalline ALBP, but corresponding amino acids present in the other crystalline proteins are also listed. Two locations for fatty acids can be seen, one where fatty acid is bound to ALBP, P2, MFB2, and HFABP; the other where IFABP binds fatty acids. The position of bound retinol in CRBP and CRBPII is also shown, although because the p-ionone ring is edge-on, it is not readily distinguished from a fatty acid. Note that the polar residues can be either ionizable or polar (Arg or Gin and Tyr). Further details of structural factors in the three motifs are given in the text.

manner because they share such high sequence identity, but it is interesting and not yet explainable that MFB2 should also bind fatty acids in the same manner. The other fatty acid-binding motif is the IFABP motif. As shown in Fig. 14, it is characterized by the carboxylate of the fatty acid making a salt bridge with Arg- 106. This leads to notably different positions for the ligand carboxylate. Interestingly, the iLBPs that bind fatty acids all appear to have arginines at positions 106 and/or 126 (Jones et al., 1988). IFABP has a phenylalanine at position 128, which may, in part, prevent it from using the P2 motif. However, at residue 34 there is a buried aspartic acid residue whose carboxylate interacts with the guanidino group of Arg-126. This aspartic acid also occurs in MFBS. In MFB2, the interaction is bridged by a water molecule. IFABP is the only known iLBP that uses this motif for interacting with the fatty acid carboxylate.


125

The third binding mode is found in CRBP and CRBPII, and is used to bind retinoids. Like the IFABP motif, it uses an amino acid at position 106 to interact with the bound ligand. In the CRBP motif, this amino acid is glutamine. The hydroxyl oxygen atom of retinol is hydrogen bonded to the side-chain nitrogen or oxygen atom of Gln-108. In the two CRBPs, the amino acid at 126 is a glutamine and at 128 it is phenylalanine. However, as can be seen in Figs. 13 and 14, the bound retinoids occupy nearly the same location as the fatty acids. The ionizable side chain of Lys-40 is also within van der Waals distance of the bound retinol, and it is probably interacting with the 7r electrons of the isoprene chain. It is unknown whether this residue is necessary for retinol binding but it is only present in the binding proteins that specifically bind retinol. In the retinoic acid-binding proteins, for example, it is a valine. There are almost certainly uncharacterized binding motifs. NMR experiments have shown that the carboxylate of a fatty acid bound to rat LFABP is solvent exposed, whereas in the three known binding motifs the polar portion of the ligand is always buried in the center of the protein. As noted above, in IFABP this has been confirmed by "C NMR (Cistola el al., 1989). Although arginines and glutamines are always conserved at position 126, this is not the case at position 106. T h e amino acid sequences near amino acid 106 of gastrotropin, IFABPS, and LFABP vary considerably from the typical sequence, and so it is likely that there is at least one other undescribed binding motif. Also, some of the proteins have multiple specificities. P2 can bind retinol and retinoic acid in addition to fatty acid. Because P2 does not have the glutamine available at position 106 like CRBP, it cannot use the CRBP binding motif to interact with the retinol. In view of the ligand-protein interactions for both fatty acids and retinoids, it is certain that position 106//3H plays an important role in determining the ligand binding affinity and specificity in the iLBPs. Although the second Arg/Gln at position 126 is not involved in direct interaction with the ligand in the case of CRBP, CRBPII, and IFABP, it is in the rest of the fatty acid-binding proteins. Furthermore, it is well conserved throughout the iLBP family. In fact, CRBP and CRBPII are the only two binding proteins without a positively charged arginine at that position. The accommodation of the hydrocarbon tail is the same in all but the insect form of the protein, MFB2, as shown in Fig. 13. If viewed in stereo, it should be apparent that all of the ligands are bent. Retinol has less curvature than the fatty acids. Although never analyzed in detail, the curvature occurs near the C-9 and (2-10 positions of the fatty acid. The binding site therefore appears to be premade for unsaturated fatty acids.

126


Saturated fatty acids apparently can easily assume the same conformation (Xu et al., 1993). In ALBP, the first few carbon atoms of the bound fatty acid are in van der Waals contact with a number of water molecules. In addition to water molecules, this network includes the specificity side chains on /3H and PJ. Higher numbered carbons in the bound fatty acids are generally in contact with nonpolar atoms (methylene carbons) belonging to side chains that line the cavity. IX. MEMBERSOF iLBP FAMILY WITH KNOWNCRYSTAL STRUCTURE A . Myelin P2 Protein P2 is a component of myelin from the peripheral nervous system. It is localized on the cytoplasmic side of Schwann cells where it behaves as peripheral membrane protein, although a small amount is found in the cytoplasm (Trapp et ad., 1984). Like the other iLBPs, the exact biochemical role of P2 is unknown. Its cellular localization and its ability to bind different fatty acids and retinoids (Uyemura et al., 1984) suggest that it may function in fatty acid trafficking. It would therefore play a major role in the movement of fatty acids between the site of uptake and that of esterification during the massive phospholipid synthesis phase of myelinating Schwann cells. Myelin P2 was the first protein in the iLBP family whose structure was determined (Jones et al., 1988). The crystals contain three copies of the molecule in the asymmetric unit and the structure has now been refined at 2.7 A (Cowan et al., 1993). Because the crystals do not diffract beyond 2.7 A, the precision of the model is not as high as in the other studies and this factor could affect precise hydrogen bonding measurements. In an attempt to overcome this problem, the three molecules were restrained to be similar during the refinement. Only well-determined water molecules were included in the final model. N o fatty acid was added to the sample, but later mass spectroscopy measurements indicated that the predominant ligand was oleic acid, which also agreed with the shape of the electron density. This density could not be accounted for by the protein model. The oleic acid is located in the internal cavity described above. It also has the prototypical fatty acid-binding site, the P2 motif, which includes the /3J and /3H arginines and the PJ tyrosine (see Fig. 14). In this type of binding site, the carboxylate group of the fatty acid interacts with three polar groups: the guanidinium groups of Arg-106 and Arg- 126, and the hydroxyl group of Tyr-128. As shown in Fig. 14, both carboxylate oxy-


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gens of the ligand are within hydrogen bonding distance of the tyrosine hydroxyl; in addition, one carboxylate oxygen hydrogen bonds to Arg126 and the other to Arg-106. Arg-126 is also found to have its planar guanidinium group stacked directly over the center of the ring of Phe- 16 at a separation of 4.0 A. Several hydrophilic residues project into the p barrel and form a network of hydrogen bonds and salt links, clustered around the carboxylate groups of the ligand. At the current crystallographic resolution, the network has been restricted to include only five water molecules, all of which are in the barrel. Other side chains create a hydrophobic environment around the fatty acid tail. The complementarity of ligand and protein appears relatively poor and the fatty acid still has an accessible surface area of 33 A2.This may be related to the lack of specificity for ligand that has been reported by Uyemura et al. (1984); they report that oleic acid, retinoic acid, and retinol are able to bind to P2. The bound fatty acid has a U-shaped hydrocarbon tail that extends toward the hydrophobic lining formed by the two helices, a1 and aII. Access from the external milieu to the cavity appears to occur around the side chains of Thr-29, Leu-32, Phe-57, and Lys-58. In the crystalline state, one of the three molecules in the asymmetric unit has the portal closed due to a different conformation of Phe-57 (Cowan et al., 1993). In P2, the conformation of Lys-58 also appears to have an effect on the size of the portal. Last of all, it is noteworthy that myelin P2 is a basic protein. With added positively charged groups on its surface, it seems reasonable to assume that the side chains on the outside surface of the molecule are available to interact with phospholipid head groups in the myelin membrane. P2 and ALBP have tyrosine at position 19 (helix d). As will be described in the next section, phosphorylation by tyrosyl kinases occurs at this position. In terms of structure/function relationships, the phosphorylation of Tyr-19 is difficult to explain. If we assume a static picture for the protein as derived from the crystal structure, the tyrosine is buried and inaccessible to the protein kinase. A more detailed description of Tyr-19 will be given in Section IX,B. B. Adipocyte Lipid-Binding Protein The adipocyte member of this family was named “lipid-binding protein” because it was found to bind both long-chain fatty acids and retinoic acid (Matareseand Bernlohr, 1988).It is also called adipocyte P2 (Hunt et al., 1986) because of its high sequence similarity to the bovine myelin P2 protein (67%). The protein was suggested to play a role

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similar to that of other members of the family, that is, to solubilize and transport the cellular free fatty acids. For fat cells, the intracellular trafficking of fatty acids is bidirectional. Fatty acid flux into the adipocyte occurs during lipogenic (feeding) conditions and out of the cell during lipolytic (fasting) conditions. Presumably the protein fulfills the trafficking roles in both directions, but this point is untested. The crystal structure of ALBP as described in the comparative studies above shows that the iLBP properties are again present. They include (1) /3 barrel with a cavity, (2) no major conformational difference between the apo and holo forms, (3) specificity generated by interaction of the carboxylate of the fatty acid with two arginines and a tyrosine, and (4) similar conformations for both bound unsaturated (oleic acid) and saturated (stearic acid) ligands (Xu et al., 1992, 1993). T h e internal carboxylate group of the lipid interacts directly with the guanidinium group of Arg-126 and the hydroxyl group of Tyr-128. Unlike myelin P2, the guanidinium group of Arg-106 interacts with the carboxylate indirectly through an ordered water molecule. This may be due to the residue at position 40, a methionine in ALBP, and a valine in P2, which would result in a close contact to the guanidino group of an arginine having the P2 conformation. Compared to P2 and HFABP, the fatty acid, when bound to crystalline ALBP, is slightly more extended. Somewhat unusual in the crystal structure of ALBP is the position of Met-35. The hydrophobic side chain of this residue is on the surface of the molecule pointing into the solvent. Together with Val-32 and Phe-57, they form a small hydrophobic patch on the outside of ALBP. Note that this patch is adjacent to what is believed to be the portal for ligand binding described earlier. At this point, the molecular basis or need for this somewhat unusual region is unknown. One of the most interesting feature of ALBP is that the protein is phosphorylated in 3T3-Ll adipocytes and in vitro by the activated insulin receptor kinase (Bernier et al., 1987; Hresko et al., 1988; Buelt et al., 1991). Phosphoamino acid analysis revealed that the phosphorylation was exclusively on Tyr-19. Interestingly, myelin P2 can also be phosphorylated by the insulin receptor (Buelt et al., 1993). Ligand binding and phosphorylation appear to be interrelated. Lipid binding by the protein accelerates the phosphorylation by lowering the K, of the reaction, whereas the phosphorylation of the protein appears to inhibit the lipid exchange (Buelt et al., 1991, 1992). Inspection of the crystal model shows that the phosphorylated Tyr-19 is buried about 10 A away from the protein surface. This can be seen in the stereo diagram found in Fig. 15. Simple rotation of the aromatic ring around the torsional angles, xI and x 2 , will not bring the tyrosine to the protein surface, and as already noted, a similar case exists in P2.


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FIG. 15. A stereo diagram of the Ca model of ALBP, the bound fatty acid, and the position of Tyr-19, including a Ca representation of the crystal structure of ALBP along with its bound fatty acid. Also shown is Tyr-19, the amino acid that is phosphorylated in uiuo in both ALBP and P2. The amino acids are numbered according to the ALBP sequence.

Clearly this is a structure/function problem for which the crystallographic data do not give a full explanation. The present position of Tyr- 19 leaves unanswered questions about how this tyrosine is recognized by the receptor kinase. Furthermore, even after recognition, the phosphorylation event must be accompanied by a major conformational change in the iLBP. The location of Tyr-19 is on the first helix, aI.From a purely speculative point of view, perhaps the most obvious conformational change for accommodating the phosphorylation of Tyr-19 would be the relative movement of the a helices to the /3 barrel. One hypothetical mechanism for the modification is a sort of “lifting of the helical lid,” which would bring Tyr- 19 to an accessible position. To add to the dilemma, phosphorylated protein has been shown to be structurally similar, but not identical, to the native protein by circular dichroism (Buelt et al., 1992). If indeed a major conformational change accompanies modification by the protein kinase, it could only happen in some temporal dynamic state. Finally, it is interesting that the tyrosylphosphorylated protein is readily recognized by antiphosphotyrosine monoclonal antibodies. Moreover, the phospho-ALBP is readily de-

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phosphorylated in vitro and in situ, suggesting time-averaged accessibility of the phosphotyrosine to the phosphatase. When phosphorylated, two properties are altered. The holo form cannot release bound fatty acid and the apo form will not bind fatty acid (Buelt et al., 1992). One possibility is that the phosphorylated tyrosines will interact with the arginines, preventing binding of any fatty acid (Cowan et al., 1993). Calculation of the electrostatic potential for ALBP shows that the lipid binding cavity is positive. The inclusion of a phosphate moiety on Tyr-19 changes the electrostatic potential of the cavity significantly, going from positive to negative. Until crystals of phosphorylated protein are obtainable, it may be difficult to explain these modifications.

C. Heart Fatty Acid-Binding Protein In the cardiac myocyte, the bulk of the ATP necessary to drive the rhythmic beating process is derived from oxidative phosphorylation (Oram et al., 1973). T h e acetyl-CoAs necessary for such a process are derived from fatty acid degradation. T h e trafficking of fatty acids from their site of uptake (plasma membrane) to utilization (mitochondria) is a function presumably filled by the heart FABP. The crystal structure of HFABP was determined independently by two laboratories. The first crystal structure, bovine HFABP, was determined to medium resolution and confirmed that HFABP indeed had the characteristic iLBP p barrel (Muller-Fahrnow et al., 1991). It was not until the high-resolution structure of human HFABP was determined that the bound ligand was observed in the binding cavity (Zanotti et al., 1992). HFABP binds fatty acid using the P2 motif. T h e chemical nature of the fatty acid bound in the crystalline state is unknown but it has been modeled as a palmitate. The carboxyl moiety of the fatty acid makes hydrogen bonds to Arg-126, Tyr-128, and to a water, which in turn is hydrogen bonded to Arg- 106. The positioning of the hydrocarbon tail of the palmitate bound to HFABP is similar to that bound in P2 in that it turns back toward the bound carboxylate in such a way that the fatty acid appears as a U-shaped structure. T h e temperature factors of the final two carbons of the bound fatty acid are quite high, 100 A*, and so they could be highly mobile. The U shapes of P2s (CIS)and HFABPs (c16)are to be contrasted to the bound oleate (CIS) of ALBP, whose hydrocarbon tail is relatively straight and projects past Phe-57, out of the protein into the solvent.


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D. Manduca sexta Fatty Acid-Binding Protein 2 Manduca sexta fatty acid-binding protein 2 is unique among the iLBPs of known crystal structure in that it is from an insect; all of the others are from vertebrates. This makes the evolutionary distance greater, and comparisons to the mammalian proteins are valuable for ascertaining which amino acids are absolutely necessary for fatty acid binding. MFB2 is expressed in the midgut of Manduca sexta larvae, where it comprises about 12% of the total protein. Another fatty acid-binding protein, MFB 1, is also expressed in this tissue, and it makes up about 2% of the total protein (Smith et al., 1992). The crystal structure of MFB2 was determined with a bound palmitate and it has all of the conformational properties of the iLBP family (Benning et al., 1992). The carboxylate is bound in the P2 motif using the two arginines, but the placement of the hydrocarbon tail is different from the other iLBPs as shown in stereo in Fig. 13. The binding cavity has a different shape than the other iLBPs, and consequently the lipid is bound in a different conformation. The cavity of MFB2 is closest to the surface of the protein in the space between the loop connecting strands BE and PF, the loop between strands PG and PH, and the turn between the two (Y helices. In fact, there is an opening of about 4.5 8, in diameter into the binding cavity at this location. This is different from the portal described for the mammalian iLBPs. The MFB2 portal lies between residues 23,24, 77, 78, 97, and 98. Surprisingly, the tail of the fatty acid does not point toward this opening, rather it lies between aI,the loop between strands PG and PH, and the loop between strands PI and PJ. The reorientation of the ligand in MFB2 appears to be due to sequence differences. The conformational differences can be traced to changes in cavity shape. This in turn is related to residues 22,33,78, and 97 (ALBPI P2 numbering). All but 97 are illustrated in Fig. 16. The differences in cavity shape between MFB2 and ALBP appear to be due to the exchange of large for small side chains. For example, in MFB2, residue 32 is a leucine. In P2, pointing in that direction is Ala-33; the result is the extra space created by the three missing carbons. A similar arrangement occurs near Ala-22 in P2, which corresponds to leucine 23 in MFB2. To create extra space in the other direction, Trp-97 in P2 occupies space which occurs near Gly-96 in MFB2. Similarly, Arg-78 in P2 is near where Ser-78 and Asp-77 are to be found in MFB2 (see Fig. 16). Therefore, the difference in conformation of the bound ligand as found in the P2 family, as compared to MFB2, appears to be a direct result of the cavity shape and the replacement or reorientation of large versus small side chains.

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FIG. 16. Differences in the binding cavity in vertebrate and insect iLBPs. The stereodiagram illustrates the differences in the conformation of the fatty acids bound to crystalline vertebrate and insect iLBPs. The reader should also refer to Fig. 13. The differences in conformation of bound ligand are likely to be attributable to amino acids 22, 32. and 78 of ALBP and the corresponding side chains in crystalline MFB2. The diagram was produced from the coordinates after least-squares alignment as described in Fig. 5. The atoms of MFB2 are represented with bold lines.

As mentioned earlier, despite the unique behavior of the hydrocarbon tail of the bound fatty acid, the carboxylate binds in the P2 motif. As in the other proteins with this specificity site, the carboxyl group is within hydrogen bonding distance of Arg-128 and Tyr-130. What is unique about MFB2 is the presence of a putative sulfate at the carboxylatebinding site (Benning et al., 1992). T h e electron density map of MFB2 contained electron density large enough to accommodate either a phosphate or sulfate ion. During refinement, a sulfate ion was placed in this density. The ion had a final average temperature factor of 75 81'. This sulfate makes hydrogen bonds to the bound fatty acid, Lys- 105 (MFB2 numbering), a tyrosine, and a histidine. It should be noted that in MFB2, the separation of P strands D and E is greater in this protein and allows for a direct entry in the cavity via the gap. MFBS is the first structure to be determined wherein the highly conserved tryptophan at position 8 is not conserved. In MFB2, the spatially equivalent residue is a tyrosine. The N terminus is two residues shorter than in ALBP, and to align the strictly conserved Gly-6 requires an insert in amino acid sequence (Table 111) between the usual G-X-W motif. In the MFBS structure, the N terminus differs somewhat from the other structures in the family and is the outlier in Fig. 5. Lys-5 (MFB 1 numbering) is in fact spatially equivalent to the position of Gly-6 (ALBP/P2


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numbering) in the other members of the family. This lysine takes the place of the basic residue normally occurring at the C terminus of other members of the family (Arg-130 in ALBP/P2) which in MFB2 is an alanine residue. The residue at the position normally occupied by Trp-8 is, we believe, important for folding because of its interaction with residues at the C-terminal end of the molecule. In MFBP, this interaction is still achieved by three tyrosine residues: 6,8, and 108 in close contact. Although the insect lipid-binding protein has the same structural motif as other iLBPs, the side chains used to define the specificity site are somewhat different and appear to include an additional anion (Benning et al., 1992). In addition, the ligand has a different conformation and the overall structure suggests that a different portal might be favored in this protein. E . Intestinal Fatty Acd-Binding Protean Intestinal fatty acid-binding protein is a representative of the iLBP family that binds fatty acids exclusively; it will not bind retinoids. IFABP is presumably part of the gut absorptive process. The protein is synthesized in the polarized columnar epithelial cells of the small intestine (Shields et al., 1986; Sweetser et al., 1988) and contains a single site for the noncovalent binding of long-chain (C16 to (222) saturated and unsaturated fatty acids. T h e K d values for these ligands range from 0.2 to 4 f l (Lowe et al., 1987; Miller and Cistola, 1993). Crystal structures of both apo- and holo-IFABP have been determined and refined at 1.2 and 2.0 A, respectively (Scapin et al., 1992; Sacchettini et al., 1989). Because of the high resolution, the 1.2-A structure obtained from the apo-IFABP crystals is the best determined of any iLBP family member to date. The crystal structure of the holo form has a unique feature, the IFABP binding motif. In IFABP, the carboxylate group of the bound fatty acid forms a salt bridge with an internal arginine, Arg106. Arg-106 also hydrogen bonds to Gln-115, which in turn interacts with the fatty acid through two water molecules, forming a five-member “quintet” (Sacchettini et al., 1989). Hydrogen bond interactions are also observed among other internal side chains, one of which forms a similar quintet. The hydrocarbon chain of the bound palmitic acid is extended with a slight left-handed helical twist (Sacchettini et al., 1989). T h e chain is mostly surrounded by the side chains of aromatic and hydrophobic amino acid residues. There are seven ordered water molecules located within the cavity, along with the fatty acid chain. These solvent molecules are hydrogen bonded to polar side chains in the cavity.

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The ligand-binding cavity of IFABP is different from those of other proteins in the family. The amino acid side chains that comprise the cavity are predominantly hydrophobic (23 hydrophobic residues versus 11 polar residues). Also, the residues that are in close contact with the bound ligand are quite different from those of other proteins whose crystal structures are known. There is a cluster of residues close to the ligand uniquely located on BE and PF in IFABP. Superimposition of the apo and holo structures of IFABP indicates that the overall agreement for main-chain atoms is 0.42 8, (Scapin el al., 1992). The side chains of residues located in the interior cores of both models occupy similar positions, including Arg- 106. The 24 ordered water molecules located inside apo-IFABP are defined in the 1.2-8, structure. Six of the water molecules in the binding region have positions similar to the positions of fatty acid atoms in the holoprotein. Among the remaining waters, six have been located in positions nearly identical to those of the internal waters in the holo protein. The observation suggests that the ligand-binding cavity of the protein does not undergo significant change on removal of the bound ligand. Rather, the space left by the ligand is largely filled up by both unordered and ordered water molecules. These water molecules presumably stabilize the structure by forming hydrogen bonds with internal polar side chains. F. Cellular Retinol-Binding Protein Cellular retinoid-binding proteins have been postulated to mediate the uptake and distribution of cytosolic retinoid compounds. It has been suggested that CRABP is critical in controlling the amount of retinoic acid available to bind to the receptors (Dolle et al., 1990) and therefore indirectly to regulate the expression of retinoid-regulated genes. Ligandbinding studies have shown that both CRBP and CRBPII bind retinol and retinal (Levin et al., 1988; Li et al., 1991). CRBP, however, shows a strong preference for binding retinol, while CRBPJI binds retinol and retinal equally well. Neither CRBP nor CRBPII binds retinoic acid, whereas CRABP binds retinoic acid exclusively (Fiorella and Napoli, 1991). The differences in ligand specificity between the three proteins clearly suggests functional diversity. CRBP and CRBPII have been shown to interact with the enzyme lecithin (phosphatidy1choline)-retinol acyltransferase, donating the bound retinoid to the enzyme for acylation with a fatty acid derived from the sn-1 position of phosphatidylcholine (Herr and Ong, 1992). CRBP and CRABP are distributed in many different tissues but CRBPII is localized primarily in intestinal cells. The structures of CRBP and CRBPII have been determined and refined to 2.1 and 1.9 A, respectively (Cowan et al., 1993; Winter et al., 1993).


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N o obvious portal to the retinoid-binding cavity of CRBP was suggested by the crystallographic studies (Cowan et al., 1993). Hence the mechanism by which the ligand enters the binding cavity is obscure, although it is reasonable that the portal would be similar to that mentioned above for the fatty acid-binding proteins. The bound retinol molecule is extended along the barrel axis in a manner similar to fatty acids but shows better complementarity with the protein side chains. The retinoid hydroxyl group is in the interior of the protein and the hydrophobic p-ionone ring is close to the protein surface similar to the hydrocarbon chain of fatty acids in the FABPs. Overall, the retinol is in a nearly planar conformation with the double bond of the p-ionone ring trans to the isoprene tail. The hydroxyl group or hydrophilic end of the retinol hydrogen bonds to Gln-106, which in turn is able to make a hydrogen bond to the phenyl ring of Phe-4. Th e other interesting pol'ar interaction involves Lys-40. The positive charge on the side chain of Lys-40 appears to interact with the isoprene tail of the retinol, with the NZ atom located above the plane of the conjugated r-electron field. The p-ionone ring is located in a hydrophobic environment formed by the two helices and PC-PD and PE-PF turns. Eight water molecules were also identified in the cavity. All of them make hydrogen bonds to internal polar side chains o r water molecules. G . Cellular Retinol-Binding Protein II The crystal structure of CRBPII is closest to that of CRBP. T h e C a positions agree within 0.7 A, as is apparent in Table 11. Unlike bound retinol in CRBP, the dihedral angle between the plane of the p-ionone ring and the chain is not an ideal cis or trans (Winter et al., 1993). In the report of the crystal structure, the complementarity between the bound retinol and protein was examined and appeared fairly good (Winter et al., 1993). In the holo protein, within the cavity there are five empty spaces that are not occupied by retinol atoms. Most of these subcavities contain water molecules, which appear to be an integral part of the protein structure because they occupy similar positions in the apo structure. Twenty two amino acids come within 5.1 8, of the bound retinol molecule. Five of them are within 3.6 A-Lys-40, Thr-53, Arg-58, Trp-104, and Gln-106. In fact, the polar side chains of these residues are close to the bound retinol and again it seems that at least part of the lipid-protein interactions occur through polar atoms. Of particular interest are Lys-40 and Gln-106. As in CRBP, Lys-40 is located above the isoprene tail of the retinol and could interact with the welectron field of the ligand. Gln- 106, on the other hand, makes a hydrogen bond with the only polar group, the hydroxyl group of the retinol molecule.

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As indicated in Table VII, different retinoids bind to CRBP and CRBPI1 with different binding affinities. The structure determination of CRBP and CRBPII allows one to analyze the molecular basis for the variability in ligand binding affinity and specificity. T h e binding of retinal to CRBP requires Gln-106 in such an orientation that the favorable interaction between Gln-106 and Phe-4 is lost, or the hydrogen bond to the ligand is lost, as shown in Fig. 17a and b. In CRBPII, Phe-4 is replaced by a glutamine. The ability of CRBPII to differentiate retinol from retinal is therefore eliminated. The loss of the phenylalanine also results in some local rearrangement around Gln- 106 when compared to CRBP. The geometry of the hydrogen bond between the hydroxyl group of the bound retinol and Gln-106 is not optimal. This may be reflected in the reduced retinol binding affinity of CRBPII.

a

ILE/THR 51

c*" I!!

PHE/CW 4

,

F'HE/CLN 4

FIG.17. Specificity of retinoid binding in the iLBPs. (a) The stereodiagram shows stick models of the bound retinols, and nearby amino acids 4, 40, 51, and 108 of CRBP and CRBPII (CRBP numbering). The amino acids are given, with CRBP first; that is, Phe/Gln-4 means the Phe side chain of CRBP versus the Gln side chain of CRBPII at position 4. Nearby water molecules found in the crystal structure are displayed as crosses. (b) A schematic drawing of the hydrogen bonding between the bound retinoid, and amino acids 4 and 108 of CRBP and CRBPll using the CRBP numbering scheme. The drawing offers a

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X. MEMBERSOF eLBP FAMILY WITH KNOWNCRYSTAL STRUCTURE A . Serum Retinol-Binding Protein

Serum retinol-binding protein (RBP) is a retinoid-binding protein found in the circulatory system. The protein is synthesized in the hepatocytes, where it accumulates until the presence of its ligand in the circulation triggers the secretion of the complex (Ronne et al., 1983; Good-

b

CRBP:Retinaldehyde

Phe-4

Gln-108

Retinaldehyde

........................................................................................................................... CRBP:Retinol

Phe-4

Gln-108

Retinol

.............................................................................................................................. CRBPI1:Retinaldehyde

Gln-4

Gln-108

Retinaldehyde

possible explanation as to how CRBP and CRBPII bind retinoids differently despite their close structural and sequence homology. The hypothesis, described more fully in the text, implicates the hydrogen bonding possibilities for the polar recognition unit Gln-108. In CRBP, the potential formation of a Phe hydrogen bond orients the recognition unit Gln-108 in a way that does not support a hydrogen bond to retinaldehyde.

138


man, 1984). RBP circulates bound to transthyretin. The transthyretin tetramer contains a single high-affinity site for iodothyronine (T4) and a single site for RBP, yielding a RBP-(transthyretin)4-T4 complex (Goodman, 1984). The formation of the protein complex prevents RBP from being filtered through the kidney glomeruli (Kanai et al., 1968). On delivery of the retinol, the protein complex dissociates and the RBP is filtered and degraded (Rask et al., 1980).The crystal structure of human serum retinol-binding protein was initially determined by Newcomer et al. (1984), and was refined to 2.0 A by Cowan et al. (1990). RBP consists of a single globular domain composed of eight antiparallel p strands. Near the center of the p barrel, these strands enclose a cavity that is the retinol-binding site, as was shown in Fig. 2a. The rest of the secondary structure includes a short a helix and an additional /3 strand at the C terminus. The four-turn a helix packs near the p strands at an angle of about 15". The side-chain interactions between the helix and p sheet are very extensive and involve mostly hydrophobic residues. The bound retinol molecule is completely encapsulated by the p barrel, with the p-ionone ring in the center and the isoprene chain stretching along the barrel axis almost to the protein surface. The complementarity between the retinol and the protein is high, indicated by the 1 A' accessible surface area of retinol in holo-RBP. From the structure, it is obvious that retinol can enter the protein only from one end of the barrel. The side chains that are in contact with the retinol are predominantly nonpolar or aromatic. A salt bridge between Arg-12 1 and Asp- 102, and Gln-98 and Leu-35, seems to control access of the binding site to solvent. The retinol hydroxyl group, however, is surrounded by three leucines and makes a single hydrogen bond to an internal water molecule. The dihedral angle between the p-ionone ring and the isoprene tail is 62", which is one of the frequent conformations observed in retinoid crystal structures (Cowan et al., 1990). Attempts to crystallize apo-RBP were unsuccessful. A molecular dynamics study (8iqvist et al., 1986)of the conformational changes induced by removal of the retinol molecule suggests a change in the conformation in the entrance loops of the apo form. Because it has been suggested that these loops make up part of the transthyretinbinding site (Cowan et al., 1990), the study is consistent with the observation that after removal of retinol, the RBP-transthyretin complex breaks up. An independent study of the human RBP molecule has recently been published (Zanotti et al., 1993). Zanotti and co-workers obtained a new crystal form and succeeded in crystallizing holo- and apo-RBP. Within the limits of the study (2.5 A), the holo structure was identical to that


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described by Newcomer et al. (1984) and Cowan et al. (1990). In the apo-RBP structure, changes were observed in one of the entrance loops, 32-37. Particularly Leu-35 and Phe-36 appeared involved. In a separate study using bovine RBP, Zanotti and co-workers (1993) have solved holo and apo structures to high resolution (1.9 and 1.7 A, respectively) with similar results. T h e space group of the latter study is very similar to that of the human RBP study of Newcomer et al. (1984) and, not surprisingly, the structures are similar. However, the retinol conformation is different. In the bovine RBP study, the retinol is observed in the trans conformation. Given the assumptions of the molecular dynamics study, in particular ignoring surrounding water molecules, the theoretical and structural studies are in surprisingly good agreement.

B . @-Lactoglobulin /?-Lactoglobulin (P-LG) is a very abundant protein found in the milk of mammals (McKenzie, 1971; Liberatori, 1977). The protein has been studied for decades and is considered one of the classical markers for milk proteins. It has been shown by a number of investigators that bovine P-LG can form a complex with retinol. However, the exact in vivo function of the protein is still not known. The monomer of bovine P-LG has a molecular weight of 18,000, corresponding to a chain of 162 amino acid residues. There are two genetic variants, commonly known as P-LG A and P-LG B (Braunitzer et al., 1973). The differences between the two variants are located at two positions where an Asp and a Val in variant A are substituted by a Gly and by an Ala in form B (McKenzie et al., 1972). Crystal forms of P-LG A, P-LG B, and P-LG A-retinol complex have been obtained. The structures have been determined for the apo structure at 2.5 8, (Papiz et al., 1986) and for a retinol complex (Monaco et al., 1987). The overall folding of P-LG is remarkably similar to that of human plasma retinol-binding protein (Papiz et al., 1986). The core is made u p of an eight-stranded antiparallel P sheet, which forms a P barrel. There is also a short (Y helix at the C terminus. As described by Monaco et al. (1987), the interior of the@barrel is essentially hydrophobic. The conformational difference between the A form and B form of the protein is not significant. The most interesting observation is that the bound retinol molecule interacts with the protein in a way completely different from that of serum retinol-binding protein. The calculated difference map showed that the molecule is not located in the central P barrel but binds to the a-helixl/?-barreI interface delimited by about 15 residues. Most of the

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LEONARD BANASZAK ET AL

pocket residues are hydrophobic with the exception of Lys-14 1. Further experimental evidence suggests that the retinol molecule bound to P-LG is in a much more open position and subject to chemical or enzymatic attack. It seems that two proteins, P-LG and RBP, share a similar structure, performing a possibly analogous function in a very different way. Despite many similarities, there is much more structural variation between pairs of related molecules in the extracellular protein family than in the intracellular proteins discussed above. This is apparent in the lengths of the p strands, in the conformation of loops, and in the movement of the a helix as a rigid body on the surface of the P barrel (Cowan et al., 1990). In spite of these variations, the /3 strands forming the barrel are directly superimposable. C . Bilin-Binding Proteinsllnsectacyanin

Bilin-binding protein (BBPs), or insecticyanin, are hydrophobic ligand-binding proteins that bind bile pigments. Together with carotenoid-protein complexes, they are thought to play a major role in coloration associated with insect camouflage. The proteins ( 189 amino acids) generally bind a biliverdin compound whose chromophore gives rise to the intense blue coloration of the protein. Two independent crystallographic studies have described the structures of BBPs from a butterfly (Huber et al., 1987a,b) and from the tobacco hornworm (Holden et al., 1987). The protein fold for both proteins is very similar to the RBP structure described above and shown earlier in Fig. 2a. Biliverdin, the bound chromophore, consists of four pyrrole rings and is located within the /3 barrel in a position almost identical to that of the retinol in RBP. Extensive interactions are made with hydrophobic residues lining the interior of the barrel.

D. Rodent Pheromone-Binding Urinary Proteins: A2U and M U P Rat an-globulin (A2U) and the homologous mouse protein (major urinary protein, MUP) are the principal proteins secreted in the urine of male rodents. Because male urine affects the behavior and sexual response of females, it has been suggested that these proteins function as pheromone-binding proteins. The crystal structures of both proteins have been determined (Bocskei et al., 1992). As expected (Cowan et al., 1990), both structures have the RBP fold. T h e Ca coordinates of the /3 barrel agree with an RMS difference of approximately 1.2 A. Both contain a ligand in the center of the barrel, where extensive interactions are made with the hydrophobic residues lining the barrel. For the MUP


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protein (resolution 2.4 A), the ligand is thought to be 2-(sec-butyl)thiazoline, but for the a2-globulin, the ligand is unknown, but is visible in a 2.8 8, resolution electron density map.

XI. FACTORS INVOLVED IN LIPIDBINDING IN iLBPs A. Site-DirectedMutagenesis of iLBPs Site-directed mutagenesis has been carried out on a number of iLBPs to investigate the role of individual residues identified in the crystallographic studies as participants in lipid binding. Side chains involved in directing specificity have received the most attention. However, much additional work needs to be done to study other conformational factors in ligand binding. This should include the hydrophobic side chains and cavity residues involved in forming water networks. These residues may be equally important in determining the ligand specificity, defining lipid conformation, and contributing to the overall binding energy. One other point is noteworthy. Although there are crystal structures for several iLBPs, no structures have been determined for any of the site-directed mutants. One can argue that because there is a wide degree of sequence variability within the entire gene family, a single mutation in the interior of the protein will not affect the overall folding of the protein. Whether this is true needs to be tested. With the paucity of crystallographic or NMR studies on the mutants, the results of sitedirected mutagenesis based on the structural data of wild-type proteins need to be interpreted with caution. Because the specificity motifs within the iLBP family seem to be characterizable by amino acids 106, 126, and 128 (or their homologous site equivalents relative to the ALBPIPB numbering system), these amino acids have been most extensively studied. By mutating a glutamine at position 106 into an arginine, one would expect to observe the CRBP specificity. When the Ql06R CRBP mutant was made, the affinity for retinol decreased by a factor of three and the site-specific change enabled the protein to bind all-tram-retinoic acid, 13-cis-retinoicacid, and retinal, in addition to all-tram-retinol (Stump et al., 1991). Two different mutations in CRBPII, Ql06R and Q126R, showed dramatic decreases in all-tram-retinol and retinal binding, but no increase in retinoic acid binding. Ql06R but not Ql26R produced a dramatic increase in palmitic acid binding (Cheng et a f . , 1991). Alternatively, a mutation (R106Q) in IFABP should affect fatty acid binding. This mutation caused a 20-fold decrease in fatty acid binding (Jakoby et al., 1993).

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In addition, the carboxylate group of bound fatty acid was in a solventaccessible environment, suggesting a lipid-binding mode different from that of wild-type protein. Furthermore, this mutant had an affinity for retinol when the wild-type IFABP had none (Jakoby et al., 1993). These experiments show that it is possible to switch the binding specificity of iLBP proteins by mutating individual amino acids associated with the specificity motif. These results d o not, however, indicate that the mutated positions are singularly responsible for ligand selectivity. Substitution of other cavity residues may have similar or contributory effects. Other site-directed mutagenesis studies have been undertaken in this family. R126Q or Y 128W in ALBP resulted in at least a 20-fold decrease in cis-parinaric acid binding (Sha et al., 1993). T h e loss of binding for the tryptophan mutant is probably due to steric effects in that the larger side chain blocks the wild type carboxylate-binding site. In CRABP, mutating the arginines to glutamines affected retinoic acid binding, but did not result in a retinol-binding protein (Zhang et al., 1992). All these mutational results suggest that the polar interactions, including hydrogen bonding and electrostatic interactions, play important roles in lipid binding to iLBPs. B. Thermodynamics of Lipid Binding Crystallographic studies of the iLBPs show that the lipid-binding cavities are not all hydrophobic, as might initially be anticipated. As noted in the preceding section, specificity appears to arise from polar interactions. But equally noteworthy is the fact that there are other polar, even charged, side chains within the ligand-binding cavity. The reason for their presence is unclear, although the structural data may offer some clues. Amphipathic lipid molecules at first glance suggest that a proteinbinding site would require a polar and hydrophobic counterpart. What is unusual about lipid binding in the iLBPs is that generally the carboxylate or hydroxyl group of lipid is located in the innermost part of the cavity rather than near the protein surface, as observed in serum retinolbinding protein. There is evidence suggesting that rat liver fatty acidbinding protein might be an exception (Cistola et al., 1989). Equally unexpected is the fact that the hydrophobic part of the fatty acid or retinoid is not in a totally hydrophobic portion of the P-barrel cavity. Titration calorimetry has been used to study the binding of ligands to the iLBPs (Jakoby et al., 1993; Miller and Cistola, 1993). This has shed new light on the thermodynamics of the association reaction permitting the Gibbs binding energy to be understood in terms of enthalpic and

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entropic contributions (Jakoby et al., 1993). For example, the binding of oleate to CRBPII and IFABP as well as to the mutants CRBPII (QlOSR) and IFABP (R106Q) has been studied in this fashion. T h e results are given in Table VIII (Jakoby et al., 1993). Here it is important to note that except for the IFABP (R106Q) mutant, most of the binding energy derives from enthalpic contributions. This then emphasizes the polar interactions seen in the crystallographic models. .4s seen in Table VIII, studies with IFABP indicate that almost 90% of the binding energy derives from enthalpic contributions. Trying to correlate this with the crystallographic model suggests that the formation of the ion pair between Arg-106 and the fatty acid carboxylate is of primary importance. The formation of a single ion pair between protein and fatty acid may not be the entire account of polar interactions. Noteworthy is that the ion pair should have a larger effect in ALBP, HFABP, MFB2, and P2 when two arginines and a tyrosine are involved. This does not seem to be the case, because the K d values appear to be about the same for oleate binding to the proteins with the double-arginine motif. For technical reasons involving solubilities of the retinoid, similar calorimetric measurements could not be made with retinol (Jakoby et al., 1993). In the broadest sense, the lipid-protein interface in the iLBPs is not constructed solely from protein atoms but includes ordered and disordered cavity water molecules. Because the water molecules are only weakly bonded to each other and to protein atoms, there is internal flexibility within the binding cavity. The protein may have to change water positions to accommodate the lipid ligand and the changes necessary for this accommodation may be different for different ligands. T h e water effect along with weak van der Waals interactions between methylene carbons may all make a significant contribution to the enthalpy associated with ligand binding and the lack of specificity in lipid binding found for some of the proteins in the family. TABLE VIII Titration Calorimetry: Binding of Oleate to U P S Protein Thermodynamic parameter

CRBPl 1 (Q109R)"

(W)

0.20 -38.1 -25.2 13

Kd

AG (kJ/mol)

AH (kJ/mol) TAS (kl/mol)

IFABP 0.20

-39 -34 4

IFABP (R106Q) 4.2 -30.7 -4 26

a Wild-type CRBPII has no detectable affinity for fatty acids. Data from Jakoby et al. (1993).

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Although not always a general rule, it appears that retinoid compounds have greater binding affinity for their binding proteins than do fatty acids, i.e., nanomolar range versus micromolar range (MacDonald and Ong, 1987; Lowe et al., 1987). One suggestion is that the retinol molecule will have the same conformation in the bound and unbound states. For example, the bound form of retinol is conformationally very similar to its crystal structure. Because of the presence of conjugated double bonds in the retinol molecule, the flexibility of the molecule is highly restricted. Fatty acids, on the other hand, have a large degree of rotational freedom around single C-C bonds. Such being the case, there would be a relatively more negative entropic contribution to the free energy of fatty acid binding compared to retinol binding. As a last note, serum retinol-binding protein binds retinol with high affinity. The ligand is located in the binding cavity with its hydroxyl group close to the protein surface. T h e complementarity between the retinol molecule and the protein resembles that of an enzyme-substrate complex. The cavity is very hydrophobic and there is no room for water molecules inside the cavity. T h e protein makes no direct polar interaction, even for the hydroxyl group of the retinol; rather, the hydroxyl group makes a hydrogen bond to a water, which in turn makes a hydrogen bond to the protein. It seems that the lipid binding in RBP is dominated by the hydrophobic effect, sharply contrasting with what is seen in the iLBPs. C . Protein Stability and Folding

Intracellular lipid-binding proteins present an interesting model to study protein folding. Although the sequence identity ranges from less than 20% to over 80% among the members of the family, overall the folds of the proteins are essentially the same. The structures of the proteins are unusual in that the hydrophobic core is relatively small and many hydrophilic side chains project into a large, solvent-filled interior cavity. The proteins are stable between pH 3 and pH 12 and to a temperature >70°C at pH 7.2 and low ionic strength (Ropson et al., 1990). Equilibrium studies of the folding reaction of apo-IFABP and apoALBP shows that the AG,,, values are 5.2 and 3.9 kcal/mol, respectively, using guanidinium hydrochloride as a denaturant (Ropson el al., 1990; Buelt et al., 1992). Given the small size of the protein hydrophobic core and no disulfide bond in the structure, the stabilities of these proteins are incredible. Kinetic studies of guanidinium hydrochloride-induced IFABP unfolding and refolding reactions showed both processes were rapid and complex, indicating the presence of intermediates in the fold-


145

ing process. The unfolding process can be interpreted as a two-phase process, suggesting that there is only one intermediate along the pathway. The refolding process, on the other hand, showed at least two intermediates. The changes in amplitudes of the reaction also indicated the existence of multiple pathways for the refolding (Ropson et al., 1990). Holo-IFABP shows increased stability with an additional 2.5 kcal/mol in AG,,, using both guanidinium hydrochloride and urea as denaturant (Ropson et al., 1990). Similar results have been obtained by comparing the guanidinium hydrochloride denaturation of apo- and holo-ALBP (V. Matarese and D. A. Bernlohr, unpublished). Comparison of the apo- and holo-IFABP and ALBP structures indicates the difference between the two structures is minimal (Scapin et al., 1992; Xu et al., 1992).The more negative AG can be partly attributed to the coulombic interaction between the carboxylate group and Arg- 106 and to the van der Waals interactions between the hydrophobic chain of the ligand and the protein cavity. In an analysis of the thermostability of CRABP and a number of arginine mutants (Zhang et al., 1992),retinoic acid binding was found to increase the melting temperature by 17.3"C, corresponding to a free energy change of 5.6 kcal/mol. Furthermore, the R106Q mutant stabilized the structure by 3.0 kcal/mol, R126Q by 3.7 kcal/mol, and the double mutant by 4.7 kcal/mol. T h e fact that the iLBPs do not have the typical hydrophobic core make them of special interest in folding studies. Furthermore, they are conformationally related to many other P-barrel proteins and a number of different forms are known (see Table 111). However, the mechanism of folding is still unknown, and as with all proteins, such determinations are difficult. With a growing knowledge of the invariant residues, the iLBPs may be an important model system.

XII. CONCLUDING REMARKS One of the reasons for studying lipid-binding proteins is that by examining the structural aspects of protein-lipid interaction one may, in effect, be able to infer physiological function. This has not been the case. The results presented within this review are far more informative as they pertain to the chemistry of lipid-protein interactions than they are about telling us the cellular function(s) of the iLBP multigene family. Nonetheless, the scientific community is now armed with a wealth of structural information regarding these P-barrel proteins. In comparison to other known protein structures, although a number of others contain some polar residues within their core, few are typified by a large cavity. And it was unexpected by many that a cavity designed to accommodate a hydro-

146


phobic ligand would appear to be filled with water molecules in the apo form. It seems equally unusual that a relatively large ligand such as a fatty acid o r retinoid could be accommodated inside a protein without detectable conformational changes. Furthermore, although a portal has now been postulated, the binding cavity in the iLBPs is essentially sealed from the milieu. Armed with the crystallographic data, site-directed mutagenesis has demonstrated that part of the specificity of lipid-binding resides in several related motifs that interact in a polar fashion with the hydrophobic ligand. Furthermore, the binding of fatty acids can be obliterated by a Tyr to T r p mutation in the binding cavity. Calorimetric measurements have suggested that much of the free energy of binding derives from enthalpic effects, which should translate into polar interactions. T h e iLBPs represent an especially good system for protein engineering and the authors hope that the next review will contain information not even imagined herein. In reviewing the crystallographic results concerning the FABPs, one might conclude that the proteins serve as nothing more than intracellular fatty acid buffers, establishing a pool of soluble lipid for metabolic utilization. Simply stated, the FABPs bind their respective ligands with roughly similar affinities and specificities. As shown in this review, the atomic details of such interactions vary considerably. Moreover, they do not appreciable alter their tertiary structures in response to ligand binding as one might imagine if the proteins were to interact with cellular enzymes. Despite the remarkable similarity in overall protein structure between family members, nature has presented a compelling argument that the hydrophobic ligand-binding proteins do serve separate and discrete functions. Perhaps the most cogent argument is the simple appreciation that the proteins are members of a multigene family. If the hydrophobic ligand-binding proteins serve to simply solubilize a fatty acid or a retinoid, why does nature choose to utilize different proteins in different cell types? If the iLBPs were to associate with other proteins in the cell, then the evolutionary driving force behind the expansion of the family is more obvious. In the case of nearly all of the iLBP family members, the search for such specific interactions has proved generally disappointing. An exception to this is the recent work from Herr and Ong (1992), who have detected subtle differences in the hydrodynamic radius of the cellular retinol-binding proteins on ligand binding. Such physical changes in protein structure may translate into differential interaction with cellular enzymes. Consequently, one might be able to rationalize the expansion of


147

the retinoid-binding proteins by pointing to specialized proteins with which they may or must interact. A second consideration may be that the differences in protein sequence translates into differences in ligand affinity, thereby producing a gradient of high-affinity and low-affinity binding proteins (Cistola et al., 1989). In a cellular context, this would lead to multiple accessible lipid pools. The pools in turn may affect lipid metabolism uniquely in different cells. Again, based upon in vitro analysis, the binding affinities for the FABPs for fatty acids are generally in the micromolar range, whereas those for retinoids are closer to 10 nM. Consequently, the binding data suggest that there is little significant difference in affinity for a ligand between subgroups of family members. One typical argument for necessity in diversity of the family is that each cell type possesses a unique metabolism and that a specialized binding protein is necessary to facilitate such metabolism (Matarese and Bernlohr, 1988). However, one must keep in mind that many of the compounds that hydrophobic ligand-binding proteins associate with are biochemically inert. They must be activated for further utilization. For example, the two common cellular enzymes that esterify fatty acids, the microsomal ATP-dependent acyl-CoA synthetase (or fatty acid : CoA ligase) and the GTP-dependent organellar synthetase, both produce an acyl-CoA (Bar-Tana et al., 1975). Therefore, virtually all cells utilize fatty acids in the same fashion-they esterify them with CoA. Cells utilize acyl-CoAs in vastly different fashions. A class of acyl-CoAbinding proteins has been characterized (Knudsen, 1990). It appears as though there is a single functional acyl-CoA-binding protein expressed in a variety of cells. If this is the sole CoA carrier, all cells would utilize the same acyl-CoA-binding protein as the cellular donor of acyl-CoAs for metabolism. In a metabolic sense one might argue that the unique metabolism of specialized cells should be exemplified by the evolution of a single hydrophobic ligand-binding protein expressed in all cell types, with a multigene family of acyl-CoA-binding proteins each funneling its ligand into the specialized pathways pursuant to each cell. Last of all, the need for the iLBPs could derive from the need to maintain low levels of amphipathic compounds within the cytoplasm. This passive role is related to protecting against the potential detergentlike character of fatty acids and retinoids. If this is one of their physiological roles, an alternate possibility is that the hydrophobic ligand-binding proteins do not interact with unique protein components of the cell but with other cellular structures, such as intracellular membranes. Experiments (Wootan et al., 1993) have suggested that local membrane composi-

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tion may affect the kinetics of a fatty acid-binding protein to either accept or donate a ligand. Because different cell types have different bilayer surfaces, the need for specialized proteins could be related to their ability to interact with their membranes and buffer the presence of amphipathic lipids. Clearly, the structural data presented here and in the accompanying references allow for a variety of experiments that probe structure/ function relationships of the lipid-binding proteins. Techniques such as X-ray crystallography and NMR, coupled with the availability of native and engineered mutants, make mechanistic studies of the lipid-binding proteins an interesting and exciting field of study. We hope that the information presented here will provoke new generations of experiments designed to address the physical, chemical, and biological role(s) for these proteins. ACKNOWLEDGMENTS The authors would like to thank the members of the Banaszak. Bernlohr, and Jones laboratories for their helpful comments during the preparation of this manuscript. In addition, we would like to thank our colleagues who allowed us to include unpublished and “in press” information as part of this review. Last, the authors acknowledge support from the National lnstitutes of Health (GM 13925 to L.J.B.), the National Science Foundation (DMB9118658 to D.A.B. and L.J.B.), and the Swedish Natural Science Research Council and Uppsala University (to T.A.J.).

REFERENCES Aqvist, J., Sandblom, P., Jones, T. A., Newcomer, M. E., van Gunsteren, W. F., and Tapia, 0. (1986).J. Mol. Biol. 194,593-604. Bar-Tana,J., Rose, G., and Shapiro, B. (1975).In “Methods in Enzymology”(J.Lowenstein, ed.), Vol. 35, pp. 117-122. Academic Press, New York. Bass, N. M. (1988). Inf. Rev.Cylol. 111, 143-184. Baxa, C. A., Sha, R. S., Buelt, M. K., Smith, A. J., Matarese, V., Chinander, L. L.. Boundy, K. L., and Bernlohr, D. A. (1989). B b c h i s t v 28,8683-8690. Benning, M. M., Smith, A. F., Wells, M. A., and Holden, H. M. (1992).J. Mol. Biol. 228, 208-219. Bernier, M., Laird, D. M., and Lane, M. D. (1987). Proc. Natl. Acad. Sci. U.S.A. 84, 18441848. Biicskei, Z., Groom, C. R., Flower, D. R., Wright, C. E., Phillips, S. E. V., Cavaggioni, A., Findlay, J. B. C., and North, A. C. T. (1992). Nature (London) 860, 186-188. Bohmer, F.-D., Kraft, R., Otto, A., Wernstedt, G., Hellman, U., Kurtz, A., Muller, T., Rhonde, K., Etzold, G., Lehmann, W., Langen, A., Heldin, C.-H., and Grosse, R. (1987).J. Biol. C h . 262, 15137-15143. Branden, C., and Tooze, J. (1991). “Introduction to Protein Structure.” Garland Publ., New York. Braunitzer, G., Chen, R., Schrank, B., and Stangl, A. (1973). Hoppe-Seyler‘s Z. Physiol. Chem. 354,867-878.


149

Buelt, M. K., Shekels, L. L., Jarvis, B. W., and Bernlohr, D. A. (1991).J.Biol. Chem. 266,

12266-12271. Buelt, M. K., Xu, Z., Banaszak, L. J., and Bernlohr, D. A. (1992).Eiochnnishy 31, 3493-

3499. Buelt, M. K., Bernlohr, D. A., and Tennekoon, G. (1993).In preparation. Cheng, L., Qian, S., Rothschild, C., Avignon, A., Lefkowith, J., Gordon, J. I., and Li, E. (1991).J.Biol. Chem. 466,24404-24412. Chytil, F., and Ong, D. E. (1987).Annu. Rev. Nutr. 7,321-335. Cistola, D. P., Sacchettini, J. C., Banaszak, L. J., Walsh, M. T., and Gordon, J. I. (1989). J . Biol. Chem. 264,2700-2710. Cowan, S. W., Newcomer, M. E., and Jones, T. A. (1990).Prolcinc 8,44-61. Cowan, S . W., Newcomer, M. E., and Jones, T. A. (1993).J.Mol. Bwl.230,1225-1246. Dolle, P., Ruberte, E., Leroy, P., Morriss-Kay, G., and Chambon, P. (1990).Deuelopmmt (Cambridge, UK)110,1133-1151. Fiorella, P. D., and Napoli, J. L. (1991).J. Eiol. Chem. 266, 16572-16579. Gantz, I., Northwehr, S. F., Lucey, M., Saccettini,J. C., DelValle,J., Banaszak, L. J., Naud, M., Gordon, J. I., and Yamada, J. H.(1989).J.B i d . Chem. 264,20248-20254. Giguere, V., Lyn, S., Yip, P., Siu, C., and Amin, S. (1990).Proc. Natl. Acad. Sci. U.S.A. 87,

6233-6237. Glatz, J. F. C., and Veerkamp, J. H. (1985).1nt.J. Eiocha. 17,13-22. Goodman, D. S. (1984).In “The Retinoids” (M. B. Sporn, A. B. Roberts, and D. S. Goodman, eds.), Vol. 2,pp. 41-88.Academic Press, New York. Hendrickson, W.,Pahler, A., Smith, J., Satow, Y., Merritt, E., and Phizackerley, R. (1989). Proc. Natl. Acad. Sci. U.S.A. 86,2190-2194. Herr, F. M., and Ong, D. E. (1992).Biochemistry 31,6748-6755. Holden, H., Rypniewski, W., Law, J., and Rayment, I. (1987).EMBOJ. 6, 1565-1570. Hresko, R. C., Bernier, M.,Hoffman, R. D., Flores-Riveros,J. R., Liao, K., Laird, D. M., and Lane, M. D. (1988).Proc. Natl. Acad. Sci. U.S.A. 85,8835-8839. Huber, R., Schneider, M.,Epp, O., Mayr, I., Messerschmidt, A., and Pflugrath, J. (1987a). J.Mol. BWl. 195,423-434. Huber, R., Schneider, M., Mayr, I., Muller, R., Deutzman, R., Suter, F., Zuber, H., Falk, H., and Kayser, H. (1987b).J.Mol. Biol. 198,499-513. Hunt, C. R., Ro, J. H.-S., Dobson, D. E., Min, H. Y., and Spiegelman, B. M. (1986).Proc. Natl. Acad. Sci. U.S.A. 89,3786-3790. Jakoby, M. G., Miller, K. R., Toner, J. J., Bauman, A., Cheng, L., Li, E., and Cistola, D. P. (1993).BioChemistty 32,872-878. Jones, T. A., Bergfors, T., Sedzik,J., and Unge, T. (1988).EMBOJ. 7, 1597-1604. Kanai, M., Raz, A., and Goodman, D. S. (1968). J. Clin. Invest. 47,2025-2044. Kanda, T., Odani, S., Tomoi, M., Matsubara, Y., and Ono, T. (1991).Eur. J.Biochem. 197,

759-768. Kim, H. K., and Storch, J. (1992a).J.Biol. Chem. 267,77-82. Kim, H. K.,and Storch, J. (1992b).J. Eiol. Chem. 267,20051-20056. Knudsen, J. (1990).Mol. Cell. Biochem. 98,217-223. Krieg, P., Feil, S., Fiirestenberger, G., and Bowden, G. T. (1993).J. Bwl. Chem. 268,

17362- 17369. Levin, M. S., Locke, B., Yang, N.-C. C., Li, E., and Gordon, J. I. (1988).J. Biol. C h a . 263,

I7715-17723. Levitt, D. G., and Banaszak, L. J. (1992).J. Mol. Graphics 10,229-234. Li, E., Qian, S.-J., Yang, N.-C. C., d’Avignon,A., and Gordon,J. I. (199O).J.Biol. Chem. 265, 1 1549- 1 1554.

150


Li, E., Qian, S.-J.,Winter, N. S.. d’Avignon, A., Levin, M. S.,and Gordon,J . 1. (1991).J.Biol. Chem. 266,3622-3629. Liberatori, J . (1977). Folio Vet. Lat. 7,205-222. Lipman, D. J., Altschul, S. F., and Kececioglu,J . D. (1989). Proc. Natl. Acad. Sci. U.S.A. 86, 44 12-44 15. Lowe, J. B., Sacchettini,J. C., Laposata, M., McQuillan,J. J., and Gordon,J. 1. (1987).J.Mol. Biof. 197,695-706. MacDonald, P. N., and Ong, D. E. (1987).J.Biof. Chem. 262, 10550-10556. Matarese, V., and Bernlohr, D. A. (1988).J.Biol. Chem. 263, 14544-14551. McKenzie, H. A. (1971). In “Milk Proteins: Chemistry and Molecular Biology” (H. A. McKenzie, ed.), Vol. 2, pp. 257-330. Academic Press, New York. McKenzie, H. A., Ralston. G. B., and Shaw, D. C. (1972). Biochemistry 11,4539-4547. McRee, D., Tainer, J.. Meyer, T., Van Beeumen, J., Cusanovich, M., and Getzoff, E. (1989). Proc. Natl. Acad. Sci. U.S.A. 86,6533-6537. Medzihradszky, K., Gibson, B., Kaur, S., Yu,Z., Medzihradszky. D., Burlingame. A. L.. and Bass, N. M. (1992). Eur.1. Biochm. 203,327-339. Miller, K. R., and Cistola, D. P. (1993). Mof. Cell. Biochem. (in press). Mishkin, S., Stein, L., Gatmaitan, Z., and Arias, 1. M. (1972).Biochem. Biophys. Res. Commun. 47,997-1003. Monaco. H.L., Zanotti, G., Spadon. P., Bolognesi, M.. Sawyer, L., and Eliopoulos, E. E. (1987).J.M o f . Biof. 197,695-706. Moser, D., Tendler, M., Griffiths, G., and Klinkert, M. (1991).J. Biol. Chem. 266, 84478454. Muller-Fahrnow, A., Egner, U., Jones, A.. Rudel, H., Spener, F., and Saenger, W. (1991). Eur. J.Biochem. 199,27 1-276. Musacchio,A., Noble, M., Pauptit, R., Wierenga, R., and Saraste, M. (1992).Nature (London) 359,851-855. Newcomer, M. E., Jones, T. A., diqvist,J., Eriksson, U.. Rask, L., and Peterson, P. A. (1984). E M B O J . 3, 1451-1454. Ockner, R. K., Manning, J . A., Poppenhausen, R. B.. and Ho. W. K. L. (1972).Science 177, 56-58. Oram, J. F., Bennetech, S. L., and Neeley, J. R. (1973).J. Biof. Chem. 248,5299-5309. Papiz, M. Z., Sawyer, L., Eliopoulos, E. E., North, A. C. T., Findlay,J . B. C., Sivaprasadarao, R.. Jones, T. A., Newcomer, M. E., and Kraulis, P. J. (1986). Nature (London) 324, 383-385. Peeters, R. A., Veerkamp, J . H., Van Kessel, A. G., Kanda, T., and Ono. T. (1991).Biochem. J. 276,203-207. Raag, R., Appelt, K., Xuong, N.-H., and Banaszak, L. J . (1988).J.Mol. Biol. 200,553-569. Rask, L., Anundi, H., Bohme, J., Eriksson, U., Fredriksson, A., Nilsson, S. F., Ronne, H., Vahlquist, A., and Peterson, P. A. (1980).J. Cfin. Lab. Invest. 40,45-61. Richardson, J. S., and Richardson, D. C. (1989).In “Prediction of Protein Structure and the Principles of Protein Conformation” (G. D. Fasman, ed.), pp. 1-98. Plenum, New York. Ronne, H.,Ocklind, C., Wiman, K., Rask, L., Obrink, B., and Peterson, P. A. (1983).J.Cell Biol. 96, 907-910. Ropson, 1. J., Gordon, J. I., and Frieden, C. (1990). Biochemistry 29,9591-9599. Sacchettini, J. C., and Gordon, J. I. (1993).J.Biof. Chem. 268, 18399-18402. Sacchettini, J. C., Gordon, J . I., and Banaszak, L. J. (1987). J . Biof. Chem. 263, 58155819. Sacchettini, J. C., Gordon, J. I., and Banaszak, L. J.(1989).J . Mol. Biof. 208,327-339.


151

Sacchettini, J. C., Hauft, S. M., Van Camp, S. L., Cistola, D. P., and Gordon, J. I. (1990). J. Biol. Chem. 265, 19199-19207. Scapin, G., Spadon, P., Mammi, M., Zanotti, G., and Monaco, H. (1990).Mol. Cell. Biochem. 98,95-99. Scapin, G., Gordon, J. I., and Sacchettini, J. C. (1992).J.Biol. Chem. 267,4253-4269. Sha, R. S., Kane, C. D., Xu, Z., Banaszak, L. J., and Bernlohr, D. A. (1993).J.BioLChem. 268, 7885-7892. Shields, H. M., Bates, M. L., Bass, N. M., Alpers, D. H., and Ockner, R. K. (1986).J.Lipid Res. 27,549-557. Smith, A. F., Tsuchida, K., Hannernan, E., Suzuki, T. C., and Wells, M. A. (1992).J. Biol. Chem. 267,380-384. Stump, D. G., Lioyd, R. S., and Chytil, F. (1991).J.Biol. Chem. 266,4622-4630. Sundelin, J., Das, D., Ericksson, U., Rask, L., and Peterson, P. (1985).J. Biol. Chem. 260, 6494-6499. Sweetster, D. A., Heuckeroth, R. O., and Gordon, J. I. (1987).Annu. Rev. Nutr. 7,337-359. Sweetser, D. A., Birkenmeier, E. H., Hoppe, P. C., Mckeel, D. W., and Gordon, J. I. (1988). Genes Dev. 2, 13 18- 1332. Trapp, B. D., Bubois-Dalcq, M., and Quarles, R. H. (1984).J.Neurochem. 43,944-948. Tsonis, P. A., and Goetinck, P. F. (1988). Biochem. J. 249,933-934. Uyernura, K., Yoshinura, K., Suzuki, M., and Kitamura, K. (1984). Neurochem. Res. 9, 1509- 15 14. Vainshtein, B. K., Melik-Adamyan, W. R., Barynin, V. V., Vagin, A. A., Grebenko, A. I., Borisov, V. V., Bartels, K. S., and Rossmann, M. G. (1986).J.Mol. Biol. 188,49-61. Veerkamp, J. H., Peeters, R. A., and Maatrnan. R. G. H. J. (1991). Biochim. Biophys. Acfu 1081,l-24. Weiss, M. S., Kreusch, A., Schiltz, E., Nestel, U., Welte, W., Weckesser, J., a n d Schulz, G. E. (1991).FEES Lett. 280,379-382. Winkler, F., Hunziker, W., and DArcy, A. (1990).Nufure (London) 343,771-774. Winter, N. S., Bratt, J. M., and Banaszak, L. J. (1993).J.Mol. Biol. 230, 1247-1259. Wootan, M. G., Bernlohr, D. A., and Storch, J. (1993).Biochemistry in press. Xu, Z., Bernlohr, D. A., and Banaszak, L. J. (1992).Biochemistry 31,3484-3492. Xu, Z., Bernlohr, D. A., and Banaszak, L. J. (1993).J.Biol. Chem. 268,7874-7884. Zanotti, G., Ottonello, S., Berni, R., and Monaco, H. (1993).J.Mol. Biol. 230,613-624. Zanotti, G., Scapin, G., Spadon, P., Veerkarnp, J. H., and Sacchettini, J. C. (1992).J. Biol. Chem. 267,18541-18550. Zhang, J., Liu, Z.-P., Jones, T., Gierasch, L., and Sambrook,J. (1992).Proteh: S t n u t . Funct., Genet. 13,87-99.


By DANIEL C. CARTER and JOSEPH X. HO Space Sclence Laboratory, Blophyslcs Branch, Natlonal Aeronautics and Space Admlnlatratlon, Marshall Space Fllght Center, Huntsville, Alabama 35812

1. Introduction ....................................................... 11. Albumin Structure ..................................................

A. Primary Structure .............................................. B. Early Concepts of Albumin Structure ............................. C. Three-Dimensional Structure .................................... 111. Nature of Ligand Binding ........................................... A. Small Organic Compounds: Sites I and I1 ......................... B. Long-Chain Fatty Acids: Sites Ill and IV .......................... C. Metals: Sites V and VI ........................................... IV. Evolution of Albumin Structure ...................................... V. Summary and Future Directions ...................................... References .........................................................

153 155 155 161 166 176 181 186 188 190 194 196

I. INTRODUCTION Often utilized as a substitute for a typical protein, albumin needs no introduction to the protein chemist. Because of its availability, low cost, stability, and unusual ligand-binding properties, serum albumin has been one of the most extensively studied and applied proteins in biochemistry. However, as a protein, albumin is far from typical, and the widespread interest in and application of albumin have not been balanced by an understanding of its molecular structure. Indeed, for more than 30 years structural information was surmised based solely on techniques such as hydrodynamics, low-angle X-ray scattering, and predictive methods. Serum albumin was recognized as a principal component of blood as early as 1839 (Ancell, 1839). Early researchers generally referred to protein as “albumen” stemming from Latin albus after the white color of flocculant precipitates produced by various proteins. Today, one should be aware that several proteins share this name but they are structurally and functionally unrelated to serum albumin, e.g., ovalbumin and prealbumin. Hence, care should be taken to distinguish ablumen, which refers to egg whites, from albumin or serum albumin. As the most abundant protein in the circulatory system and with typical blood concentrations of 5 g/100 ml, albumin contributes 80% to colloid osmotic blood pressure. In addition, it has now been determined that albumin is chiefly responsible for the maintenance of blood pH (Figge et ADVANCES IN PROTEIN CHEMISTRY, Vol. 45

153

Copyright 01994 by Academic Press, lnc. All rights of reproduction in any form reserved.

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DANIEL C. CARTER A N D JOSEPH X. HO

al., 1991). It is located in every tissue and bodily secretion, with the extracellular protein comprising 60% of total albumin. In mammals albumin is synthesized by the liver (Peters and Anfinsen, 1950) and possesses a half-life in circulation of 19 days (Waldmann, 1977). Despite the major role of albumin in circulating plasma, the absence of albumin, or “analbuminemia,” though rare, has been observed in several individuals (Russi and Weigand, 1983) and one strain of rats (Nagase et al., 1979). Still, albumin is expressed in these individuals, but at a much lower level. Clinical manifestations of analbuminemia are not completely understood, but include chronic fatigue, hyperlipidemia, and, in rats, an increased susceptibility to cancer (Kakizoe and Sugimura, 1988). Perhaps, the most outstanding property of albumin is its ability to bind reversibly an incredible variety of ligands. (See later, Table V, which provides a selective listing of the extensive literature regarding ligand binding to albumin; see also Section 111.) Befitting its inclusion in this volume, albumin is the principal carrier of fatty acids that are otherwise insoluble in circulating plasma. But albumin performs many other functions as well, such as sequestering oxygen free radicals and inactivating various toxic lipophilic metabolites such as bilirubin (Emerson, 1989). Although albumin has a broad affinity for small negatively charged aromatic compounds, it has high affinities for fatty acids, hematin, and bilirubin. Additionally, it forms covalent adducts with pyridoxyl phosphate, cysteine, glutathione, and various metals, such as Cu(11), Ni(II), Hg(II),Ag(II), and Au(1). The participation of albumin as the key carrier o r reservoir of nitric oxide, which has been implicated in a number of important physiological processes, including neurotransmission, serves to illustrate further the continuing recognition of the utility of albumin (Stamler et al., 1992). Without question, albumin is the most multifunctional transport protein known to date. Albumin belongs to a multigene family of proteins that includes afetoprotein (AFP) and vitamin D-binding protein (VDP),also known as G complement (Gc) protein. Although AFP is considered the fetal counterpart of albumin, its binding properties are distinct and it has been suggested that AFP may have a higher affinity for some unknown ligands important for fetal development. VDP plays an important role in calcium regulation. Additionally, AFP and VDP both interact with the class I1 major histocompatibility complex. These and other data suggest that these proteins may play an important role in modulating the immune system (Oers et al., 1989). Albumin, although homologous in structure, shares no immunological properties with these proteins. The occurrence of AFP in adult serum is usually associated with disease (Adinolfi et al., 1975). A high degree of conformational flexibility of albumin has been recog-

SERUM ALBUMIN STRUCTURE

155

nized for many years. Foster (1960) documented several isomeric forms of albumin that can be induced reversibly as a function of pH (see Section II,C,2,d). Although there has been speculation about the possible function of one of the transitions, the physiological significance of these isomeric forms remains in question. Depsite the conformational adaptability of albumin, it is not readily denatured and survives heat pasteurization at temperatures of 60°C for 10 hr. without deleterious effects (Shrake et al., 1984). In addition to potential conformational isomerization, other forms of albumin microheterogeneity occur as well. In circulating plasma approximately 30% of free sulfhydryl, Cys-34, is oxidized by cysteine and glutathione (Peters, 1985). Additional forms of albumin impurities arise from its high affinity for fatty acids, bilirubin, hematin, and metals, such as nickel and copper. Moreover, microheterogeneity can be introduced by the isolation procedures. For example, some albumin preparations contain up to 20% of dimerized albumin. The percentage of this dimer increases with the age of the protein unless Cys-34 has been blocked with cysteine or glutathione. Albumin for which Cys-34 is free is referred to as “mercaptalbumin.” Furthermore, 7 to 10% of normal human albumin in circulation is glucosylated and a much higher percentage is observed for individuals with diabetes (Guthrow et al., 1979). Over the years there have been many outstanding reviews on albumin (Hughes, 1954; Foster, 1960; Kragh-Hansen, 1981; Fehske et al., 1981; Brown and Shockley, 1982; Peters, 1980, 1985, 1992; Rothschild et al., 1988) and several other reviews that describe more specialized studies. Because of the extensive nature of the literature regarding albumin, the authors have relied heavily on these reviews to provide many of the key works on albumin. They were essential both during the structure determination process and in the preparation of this review. It is the aim of this review to correlate many of the findings of past significant papers with the current knowledge of the recently determined X-ray structure (He and Carter, 1992). Clearly, such an immense body of literature predating the structure determination provides an opportunity and privilege afforded few crystallographers. 11. ALBUMIN STRUCTURE

A. Primary Structure

In recent years there has been a flurry of albumin sequence information, and now amino acid sequences have been deduced for several albumins as well as other members of the multigene family (Table I). The

156

DANIEL C. CARTER AND JOSEPH X. HO

TABLE I Determindion of Amino Acid Seguences Protein and source

Abbreviation

Ref.

Albumin Human

HSA

Bovine

BSA

Behrens et al. (1975),Meloun et al. (1975),Lawn et al. (198l),Dugaiczyk et al. (1982) Brown (1975),Holowachuk (1991) Ho et al. (1993) Sargent etal. (1981) Minghetti et al. (1985) Weinstock and Baldwin (1988) Brown et al. (1989) Haefliger et al. (1989) Byrnes and Gannon (1990) Gray and Doolittle (1992)

Equine Rat Mouse Pig Sheep Frog Salmon Lamprey a-Fetoprotein Human

ESA RSA MSA PSA OSA XSA SSA LSA

Rat Mouse Vitamin D-binding protein Human

-

Rat Mouse

AFP

VBP

-

Law and Dugaiczyk (1981), Morinaga el al. (1983) Jagodzinski et al. (1981) Gorinetal. (1981) Yang et al. (1985),Schoentgen et al. (1986) Cooke and David (1985) Yang et al. (1990)

sequences of all albumins are characterized by a unique arrangement of disulfide double loops that repeat as a series of triplets. Mammalian albumins have nine such loops formed by 17 disulfides. From observations of these loops and other homologies, albumin was deduced to be the product of three homologus domains (I, 11, III), a result of some early gene fusion event (Brown, 1977; McLachlan and Walker, 1977). Interestingly, the only known exception to the three-domain structure is lamprey albumin, which has seven domains. Brown, studying bovine serum albumin (BSA), further divided these domains into three subdomains, A, B, and C, after the three disulfide loops, and utilized this nomenclature to discuss hypothetical structure and ligand-binding properties. Other investigators have preferred to associate ligand-binding activities with the individual loops (Ll-Lg) (Peters, 1985) (Fig. 1). From examinations of


157

11 12 DOMAIN1

13

14 15

DOMAIN I1

16

17

18

DOMAIN 111

19

FIG. 1. Illustration of the loop-link-loop pattern in the primary structure of serum albumin. The three domains ( I , 11, and 111) are shown; loops are numbered 1 to 9 (LI-Lg).

TABLE I1 Aligned Albumin Sequences Illustrating Conserued Amino Acids and Invariant ResidueP SEQUENCE HOMOUXSY: HOMAN/BOVINE/EQUINE/OVINE/RAT/FRW/SALMON

TABLE I1 (continued)

Light shading indicates conserved amino acids; dark shading indicates invariant residues. The approximate positions of the 28 helical segments of HSA are also indicated; *, deleted.

159

160


the internal sequence homology of BSA, McLachlan and Walker (1977) correctly deduced that albumin is composed of six subdomains, which were later verified by the crystal structure (Carter et al., 1989; Carter and He, 1990; He and Carter, 1992). A modified nomenclature was adopted to reflect the domain structure observed in the three-dimensional structure better, and it should not be confused with the earlier, similar nomenclature of Brown (see Section II,C,2,a). It should be also be noted that throughout the remainder of this review all albumin sequences (see, e.g., Table 11) are normalized to that of HSA. T h e amino acid sequence of albumin is further characterized by unusually high percentages of Cys (35%) and charged amino acids, and low percentages of tryptophan, glycine, and methionine (Brown and Shockley, 1982; Peters, 1985). The primary structure of albumin is unusual among extracellular proteins in possessing a single free sulfhydryl (Cys34) and in having no sites for enzymatic glycosylation. However, this is not the case for other albumin family members, such as a-fetoprotein (Law and Dugaiczyk, 1981; Morinagaetal., 1983), VDP (Yangetal., 1985; Schoentgen et al., 1986), and lamprey and frog albumins, which are glycoproteins. The sequence homologies are high for all albumins determined thus far except for salmon, frog, and lamprey (Table 11). Still, all conserve the characteristic repeating disulfide pattern. High percentages of sequence identities have been noted between HSA and BSA (76%) (Peters, 1985), and the recently determined amino acid sequence deduced from cDNA for horse serum albumin (ESA), which reveals sequence identities between ESA and HSA, BSA, and RSA for 76, 73, and 76%, respectively (Ho et al., 1993). HSA and human AFP exhibit 39% homology (Morinaga et al., 1983). One of the major differences between AFP and serum albumin is the absence of a double disulfide bridge located in L6. Similarly, VDP lacks L9 in its entirety, and overall VDP proteins exhibit much lower homology with albumin, suggesting an early divergence of this gene. Homology among VDPs, however, is quite high (Gray and Doolittle, 1992). Gray and Doolittle (1992) recently constructed a phylogenetic tree for the albumin superfamily based on their work with the more distantly related lamprey serum albumin (LSA).From the sequence they deduce that LSA, like all other members of the albumin family, is composed of 190 amino acid repeats, suggesting, “that the invention of albumin antedates the vertebrate radiation.” From these data and comparative sequence alignments with other members of the albumin multigene family, they place the gene duplication event that gave rise to other members of the albumin three-domain structure prior to the appearance of vitamin D-binding protein, approximately 450 million years ago. It has


161

been theorized that the 190-amino acid “protoalbumin” originated by an incomplete gene fusion event involving some primordial globin or globin precursor (Brown, 1976). A more detailed discussion of the structural evolution of albumin is presented in Section IV. Polymorphisms in the sequences of human albumin are quite rare, but are relatively common in cattle, horses, and sheep (Tucker, 1968). T o date, the positions of 43 single-site point mutations in HSA have been determined, largely due to the efforts of F. W.Putnam, and these data are presented in Table 111. T h e majority of the variants have been identified by anomalous electrophoretic migration (Tarnoky, 1980) or, in one case, by the unusually high affinity for thyroxin (Borst et al., 1983). Therefore, typical commercial preparations of HSA produced from outdated, pooled blood can be considered essentially homogeneous in amino acid sequence.

B . Early Concepts of Albumin Structure Over the years, albumin has been probed with diverse experimental methods, including hydrodynamics, low-angle X-ray scattering, fluorescence energy transfer, electrophoretic methods, and NMR, IR, UV, mass, and Raman spectroscopies. Virtually every measurable property has been determined more than once (Table IV), with each investigator seeking to establish important insight into albumin structure and chemistry. Based largely on hydrodynamic experiments (Hughes, 1954; Squire et al., 1968; Wright and Thompson, 1975) and low-angle X-ray scattering (Bloomfield, 1966), the early conception of serum albumin began to converge on the shape of an oblate ellipsoid having dimensions or axes of 140 x 40 A (Fig. 2). Experiments have continued to support these dimensions (Bendedouch and Chen, 1983; Feng et al., 1988). Brown and Shockley ( 1982), with their demonstrated knowledge of albumin, constructed a “cigar-shaped” three-domain model of BSA that complied with an incredibly diverse variety of data. In the absence of an X-ray structure, this model has served as a frequently referenced conceptual model, albeit an incorrect one, for the past decade. In fact, in the early stage of crystallographic analysis of HSA at low resolution, due to the special packing arrangement of the molecules in the crystal, it seemed to us that the cigar-shaped model was plausible (Carter et al., 1989; Carter and He, 1990). Despite the general acceptance of the cigar-shaped structure of serum albumin (Fig. 2), several research groups provided evidence otherwise. Electron microscopy indicated that BSA was a roughly spherical molecule

TABLE I11 Amino Acid Substitutions and Codon Chunges fm Albumin Cenctu Variant9

Substitution -2 Arg+ His - 2 Arg + Cys - 1 Arg + Gln - 1 Arg -+ Pro - 1 Arg+ Leu 1 Asp+ Val

& r4

Variant name

60 Glu + Lys 63 Asp + Asn 82 Glu -+ Lys 114 Arg + Gly 119 Glu -+ Lys 128 His + Arg 240 Lys + Glu 268 Gln --* Arg 269 Asp + Gly 313 Lys+ Asn 318 Asn- Lys 320 Ala + T h r

Lille, etc. Malmo I, etc. Christchurch, etc. Takefu, etc. Jaffna Iowa City-2 Blenheim Nagasaki-3 (exonic borders = I ) Torino Malmo (#95) Vibo Valentia Yanomama Nagoya Komagome-2 Herborn Malmo (# 10) Nagasaki- 1 Tagliacozzo, etc. Malmo (#47) Redhill

321 Glu + Lys

Roma

3 His+Gln

Codon changeb

Minimum change

Ref.

TICST + CAT TIGGT + I G T TlCGA -+ CAM TlCGA + CCA TlCGA + C I A AlGAT -+ G P

G+A C+T G-A G+C G-T A-T

Abdo et al. (1981). Arai et 01. (1990), Takahashi el al. (1987b) Brennan el al. (1990) Arai et al. (1990), Brennan and Carrel1 (1978) Arai etal. (1990), Takahashi et al. (1987b) Galliano et of. (1989) Brennan et al. (1989)

CAGIA + C A a or CJ (CIGT + AGMCIA)

C + AlG

Arai et 01. (1989b), Takahashi et al. ( 1 9 8 7 ~ )

TIGAA + -MA TISAC -MC TlGAA + -MA ClGGA + -aA AIGAG + -MG TICAT + CGT Cl-MA -+ GAA T1C-A + CGA AlGAT + GGT TlAAG A A U or C) A A s l T A A U or CJ TlWT ACT

G-A G+A G+A C+G G+A A+G A-+G A-G A+G G .--, TIC C AlG G+A

TIGAG

G-A

Galliano et al. ( 1990) Carlson ef af. (1992) Galliano et al. ( 1990) Takahashi et al. ( 1 9 8 7 ~ ) Arai et al. (1990) Madison et al. ( 1991) Minchiotti el al. (1993) Carlson et al. ( 1 992) Arai el al. (1989b). Sugita et al. (1987), Takahashi et al. ( 1 9 8 7 ~ ) Galliano ef al. (1986, 1990) Carlson ef al. (1992) Brand et al. (1984). Hutchinson and Matejtschuk (1985), Brennan et al. (1990b) Galliano et al. (1988, 1990)

-

--

AAG

-

---

333 Glu Lys 354 Glu Lys 358 Glu Lys 365 Asp + His 365 Asp Val 372 Lys + Glu 375 Asp- Asn 376 Glu Lys 376 Glu Gln 382 Glu + Lys 479 Glu + Lys 494 Asp + Asn 501 Glu --* Lys 505 Glu Lys 536 Lys + Glu 541 Lys + Glu 550 Asp Gly 550 Asp- Ala 563 Asp Asn 565 Glu + Lys 570 Glu + Lys 573 Lys + Glu 574 Lys-, Asn

--

Sondrio Hiroshima-I Coari-I, etc. Parklands Iowa City-I NaskapilMersin, etc. Nagasaki-2 Tochigi, etc. Malmo (#5, etc.) Hiroshima-2 Dublin, etc. Casebrook Vancouver, etc. Ortonovo Castel di Sangro Maku Mexico Malmo (#61 and 96) Fukuoka- 1, etc. Osaka-1 B-tYpe Gent (MilFg) Vanves

----------

T&AA -MA TIGAA- AAA AIGAG -AG AIGAT + CAT AIGAT G T T Cl-AA GAA CISAT -AT TIGAA -MA TIGAA CAA GIGAA -MA AlGAA -MA CISAT -AT AIGAG -MG TIGAA -MA CI-MG GAG A/-MA GAA TIGAT GGT TIGAT + C&T CISAT + -&4T GIGAG -&4G CIGAG -AG TI-MA GAA AIAAB + AA(T or C J

G- A G-A G- A G-C A-T LA- G G- A G- A G+C G-A G- A G- A G- A G+ A A-G A+G A+G A+C G-A G-A G-A A+G A + TIC

Minchiotti et al. (1992) Arai et al. (1989b), Takahashi et al. ( 1 9 8 7 ~ ) Arai et al. (1989a) Brennan (1985) Madison el al. (1991) Takahashi et al. (1987a) Arai et al. (1989b), Takahashi et al. ( 1 9 8 7 ~ ) Arai et al. (1989b) Carlson et al. ( 1992) Arai et al. (1989b), Takahashi el al. ( 1 9 8 7 ~ ) Sakamoto et al. (1991) Peach and Brennan (1991) Huss et al. (1988) Galliano et al. ( 1993) Minchiotti et 01. (1990) Takahashi et al. ( 1 9 8 7 ~ ) Franklin et al. (1980) Carlson et al. ( 1992) Arai et al. ( 1990) Arai el al. ( 1990) Winter et al. (1972) ladarola et al. (1985) Minchiotti et al. (1987)

-

Positions of currently known single-site substitutions in human serum albumin. Provided in advance of publication as a courtesy by F. W. Putnam. Except for proalbumin Malmo I, none of the above represents a C + T change (even in the absence of a CG), i.e., CG TG. Chain termination mutants not listed above: Catania, Rugby Park, and Venezia. Diagonal lines on either side of a codon are used to separate bases from a preceding or following codon. This facilitates identification of CpG dinucleotides overlapping two codons.

’

164


TABLE IV Physical Meczsuremenls of Serum Albumin Value

Parameter

Source

Ref.

Diffusion coefficient &,,w Partial specific volume Sedimentation coefficient

6.0 x lo-' dcm2/sec 0.733cmS/g 4.5 x 10-13s

HSA HSA HSA

Oncley el al. (1947) Charlwood (1961) Oncley et al. (1947)

Intrinsic Viscosity Specific absorbance

0.042 (N), 0.16 (F) 0.51 (HSA), 0.667 (BSA) (liters g-l cm-')

HSA

Charlwood (1961) Peters (1975)

Molecular dimensions" Equilater triangle side depth Molecular volume" Molecular surface area Isoelectric point, pH -5.3 (defatted) Radius of gyration"

80 A 30 A 88248.9 A3 28202.1 A2 -4.7 (1-2 fatty acids)

26.7 A

Calculated from the atomic coordinates of human serum albumin (He and Ca;ter. 1992).

with a height of 60 A and a depth of 45 8, (Slayter, 1965). Stokes radii of BSA determined by diffusion data were approximately 37 8, (Longsworth, 1954). Perhaps the most interesting of these data are the lowresolution dark-field electron micrographs of the homologous human and bovine AFP (Fig. 3) (Luft and Lorscheider, 1983), which revealed AFP as a U-shaped molecule having dimensions of approximately 80 8,. However, at that time, any of its similarity in shape to albumin was dismissed because AFP, unlike albumin, lacked a pair of disulfide bridges at L6, which it was believed would allow the AFP molecule to fold in this Serum Albumin 041-k

4 - A

T 'Oi FIG. 2. Classical preception of the structure of serum albumin. Reproduced with permission from Peters (1985) and Academic Press.


A

165

B

FIG. 3. Digitized matrices of mass-scattered electrons from single-average molecular images of human a-fetoprotein (A) and bovine a-fetoprotein (B) with respective contoured mass maps (C and D). Reproduced with permission from Luft and Lorsheider (1983). OCopyright 1983 American Chemical Society.

manner. Other interesting conflicting data were presented by Hagag and colleagues ( 1983).They predicted nearly equal distancesbetween Cys-34, Trp-214, and Tyr-411 (-25 A) using a covalently attached fluorescent probe on Cys-34 and fluorescence energy transfer methods, and concluded that albumin must be folded more in accordance with a U-shaped model, hereafter referred to as “heart-shaped.” Subsequent studies on the N-B transition (see Section II,C,2,d) of HSA using ‘HNMR indi-

166


cated that an oblate elipsoid structure of HSA was unlikely; rather, it was proposed that HSA was folded into the heart-shaped structure similar to AFP (Bos et al., 1989). Overall, discrepancies with the cigar-shaped model have largely been dismissed in previous albumin reviews (Peters, 1985, 1992; Kragh-Hansen, 1990; Brown and Shockley, 1982).

C . Three-Dimensional Structure

I . Crystalliuztion Detailed structural knowledge of a protein molecule can only be precisely determined by X-ray crystallographic methods. A descriptive history of albumin crystallization is therefore presented, owing to its underlying importance to both the determination of the three-dimensional structure and to the bulk purification of albumin. Crystallized preparations of albumin were known as early as 1894 (Giirber, 1894), long before the advent of X-ray crystallography. Much later McMeekin (1939) gave a more detailed description of the preparation of the ESA crystals used by Giirber, showing that preparations of albumin with improved homogeneity and low carbohydrate content could be produced. These hexagonal crystals are shown in Fig. 4D (see color insert). Today, the majority of the commercial albumin preparations owe their origin to the fractionation methods developed by Cohn et al. (1947). These methods were aimed at producing stable human blood substitutes for battlefield applications using BSA purified by cold ethanol fractionation. Crystals of human serum albumin could also be produced by this method by the addition of cofactors such as decanol o r caprylic acid. Expounding on the same techniques of Cohn, Lewin (195 1) published an extensive list of derivatized human and bovine albumin crystals. Similarly, although originally noted in 1947, Hughes and Dinitz (1964) reported details of the crystallization of a mercury dimer of HSA and discussed this technique as a method for producing mercaptalbumin. Selected forms of human albumin were first studied by Low and Weichel (195 l ) , followed by more detailed studies using X-ray diffraction methods (Low and Richards, 1954). Their work primarily focused on the hydration of protein crystals and determination of various protein properties such as molecular dimensions and weight. Although these studies were conducted prior to the first successful structure determinations for proteins (Perutz et al., 1960; Kendrew et al., 1960), they included some of the first observations that protein crystals contain large percentages of aqueous solvent. Modern crystallographic studies of albumins were begun in the early 1970s and resulted in the preliminary characterization of several additional crystal forms, including the ESA crystals


167

originally reported by Giirber, the HSA crystals of Kendall (Kendall, 1941; McClure and Craven, 1974), and one additional new crystal form of HSA (Rao et al., 1976). These attempts to provide structural information were unsuccessful, presumably due to the lack of reproducibility or other undesirable properties of the crystals (Peters, 1985). Progress was no doubt further impeded by the available technology of the time. It is presumed that many of the crystal forms that can be produced from various albumin preparations have gone unreported because of the very poor diffraction qualities of these crystals. For example, ESA crystals grown from solutions of polyethylene glycol, although quite large and optically beautiful, exhibit very poor diffraction quality (Fig. 4L,see color insert). In 1989 we reported a new crystal form of human serum albumin that could be grown reproducibly and that eventually proved suitable for structure determination (Carter et al., 1989). These crystals (Fig. 4A, see color insert) are unusual in possessing large, continuous solvent channels that have cross-sectional dimensions of approximately 100 A and an overall solvent content of 78% (Fig. 5). A markedly better crystal form was later produced from rHSA (D. C. Carter, B. Chang, K. Keeling, Z. Krishnasami, and J. X. Ho, unpublished results; He and Carter, 1992) and subsequently also by using wild-type HSA. Currently, work in our laboratory has produced several new crystal forms and the observation of improved quality for crystals grown in the microgravity environment (Miller et al., 1992). Indeed, HSA was among the very first proteins crystallized in space (DeLucas et al., 1989). Several additional high-quality crystal forms are now grown routinely in our laboratory for structural studies (Fig. 4, see color insert). These crystals include canine serum albumin (CSA) (Fig. 4C and E), which exhibits diffraction to 2.0 A, and a crystal produced from turkey albumin (TSA) (Fig. 4J), which diffracts to 1.7 A. In addition, several diffraction-quality crystal forms of HSA and canine serum albumin (CSA) containing homogeneous preparations of various long-chain fatty acids have now been realized. Crystallographic data have been collected on each of the crystal forms shown in Fig. 4. Most of these crystal structures have now been solved by the molecular replacement method (Rossman and Blow, 1962) and are in the refinement stage. 2. X-Ray Structure of Albumin A discovery is like falling in love and reaching the top of a mountain after a hard climb all in one, an ecstasy induced not by drugs but by the revelation of a face of nature that no one has seen before and often turns out to be more subtle and wonderful than anyone had imagined.

Max F. Perutz (1989), in “Is Science Necessary”

168


FIG.5. The arrangement of the electron density in a tetragonal crystal of human serum albumin. Prominent features of the molecular packing arrangement are large (90 x 90 A) solvent channels (shown in white) that pass through the crystal parallel to the crystallographic c axis. The unit cell and symmetry operations parallel to the c axis are illustrated. Reproduced with permission from Carter et al. (1989); 0 American Association for the Advancement of Science (AAAS).

Although the structure determination of albumin was far from the thrilling experience shared by Max Perutz and John Kendrew when observing for the first time the atomic configuration of a protein molecule so eloquently described above by Perutz (Perutz, 1989), the albumin structure was quite different both in outline and detail, and clearly “more subtle and wonderful than anyone had imagined.” It is of interest to note that the first glimpses of albumin structure (Fig. 6, see color insert) marked exactly 150 years of albumin research. To date there have been several albumin structures determined by crystallographic methods in our laboratory, including human wild-type (HSA), recombinant human (rHSA), equine, and canine albumins. Currently, the most detailed infor-


169

mation is available for human and equine serum albumins, therefore much of the following discussion will be limited to these. Rather than the prolate ellipsoid predicted by so many studies, as discussed previously (Section II,B), the crystal structure of albumin reveals a heart-shaped molecule that can be approximated to an equilateral triangle with sides of -80 8, and a depth of -30 8, (Fig.7, see color insert). This observation is virtually identical with the low-resolution dark-field micrographs of the 39% homologous human and bovine a-fetoprotein (Fig. 3) (Luft and Lorschieder, 1983), and is in agreement with the spectroscopically determined distances between Cys-34, Trp214, and Tyr-411 (Hagag et al., 1983). Thus albumin, under neutral conditions of pH, has an axial ratio of approximately 2.66, which agrees well with the value of 3.0 based on observations of dielectric and birefringence relation times (Moser et al., 1966). The solvent-accessible surface and molecular volume of HSA are 28,202 A' and 88,249 AS,respectively, based on the calculations using the molecular coordinates and the algorithm of Richards (1985). Similarly, the radius of gyration is 26.7 A, which compares well with the hydrodynamic radius of HSA of 26.4 8, (rotational) as measured by light scattering and electron spin resonance (ESR) (Cannistraro and Sacchetti, 1986). As proposed by spectroscopic (Jacobsen, 1972; Sjoholm and Ljunstedt, 1973; Chen and Lord, 1976) and predictive methods (McLachlan and Walker, 1977), the albumin structure is predominantly a-helical. Approximately, 67% of HSA is helical, with the remaining polypeptides occurring in turns and extended or flexible regions between subdomains (Fig. 7). The overall agreement between the percentages of observed and predicted secondary structure is quite remarkable. For example, Pearson (1990) predicted a 65% ahelix content for serum albumin based on sequence comparisons of several albumins and further estimated a P-sheet and p-turn percentages of 10% and 19%,respectively. Although there is no /3 sheet in the structure of serum albumin, -23% of the structure is in an extended chain conformation, which would be predicted as p strand, and -10% of the remaining structure exists in turns. This adds further supporting evidence for the greater accuracy of secondary structure predictions that utilize reliably aligned multiple sequence data (Zvelebil et al., 1987). a. Domain Structure. As noted in the discussion of the primary structure of serum albumin, the three-dimensional configuration is composed of three homologous domains (I, 11, and HI). Each domain in turn is the product of two subdomains (IA, IB, etc.), which are predominantly helical and extensively cross-linked by several disulfide bridges (Fig. 8,

170


see color insert). Furthermore, the six subdomains share a common helical motif (Fig. 8). This motif primarily corresponds to the amino acids encompassed within the double disulfide loops 1, 3, 4 , 6 , 7, and 9. Each motif is related by a pseudo twofold axis (I: 168", 11: 163", 111: 171") shown in Fig. 8. These data are consistent with the theory that the 190-amino acid protoalbumin was the product of an early globin gene fusion event that predates the occurrence of LSA. The association of protein domains by pseudo twofold symmetry has been observed in the crystal structures of many proteins that are products of tandem gene fusion. When the a-carbons of the individual domains are superimposed (HSA), they give an average root-mean-square (RMS) difference between a-carbons of 3.77,4.32, and 3.63 8, for domains 1-11, 1-111, and 11-111, respectively. Cross-species comparison between HSA and ESA gives an average RMS difference in a-carbons of 2.01 8, for the entire molecule, which indicates the highly conserved nature of the three-dimensional structure. Ideally, one can divide each of the domains into 10 helical segments, hl-h6 for subdomain A and h7-hlO for subdomain B. The helical nomenclature presented in Fig. 8 provides an important framework for subsequent comparisons between the homologous domains, related structures, and discussions involving con formational change. Although all subdomains share a common four-helix motif, there are distinct differences. T h e A subdomains supplement the three-helix bundle on the C-terminal side with an additional but smaller disulfide double loop (loops 2, 5 , and 8) to form a small globinlike structure that is extensively cross-linked by four disulfide bridges (Fig. 9A, see color insert). The B subdomains supplement the helical motif on the Nterminal side with a conformationally extended polypeptide to create a folding topology that closely resembles a simple up-down helical bundle (Fig. 9B). This section of polypeptide represents the evolutionary loss of homologous helix h 1, although a helical remnant remains as part of the disulfide bridge. The A and B subdomains assemble through hydrophobic helix packing interactions primarily involving h2, h3, and h8. In addition, the subdomains are linked together by a presumed flexible extension of polypeptide encompassing residues Lys- 106 to Glu- 119, Glu-292 to Val-315, and Glu-492 to Ala-511 in the three domains, I, 11, and 111, respectively. These linkages do not appear crucial for the conformational stability at neutral pH and show the greatest differences in conformation when comparing related three-dimensional structures. Domains I and I1 and domains I1 and 111 in turn are connected through helical extensions of hlO(1)-hl(I1) and hlO(I1)-hl(III), creating the two longest helices in HSA. Consequently, the actual number of helices in the structure is 28 rather than 30. The positional occurrence of these helices


171

in the primary structure of HSA is presented in Table 11. Because this helical extension restricts the potential packing arrangement between domains, they do not associate by simple twofold rotation. Instead, quite surprisingly, they appear to be related by a pseudo twofold screw-axis symmetry with a rotation angle of 165" for 1-11 and 167" for 11-111. b. Nature of DzSulfides in Albumin. Disulfide pairings in albumin occur as predicted by Brown (1975) based on amino acid sequences of proteolytic fragments and steric considerations. In further agreement with Raman spectroscopic studies (Akoi et al., 1973), the conformations of the disulfides are primarily gauche-gauche-gauche and C/~~-SI-SP-CPO, with torsion angles clustering around +80". The disulfide pairings in HSA are located almost exclusively between helical segments. Such disulfide pairings are rarely observed in protein structure and usually involve isolated occurrences, e.g., crambin (Teeter and Hendrickson, 1979). Albumin is the first observed case wherein an entire protein folding topology is based on this theme. Given this interesting feature, one is better able to appreciate the stability of albumin under a variety of experimentally harsh conditions. For example, albumin can be heated to 60°C in the presence of caprylic acid, for 10 hr without deleterious effects (Shrake et al., 1984). The helical motif of albumin undoubtedly provides a natural framework that restricts mixed disulfide pairing. Katchalski et al. (1957) concluded that the disulfides in albumin were protected at neutral pH from reducing agents. This is also apparent in the structure, which shows that the majority of disulfides are well protected and most are not readily accessible to the solvent. It is well known that blocking of the free sulfhydryl, Cys-34, with iodoacetamide, cysteine, or glutathione prevents the occurrence of mixed disulfides in aged albumin, as well as the occurrence of the albumin dimer (Peters, 1985). In the structure, Cys-34 is not in close proximity to any disulfide, consequently, it is difficult to imagine its intramolecular participation without substantial conformational change of the albumin molecule. However, it could perhaps participate in the formation of mixed disulfides by catalyzing the reaction via intermolecular interaction, because Cys-34 is known to have an unusually low p K s ~ (Pedersen and Jacobsen, 1980) (see Section I K C , 1). Surface Charge Distribution. As a highly soluble protein that can be prepared at concentrations as high as 30% (w/v),albumin is ideally suited for its role as the major plasma protein. This amazing property is no doubt related to its high negative charge at neutral pH. In this regard, albumin is known to have an interesting asymmetric charge distribution c.

172


within the primary structure. At neutral pH, Peters (1985) calculated a net charge of - 10, -8, and 0 for domains I, 11, and 111 of HSA, respectively, and similar distributions are present in other albumins. The surface charge distribution of human serum albumin is shown in Fig. 10 (see color insert). Unlike the asymmetric charge distribution expected based on the primary structure, the distribution seems fairly uniform. There does not appear to be any significant pattern of basic versus acidic residues, except for a group of invariant basic amino acids on the surface of domain I (see Section IV). There are, however, distinct areas of amino acids with neutral charge that could potentially be important for interactions with long-chain fatty acids. There have been 43 point mutations identified for human serum albumin, as shown in Table 111. These variants were usually discovered because of their anomalous electrophoretic migration (see, for example, Tarnoky, 1980). The specific point mutations have been determined primarily through the efforts of F. W. Putnam and co-workers. With the exception of the point mutation Ala-320 to Thr, which is partially internalized in the structure to pack with other lipophilic residues, all of the remaining substitutions in Table 111 are located on the surface of HSA and are exposed to solvent.

d. Conformational Flexibility. The ability of albumin to undergo a major reversible conformational isomerization with changes in pH was first demonstrated by Luetscher in 1939. Foster, noting these early studies involving electrophoretic heterogeneity, reinvestigated this fascinating phenomenon in much greater detail (Foster, 1960, 1977). The only other known major conformational changes that occur in albumin are induced by the interaction of albumin with fatty acids (Peters, 1985). Foster classified the pH-dependent forms as “N,” for normal form, which is predominant at neutral pH; “B,” for the basic form occurring above pH 8.0; “F,” for fast migrating form produced abruptly at pH values less than 4.0; “E,” for expanded form at pH less than 3.5; and “A,” for aged form occurring with time at pH values greater than 8.0. Perhaps the most interesting of these isomers are the B and F pH transitions, and fatty acid-induced, conformational isomers. i. N-F transition. As early as 1960, Foster clearly recognized that the formation of the F form involved an abrupt opening of the molecule. Much later Geisow and Beaven (1977) proposed that the N-F transition involved the unfolding of domain 111 from the rest of the molecule, and this was later verified by Khan (1986) using proteolytic fragments of BSA encompassing residues 377-582, the so-called “T-A fragment” (Peters and Feldhoff, 1975). It had been previously shown that no conforma-


173

tional changes occur during the N-F transition for fragments containing domains I and I1 (Khan and Salahuddin, 1984). Recent experiments in our laboratory using proteolytic fragments of BSA and turkey serum albumin (TSA) confirm the conclusions of previous studies and further illuminate the nature or “structure” of the N-F transition. Using limited peptic cleavage at residue 306 to produce the two fragments, A and B, which constitute the two halves of albumin (King, 1973),we undertook to test our hypothesis that during the N-F transition the albumin molecule expands by the dissociation of the C-terminal half, or “tail,” from the “head” of albumin previously described in Section II,C,2. Figure 11 illustrates the retention times on fast protein liquid chromatography (FPLC)columns of pepsin-treated BSA (BSAp) and TSA (TSAp) under a variety of pH conditions. As can be seen, BSAp elutes with a retention time identical to that of BSA until pH 4.0, where both TSA and BSA abruptly split into two fragments of equal size. This coincides exactly with the pH known to induce the F conformational state of albumin (D. C. Carter, P. D. Twigg, and K. Keeling, 1993). Furthermore, as demonstrated earlier by King (1973), these fragments recombine when the pH is returned to neutral. These findings also provide additional supporting evidence for the predominance of the heart-shaped conformation in solution in the pH range from 4.5 to 8.0, because the fragments never dissociate during column chromatography at pH greater than 4.0. Additionally, it is interesting to note that the F transition is conserved even in distantly related species such as birds (Fig. 11) (Feldhoff and Ledden, 1983).T h e change in hydrodynamic ratio indicating an increase in volume for BSA of 1 1% is consistent with this observation (Leob and Scheraga, 1956), as are the dimensions of roughly 40 X 129 8, predicted by Victor Bloomfield (1966) for BSA at low pH (3.5) deduced from low-angle X-ray scattering experiments. The F form is further characterized by a dramatic increase in viscosity, much lower solubility, and a significant loss in helical content (Foster, 1960). T h e proposed structures of the F and E conformational isomers are illustrated in Fig. 12. Structurally, the interface between the two halves of the molecule is held together by both hydrophobic and salt bridge interactions. Among these residues there are potential salt bridges between Lys-205 and Glu465, Asp- 187 and both Lys-432 and Arg-52 1, Arg-2 18 and Asp-45 1, and Lys-190 and Glu-425, which should be more clearly defined in a higher resolution crystal structure. Hydrophobic interactions that associated IA, IB, and IIA, with IIB, IIIA, and IIIB include a major interdomain cluster involving Phe-206, Leu-48 1, Ala-482, Trp-2 14, Leu-347, Val-344, Val-343, Leu-33 1, Ala-2 17, and Tyr-452.

174


tSA

BSA Elution

1

Vdunn 13.34 ml

2

14.65 ml

IL,

uutkn

-vdunw, 1

13.10ml

2

14.6 ml

1( 13.15ml

1

12.95 ml

2

2

14.32 mi

14.38ml

mlml 0 10 20 30 Elution Volume, mls

1

13.01 ml

2

14.24ml

1

21.03ml

1

20.7ml

2

23.68ml

2

23.59ml

0 1 0 2 0 3 0 Elution Volume, ml

FIG.11. Chart of retention times for bovine serum albumin (BSA) and turkey serum albumin (TSA), which have been proteolitically treated with pepsin. For pH values 7.0-5.0, peaks 1 and 2 represent the albumin dimer and the proteolitically nicked monomer, respectively, at different pHs (N form). At pH 4.0 there is an abrupt dissociation of the two halves of albumin to yield peaks 1 and 2, which represent the intact monomer (incomplete digestion, F form) and the two equal molecular weight halves of albumin, respectively. (D. C. Carter, P. D. Twigg, and K. Keeling, Gnpublished results).

It is interesting to speculate whether the F conformation is of physiological significance given the lower pH measured on the membrane surfaces of several tissues (Wilting et al., 1982).Additionally, it is possible that this is the form that is bound to the surfaces of highly oriented pyrolytic graphite as revealed by a scanning tunneling microscopy study


175

FIG. 12. Ribbon diagrams of serum albumin in its N form, and in its proposed F and E forms. Figure drawn using program RIBBONS (Carson, 1987).

176


by Feng et al. (1988), although it was assumed in their paper to be the normal conformation. Perhaps this conformation can also be induced by the surface binding process. T h e conservation of the N-F transition and the p H of the transition among various diverse species clearly suggest a physiological role for this transformation. One could speculate that the affinity for ligands is reduced in the F conformation, perhaps facilitating ligand off-loading at various tissue interfaces. T h e observation that the sulfhydryl becomes more solvent exposed under both N-F and N-B transitions would also be consistent with this supposition. ii. F-E transition.At pH values lower than 4.0 albumin undergoes another expansion, and electron microscopy reveals the molecule as a series of balls and strings with approximate dimensions of 21 x 250 8, (Harrington et al., 1956). It seems quite reasonable to expect that this isomeric form corresponds to the sequential subdomain-subdomain dissociation concomitant with the loss of the intradomain helices h lO(1)hl(I1) and hlO(I1)-hl(II1) (Fig. 12). iii. N-B transition.At pH 9.0 albumin undergoes another conformational change, albeit a less dramatic one, the basic form (B). This form was originally identified by its slower anodic migration during electrophoresis. Structurally, the conformational change occurs more slowly than the F transition and there is evidence that discrete steps are associated with this transition (Hart et al., 1986). There is a decrease in helical content and increase in affinity for selected ligands (Zurakowski and Foster, 1974). At the present time, little structural information is known about this isomer. 111. NATURE OF LIGAND BINDING

Th e most prolific area of albumin research has involved binding studies. An appreciation of the extraordinary binding chemistry of albumin can be obtained by examining a small subset of the binding studies performed to date (Table V). Most ligands are bound reversibly, and typical association constants (K,)range from lo4 to lo6 M - ' . Because of the incredible diversity of ligands bound by albumin, early researchers saw ligand binding to serum albumin as nonspecific in nature and did not recognize that there were discrete sites per se. Instead they envisaged the ligands as randomly attached to the surface, somewhat like a sponge. This rather uninteresting view of albumin has changed over the past years, and now it is generally recognized that there are a small number of distinct binding locations. These binding sites have been studied primarily by equilibrium dialysis or spectroscopic methods. Data from such studies are generally analyzed by the Scatchard method (1949) and this

Color Plates

FIG.4. Crystals of serum albumin. (A) Tetragonal crystals of HSA, (B) monoclinic crystals of baboon serum albumin, (C) triclinic crystals of CSA, (D) hexagonal crystals of ESA, (E) monoclinic crystals of rHSA complexed with lauric acid, (F) monoclinic crystals of HSA complexed with cysteine, (C) monoclinic crystals of rHSA, form 2, ( H ) crystals of feline serum orthorhombic crystal of TSA, (K) albumin, (I) monoclinic crystals of rHSA, form 1, tetragonal crystals of BSA, and (L) hexagonal crystals of ESA.

u)

FIG.6. Section of the 6.0-A electron density of subdomain IIIB. Helical rods of density 8.0 to 10 A in diameter, indicative of the a-helical structure of serum albumin, were the dominant features of the electron density. Reproduced with permission from Carter el al. (1989) and the American Association for the Advancement of Science (AAAS). FIG.7. Stereo view of human serum albumin illustrating the overall topology and secondary structure. The positions of the 17 disulfides and the side chain of Cys-34 are shown in red. Structurally, HSA consists of 28 helices, which range in size from 5 to 31 amino acids in length and which can be grouped into 10 principal helices within each domain. The positions of the two major binding sites of HSA, located within subdomains IIA and IIIA, are illustrated with the ligand 2,3,5triiodobenzoic acid, shown in white. Figure drawn using program RIBBONS (Carson, 1987). FIG.8. Cylindrical representation of a typical domain (11) structure. Note that helices h l , h2, h3, and h4 of subdomain A are related to helices h7, h8, h9, and h10 of subdomain B by a pseudodiad (pointing toward the reader). Figure drawn using program RIBBONS (Carson, 1987). FIG.9. Stereo view of the IIA (A) and IIB (B) subdomains. Figure drawn using program RIBBONS (Carson, 1987). FIG.10. Space-filling model of serum albumin molecule with basic residues colored in blue, acidic residues in red, and neutral ones in yellow. (A) Front view, (B) back view, (C) left side, and (D) right side. Figure drawn using program RIBBONS (Carson, 1987).

FIG.13. Stereo view showing the homology between the triiodobenzoic acid-binding sites within subdomain IIA (A) and subdomain IIIA (B); the subdomains have been superimposed by the method of least squares. The protein components are shown in orange and green for ESA and HSA, respectively, and the positions of bound triiodobenzoic acid are shown in red and yellow for ESA and HSA, respectively. It should be noted that many of the side-chain conformations illustrated in Fig. 6 can be determined with confidence only at higher resolution. In addition, the complex has not been refined. Reproduced with permission from Ho el al. (1993). FIG.14. Dotted surface diagram showing the major binding pocket inside subdomain IIIA (yellow ribbons). Figure drawn using program RIBBONS (Carson, 1987). Fiti. 15. Electron density of bound 2,3,5-triiodobenzoic acid (TIB) at 3.0 A produced from (FI--FN)a,where F,and F, refer to measured diffraction data from the TIB cocrystal complex and native data, respectively, and the phases, a,are from the native model. Atoms: red, oxygen; yellow, carbon; orange, iodine. Yellow and blue contour levels at 2.0 ci and 4.0 0, respectively. Reproduced with permission from Ho el aL (1993). FIG.16. Conformational difference between native rHSA (cyan) and rHSA bound with fatty acid-lauric acid (yellow). Note that the largest displacement occurs at the two terminus subdomains, i.e., subdomains IA and IIIB. Figure drawn using program RIBBONS (Carson, 1987;J. X. Ho, D. Carter el al., unpublished results).

FIG 17. Stereo ball-stick model of serum albumin structure at the region around residue Cys-34. Red, oxygen; yellow, carbon; blue, nitrogen; green, sulfur. Figure drawn using program Turbo FRODO (Cambillaux and Horjales, 1987). FIG.19. Ribbon diagram showing helices h l , h2, h3, and h4 of subdomain IIIA of rHSA (cyan) superimposed on helices E, F, H, and G of myoglobin (yellow). Note that the polarity of h3 and h4 is opposite to that of G and H, although the helices are packed similarly; N, the two N termini. Figure drawn using program RIBBONS (Carson, 1987).

FIG.6

FIG.7

FIG.8

\

FIG.9A

FIG.9B

.,

FIG. 10A (top) & B (bottom!

FIG.1OC (top) 8c D (bottom)

FIG.13A

FIG.13B

FIG.14

FIG.15

FIG.16

FIG.17

FIG.19

TABLE V Seleckd Binding Conrtuntsfor Endogenous and Exogenous Ligandr with Serum Albumina Ligand

K , , W-')

K,, Of-')

Albumid

Endogenous Substances Aldosterone Arachidonate Bilirubin Bilirubin Bilirubin Chenodeoxycholate Cholate Cholate Cortisol Estradiol Hemin Hemin Hematin Linoleate Linoleate Lithocholate Lysolecithin Oleate Oleate Oleate Oleate Oleate Palmitate Palmitate Progesterone Prostaglandin El

3.2 x 3x 1.4 X 5.5 x 5x 2x 3.2 x 5.5 x 5.0 x 1.0 x 5.0 x 1.1 x 1.1 x 1.3 x 7.9 x 9x 4.3 x 1.1 x 26.0 x 9.4 x 2.9 x 2.1 x 6.0 x 6.2 x 3.6 x 7x

105 107 10' 107 107 105 104 104 103 105 107 108 108 107 107 107 104 108 107 107 107 107 107 107 105 104

Ref. ~

~~

5 x 105 4.4 x 106 2 x 103

-

-

2.5 X lo6 -

4.0

X

-

lo6

3.0 X lo6 6 X lo3 -

HSAf HSA HSAf HSAd HSA HSA HSA~ HSA HSA~ HSAf HSA~ HSA~ HSA HSAd HSA HSA BSAf HSAd HSA HSA HSA HSA HSAd HSA HSAd HSA

~

Richardson et al. (1977) Saw et al. (1981) Jacobsen (1969) Brodersen (1979) Jacobsen (1977) Roda et al. (1982) Burke etal. (1971) Roda et al. (1982) Yates and Urguhart (1962) Daughaday (1959) Beaven et al. (1974) Adams and Berman (1980) Adam and Berman (1980) Goodman (1958) Spector and Fletcher (1978) Roda et al. (1982) Klopfenstein (1969a,b) Goodman (1958) Spector and Fletcher (1978) Spector and Fletcher (1978) Spector and Fletcher (1978) Spector and Fletcher (1978) Goodman (1958) Spector and Fletcher (1978) Westphal and Harding (1973) Unger (1972) (continued)

TABLE V (continued)

K,, W-')

Ka, w-')

Albuminb

Stearate Taurocholate Testosterone

15.0 x 107 1.2 x 104 2.38 x 104

-

HSA BSAd HSA~

Testosterone L-Thyroxine L-Tryptophan

4.2 x 104 1.6 X lo6 1.6 x 104

Ligand

=;

6.3 x 104 L-Tryptophan Negatively charged and electrostatic neutral drugs 1.96 x 105 Acenocoumarin 8.0 x lo6 Camptothecin Chlorazepate 1.3 x 104 1.3 x 105 Chlorophenoxyisobutyrate 3.3 x 105 Chlorophenoxyisobutyrate 3.07 x 104 Chlorothiizide Chlorpropamide Cinchophen Clofibrate Dansylglycine Diazepam Dicoumarol Dicoumarol Dicoumarol Dicoumarol Digitoxin Digitoxin Furosemide

4.5 x 1.4 x 2.5 x 4.6 x 4.9 x 2.9 X 2.2 x 7.7 x 3.5 x 4.3 x 6.9 x 1.7 x

104 105 104 105 105 lo6 lo6 105 105 104 104 105

6

X

-

lo4

6.0 x 103 -

1.5 x 103 3.9 x 103 1.7 x 10'

-

4.7 x 102 -

1.8 x 105 1.3 x 104 -

9.6 X lo3

HSA HSAd HSAf HSA HSA~ HSA~ HSAf HSA~ HSA~ HSAf

HSA~ HSAf HSAf HSAf HSAf HSAf HSA~ HSAf HSA~ HSAf HSAf HSAf

Ref. Spector and Fletcher (1978) Green et al. (1971) Vermeulen and Verdonck ( 1968) Pearlman and Crepy (1967) Steiner et al. (1966) McMenamy and Oncley ( 1958) Sollene et al. (1981) Tillement et al. (1974) Chignell(l973) Coassolo et al. (1978) Spector et al. (1973) Tillement et ~ l(1974) . Breckenndge and Rosen (1971) Crooks and Brown (1974) Mudge et al. (1978b) Nazareth et ~ l (1974) . Chignell(l969) Miiller and Wollert (1973) Perrin et ~ l (1975) . Garten and Wosilait (1972) Chignell(l970) Choetal. (1971) Brock (1975) Lukas and De Martino (1969) Sebille et al. (1978)

c (

4 (D

Fusidic acid Glibenclamide Halofenate Ibuprofen Indomethacin Indomethacin Iopanoate Iopanoate Iophenoxate Novobiocin Phenylbutazone Phenylbutazone Phenylbutazone Phen ylbutazone Salicylate Salicylate Salicylate Sulfaethidole Sulfaphenazole Tolazamide Tolbutamide Warfarin Warfarin Warfarin Warfarin Positively charged drugs Chlorpromazine Chlorpromazine Desipramine Imipramine

7.8 x 7.7 x 1.6 x 2.73 X 1.0 x

104 105 105

lo6 106

3.0 x 105

6.7 X 7x 7.7 x 5.5 x 2.37 X 1 x 2.5 x 2.3 x 7.1 x 2.2 x 1.3 x 1.5 x 9.2 x 8.7 x 2.2 x

lo6 105 107

105

1V 105 105 105 104 105 105 105 104 104 105

2.5 x 105 8.9 x 104 1.5 x 105 2.3 x 1P 4.2 x 104 1.9 x 105 7.02 x 104 2.39 x 104

Guttler et al. (1971) Crooks and Brown (1974) Spector et al. (1973) Whitlam et al. (1979) Hultmark et al. (1975) Mason and McQueen (1974) Mudge et al. (1978a) Lang and Lasser (1967) Mudge et al. (1978a) Brand and Toribara (1975) Rosen (1970) Chignell(l969) Brown and Crooks (1976) Tillement et al. ( 1974) Keresztes-Nagy et al. (1972) Brown and Crooks (1976) Hultmark et al. (1975) Janssen and Nelen (1979) Brown and Crooks (1976) Crooks and Brown (1974) Brown and Crooks (1976). Crooks and Brown (1974) Sudlow et al. (1975) Garten and Wosilait (1972) Brown and Crooks (1976) Tillement et al. (1974)

7.0 x 103 1.95 x 104 1.0 x 105

1.4 x 1.5 X 9x 3.8 x 3.7 x 4.56 x 4x 1.3 x 5.6 x 3.3 x 1.6 x 2.9 x 1.6 X

104

lo6 102

105 104 104 104 103 103 103 103 103

los

1.1 x 103 1.5 x 105

1.7 X

lo2

1.1 x 104 6.7 x 105 1.5 x 103 5.9 x 105

HSA~

Gabay and Huang (1974), Huang and Gabay (1974) Sharples (1975) Sharples (1975) Shardes (1975)

(continued)

TABLE V (continued)

Ligand Lidocaine Mepivacaine Pamaquine Procaine Promazine Quinidine Quinine Tnflupromazine Inorganic ions

c11-

SCNZn2+ Cd2+ Mn2+ cu2+ cu2+ Ni2+ co2+ Ca2 Ca2+ Ca2+ +

Mi?+ a

K,, W - ' ) 1.3 x 2.5 x 6.4 x 3.1 x 8.5 x 1.4 x 7.5 x 5.5 x

105 105 107 103 104 103 103

7.2 X 6.15 x 3.35 x 5.7 x 1.3 x 2.4 x 9x 1.6 X 3x 6.5 x

lo2

104

103 104

lo2 103 104 106 10l6

105 103

1o2 9 x 102 8.3 X 10' 1o2

Adapted and modified from Kragh-Hansen (1981).

f, Serum albumin with fatty acid; d, defatted serum albumin.

Ka*

of-')

-

-

-

6.1 x 10' 6.7 X 10' 7.8 x lo2 5.0 x 10'

1.6 x lo2

-

-

Albuminb

Ref.

HSAf HSA~ BSAf HSAf HSA~ HSAf HSAf HSAd

Sawinski and Rapp (1963) Sawinski and Rapp (1963) Naik et al. (1975) Sawinski and Rapp (1963) Sharples (1975) Nilsen and Jacobsen (1976) Paubel and Niviere (1974) Huang and Gabay (1974)

HSAI HSAf HSAf HSAf BSA~ HSA~ BSAf HSA HSA~ HSAf HSAI HSA HSA HSAf

Scatchard and Yap (1964) Scatchard and Yap (1964) Scatchard and Yap (1964) Waldmann-Meyer (1960) Waldmann-Meyer (1960) Nandedkar el al. (1973) Peters (1975) Lau, el al. (1974) Callan and Sunderman (1973) Nandedkar et al. (1972) Pedersen (1971) Fogh-Andersen (1977) Fogh-Andersen (1977) Pedersen (1971, 1972)


181

field of research has been well reviewed by Lindup (1987). Although controversy remains about the exact number of discrete binding locations on albumin, the general consensus is as follows: there are two principal binding areas for small heterocyclic or aromatic carboxylic acids; there are at least two to three dominant long-chain fatty acid-binding sites unique and separate from the binding sites for small anionic compounds (at normal physiological concentrations); and there are two distinct metal-binding sites, one involving Cys-34 and the other the N terminus. Thus, with normal ligand/albumin concentrations, there are six dominan: lreas of ligand association to albumin. New perspectives on several of these binding sites were gained from the crystal structures of albumin. The sites identified for the small heterocyclic and aromatic carboxylic acids were found to reside within specialized cavities in subdomains IIA and IIIA. These sites are consistent with Site I and Site 11 proposed by Sudlow et al. (1976). T h e two metalbinding sites have been previously identified by various spectroscopic and chemical means. The location of bound Cu(I1) and Ni(I1) involves the first three residues of the N terminus, with a His at position 3 playing a critical role (Peters and Blumenstock, 1967; Lau et al., 1974). T h e second metal-binding site involves the free sulfhydryl of serum albumin, Cys-34. Here, the amino acid cysteine and peptide glutathione are covalently bound, as well as various metals such as Cd, Au, Hg, and Ag. The locations of the two long-chain fatty acid sites remain more obscure, although Brown and Shockley (1982) have proposed that long-chain fatty acids occupy separate sites on the B subdomains. Crystallographic analyses of a complement of fatty acid analogs bound to albumin are in progress and answers to these questions should be forthcoming. The bilirubin site is often considered distinct from the other ligands. However, we believe that bilirubin is primarily bound to the site within IIA, and this does not contradict the experimental data in the literature. The following segments of this section describe in detail the current understanding of each of the six binding sites. A. Small Organic Compounds: Sites I and II

The vast majority of ligands in Table V are bound in one o r both sites within specialized cavities of subdomains IIA and IIIA. At this time the binding locations of several key compounds, historically used as markers in drug or ligand displacement interactions, have been determined at various resolutions (Table VI). Clearly, from Table VI it can be seen that IIIA appears to possess the primary binding activity for albumin whereas IIA is more specialized.

182


TABLE V1 Locations of Ligand Binding to HSA" Data set

Ligand

D

N

Rf

Observed location

HSAAlA HMRWN 1A HSAV 1A HDGXlA HCHLOlA HMRIB 1A HAZTlA HTIBlA HMRIS4A HDlSlA

Aspirin Warfarin Diazapam Digitoxin Chlofibrate Ibuprofen AZT TIB 1s DIS

4.0 5.0 6.8 5.0 6.0 6.0 4.0 4.0 4.0 4.0

7362 2555 2075 375 1 2175 2402 7548 543 1 6334 4734

0.11 0.167 0.1 18 0.137 0.138 0.215 0.124 0.12 0.19 0.20

IIA,IIlA IIA IIIA IIIA IIIA IIIA llIA IlA,lIIA IIA,IIIA IlA,IIIA

a D,Resolution or d spacing in angstroms; N , number of unique reflections; R/, fractional R-factor between native and derivative data sets. To prepare ligand-HSA complexes, crystals of HSA were placed in a stabilizing solution of 45% polyethylene glycol 400 MW (pH 6.8-7.0), which contained approximately 0.5 to 5 mM of the compound of interest, and were allowed to stand for 24 to 72 hr. X-Ray diffraction data were collected on Siemens multiwire area detector as described by He and Carter (1992). Reprinted with permission from Nature (He and Carter, 1992); copyright 1992, Macmillan Magazines Limited.

Prior to the structure determination, solution studies by Sudlow et al. (1975) led to the classification of binding sites on HSA in two categories, which were denoted Site I and Site I1 from experiments based on molecular interactions with fluorescent probes. The binding locus of large heterocyclic compounds possessing a negative charge was identified as Site I, with Site I1 showing a preference for small aromatic carboxylic acids. Spectroscopic studies have confirmed the notion of two principal binding sites, and identified Tyr-411 with Site 11, the principal active binding region of HSA. Tyr-411 has also been identified with the esterase-like activity of HSA (Sollene and Means, 1979; Ozeki et al., 1980; Kurono et al., 1983). Based on competitive inhibition studies, Lys199 interacts with drugs of Sudlow's Site I (Sudlow et al., 1975). Additionally, in the same studies, fluorescence damping of Trp-2 14 on ligand binding has also associated this residue with Site I. Consequently, it is clear that Sudlow's Site I corresponds to the hydrophobic pocket in subdomain HA, and his Site I1 to the pocket within IIIA. This realization, together with the crystallographically determined major binding locations for several key markers (Table VI), can now be utilized to explain a wealth of ligand-binding data without ambiguity. Hence, we have adopted Sudlow's nomenclature and expanded it to include all six major ligand-binding sites.


183

The mystery associated with the diverse binding chemistry of albumin can be explained in simple terms. One can, from efficiency considerations, appreciate that nature has converged on relatively few binding sites with multifunctional properties. The amino acid residues that line the cavities are quite similar in charge distribution for both IIA and IIIA, but still impart specialized selectivity. In each of the two subdomains there is an asymmetric distribution leading to a hydrophobic surface on one side and a basic or positively charged surface on the other (Fig. 13, see color insert). This explains the discriminatory affinity of albumin for small anionic compounds. The van der Waals surface of the binding pocket in IIIA is shown in Fig. 14 (see color insert). The region is an elongated sock-shaped pocket wherein the foot region is primarily hydrophobic and the leg is primarily hydrophilic. The opening to the pocket is clearly accessible to the solvent. Albumins of several species possess interesting enzymatic properties. For example, Tyr-4 11 has been associated with weak esterase activity, reacting specifically and rapidly with p-nitrophenyl acetate and many other activated esters, including those of selected fatty acids (Koh and Means, 1979). Human serum albumin is the most reactive of the species studied thus far. In contrast, ESA exhibits little or no reactivity (Elkarim and Means, 1988). In the crystal structure of HSA, the reactive hydroxyl of Tyr-411, located in the IIIA binding pocket in close proximity to bound ligands, is -2.7 8, from the nitrogens of Arg-410, which may explain the unusual reactivity of this residue toward nucleophilic substitution. However, in the recently determined structure of ESA, no obvious explanation can be offered for its lack of esterase activity. Previously, this difference in activity was explained by the amino acid substitution of Tyr-411 by Leu (Chincarini and Brown, 1976), based on partial amino acid sequence determinations. However, the recently determined cDNA sequence of ESA clearly indicates that Tyr-411 is conserved (Ho et al., 1993).Perhaps the differences observed are due to polymorphism, which has now been demonstrated for several hooved species. Other chemical reactions of HSA associated with Site I and Site I1 involve the participation of Lys-199. This residue, which can be acetylated by aspirin (Walker, 1976), appears to be a site of nonenzymatic glycosylation of HSA (Day et al., 1979), and is considered to be the major site of conjugation to benzoylpenicillin groups (Yvon and Wal, 1988). Interestingly, the covalent adduct produced by the reaction of Lys-199 with penicillin is the principal antigenic determinant of the allergic reaction to this drug (Ahlstedt and Kristofferson, 1982). The reactivity of Lys-199 may be attributed to its unusually low pK, of 7.9 (Gerig and Reinheimer, 1975), which can be rationalized based on the close interaction of Lys- 199 with His-242.

184

DANIEL C. CARTER AND JOSEPH X. H O

As one might predict, there is enantiomorphic selectivity with regard to various chiral ligands. For example, albumins in general have a 100fold higher affinity for L-tryptophan (King and Spencer, 1970; Lagercrantz et d.,1981) and stereoselectivity is also known for many pharmaceuticals such as d-oxazepam hemisuccinate (Muller and Wollert, 1975). Much of the chemistry of albumin can be understood from detailed observations of the 2,3,5-triiodobenzoic acid (TIB) complexes with albumin. The crystalline complex of TIB with HSA and ESA has been determined with resolution sufficient to position the molecule within the binding pocket unambiguously and to identify the chemistry of interaction (Fig. 15, see color insert). This ligand has a moderate and equal affinity for IIA and IIIA in both HSA and ESA. Association constants were estimated by Scatchard analysis (Scatchard, 1949) using direct linear plots (Eisenthal and Cornish-Bowden, 1974) to be 2.2 x lo5 M-' and 8.3 X lo4 M - ' for HSA and ESA, respectively. Overall, ESA and HSA exhibit a similar but distinct binding chemistry with TIB (Fig. 13). However, it is the similarities, not the differences, that are striking. Th e major binding differences occur when comparing the two IIA subdomains. In IIA the carboxyl of ESA-bound TIB is oriented primarily toward Arg-257, but also shares this interaction with His-242 (not Lys-199 as in HSA). This is a result of a shift of TIB in ESA by a few angstroms toward helix h6(II) (Fig. 13A). T he binding of TIB in subdomain IIIA is quite similar for both ESA and HSA (Fig. 13B). Altogether, they share 18 out of 19 equivalent residues in close proximity to TIB in IIA and 15 out of 16 for IIIA (Table VII). When one considers only the residues in immediate contact with ligand (approximately 10 to 11 residues in each subdomain), 8 or more are strictly conserved in all mammalian albumins. Observations of the amino acid sequences of all albumins in Table I1 reveal that several of the residues are invariant in subdomain IIIA, whereas relatively few are conserved in IIA (Section IV). This agrees with the observed similarity in binding chemistry with TIB in subdomain IIIA of ESA and HSA. Knowledge of these residues should provide important information in guiding future studies of the underlying chemistry of albumin. T h e absence of a similar binding chemistry in analogous subdomain IA can be explained by the crystal structure. A nonhelical section of polypeptide, which includes Cys-62 of the first disulfide, allows helix h3 to markedly increase its packing angle with h4 (normally near O"), effectively eliminating the binding pocket within IA. This slight but significant difference in folding toplogy among domains can be readily seen in Fig. 7 (see color insert).

-

TABLE VII Amino Acids Involved in Licand Binding' 11A

K( 199)

Equine Human

Y(149) Y Y

Y Y

F(211) Y(V)W Yw

W(214)

L(234) Y(I) Y

L(238)

H(242)

R(257)

Equine Human

Y Y

Y Y

Y Y

Y Y

S(2 15) Y(A)w Yw

R(2 18) Y Yf

L(219) Y Y

R(222) N(K) Y

F(223)

L(260) Yw Yw

A(261) Yw Yw

l(264)

l(290)

A(291)

Y Y

Y Y

Y Y

F(395) Nw Yw

R(4 10)

Y(411)

L(430)

Y Y

Y Y

Y Y

Y Y E(292) Yw Yw

IIlA P(384)

L(387)

I(388)

Equine Human

Y Y

Y Y

Y Y

N(391) Y Y

C(392) Yw Yw

V(433) Y Y

C(438) Yw Yw

A(449) Y(S) Y

E(450)

L(453)

R(485)

Equine Human

Y Y

Y Y

Y Y

S(489) Y Y

Amino acids that are involved in binding in ESA and HSA are marked Y for yes, N for none. Boldface Y indicates that the amino acids are conserved in both ESA and HSA. Amino acids that are more remote but still form important sides of the binding region are designated w for wall and f for far, as additional identifiers. Reproduced with permission from Ho et al., 1993.

186


B . Long-Chain Fatty Acids: Sates III and IV Albumin has a well-known affinity for a variety of saturated and unsaturated fatty acids of varying chain lengths (Table V). It is recognized as the principal transport protein for fatty acids and other lipids that would otherwise be insoluble in the circulating plasma. The total fatty acid capacity of albumin varies with fatty acid chain length, but averages approximately six fatty acids per albumin molecule. Under normal physiological conditions, albumin carries one or two fatty acids (Peters, 1985). This seems to agree with a number of studies indicating that long-chain fatty acids bind to separate and distinct sites, compared to ligands that bind to Site I and Site I1 (Koh and Means, 1979; Sollene and Means, 1979). Reed (1986) and Parks et al. (1983) also find that there are two to three dominant long-chain fatty acid sites. A method has been developed by Brodersen et al. (1990) to compensate for the inherent solubility problems associated with long-chain fatty acids in aqueous media. In this method, which should have broad applicability, the fatty acids are transported from solutions of HSA containing fatty acids, across membranes to albumin solutions devoid of fatty acids. Conclusions from these experiments are in agreement with earlier classical studies and imply that there are two high-affinity long-chain fatty acid-binding sites and four with lesser affinity. Despite this consensus, one earlier study suggests that there are no major sites that dominate; rather, it is proposed that all fatty acids are distributed evenly over six to nine sites (Spector and Fletcher, 1978). Hamilton et al. (1991), using BSA and '%-enriched oleic acid, determined one primary site in domain IB and two additional primary sites in domain 111. Although little is known about the precise locations of these bound long-chain fatty acids, Reed (1986) asserts that three lysine residues are primarily associated with the carboxyls of the fatty acids, namely Lys-116, Lys-349, and Lys-473. On the other hand, glucosylation of albumin at Lys residues 525, 199, 281, and 439 greatly inhibits the binding of cis-parinaric acid (Shaklai et al., 1984; Iberg and Fliickiger, 1986) and Shockley and Brown (1980) identify residues His-145, Lys-220, Cys-34, Lys-412, and His-336 as reactants with N-danzylaziridine, which inhibits binding of stearate, palmitate, and oleate. Further experiments by Brown and Shockley (1982) revealed that stearate and palmitate, but not bilirubin or octanoate, can inhibit the reaction with His-145. Hence, there is no clear consensus within the current scientific literature on the exact location of bound long-chain fatty acids, except that their locations do not correspond to Site I and Site 11. Additionally, there appears to be general agreement that the major sites reside in domains I and 111 (Peters, 1985), possibly in the B subdomains (Brown and Shockley, 1982).


187

The binding chemistry for fatty acids shifts abruptly for fatty acids with chain lengths of 10 carbon atoms or less. Smaller fatty acids such as these compete for a common binding site with tryptophan and diazepam (Kragh-Hansen, 1983). Thus, one of the primary binding sites for these medium-length fatty acids can be assigned with confidence to the binding pocket within IIIA. Crystallographicstudies of fatty acid binding to serum albumin are in progress in our laboratory as this manuscript finds its way to the publisher. However, some interesting preliminary results can be discussed here. We have found from the study of cocrystals of rHSA and HSA with long-chain fatty'acids of laureate or palmitate that significant conformational changes have taken place. The binding of three or more fatty acids produces a slight opening of the interface between the two halves of the molecule and a rotation of domain I (Fig. 16, see color insert). This is contrary to previous fluorescence and absorption spectroscopic studies that propose that albumin becomes more compact with the addition of long-chain fatty acids (Honor6 and Pedersen, 1989). In addition, it appears that the environment surrounding Cys-34 becomes more exposed to solvent. The latter findings are in excellent agreement with the observed increase in oxidization of Cys-34 as a function of fatty acid binding (Takabayashiet al., 1983).Conversely, the same study found that oxidation of the free sulfhydryl increased the affinity of albumin for fatty acids. Recent diffraction experiments wtih HSA cocrystallized separately with lauric acid and a novel iodinated lauric acid analog have revealed three major sites for the iodinated fatty acid. One located in IB, one in the binding pocket of IIIA, and the other in IIIB. These findings, although based on a derivatized fatty acid, agree well with previous studies that indicate the presence on albumin of two to three high affinity fatty acid binding sites (Koh and Means, 1979; Sollene and Means, 1979; Reed 1986; Parks et al., 1983). Furthermore, they are in agreement with the number and location of fatty acid sites proposed by the "C cleic acid studies of Hamilton et al. (1991). Based on the previous findings of Means and co-workers one might assume that the two highest affinity long chain fatty acid sites are those located on the two B subdomains. Additionally, these two high affinity sites are located at or near the surface of the molecule which may explain why, despite the high association constants of fatty acids (lo' M-'),they can be readily exchanged between albumin molecules in solution (Peters, 1985). In due course of the structure refinement, with an improved model and diffraction data, we hope a more detailed description of the binding process can be made.

188


C . Metals: Sites V and VI Albumin has a high affinity for Cu(II), Ni(II), Hg(II), Au(I), and Ag(II), and weaker affinities for calcium and zinc. Many individuals have spent significant portions of their professional careers studying the particular chemistry associated with each metal-binding site. For example, Sarkar and co-workers have extensively examined the Cu and Zn binding chemistry associated with the N terminus (Dixon and Sarkar, 1974), and Shaw and co-workers studied the chemistry associated with Au complexes with Cys-34 (Shaw, 1989). There have been fewer studies characterizing the affinity for Zn. However, Lakusta et al. (1980) associate this binding with the Cu(I1)-binding site. Consequently, albumin possesses two major binding sites for metals, which we denote as Site V and Site VI in the detailed discussion that follows. 1 . Cys-34: Site V

Albumin is responsible for the largest fraction of free sulfhydryl (Cys34) in blood serum, and studies have shown that it is also the most reactive sulfhydryl. The chemical reactivity of Cys-34 is reported to have an unusually low pKsH of 5 compared with 8.5 and 8.9 for cysteine and glutathione, respectively (Pedersen and Jacobsen, 1980). In most preparations of albumin, 30-35% of Cys-34 is occupied by cysteine or glutathione. Blocking of Cys-34 with cysteine, glutathione, or other chemicals such as N-idosuccinimide stabilizes albumin against dimer formation (Peters, 1985). Presumably, Cys-34 plays a direct or catalytic role in this process. Cys-34 also binds Au, Ag, Hg, Cd, and, to a lesser extent, Cu. Major interests in the unusual chemistry at this site include understanding the nature of bound pharmaceuticals, such as the antiarthritic auranofin, or other gold(1)-containing thiolates (Shaw, 1989). Other pharmaceutical interests involve reactions with non-metal-containing antibiotics and the recently identified complexation with nitric acid (Stamler et al., 1992). Examinations of the crystal structures of HSA and ESA in this vicinity indicated that Cys-34 is located in a crevice on the surface of the protein and that the reactive sulfur is somewhat protected by several residues (Fig. 17, see color insert). In HSA, Cys-34 is in close proximity to Glu-82 and His-39. In ESA the Cys environment is also protected but, with the exception of His-39, which (along with Cys-34) is conserved in mammalian albumin sequences (Table 11),it involves contributions from different amino acids. Thus, one may expect that His-39 plays a major role in the enhanced reactivity of the free sulfhydryl. Interestingly, in the crystal structure of albumin complexed with three or more long-chain fatty acids,


189

the crevice containing Cys-34 has opened significantly, exposing the sulfhydryl and increasing its distance from its nearest neighbors, including His-39. It is noteworthy that in salmon albumin Cys-34 is replaced by Ser, and both salmon and frog albumins have His-39 replaced by Leu. As a result, one would predict the free sulfhydryl of Xenopur albumin to be less reactive. 2. N Terminus: Site VI

Albumin possesses for Cu(I1) and Ni(I1) one distinct high-affinity site, which has been well characterized by several research groups. Binding of Cu by albumin was perhaps first described by Fiess and Klotz in 1952 and was identified with the N terminus by Peters and Blumenstock in 1967. They surmised that the copper atom was coordinated by the N-terminal nitrogen, the next two peptide nitrogens, and the NE of His-3. Camerman et al. (1976) demonstrated similar copper binding chemistry with the tripeptide crystal structure of glycylglycyl-L-histidine-N-methylamide, a functional analog of the N-terminal tripeptide (Fig. 18). Albumins that lack histidine at position 3, such as canine albumin, have a much lower affinity for Cu(I1) and Ni(II), which presumably explains the greater susceptibility of dogs to copper poisoning (Peters, 1984). In the crystal structures of HSA, rHSA, and ESA, the first two to three residues are

FIG. 18. Structure of the copper-binding peptide, glycylglycyl-L-histidine-Nrnethylarnideillustrating the coordination state and peptide conformation. Figure modified from Carnerrnan et al. (1976).

190

DANIEL

C. CARTER

AND JOSEPH X. HO

disordered, implying a much greater degree of flexibility for the N terminus. This is understandable because substantial conformational changes would be required to accommodate the square-planar bipyramidal coordination of the copper or nickel atom.

IV. EVOLUTION OF ALBUMIN STRUCTURE A discussion of the structure of albumin would not be complete without an examination of the conserved primary structure of albumin. All the present lines of evidence point toward the evolution of albumin from a precursor one-third the current size. This was originally suggested by Brown (1976), and is now verified based on the internal sequence homology and the shared topology of each of the three domains. T h e observation that, in the albumin structure, six subdomains exist that share a common topology and are related by pseudo twofold symmetry within each domain further indicates that the current domain of albumin evolved from an incomplete gene duplication event, as originally suggested by Brown (1976). That is, the B subdomains are missing the additional smaller disulfide double loop. Consequently, it is the A (Fig. 13A) subdomains that represent the true protoalbumin, as well as the principal binding properties of albumin. Whether the B subdomains have evolved specialized functions with cell surface receptors or fatty acid binding remains undetermined. When considering the evolutionary significance of a series of the albumin sequences in Table 11, it is interesting to examine the globin family of proteins, for which a great deal of research has been amassed (Dayhoff, 1978). Comparisons of the structures of hemoglobin and myoglobin families revealed a number of residues that remained invariant (14%). Many of these were directly related to function either by association with the heme pocket or by participation in the interfaces responsible for the allosteric effect. Still others seemed important for stabilization of the protein (Dickerson and Geis, 1983). In albumin there are 332 (57%) invariant residues in the sequences of mammalian albumins and 95 (16%) invariant residues conserved, including frog and salmon albumins. In both of the above comparisons, 34 of the conserved residues are cysteines. A useful matrix illustrating the sequence identities between the known sequences of albumin is shown in Table VIII. T h e greatest level of sequence identity occurs, not surprisingly, between bovine and ovine albumins, which are 92% conserved. T h e evolution of albumin and the construction of a phylogenetic tree has been revisited by Gray and Doolittle (1992). Th e time for the gene duplication event leading to the three-domain structure shared by all fish, tetrapods, and mammals is

191


TABLE VIII Matrix Indicating Sequence ldentily between Presently Known Segunces of Serum Albumin9 Source

HSA

BSA

ESA

OSA

RSA

FSA

BSA ESA OSA RSA FSA SSA

44 1 442 435 426 22 1 161

430 539 409 218 170

438 422 222 153

404 216 165

225 159

154

a

With the exception of lamprey serum albumin. See Table I for albumin abreviations.

placed at approximately 450 million years (MY)ago, much later in evolutionary time than the 700 MY estimated by Brown and Shockley (1982). In principle, the estimate by Gray and Doolittle is more in line with those proposed for globin sequences (Dickerson and Geis, 1983). As in hemoglobin, one would expect that the evolutionary pressure to conserve residues in albumin would impart, together with the knowledge of the atomic structure, some new insight into the chemistry and function of albumin. This is clearly the case. Examination of the amino acids known to participate in the binding process within IIA and IIIA reveals a somewhat surprising result. Only two residues are invariant in the binding pocket of IIA, Arg-257 and Leu-260, whereas there are seven in IIIA Asn-390, Cys-391, Arg-4 10, Tyr-4 11, Thr-4 12, Val-432, Cys-437, Glu450, Leu-452, and Arg-484. If one considers additionally the amino acids that, with only one exception, are conserved, this brings the total to 10 in IIIA and 3 in IIA. Clearly evolutionary pressure has operated to conserve the binding chemistry of the IIIA region of the serum albumin molecule. Let us therefore consider another binding region of serum albumin, Cys-34, which is conserved in all albumin sequences given in Table 11, except salmon albumin. Examination of the residues that surround the cleft containing the free sulfhydryl reveals no conserved pattern. However, under the surface of the cleft is the highly conserved His-39, which is in close proximity to Cys-34. One must then conclude ~ this residue that it is this imidazole that imparts the unusually low p K s of and further reflects its importance in sequestering various metals, cysteine, and glutathione. His-3, a requirement for the Cu- and Ni-binding properties of albumin, is again conserved in all of the sequences given in Table I1 with the exception of those from salmon and dog (Dixon and Sarkar, 1974). As mentioned previously, this may explain the unusual sensitivity of dogs to copper poisoning (Peters, 1984), and one wonders

192


whether the problem may be compounded in fish to impart an additional sensitivity to exposure to Hg and Ag, for example. Brown (1976) noted a similarity in the long disulfide loops of BSA and the G-H region of sperm whale myoglobin, basing this observation on the match of two prolines and a similar distribution of hydrophobic residues. This led to his suggestion that albumin evolved from a primordial globin. Indeed, pori;ons of the structure of albumin strongly resemble the myoglobin/hemoglobin fold. T h e helices h 1, h2, h3, and h4 superimpose on the last four helices E, F, G, and H of hemoglobin. The striking similarity can be seen in Fig. 19, (see color insert). Although the positions of the helices are conserved, the polarity is reversed in serum albumin helices h3 and h4 compared to the corresponding G-H pair in hemoglobin. Surprisingly, not only are the general helical features consistent, the numbers of amino acids in each helical segment are within the variation seen for myoglobin as well. This resemblance takes on added meaning when one considers that the albumins of primates are known to bind hematin with high affinity (Beaven et al., 1974). Moreover, apomyoglobin is known to have a similar diverse affinity for a variety of small molecules. Whether the albumin and globin families are actually related by a common ancestor or represent yet another example of convergent evolution is not clear. McLachlan and Walker (1977), who used an extensive statistical analysis of three serum albumins to support the latter, stated “our tests do not support the idea that serum albumin evolved from myoglobin. It is more likely that parts of these proteins independently acquired similar helical structures.” In this regard it is interesting to note that the “myoglobin fold” has recently been identified in a number of functionally unrelated proteins that lack significant sequence homology (Holm and Sander, 1993). In addition to the pseudosymmetry observed between subdomains, a pseudo twofold screw axis relates two consecutive domains (see Section 11,C,2). The orientation of the axis relating domain I and I1 is slightly different from that relating I1 and 111. This observation has interesting implications regarding other multidomain structures, such as in lamprey albumin (Gray and Doolittle, 1992). If the domain-domain packing arrangements are conserved in albumins as distantly related as lamprey albumin, then after applying the pseudosymmetry operator relating domains I1 and 111 to the extra four domains of lamprey serum albumin (LSA), the structure can be described as a straight helical coil with a diameter of 80 X 40 A and a pitch of 55 (Fig. 20A). Otherwise, if the pseudosymmetry operator relating domains I and 11, and the same relating I1 and 111, are applied, the structure will be a curved helical coil, or partial superhelical coil (Fig. 20B).It is reasonable to assume that the

FIG.20. Conceptual model of seven-domain structure of lamprey serum albumin. In the HSA structure, there exist pseudo twofold screw axes between domains I and I1 and between domains I 1 and I l l , whereas the symmetry axes are not quite parallel to each other. In addition, the connections between two successive domains are made by the long, continuing helix (consisting of the C-terminus helix h10 and the N-terminus helix h l of the succeeding domain h 1). This arrangement greatly restricts the way two successive domains can be packed together. If the lamprey serum albumin forms a structure similar to that of HSA, with its extra four domains, the lamprey serum albumin structure should look like A, assuming a single pseudo symmetry repeat, or like B, assuming two alternate-diad repeats. The structure shown in A forms a straight zig-zag,or helical, shape; that shown in B forms a curved zig-zag, or super-helical shape. Figures drawn using program RIBBONS (Carson, 1987).

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actual structure will be slightly more compact to minimize the molecular surface. Furthermore, a cursory survey of the sequence of LSA suggests that among the subdomains of LSA, its IA subdomain has the highest homology to the IIIA subdomain of a “three-domain’’ albumin, based on the distribution of previously determined invariant residues in Table 11. One might postulate that it may also function likewise.

V. SUMMARY AND FUTUREDIRECTIONS Albumin is clearly an extraordinary molecule of manifold functions and applications. Although the exact function of albumin has been debated, much of the present data support the notion that the principal role of albumin in the circulatory system is to aid in the transport, metabolism, and distribution of exogenous and endogenous ligands. T h e ability of albumin to act as an important extracellular antioxidant (Halliwell, 1988) or impart protection from free radicals, and other harmful chemical agents (Emerson, 1989) agrees well with the increased susceptibility of analbuminemic rats to cancer (Kakizoe and Sugimura, 1988). T h e expression and delivery of albumin to the circulatory system by the liver therefore seem appropriate. An overview of the prolific ligand-binding properties of albumin is summarized in Fig. 2 1. The positions of known binding sites for important pharmaceutical markers such as diazepam, ibuprofen, aspirin, and warfarin are illustrated. In addition, the important endogenous markers tryptophan, octanoate, and bilirubin are also shown. With the exception of the definitive positions of the long-chain fatty acids, most albumin-ligand chemistry can now be explained by the atomic coordinates derived from crystal structures. Knowledge of the atomic structure coupled with the current applications of genetic engineering, such as site-directed mutagenesis, promises to provide an even greater understanding of albumin chemistry. It is widely accepted in the pharmaceutical industry that the overall distribution, metabolism, and efficacy of many drugs can be altered based on their affinity to serum albumin. In addition, many promising new drugs are rendered ineffective because of their unusually high affinity for this abundant protein. Obviously, an understanding of the chemistry of the various classes of pharmaceutical interactions with albumin can suggest new approaches to drug therapy and design, placing albumin in its rightful place as the “second step in rational drug design.” Application of albumin in other therapeutic approaches is widely known. Some studies have suggested that modified serum albumin may be used as a selective contrast agent for tumor detection and/or therapy (Sinn et al., 1990). Other studies have demonstrated that albumin may be used to


195

FIG. 2 1 . Illustration summarizing the various ligand-binding sites on serum albumin. The asterisks denote binding sites that can be inferred; all others have been determined crystallographically.

deliver toxic compounds for elimination of Mycobacterium tuberculosis via receptor-mediated drug delivery (Majumdar and Basu, 1991). Recently, chimeric albumin molecules such as HSA-CD4 (Yeh et al., 1992) and HSA-Cu,Zn-superoxide dismutase (Ma0 and Poznansky, 1989) have been utilized to increase the half-life and distribution, and reduce the immunogenicity, of these potential protein therapeutics. Albumin has now been cloned and expressed in several bacterial and fungal systems. The primary motivation for many of these studies has been the potential of recombinant albilmin to serve as a serum replacement product that is free from unwanted viral contaminants, e.g., hepatitis, herpes, and human immunodeficiency virus (HIV). The most successful production has been achieved by extracellular expression in yeast (Etcheverry et al., 1986; Hinchcliffe and Kenney, 1986; Kalman et al., 1990; Okabayashi et al., 1986; Quirk, et al., 1989; Sijmons et al., 1990; Sleep et al., 1991). Clearly, future scientific and therapeutic applications of albumin appear limitless.

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In conclusion, albumin may be unique among proteins in that so many scientists have spent the largest portion of their professional careers studying very specific aspects of this protein. New appreciation for the complexity and potential applications presented by the structure of albumin promises to consume the careers of many more scientists.

ACKNO w LEDGMENTS The authors are indebted to several individuals, for their generous support during the course of preparing this manuscript. In particular, we are grateful to Pam Twigg for help in preparing many of the tables and references, Teresa Miller for proofreading the manuscript, and Jewel1 Reynolds, Kim Keeling, Brenda Barnes, Tongi Shavers, and Mike Carson for assisting with the preparation of the figures. We thank F. W. Putnam for providing the tabulation of point mutations of albumin in advance of publication. Reproduction of the color figures was provided as a courtesy of the National Aeronautics and Space Administration, Marshall Space Flight Center.

REFERENCES Abdo, Y., Rousseaux, J.. and Dautrevaux, M. (1981). FEES Lett. 131, 286-288. Adams, P. A., and Berman, M. C. (1980). Biochem. J. 191,95-102. Adinolfi, A., Adinolfi, M., and Lessof, M. H. (1975).J. Med. Genet, 12, 138-151. Ahlstedt, S., and Kristofferson, A. (1982). Prog. Allergy 30,67. Akoi. K., Sato, K., Nagoaka, S., Kamada, M.,and Hiramatsu, K. (1973). Biochim. Biophys. Acta 328,323-333. Ancell, H. ( 1 839). Lancet 1,222-23 1 . Arai, K., Huss, K., Madison, J., Putnam, F. W., Salzano, F. W., Franco, M. H. L. P., Santos, S. E. B. and Freitas, M. J. M. (1989a). Proc. Natl. Acad. Sci. U.S.A., 86, 1821-1825. Arai, K., Madison, J., Huss, K., and Putnam, F. W. (1989b).Proc. Natl. Acad. Sci. U.S.A. 86, 6092-6096. Arai, K., Madison, J., Shimizu. A., and Putnam, F. W. (1990). Proc. Natl. Acad. Sci. U.S.A., 87,497-501. Beaven, G. H., Chen, S.-H., DAlbis, A., and Gratzer, W. D. (1974). Eur. J. Biochem. 41, 539-546. Behrens, P. Q., Spiekerman, A. M.,and Brown, J. R. (1975). Fed. Proc., Fed. Am. SOC.Exp. Biol. 34, 591. Bendedouch, D., and Chen, S. H. (1983).J. Phys. Chem. 87,1473-1477. Bloomfield, V. (1966).Biochemistty, 5,684-689. Borst, G . C., Eil, C., and Burman, K. D. (1983).Ann. Intern.Med. 98,366-378. Bos, 0.J. M., Labro, J. F. A., Fischer, M. J. E., Witling, J., and Janssen, L. H. M . (1989). J . Biol. Chem. 264,953-959. Brand, J. G., and Toribara, T. Y. (1975).Arch. Biochem. Biophys. 167,91-98. Brand. S., Hutchinson, D. W., and Donaldson, D. (1984). Clin. Chim. Acta 136, 197-202. Breckenridge, A., and Rosen, A. (1971).Biochim. Biophys. Acta 229,610-617. Brennan, S. 0.(1985).Biochim. Biophys. Acta 830,320-324. Brennan, S. O., and Carrell, R. W. (1978).Nature (London) 274,908-909. Brennan, S. O., Peach, R. J., and Boswell, D. R. (1989).Biochim. Biophys. Acta 993,48-50. Brennan, S . O., Arai, K., Madison, J., Laurell, C.-B., Galliano, M., Watkins, S., Peach, R., Myles, T., George, P., and Putnam, F. W. (1990a). Proc. Natl. Acod. Sci. U.S.A. 87, 3909-3913.


197

Brennan, S. 0..Myles, T., Peach, R. J.. Donaldson, D., and George, P. M. (1990b).Proc. Natl. Acad. Sci. U.S.A. 87,26-30. Brock, A. (1975).Acta Phurmucol. Toxicol. 36, 13-24. Brodersen, R. (1979).J.Eiol. Chem. 254,2364-2369. Brodersen, R., Andersen, S., Vorum, H., Nielsen, S. U., and Pedersen, A. 0.(1990).Eur.J. Biochem. 189,343-349. Brown, J. R. (1975).Fed. Proc., Fed. Am. SOC.Exp. Eiol. 34,591. Brown, J. R. (1976).Fed. Proc., Fed. Am. SOC.Exp. Biol. 35,2141-2144. Brown, J. R. (1977).In “Albumin Structure, Function and Uses”(V. M. Rosenoer, M. Oratz, and M. A. Rothschild, eds.), pp. 27-51. Pergamon, Oxford. Brown, J. R., and Shockley, P. (1982).In “Lipid-Protein Interactions” (P. Jost and 0. H. Griffith, eds.), Vol. 1, pp. 25-68. Wiley, New York. Brown, K. F., and Crooks, M. J. (1976).Biochem. Phurmacol. 25, 1175-1178. Brown, W. M., Dziegielewska, K. M., Foreman, R. C., and Saunders, N. R. (1989).Nwleic Acids Res. 17, 10495. Burke, C. W., Lewis, B., Panveliwalla, D., and Tabaqchali, S. (1971).Clin. Chim. A c h 32,

207-2 14. Byrnes, L., and Cannon, F. (1990).DNA Cell Eiol. 9,647-655. Callan, W. M., and Sunderman, F. W., Jr. (1973).Res. Commun. Chem. Pudol. Phurmacol. 5,

459-472. Cambillau, C., and Horjales, E. (1987).J.Mol. Graphics 5, 174. Camerman, N., Camerman, A., and Sarkar, B. (1976).C a n . ] . C h . 54,1309-1316. Cannistraro, S.,and Sacchetti, F. (1986).Phys. Rev. 33,745-746. Carlson, J. Sakamoto, Y., Laurell, C.-B. Madison,J., Watkins, S., and Putnam, F. W. (1992). Proc. N a L Acad. Sci. U.S.A. 89,8225-8229. Carson, M. (1987).J.Mol. Graphics, 5, 103-106. Carter, D. C.,and He, X.-M. (1990).Science 249,302-303. Carter, D. C., He, X.-M., Munson, S. H., Twigg, P. D., Gernert, K. M., Broom, M. B., and Miller, T. Y. (1989).Science 244, 1195-1198. Charlwood, P. A. (1961).Ei0chem.J. 78, 163-172. Chen, M. C., and Lord, R. C. (1976).J.Am. Chem. SOC.98,990-992. Chignell, C. F. (1969).Mol. Phurmucol. 5,244-252. Chignell, C. F. (1970).Mol. Pharmucol. 6,1-12. Chignell, C. F. (1973).Ann. N . Y . Acud. Sci. 226,44-59. Chincarini, C. C., and Brown, J. B. (1976).Fed. Proc., Fed. Am. SOC.Exp. Eiol. 35, 1621. Cho. M.J., Mitchell, A. G . , and Pernarowski, M. (1971).J.Phunn. Sci. 60, 196-200. Coassolo, P., Briand, C., Bourdeaux, M., and Sari, J. C. (1978).Eiochim. Biophys. A c h 538,

5 12-520. Cohn, E. J., Hughes, W. L., Jr., and Weare, J. H. (1947).J. Am. Chem. SOL. 69, 1753-

1761. Cooke, N. E., and David, E. V. (1985).J . Clin. Invest. 76,2420-2424. Crooks, M. J., and Brown, K. F. (1974).J.Phurm. Phurmacol. 26,304-31 1. Daughaday, W. H. (1959).Physiol. Rev. 39,885-902. Day, J. F., Thorpe, S. R., and Baynes, J. W. (1979).J.Eiol. C h m . 254,595. Dayhoff, M. 0.(1978).“Atlas of Protein Sequence and Structure,” Vol. 5. Georgetown University, Washington, DC. DeLucas, L. J., Smith, C. D., Smith, W., Senadhi, V.-K., Senadhi, S. E., Ealick, S. E., Carter, D. C., Snyder, R. S., Weber, P. C., Salemme, F. R., Ohlendorf, D. H., Einspahr, H. M., Clancy, L. L., Navia, M. A., McKeever, B. M., Nagabhushan,T. L., Nelson,G., McPherson, A., Koszelak, S., Taylor, G., Stammers, D., Powell, K., Darby, G., and Bugg, C. E. (1989).Science 246,65 1-654.

198


Dickerson, R. E., and Geis, I. G. (1983).“Hemoglobin: Structure, Function, Evolution, and Pathology.” BenjaminlCummings, Menlo Park, CA. Dixon, J. W., and Sarkar, B. J. (1974).J.Biol. C h . 249,5872-5877. Dugiaczyk, A., Law, S. W., and Dennison, 0. E. (1982).Proc. Natl. Acad. Sci. U.S.A. 79,

71-75. Eisenthal, R., and Cornish-Bowden, A. (1974).Biochem. J. 139,715-720. Elkarim, A. A., and Means, G. E. (1988).Comp. Biochem. Physiol. B. 91B,267-272. Emerson, T. E., Jr. (1989).CRC Crit. Care Med. 17,690-694. Etcheverry, T.,Forrester, W., and Hiueman, R. A. (1986).Bio. Technology 4, 720-726. Fehske, K.J., Miiller, W. E., and Wollert, U. (1981).B i o c h . Phannocol. 30,687-692. Feldhoff, R. C., and Ledden, D. J. (1983).Biochem. Biophys. Res. Commun. 114,20-27. Feng, L.,Hu, C. 2..and Andrade, J. D. (1988).J.Colloid Inlerface Sci. 126,650-653. Fiess, H. A.,and Klotz, 1. M. (1952).J.Am. Chem. SOC.74,887-891. Figge, J., Rossing, T. H., and Fencl, V. (1991).J.Lab. Clin. Med. 117,453-467. Fogh-Andersen, N. (1977).Clin. Chem. (Winston-Salem, N . C . ) 23,2122-2126. Foster, J. F. (1960).In “The Plasma Proteins” (F. W. Putnam, ed.), 1st ed., Vol. 1, pp. 179-239. Academic Press, New York. Foster, J. F. (1977).In “Albumin Structure, Function and Uses” (V. M.Rosenoer, M. Oratz, and M.A. Rothschild, eds.), pp. 53-84. Pergamon, Oxford. Franklin, S. G., Wolf, S. I., Ozdemir, Y., Yuregir, G. T., Isbir, T., and Blumberg, B. S. (1980).Proc. Natl. Acad. Sci. U.S.A. 77,5480-5482. Gabay, S., and Huang, P. C. (1974).Adv. Biochem. Psychophannocol. 9,175-189. Galliano, M., Minchiotti, L., Iadarola, P., Stoppini, M., Ferri, G., and Castellani, A. A. (1986).FEES Lett. 208,364-368. Galliano, M., Minchiotti, L., Iadarola, P., Ferri, G., Zapponi, M. C., and Castellani, A. A. (1988).FEES Lett. 233, 100-104. Galliano, M., Minchiotti, L., Stoppini, M., and Tarnoky, A. L. (1989).FEES Lett. 255,

295-299. Galliano, M., Minchiotti, L., Porta, F., Rossi, A., Ferri G., Madison, J., Watkins, S., and Putnam, F. W. (1990).Proc. Natl. Acad. Sci. U.S.A. 87,8721-8725. Galliano, M. et al. (1993).Biochim. Biophys. Actu, in press. Garten, S . , and Wosilait, W. D. (1972).Comp. Ga.Pharmacol. 3,83-88. Geisow, M.J., and Beaven, G. H. (1977).B i 0 c h . J . 163,477-484. Gerig, J. T., and Reinheimer, J. D. (1975).J.Am. Chem. SOC.97, 168. Goodman, D. S. (195q.J.Am. Chem. SOC.80,3892-3898. Gorin, M.B., Cooper, D. L., Eiferman, F., van de Rijn, P., and Tilghman, S. M. (1981). J. Biol. C h . 256, 1954-1959. Gray, J. E., and Doolittle, R. F. (1992).Protein Sci. 1,289-302. Green, H.O.,Moritz, J.. and Lack, L. (1971).Biochim. Biophys. Actu 231,550-552. Giirber, A. (1894).Sitzungsber. Phys.-Med. Ges. Wurrburg, p. 143. Guthrow, C. E., Morris, M. A., Day, J. F., Thorpe, S. R., and Baynes, J. W. (1979).Proc. Natl. Atad. Sn’. U.S.A. 76,4258-4261.

Guttler, F., Tybring, L., and Engberg-Pedersen, H. (1971). B Y . ] . Phurmacol. 43, 151-160. Haefliger, D. N.,Moskaitis,J. E., Schoenberg, D. R., and Wahli, W. (1989).J.Mol. Evol. 29,

344-354. Hagag, N.,Birnbaum, E. R., and Darnall, D. W. (1983).Biochemistry, 42,2420-2427. Halliwell, B. (1988).B i o c h . Pharmacol. 37,569-571. Hamilton, J. A., Era, S., Bhamidipati, S. P., and Reed, R. G. (1991).Proc. Natl. Acad. Sci. U.S.A. 88, 2051-2054.

Harrington, W. F., Johnson, P., and Ottewill, R. H. (1956).B i 0 c h . J . 62,569-582.


199

Hart, B. J. T., Witling, J,, and De Gier, J. J. (1986).Biochem. Phanacol. 35, 1005-1009. He, X.-M., and Carter, D. C. (1992).Noture (London)358,209-215. Hinchcliffe, E.,and Kenney, E. (1986).Eur. Pat. Appl. 201-239. Ho, J. X.,Holowachuk, E. W., Norton, E. J., Twigg, P. D., and Carter, D. C. (1993).Eu7.J. Biochem. 415,205-212. Holm, L., and Sander, S. (1993).Nature (London) 361,309. Holowachuk, E.W. (1991).G m Bank D a l a h e , Accession No. M73993. Honor6 B., and Pedersen, A. 0. (1989).Biochem. J. 258,199-204. Huang, P. C., and Gabay, S. (1974).Biochem. Phanacol. 43,957-972. Hughes, W. L. (1954).In “The Proteins” (H. Neurath and K. Biley, eds.), Vol. 2b, pp. 663-755.Academic Press, New York. Hughes, W. L., and Dinitz, H. M. (1964).J.Biol.Chem. 439,845-849. Hultrnark, D., Borg, K. 0.. Elofsson, R., and Palmer, L. (1975).Acta Pharm. Suec. 14,

259-276. Huss, K., Madison,J., Ishioka, N., Takahashi, N., Arai, K., and Putnarn, F. W. (1988).Proc. Natl. Acad. Sci. U.S.A. 85,6692-6696. Hutchinson, D. W., and Matejtschuk, P. (1985).FEBS Lett. 193,211-212. Iadarola, P., Minchiotti, L., and Galliano, M. (1985).FEES Lett. 180,85-88. Iberg, N., and Fluckiger, R. (1986).J.Biol. Chem. 461, 13542-13545. Jacobsen, C. (1972).Eur.J. Biochem. 47,513-519. Jacobsen, J. (1969).FEES Lett. 5,112-1 14. Jacobsen, J. (1977).Int. J.P e p . Protkn Res. 9,235-239. Jagodzinski, L. L., Sargent, T. D., Yang, M., Glackin, C., and Bonner, J. (1981).Proc. Natl. Acad. Sci. U.S.A. 78(6),3521-3525. Janssen, L. H. M., and Nelen, T. H. A. (1979).J.Biol. Chem. 454,5300-5303. Kakizoe, T., and Sugimura, T. (1988).Jpn.J. Cancer Res. 79,775-784. Kalman, M., Cserpan, I., and Bajszar, G. (1990).Nuleic AcidcRes. 18,6075-6081. Katchalski, E.,Benjamin, G. S., and Gross, V. (1957).J. Am. Chem. Soc. 79,4096-4099. Kendall, F. E. (1941).J.Biol. Chem. 138,97-109. Kendrew, J. C., Dickerson, R. E., Strandberg, B. E., Hart, R. G., Davies, D. R., Phillips, D. C., and Shore, C. (1960).Nature (London) 185,422-427. Keresztes-Nag, S., Mais, R. F., Oester, Y. T., and Zaroslinski,J. F. (1972).Anal. Biochem. 48,

80-89. Khan, M. Y. (1986).Bi0chem.J. 236,307-310. Khan, M. Y., and Salahuddin, A. (1984).Eu7.J. Biochem. 141,473-475. King, T. P. (1973).Arch. Biochem. Biophys. 156,509-520. King, T. P., and Spencer, M. (1970).J. Biol. Chem. 445,6134-6148. Klopfenstein, W. E.(1969a).Biochim. Biophys. Acla 181,323-325. Klopfenstein, W . E.(1969b).Biochim. Biophys. A c h 187,272-274. Koh, S.-N., and Means, G. E. (1979).Arch. Biochem. Biophys. 194,73-79. Kragh-Hansen, U.(1981).Phanacol. Rev. 33, 17-53. Kragh-Hansen, U. (1983).Biochem.J. 209, 135-142. Kragh-Hansen, U. (1990).Dan. Med. Bull. 37,57-84. Kurono, Y.,Kondo, T., and Ikeda, K. (1983).Arch. Biochem. Biophys. 227,339-341. Lagercrantz, C., Larsson, T., and Denfors, 1. (1981).Comp. Biochem.P h y s d . C . 69C,375-

378. Lakusta, H., Deber, C. M., and Sarkar, B. (1980).Can.J. Chem. 58,757-766. Lang, J. H., and Lasser, E. C. (1967).Invest. Radiol. 2,396-400. Lau, S.-J., Kruck, T. P. A., and Sarkar, B. (1974).J.Biol. Chem. 449,5878-5884. Law, S. W., and Dugaiczyk, A. (1981).Nuture (London)291,201-205.

200


Lawn, R. M., Adelman, J., Bock, S. C., Franke, A. E., Houck, C. M., Najarian, R. C.. Seeburg, P. H., and Wion, K. L. (1981).Nucleic AcidsRes. 9,6103-6114. Leob, G. I., and Scheraga, H. A. (1956).J.Phys. Chem. 60, 1633-1644. Lewin, J. (1951).J.Am. Chem. SOC.73,3906-3916. Lindup, W. E.(1987).Prog. DrugMetab. 10, 141-185. Longsworth, L. F. (1954).J.Phys. Chem. 58,770. Low, B. W., and Richards, F. M. (1954).J.Am. Chem. SOC.76,2511-2518. Low, B. W., and Weichel, E. J. (1951).J.Am. Chem. SOC.73,3911-3916. Luetscher, J. (1939).J.Am. Chem. SOC.61,2888. Luft, A. J., and Lorscheider, F. L. (1983).Biochemistry 22,5978-5981. Lukas, D. S.,and De Martino, A. G . (1969).J.Clin. Invest. 48, 1041-1053. Madison,J., Arai, K., Sakamoto, Y., Feld, R. D., Kyle, R. A,, Watkins, S., Davis, E., Matsuda, Y.-l., Amaki, I., and Putnam, F. W. (1991).Proc. Natl. Acad. Sci. U.S.A. 88,9853-9857. Majumdar, S . , and Basu, S. K. (1991).Antimicrob. Agents Chemother. 35, 135-140. Mao, G. D., and Poznansky, M.J. (1989).Biomater. Artif. Cells Artif. Organs 17(3),229-244. Mason, R. W.,and McQueen, E. G. (1974).P h m c o l o g y 12, 12- 19. McClure, R. J., and Craven, B. M. (1974).J. Mol. Biol. 83,551-555. McLachlan, A. D., and Walker, J. E. (1977).J.Mol. Biol. 112,543-558. McMeekin, T. L. (1939).J. Am. Chem. SOC.61,2884-2890. McMenamy, R. H., and Oncley, J. L. (1958).J.Biol. Chem. 233, 1436-1447. Meloun, B., Moravek, L., and Kostka, V. (1975).FEES Lett. 58, 134-137. Miller, T. Y.,He, X.-M., and Carter, D. C.(1992).J.Cryst. Growth 122,306-309. Minchiotti, L., Galliano, M., Iadarola, P., Stoppini, M., Ferri, G., and Castellani. A. A. (1987).Biochim. Biophys. Acta 916,411-418. Minchiotti, L., Galliano, M., Iadarola, P., Zepponi, E., and Ferri, G. (1990).Biochim. Biophys. Acta 1039,204-208. Minchiotti, L., Galliano, M., Stoppini, M., Ferri, G., Crespeau, Rochu, D., and Porta, F. (1992).Eichim. Biophys. Acta 1119,232-238. Minchiotti, L., Galliano, M., Zapponi, M. C., and Tenni, R. (1993).Eur. J . Biochm. 214, 437-444. Minghetti, P. P., Law, S . W., and Dugaiczyk, A. (1985).Mol. Eiol. Evol. 2,347-358. Morinaga, T., Sakai, M., Wegmann, T. G., and Tamaoki, T. (1983).Proc. Nafl. Acad. Sci. U.S.A. 80,4604-4608. Moser, P., Squire, P. G., and OKonski, T. 0. (1966).J.Phys. Chem. 70,744-756. Mudge, G . H., Desbiens, N., and Stibitz, G . R. (1978a).Drug Metab. Dispos. 6,432-439. Mudge, G . H., Stibitz, G . R., Robinson, M. S., and Gemborys, M. W. (1978b).Drug Melab. D~SPOS. 6,440-45 1. Miiller, W. E., and Wollert, U. (1973).Naunyn-SchmieLberg's Arch. P h a m c o l . 280,229-237. Miiller, W.E.,and Wollert, U. (1975).Mol. Phurmacol. 11, 52-60. Nagase, S.,Shimamune, K., and Shumiya, S. (1979).Science 205,590-591. Naik, D. V.,Paul, W. L., and Schulman, S. G. (1975).J.Pharm. Sci. 64, 1677-1680. Nandedkar, A. K. N., Basu, P. K., and Friedberg, F. (1972).Bioinmg. Chem. 2, 149-157. Nandedkar, A. K. N., Nurse, C. E.,and Friedberg, F. (1973).Int. J. Pept. Protein Res. 5, 279-28 1. Nazareth, R.I., Sokoloski,T. D., Witiak, D. T., and Hopper, A. T. (1974).J.P h r m . Sci. 63, 203-211. Nilsen, 0.G., and Jacobsen, S . (1976).Biochem. Phrmucol. 25, 1261-1266. Oers, N. S. C., Cohen, B. L., and Murgita, R. A. (1989).J. Exp. Med. 170,811-825. Okabayashi, K.,Nakagawa, Y., Hayasuke, N., et al. (1986).J.Biochem. (Tokyo) 100, 15331542.


20 1

Oncley, J. L., Scatchard, G., and Brown, A. (1947).J.Phys. ColloidChem. 51, 184-198. Ozeki, Y., Kurono, Y., Yotsuyanagi, T., and Ikeda, K. (1980). C h . Pharm. Bull. 28, 535-540. Parks, J . S., Cistola, D. P., Small, D. M., and Hamilton, J. A. (1983).J . Eiol. Chem. 258, 9262-9269. Paubel, J.-P., and Niviere, P. (1974).Eur.J. Med. Chem. 9,508-512. Peach, R. J., and Brennan, S. 0. (1991).Eiochim. Eiophys. Acta 1097,49-54. Pearlman, W. H., and Crepy, 0. (1967).J.Eiol. Chem. 242, 182-189. Pearson, W. R. (1990).In “Methods in Enzymology” (R. Doolittle, ed.), Vol. 183, pp. 63-98. Academic Press, San Diego. Pedersen, K. 0. (1971). Scand. J. Clin. Lab. Invest. 28,459-469. Pedersen, K. 0. (1972). Scand. J. Clin. Lab. Invest. 29,427-432. Pedersen, A. O., and Jacobsen, J. (1980). Eur. J. Eiochem. 106,291-295. Perrin, J. H., Vallner, J. J., and Nelson, D. A. (1975).Eiochem. Phunnacol. 24,769-774. Perutz, M. F. (1989). “Is Science Necessary?”Barrie and Jenkins, Ltd., London. Perutz, M. F., Rossmann, M. G., Cullis, A. F., Muirhead, H., Will, G., and North, A. C. T. (1960). Nature (London) 185,416-422. Peters, T., Jr. (1975). In “The Plasma Proteins” (F. W. Putnam, ed.), 2nd ed., Vol. 1, pp. 133-181. Academic Press, New York. Peters, T., Jr. (1980). “Serum Albumin-An Overview and Bibliography.” Miles Laboratories, Elkhart, IN. Peters, T., Jr. (1984). In “The Impact of Protein Chemistry on the Biomedical Sciences” (A. N. Schecter and R. F. Goldberger, eds.), pp. 39-55. Academic Press, New York. Peters, T., Jr. (1985).Adv. Protein Chem. 37, 161-245. Peters, T., Jr. (1992). “Albumin: An Overview and Bibliography,” 2nd ed. Miles Inc. Diagnostics Division, Kankakee, IL. Peters, T., Jr., and Anfinsen, C. B. (1950).J.Eiochem. (Tokyo) 86,805-813. Peters, T.,Jr., and Blumenstock, F. A. (1967).J.Eiol. Chem. 244, 1574-1578. Peters, T., Jr., and Feldhoff, R. C. (1975).Biochemtsty 14,3384-3390. Porta, F. A., Galliano, M., Rossi, A,, and Porta, F. (1990). Boll. Osp. Vurese 19, 197-210. Quirk, A. V., Geisow, M. J., Woodrow, J. R., Burton, S. J., Wood, P. C., Sutton, A. D., Johnson, R. A., and Dodsworth, N. (1989). Eiotechnol. ApPl. E i o c h . 11, 273-287. Rao, S. N., Basu, S. P., Sanny, C. G., Manely, R. V., and Hartsuck, J. A. (1976).J. Eiol. Chem. 251,3191-3193. Reed, R. G . (I986).J.Eiol. Chem. 261, 15619-15624. Richards, F. M. (1985).In “Methods in Enzymology” (H. Wyckoff el al., eds.), Vol. 115, pp. 440-464. Academic Press, Orlando, FL. Richardson, K. S.C., Nowaczynski, W., and Genest, J. (1977).J.Steroid C h . 8,951-957. Roda, A., Cappelleri, G., Aldini, R., Roda, E., and Barbara, L. (1982). J. Lipid Res. 23, 490-495. Rosen, A. (1970).Eiochem. Phurmucol. 19,2075-2081. Rossmann, M. G., and Blow, D. M. (1962). Acta Ctystallogr. 15,24-31. Rothschild, M. A., Oratz, M., and Schreiber, S. S. (1988). Hepatology 8,385-401. Russi, E.. and Weigand, K. (1983). Klin. Wochenschr. 61,541-545. Sakamoto, Y., Davis, E., Madison, J., Watkins, S., McLaughlin, H., Leahy, D. T., and Putnam, F. W. (1991). Clin. Chim. Acta 204, 179. Sargent, T. D., Yang, M., and Bonner, J. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 243246. Savu, L., Benassasyag, C., Vallette, G., Christeff, N., and Nuney, E. (1981).J.Eiol. C h . 256.9414-9418.

202


Sawinski, V. J., and Rapp, G. W. (1963).J.Dent. Res. 42, 1429-1438. Scatchard, G. (1949). Ann. N . Y . Acad. Sci. 51,660-672. Scatchard, G., and Yap, W. T. (1964).J.Am. Chem. SOC.86,3434-3438. Schoentgen, F., Metz-Boutigue, M.-H.,Joll&s,J.. Constants, J., and Joll&s,P. (1986).Biochim. Biophys. Acta 871, 189-198. Sebille, B., Thuaud, N., and Tillement, J.-P. (1978).J. Chromatop. 167, 159-170. Shaklai, N., Garlick, R. L., and Bunn, H. F. (1984).J.Biol. Chem. 259,3812-3817. Sharpies, D. (1975).J.Phurm. Phurmocol. 27,379-381. Shaw, C. F. (1989). Comments Inmg. Chem. 8(6), 233-267. Shockley, P., and Brown, J. R. (1980).Fed. Proc., Fed. Am. SOC.Ex#. Biol. 39,2069. Shrake, A., Finlayson,J. S., and Ross, P. D. (1984). VoxSung. 47,7-18. Sijmons, P. C., Dekker, B. M. M., Schrammeijer, B., Venvoerd, T. C., van der Elzen, P. J. M., and Hoekema, A. (1990). Biotechnology 8,217-221. Sinn, H., Schrenk, H. H., Friedrich, E. A., Schilling, U., and Maier-Borst, W. (1990). Nucl. Med. Biol. 17,8 19-827. Sjoholm, I., and Ljungstedt, I. (1973).5.Biol. Chem. 248, 8434-8441. Slayter, E M. (1965).J.Mol. Biol. 14,443-452. Sleep, D., Belfield, G . P., and Goodey, A. R. (1991). BiolTechnology 8,42-46. Sollene, N. P., and Means, G. E. (1979). Mol. Phurmocol. 14,754-757. Sollene, N. P., Wu, H.-L., and Means, G. E. (1981). Arch. Biochem. Biophys. 207,264-269. Spector, A. A., and Fletcher, J. E. (1978). In “Disturbances in Lipid and Lipoprotein Metabolism” M. Dietschy, A. M. Gotto, Jr., and J. A. Ontko, eds.), pp. 229-248. Am. Physiol. Soc., Rockville, MD. Spector, A. A., Santos, E. C., Ashbrook, J. D.,and Fletcher, J. E. (1973).Ann.N. Y. Acud. Sci. 226,247-258. Squire, P. G., Moser, P., and OKonski, C. T. (1968).Biochemistty 7,4261-4272. Stamler, J. S . , Singel, D. J., and Loscalzo,J. (1992). Science 258, 1898-1902. Steiner, R. F., Roth. J.. and Robbins, J. (1966).J.Biol. Chem. 241,560-567. Sudlow, G.,Birkett, D. J., and Wade, D. N. (1975).Clin. Exp. Phurmacol. Physiol. 2, 129-140. Sudlow, G., Birkett, D. J.. and Wade, D. N. (1976). Mol. Pharmucol. 12, 1052-1061. Sugita, O., Endo, N.. Yamada, T., Yakata, M., and Odani, S. (1987). Clin. Chim. A c h 164, 25 1-259. Takabayashi, K., Imada, T., Saito, Y., and Inada. Y. (1983). Eur.J. Biochem. 136,291-295. Takahashi, N., Takahashi, Y., Blumberg, B. S., and Putnam, F. W. (1987a).Proc. Nutl. Acud. Sci. U.S.A. 84,4413-4417. Takahashi, N., Takahashi, Y.,and Putnam, F. W. (1987b).Proc. Nutl. Acad. Sci. U.S.A. 84, 7403-7407. Takahashi, N., Takahashi, Y.. Isobe, T., Putnam, F. W., Fujita, M., Chiyoko, S., and Neel, J. ( 1987~). Proc. Natl. Acad. Sci. U.S.A. 84,800 1-8005. Tarnoky, A. L. (1980).Adu. Clin. Chem. 21, 101-146. Teeter, M.M., and Hendrickson, W. A. (1979).]. Mol. Biol. 127,219. Tillement, J.-P., Zini, R., DAthis, P., and Vassent, G. (1974). Eur. J. Clan. Phurmucol. 7, 307-313. Tucker, E. M. (1968). Vox Sung. 15,306-308. Unger, W. G. (1972).J.Phmm. Phurmocol. 24,470-477. Vermeulen, A., and Verdonck, L. (1968). Steroidc 11,609-635. Waldmann, T. A. (1977). In “Albumin Structure, Function and Uses” (V. M. Rosenoer, M. Oratz, and M. A. Rothschild, eds.), pp. 255-273. Pergamon, Oxford. Waldmann-Meyer, H. (1960).J.Biol. Chem. 235,3337-3345. Walker, J. E. (1976). FEBSLett. 66, 173-175.

u.


203

Weinstock, J., and Baldwin, G. S. (1988).Nwleic Act& Res. 16(18),9045. Westphal, U., and Harding, G. B. (1973).Eiochim. Ei0phys. Acta 310,518-527. Whitlam, J. B.,Crooks, M. J., Brown, K. F., and Pedersen, P. V.(1979).BiOchem. Phurmacol.

28,675-678. Wilting,J., Kremer, J. M. H., Ijzerman, A. P., and Schulman, S. G. (1982).Biochim. Eiophys. Acta 706,96-104.

Winter, W. P., Weitkamp, L. R.,and Rucknagel, D. L. (1972).Eiochnnishy 11,889-896. Wright, A. K., and Thompson, M. R. (1975).Ei0phys.J. 15, 137-141. Yang, F.,Brune, J. L., Naylor, S. L., Cupples, R. L., Naberhaus, K. H.,and Bowman, B. H. (1985).Proc. Natl. Acad. Sci. U.S.A. 82,7994-7998. Yang, F., Bergeron, J. M., Linehan, L. A., Lally, P. A., Sakaguch, A. Y.,and Bowman, B. H. (1990).Genomics 7(4),509-516. Yates, F. E., and Urquhart, J. (1962).Physiol. Rev. 42,359-443. Yeh, P., Landais, D., LeMaitre, M., Maury, I., Crenne, J.-Y., Becquart, J.. Murry-Brelier, A., Boucher, F., Montay, G., Fleer, R.,Hirel, P.-H., Mayaux, J.-F., and Klatzmann, D. (1992).PTOC.Natl. Acad. Sci. U.S.A. 89, 1904-1908. Yvon, M., and Wal, J.-M. (1988).FEES Lett. 239,237. Zurakowski, R.,Jr., and Foster, J. F. (1974).Eiochemitty 13,3465-3471. Zvelebil, M.J. J. M., Barton, G. J., Taylor, W. R., and Stemberg, M. J. E. (1987).J.Mol. Biol.

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APOLIPOPROTEIN B AND LOW-DENSITY LIPOPROTEIN STRUCTURE: IMPLICATIONS FOR BIOSYNTHESIS OF TRIGLYCERIDE-RICH LIPOPROTEINS By VERNE N. SCHUMAKER, MARTIN L. PHILLIPS, and JON E. CHATTERTON Department of Chomlrty and Blochemlstry,and the Molecular Biology Instltute, Unlverrlty of Calltornla, Lor Angeler, Lor Angeler, Callfornla BOO24

I. Introduction

.......................................................

11. Apolipoprotein B Structure .......................................... A. Gene for Human Apolipoprotein B ...............................

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B. Apolipoprotein B Message ....................................... C. Primary Sequence of Apolipoprotein B ........................... Low-Density Lipoprotein Structure ................ A. Emulsion Particle Model for Lo ins ............. B. Quantitative Molecular Model for Low-Density Lipoproteins ........ Structural Studies of Apolipoprotein B on Low-Density Lipoprotein Surfaces ........................................................... A. Lipid Extraction after Attachment of Low-Density Lipoproteins to Electron Microscope Grids ....................................... B. Mapping of Apolipoprotein with Monoclonal Antibodies C. Relating Low-Density Lipoprotein Core Circumferences to Apolipoprotein B Fragment Sizes ................................. Lipoprotein Assembly ............................................... A. Gotranslational Lipoprotein Formation in Rough Endoplasmic Reticulum ...................................................... B. Two-step Model for Assembly of Triglyceride-Rich Lipoproteins .... Summary .......................................................... References .........................................................

205 207 207 209 210 213 215 217 226 227 227 235 240 240 243 243 244

I. INTRODUCTION In this article the structure and function of apoliprotein B (apoB) will be described and an attempt will be made to relate its structure to its function. Apolipoprotein B is a moderately hydrophobic, extraordinarily large (-550 kDa) glycoprotein intimately involved in the packaging, transport, and utilization of apolar lipids, particularly cholesteryl ester and triglyceride. An apoB-like protein may be found on plasma lipoproteins from most, if not all, vertebrate species, including a “primitive” vertebrate, the hagfish [Mill and Taylaur, 1978; Goldstein et al., (1977)l. Outside of the vertebrates, apoB has not been found, although there are intriguing reports of a large (-600 kDa) apolipoprotein associated with triglyceride-rich lipoproteins present in sea urchin eggs (Marsh, 1968; Ichio et al., 1978). Apolipoprotein B plays a unique role in lipid transport ADVANCES IN PROTEIN CHEMISTRY, Vol. 45

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in the vertebrates, directing the biosynthesis of the triglyceride-richlipoproteins. In humans genetically deficient in apoB, chylomicron and very low-density lipoprotein (VLDL)levels are unmeasurable, and the two cell types that synthesize these lipoproteins, the enterocytes and the hepatocytes, respectively, accumulate droplets of unsecreted lipids. A shortened version of apoB, apoB48, directs chlomicron formation, whereas fulllength apoBlOO directs the assembly of human VLDL. ApoB plays a second metabolic role in the conversion of a portion of the VLDL remnants to low-density lipoproteins (LDLs). During this process a conformational change in apoBlOO results in the formation of a receptor-binding site (Lund-Katz et al., 1988, 1991). The recognition of this binding site on apoB by the hepatic LDL receptor is a critical step in the maintenace of a low serum LDL level and, in consequence, the delay or prevention of atherosclerosis (Goldstein and Brown, 1989). 11. APOLIPOPROTEIN B STRUCTURE A. Gene for Human Apolipopotein B

The gene for human apolipoprotein B is located on chromosome 2, and regional mapping and in situ hybridization have positioned the gene to the short arm in the p23 to p24 region (Mehrabian et al., 1986; Knott et al., 1985; Cann and Guyer, 1992). Thus, the apoB gene is unlinked to members of the gene family encoding the other major apolipoprotein species, which are dispersed on chromosomes 1, 11, and 19 (Bruns et al., 1984; Fojo et al., 1984;Jackson et al., 1984;Jeanpierre et al., 1984; Knott et al., 1985). Twenty-eight introns interrupt 29 exons (Fig. 1A) and the gene spans 43 kbp of genomic DNA, a modest stretch for a gene encodFIG. 1. The locations of significant features along the apoB polypeptide. Each line represents apoB100, with the N terminus on the left and the C terminus on the right. The horizontal axis corresponds to amino acid number. (A) Intron boundaries. (B) Sites of N-linked carbohydrate attachment. (C) Disulfide bonding pattern. The diamonds are known free cysteines; the squares are cysteines whose status is unknown. (D) The midpoints of extremely hydrophobic segments in apoB. (E)ApoB internal repeats; The taller segments are the 52-residue proline-rich repeats; the shorter segments are the 22-residue amphipathic helix repeats. (F) The domain structure proposed for apoB based on differential trypsin releasability (Yang el al., 1989). TR, Trypsin releasible; TN, trypsin nonreleasible; M, both. (G) The two sites cleaved on limited proteolysis of apoB with 12 different proteases of various specificities (Chen et al., 1989) and the location of the thrombin fragments generated by limited digestion. (H) Binding sites of monoclonal antibodies used in immunoelectron microscopy of LDLs. (I) Approximate positions of the C termini of apoB fragments generated on puromycin treatment of HepC2 cells.

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ing a protein the size of apoB. The first exon contains the 5‘-untranslated region and also codes for the signal sequence. The mature protein sequence starts with exon 2. Most of the exons range in size from 150 to 250 nucleotides, and 24 of these encode the first one-third of apoB. Exon 26 is composed of 7572 base pairs and is “by far the longest reported for a vertebrate gene” (Blackhart et al., 1986). T h e last exon, exon 29, is also very long, and contains 1906 bp. Complete sequences of all 29 exons and all but the middle regions of about a dozen large introns have been reported. T h e intron/exon boundaries have all been sequenced and conform to the standard pattern for these junctions (Blackhart et al., 1986; Carlsson el al., 1986; Higuchi et al., 1987; Wagener et al., 1987; Ludwig et al., 1987). Regulatory sequences located in the 5‘ region of the gene include a classical TATA box located 29 nucleotides 5’ of the transcriptional start site and a CAAT box 31 nucleotides 5’ of the TATA box. Two GC boxes occur on the 3’ side of the transcriptional start site within the untranslated portion of the mRNA, and therefore are “of dubious functional significance” (Blackhart et ad., 1986). Liver-specific expression is controlled by two positive elements located from -128 to -85 and -84 to -70 (Das et al., 1988). Rat liver nuclear proteins bind to these elements; thus, BRF-2 binds to the -128 to -85 region, and BRF-1 and C/EBP bind to overlapping sites at -84 to -61 and -70 to -50 (Zhuang et al., 1992). In addition, Brooks and Levy-Wilson (1992) have identified a tissue-specific transcriptional enhancer containing four distinct proteinbinding sites in the second intron of the apoB gene from +806 to +952. A C/EBP-related protein and multiple hepatocyte nuclear factors appear to compete for these four sites within the second intron. Negative regulatory sites in the 5’ region of the apoB gene have also been reported (Das et al., 1988; Paulweber et al., 1991; Paulweber and Levy-Wilson, 1991). Th e length of the chromatin loop containing the human apoB gene was determined by locating three nuclear matrix attachment sites, two at the 5’ end and one at the 3‘ end. At the 5’ end, the distal site was located between nucleotides -5262 and -4048, and the proximal site, between -2765 and -1801. At the 3’ end, a single site was found between +43,186 and +43,850. HepG2 cells, which express the apoB gene, contained all three sites whereas HeLa cells, which do not express apoB, lacked the 5’ distal site (Levy-Wilson and Fortier, 1989). A substantial number of common polymorphisms are associated with the human apoB gene, and Table I lists those which have been reported more than once. These have proved valuable in genetic (Young el al., 1986), anthropological (Rapacz et al., 1991). forensic (Butler, 1990), and medical studies (Soria et al., 1989).

209


TABLE I Common Polymorphism of Apolipoprotein B

Common coding sequence polymorphisms (observed more than once) Restriction Exon Common name Amino acids Residue no. endonuclease 1 3 14 14 23 26 26 26 26 26 26 26 27 27 29 29

Signal Seq Ag(c/g) Ag(al1d)

Leu-Ala-Leu ThrlIle Val1Ala Leullle GluIGln LeuILeu ThrlThr LeuIPro AlalAla HisIAsp GlnlArg IleIThr PhelLeu TrylPhe Serl Asn GlulLys

--14 to --16 73 59 1 618 1191 206 1 2488 2712 2822 3292 361 1 3705 3922 3937 431 1 4514

BSP12861; ApLl AluI

XbaI Mae1

MspI Secl

EcoRI

Common polymorphism located outside the coding sequence Intron Location Restriction endonuclease 5' end 2 3 4 4 20 3' end

4 kb upstream of exon 1 Nucleotide #722 92 bp 3' to exon 3 171 bp 3' to exon 4 523 bp 5' to exon 5 146 bp 5' to exon 21 Microsatellite 490 bp 3' to exon 29

A d 1

Ball HincII Pm II BalI MspI

Ref." 1 2, 3 4 5 5 5.6 7 8.9 5 5 10 597 5 5 9 7, 11, 12

Ref." 13 14 13 13 13 13 15, 16

"Key to references: (1) Boerwinkle and Chan, 1989; (2) Ma et al., 1989; (3) Young and Hubl, 1989; (4) Wang et al., 1988; (5) Ludwig et al., 1987; (6) Olofsson et al., 1987; (7) Blackhart et al., 1986; (8) Wu el al., 1991; (9) Dunning el al., 1992; (10) Xu et al., 1989; (11) Maetal., 1987; (12) Huangetal., 1986; (13) Huangetal., 1990; (14)Levy-Wilsonetal., 1991; (15) Boerwinkle et al., 1989; (16) Ludwig el al., 1989.

B . Apolipoprotein B Message

The processed mRNA transcribed from the human apoB gene contains 14,121 and 14,112 nucleotides (Cladaras etal., 1986),dependingon the presence or absence of a 9-nucleotide insertion/deletion polymorphism in the signal sequence (Boerwinkle and Chan, 1989). Messenger

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VERNE N. SCHUMAKER ET AL.

RNA is abundant in rat liver and intestine, the two organs that synthesize triglyceride-rich lipoproteins; moreover, a small amount of message is found in rat adrenal tissue and a “20-fold increase in placental apoB mRNA concentrations during the last 48 hr of pregnancy. . . suggests a specific role for this organ in maternal-fetal lipid transport immediately prior to parturition” (Demmer et al., 1986). Two different versions of the apoB protein are employed in the biosynthesis of the triglyceride-rich lipoproteins (Kane et al., 1980). T h e smaller version, called apoB48, is used for chylomicron biosynthesis by the intestine. The full-length apoB, called apoBl00, is used for VLDL biosynthesis by the liver. These two different apoB proteins are encoded by the single apoB gene. Processing occurs at the RNA level to create the message encoding apoB48 from the full-length apoB 100 message. In the human and rabbit, this processing event is tissue specific, occurring only in the enterocytes of the intestine (Powell et al., 1987), whereas in the rat, both messages are produced and expressed in the liver (Davidson et al., 1988). Processing of apoB message involves a unique mechanism consisting of the enzymatic deamination of a single cytidine at human nucleotide 6666, converting the glutamine codon 2153 (CAA) to a stop codon (UAA) (Powell et al., 1987; Chen et al., 1987; Hospattankar etal., 1987). With the formation of this stop codon, the altered message is translated to express the apoB48 polypeptide in the rat. Further processing occurs in humans and rabbits to truncate and polyadenylate the message at cryptic polyadenlyation sites located downstream from the newly formed stop codon (Powell et al., 1987). The editing reaction can be duplicated in a cell-free extract, and no metal ion cofactors, DNA or RNA cofactors, or energy requirements have been found (Garcia et al., 1992).

C. Primary Sequence of Apolipoprotein B Complete cDNA sequences of human apoB 100 have been reported by several laboratories (Knott et al., 1986; Yang et al., 1986; Cladaras et al., 1986; Law et al., 1986; Olofsson et al., 1987). Partial sequences are also available for apoBlOO from rats (Reuben et al., 1988; Matsumoto el al., 1987),swine (Maeda et al., 1988) and chickens (Kirchgessner et al., 1987). From estimation of substitution rates, it appears that apoB evolves at about twice the rate of ordinary mammalian proteins (O’hUigin et al., 1990). These authors found that the rate of substitution is not uniform across the molecule, however. There is a general increase in substitution rates going from the 5’ to the 3’ end; for example, in a comparison of the human and the rat, the 5‘-most 1155 nucleotides evolve at one-fourth the


21 1

rate of the 3’-most 1089 nucleotides. This suggests that the amino terminus of apoB may be structurally or functionally more constrained than the carboxyl terminus. In addition to a gradient in substitution rates, there are several regions, two of which have been implicated in binding to the LDL receptor, that are much more conserved than surrounding sequences (O’hUigin et al., 1990). Portions of apoB may have evolved from internal duplications, because it contains some internally homologous sequences. At least two families of related sequences are present; there are six similar 52-residue hydrophobic proline-rich sequences, unique to apoB (Fig. lE), and eight similar 22-residue sequences, which are also homologous to the peripheral apolipoproteins (Knott el al., 1986; Yang et al., 1986; De Loof et al., 1987). In addition to this self-homology, residues 3352-337 1 of apoB are similar to residues 136-155 of apoE, which contain the LDL receptor-binding site of apoE. The amino-terminal 1000 amino acids of apoB contain several long sequences homologous to the vitellogenins of vertebrates and nematodes, precursors to the egg yolk lipoproteins, the lipovitellins (Baker, 1988; Perez et al., 1991). Homologous regions to vitellogenin in apoB span approximately residues 20-280,530-600, and 800-970 (Banaszak et al., 1991). The apoB 100 sequence, deduced from cDNA sequences, consists of a signal sequence 27 or 24 amino acids long, depending on the presence or a deletion of amino acids - 14 to - 16, and the mature protein sequence of 4536 amino acids. Peptides covering approximately 90% of the sequence of the serum protein have also been sequenced directly (Yang et al., 1989). Aside from the signal sequence, no other amino acids are lost during the maturation of the protein. Ignoring the signal sequence, apoB48 consists of the N-terminal2 152 amino acids of apoB 100. Protein sequencing of the apoB48 C terminus from chylous ascites fluid indicates that in this system, Ile-2 152 has been removed, presumably by a carboxypeptidase A type activity, leaving Met-215 1 as the C-terminal amino acid (Chen et al., 1987). The protein portion of apoBlOO has a predicted molecular weight of 513,000 and that of apoB48 is 243,000. ApoBlOO contains 5 4 % carbohydrate (Lee and Breckenridge, 1976; Shireman and Fisher, 1979; Vauhkonen, 1986),so the total molecular weight of the apoB 100 glycoprotein is 540,000-550,000; that of the apoB48 glycoprotein is about 260,000. Of the 19 potential N-glycosylation sites in apoB100, 16 are actually glycosylated (Yang et al., 1989); 5 of these sites are in apoB48 (see Fig. 1B). Both high-mannose and complex forms of oligosaccharides are present. In apoB 100 there are 8-10 mol of complex type and 5-6 mol of high-mannose form (Taniguchi et al., 1989); in apoB48 there are about 4

212

VERNE

N.

SCHUMAKER ET AL.

mol of complex oligosaccharides and 1 mol of the high-mannose form (Sasak et al., 1991). It is not known whether there are O-linked carbohydrates in apoB; however, they are not present in large amounts. There are 25 cysteines in apoB; 16 of these are known to be involved in disulfide bonds, 7 are known to be free, and the remaining two, Cys-2906 and Cys-4326, are unknown (Yang et al., 1990). Only two of the free cysteines of apoB, Cys-3734 and Cys-4190, are labeled in LDL by the fluorescent probe 5-iodoacetamidofluorescein,possibly because they are exposed on the surface of the lipoprotein (Coleman et al., 1990). Of the 8 disulfide bonds in apoB100, 7 are within 1000 amino acids of the N terminus of the protein (6 in the first 500 amino acids). T h e first two disulfides, CysL2-Cys6'and C y ~ ~ ' - C ycross; s ~ ~ ,the remainder of the disulfides are between nearest neighbors. T h e number of amino acids separating the cysteines in each disulfide is relatively small, 5-49 amino acids; the exception is C y ~ ~ ' ~ ' - C wherein y s ~ ~ ~the ~ , cysteines are separated by 130 amino acids. This disulfide is apparently unique to the human sequence, because cysteines are not found in homologous positions in rat, mouse, Syrian hamster, pig, rabbit, o r chicken apoB (Law and Scott, 1990). The disulfide pattern of apoBlOO is shown schematically in Fig. 1C. The average hydrophobicity of apoB 100 is 0.916 kcallresidue, a value intermediate between that of the peripheral apolipoproteins and that of intrinsic membrane proteins (Chen et al., 1986). Aside from the signal sequence, there are no stretches of 20 or more hydrophobic amino acids corresponding to typical bilayer membrane-spanning helices. There are, however, other features of the sequence that may be related to the lipid-binding characteristics of apoB. Thus, there are 39 short sequences (5-13 amino acids, with only two sequences being more than 10 amino acids long) that have hydrophobicities typical of membrane-spanning domains (Olofsson et al., 1987). These sequences are distributed throughout the apoB sequence (see Fig. 1D). They are not uniformly hydrophobic, but are occasionally interrupted by uncharged polar or, frequently in the longer sequences, charged residues. These segments are usually predicted to have P-sheet character. T h e folding of apoB may bring some of these hydrophobic sequences together. Indeed, the first two of these sequences, residues 8-12 and 60-65, are linked by the C y ~ ' ~ - C y disulfide s~' bond. The amphipathic helix, in which residues are spaced so that the helical periodicity places hydrophobic side chains on one side of the helix and hydrophilic side chains on the other, is a common structural motif used by the peripheral apolipoproteins to bind lipid (Segrest et al., 1992); it is also a structural element present in globular proteins (Perutz et al., 1965).

213


There are many predicted amphipathic a helices in apoB. Of particular interest is a family of related sequences 22 amino acids long, which are homologous to the other apolipoproteins (De Loof et al., 1987).These are found in two regions, residues 2000-2500 and 4000-4500 (see Fig. 1E). Also noteworthy are residues 3352-337 1 of apoB, which are homologous to the LDL receptor-binding region of apoE (residues 136-155) and conform to an amphipathic a helix. The homologous sequence in apoE has been shown to be an amphipathic a helix by crystallographicanalysis of the amino-terminal domain of apoE (Wilson et al., 1991). Sequences corresponding to arnphipathic @ strands, in which predicted @ structure is combined with alternation of hydrophobic and hydrophilic amino acids, are also present in apoB. These sequences are spread throughout apoB. Although there are a few long amphipathic @-strand segments containing 10-20 residues, most are short, no more than five amino acids long with 3 hydrophobic residues. There is some clustering of amphipathic @ strands in the proline-rich repeats (see Fig. lE),with a five-amino acid consensus sequence of acidic-aromatic-polaraliphatic-proline (Knott et al., 1986).In the moderate-resolution crystal structure of lamprey lipovitellin (Raag et al., 1988),the lipid hydrocarbon chains, localized by neutron diffraction in varying D20/H20 mixtures (Timmins et al., 1992),are in a cavity lined by @ sheet. Lamprey vitellogenin (lipovitellin)displays homology with other vertebrate and nematode vitellogenins (Banaszak et al., 1991) to which apoB is homologous, and it is suggestive that homologous amphipathic @-strandsequences are present. A good example of such a segment is shown below.

apoB 48-58

R

Vitellogenins Xenopus A 1 54-64

K

N K E S \I/ \c/ \v/ \I/

R

S R R Q V \I/ \A/ \A/ \I/ \A/

C. Elegans Vit-2 54-64

\/

N

K

E

\=/ jV/ \L/

E

P

Iv/

\A/Y

I1 I. LOW-DENSITY LIPOPROTEIN STRUCTURE

Low-density lipoproteins contain a single molecule of apoB 100 (Knott et al., 1986) and almost no other protein; therefore, they are uniquely

suited to the study of the interactions between apoB and lipids. Lipoproteins have densities that are lower than the densities of plasma proteins, which do not contain lipids. This characteristic is used to purify and fractionate lipoproteins by sequential flotation centrifugation to

2 14


yield the familiar lipoprotein categories, which include the chylomicrons, the very low-density lipoproteins, the intermediate-density lipoproteins (IDLs), the low-density lipoproteins, and the high-density lipoproteins (HDLs) (Schumaker and Puppione, 1986). Traditionally, human LDLs have been defined as those lipoproteins isolated in the density interval between 1.063g/ml > LDL > 1.019g/ml. This density interval may be too broad, however, because a substantial contamination with apolipoproteins other than apoB may be present on those lipoproteins at the two extremes of this density range; the contaminating apolipoproteins are principally apoE (present on IDLs and HDLs at the low- and high-density extremes of this density interval, respectively) and apo(a) [present on Lp(a) at the high-density extreme]. This contamination was almost absent in the more tightly defined LDL fraction lying between 1.024 and 1.050 g/ml (Chapman et al., 1988). However, it is possible that loosely associated apolipoproteins are normally bound to LDLs in plasma and are subsequently lost during the multiple flotation steps involved in lipoprotein isolation (Mahley and Holcombe, 1977). The average weight percent composition of pooled human LDL is about 19% protein, 1% carbohydrate, 43% cholesteryl ester, 11% unesterified cholesterol, 4% triglyceride, and 22% phospholipid (see later, Table 11). Because there is a single molecule of apoBlOO on the LDL, a simple method of obtaining the number-average molecular weight of a lipoprotein preparation is to divide the molecular weight of the apoB 100 by the weight percent protein on the LDL. An exact value for the molecular weight of the protein portion of the apoB 100 glycoprotein was calculated from its amino acid sequence by Yang et al. (1986)to be 512,937. Thus, the number-average molecular weight of typical LDL, calculated from its protein content, is M , = 512,937/0.19= 2.7 x lo6. Moreover, assuming a spherical shape and a density of 1.030 g/ml, the anhydrous lipoprotein radius may be calculated from Eq. (1):

where d is the density and N is Avogadro's number. Substituting the values for density and molecular weight, given above, Eq. (1)yields R = 101 A. By definition, the Stokes radius is obtained as the product of the anhydrous radius and the translational frictional ratio; the translational frictional ratio has been measured for LDLs and found to be 1.1 1 (Fisher


215

et al., 1971). Thus, the average Stokes radius of pooled human LDL, as calculated from the number-average molecular weight, is 112 A.

A. Emulsion Particle Model f o r Low-Density Lapoproteins Structurally, LDLs are well described as emulsion particles. An emulsion may be defined as one liquid embedded in another and kept in solution by an emulsifying agent. For LDLs, the first liquid is a droplet of oil, largely cholesteryl ester but containing some triglyceride; the second liquid is the aqueous plasma and the emulsifying agent is a monolayer of phospholipid, unesterified cholesterol, and protein. This monolayer forms an amphipathic surface coat surrounding the oil droplet and separating the hydrophobic, liquid core of the LDL from the aqueous plasma (Bradley and Gotto, 1978). What is the evidence supporting the emulsion particle model for LDLs? One feature that may be used to distinguish an emulsion particle from a closed bilayer vesicle is that, for an emulsion particle, all of the phospholipid should be exposed to the external medium, whereas for a bilayer vesicle, somewhat more than one-half should be exposed. Enzymatic hydrolysis of LDLs by phospholipase Ag converted all of the phosphatidylcholine and phosphatidylethanolamine to their corresponding lysophospholipids (Aggerbeck et al., 1976), indicating that all of the phospholipid was located at the aqueous interface at the LDL surface. In addition, 31P NMR studies of LDLs have demonstrated that all of the phosphate was accessible to small amounts of Pr3+, a rare earth probe that should not cross a bilayer because of its ionic nature (Yeagle et al., 1978). These results agree with the emulsion particle model. The emulsion particle model also places apoB at the surface of the LDL, consistent with proteolysis studies, and with studies of the binding of anti-apoB monoclonal antibodies to LDLs. Thus, trypsin removed about 30% of the protein (Triplet and Fisher, 1978; Chapman et al., 1987; Margolis and Langdon, 1966a), leaving a collection of peptide fragments of variable lengths associated with the lipoprotein; both the trypsin-removable and trypsin-nonremovable peptides have been sequenced and found to be distributed along the length of apoB (Yang et al., 1990) (see Fig. lF), showing that trypsin cleavage was not restricted to a few domains. Many monoclonal antibodies have been generated to epitopes located along the length of apoB using LDL as an immunogen; these monoclonals also bound to LDLs, demonstrating that much of the protein must be exposed to the solvent (Pease et al., 1990). Variablecontrast neutron scattering studies have also indicated that the protein is

2 16

VERNE N . SCHUMAKER E T AL.

located far from the center of the LDL (Laggner et al., 1981). Again, the results are consistent with the emulsion particle model. The emulsion particle model places the nonpolar lipids in an oil droplet at the center of the LDL. The nonpolar lipids are observed to undergo a liquid-to-liquid crystalline transition at physiological temperatures, characteristic of cholesteryl esters (Deckelbaum et al., 1977). These workers showed that the midpoint of the temperature transition varied with the triglyceride content of the LDL. Detailed analysis of the lowangle X-ray scattering pattern strongly suggested that oscillations in the radial distribution function, with a periodicity of 30 to 40 81, and the 36-81 fringe at 10°C were “explained by two layers of cholesteryl ester molecules oriented radially in a smectic-like phase within the core of a single LDL” (Deckelbaum et al., 1977). The thickness of the surface monolayer of phospholipid, cholesterol, and protein has been estimated from the variation in lipoprotein composition with lipoprotein size, and values of 21.5 81 (Sata et al., 1972) and 20.2 81 have been estimated (Shen et al., 1977). These values seem reasonable, although they are somewhat less than one-half the thickness observed for egg lecithin bilayers. For such bilayers, the thickness of the lipid layer varies inversely with the degree of hydration, from about 36 81 for the least hydrated form to about 30 81 for the most hydrated. To these values must be added twice the distance “extending from the glycerol-3carbon to the edge of the phosphorylcholine group, when lying parallel to the plane of the bilayer, [which] gives a distance of about 8 81 perpendicular to the plane of the bilayer” (Small, 1986). Thus 8 81 + 881 + 30 81 = 46 81 for the bilayer, and one-half of this value, or 23 81, would be estimated for the thickness of a hydrated monolayer. The average LDL size and density are found to vary between individuals (Adams and Schumaker, 1969; Fisher et al., 1975; Krauss and Burke, 1982), and these studies suggest that the variation is due to both genetic and dietary factors. Figure 2 shows hydrodynamic data taken from three studies of subfractionated LDLs and IDLs isolated from different individuals, as summarized by Schumaker (1973). Subfractionation of LDLs by density has yielded particles differing substantially in molecular weights, as determined from their flotation coefficients. To compare values measured under different solvent conditions in these three studies, the flotation coefficients for the particles shown in Fig. 2 have all been corrected for solvent density and viscosity to the values they would exhibit in a KBr solvent with a density of 1.20 g/ml, and the viscosity of KBr at 25°C (Schumaker, 1973). A quantitative molecular model for LDLs consistent with these data will be developed next.


217

BUOYANT DENSIM (g/rnl)

FIG. 2. Density dependence of the flotation coefficient for LDLs and IDLs. The experimental values (0)were compiled by Schumaker (1973) and represent fractionated lipoproteins from both normal and abnormal lipidemic individuals. The calculated values, represented by the solid line, are a plot of column 2, Table 11, of this review, and demonstrate that the emulsion particle model is compatible with the observed hydrodynamic properties of these lipoproteins.

B . Quantitative Molecular Model for Low-Density Lipoproteins For molecular modeling of the lipoproteins, values for the partial specific volumes of the lipoprotein components are required. The partial specific volume of an aqueous egg yolk lecithin suspension is 0.984 ml/g (Hauser and Irons, 1972), and this provides a reasonable approximation for the partial specific volume of the phospholipid occupying the surface monolayer of a lipoprotein. The reciprocal of the density of liquid triolein (Small, 1986) yields its partial specific volume, 1.102 mVg, and provides a reasonable approximation for triglyceride dissolved in the cholesteryl ester-filled core of the LDL. For cholesterol, the partial specific volume of 1.021 ml/g measured in benzene (Haberland and Reynolds, 1973) has been employed. The value of 0.740 ml/g employed for the partial specific volume of apoBlOO was determined from its amino acid composition (Lee et al., 1987). A value of 0.60 ml/g was used for the partial specific volume of the carbohydrate moiety. One important parameter, the partial specific volume of cholesteryl ester, remains to be determined. As will be shown below, its value is estimated to be 1.058 ml/g. The emulsion particle model for LDLs developed here assumes that the particles have a spherical core consisting of cholesteryl esters and

218


triglyceride. The triglyceride content as a percentage of total core lipid was allowed to vary according to a least-squares quadratic fit between the lipoprotein radius and the compositional data of Chapman et al. (1988); this variation in the triglyceride content resulted in only a very small change in lipoprotein density, size, and shape, while bringing calculated lipoprotein composition into agreement with experimental observation. According to the emulsion particle model, the core is surrounded by a spherical shell of 20.2 A (Shen et al., 1977) consisting of apoBlOO (M 5 12,937), phospholipid, and cholesterol. T h e phospholipid and cholesterol were assumed to be present in a 1 : 1 molar ratio, close to that observed for LDLs (Chapman et al., 1988).T h e single molecule of apoB was assumed to be located at the surface of the LDL but embedded within the surface shell surrounding the core. This is reasonable, because the volume of phospholipid and cholesterol is sufficient to fill only about 70% of the volume of a 20.2-A surface shell around an LDL of average size. Conveniently, the volume of a single apoB molecule almost exactly fills the remaining 30% of this volume. From this emulsion particle model, it was possible to predict the lipoprotein composition, buoyant density, and hydrodynamic properties of the LDL as a function of lipoprotein size, given the partial specific volumes of the lipid, protein, and carbohydrate components. A crosssectional slice through the model is shown in Fig. 3 as two concentric circles, representing the hydrophobic core surrounded by a monolayer of phospholipid, cholesterol, and protein. The model parameters are given in the footnote to Table I1 and include the thickness of the shell, the

ApoB FIG.3. The emulsion particle model for LDLs. This cross section of the spherical emulsion particle shows a core of radius, r, surrounded by a shell of thickness, t, in which is embedded a single molecule of apoB. The external radius R = r + t.


219

partial specific volumes of each component, and the molecular weight of the protein. The carbohydrate, which is attached to the surface of the protein, was allowed to extend into the aqueous phase. It contributes to the molecular weight, volume, and density of the lipoprotein and to the frictional ratio of 1.1 1, but does not displace the shell lipids. Table I1 lists predicted values for the lipoprotein size, hydrodynamic properties, and composition, calculated according to this model. In order to calculate the parameters listed in Table 11, a radius was selected and the lipoprotein volume was calculated. The radius of the core was determined by subtracting the shell thickness from the lipoprotein radius; then the volume of the core was computed. From the calculated ratio of the two core lipids (from the quadratic fit between radius and the percentage of core triglycerides) and their partial specific volumes, the density of the core lipid was determined; then, given the core volume, the core weight was calculated. Once the weight of the core was known, then the weights of cholesteryl ester and triglyceride were calculated from their weight ratio. The difference between the lipoprotein volume and the core volume yielded the shell volume. From the shell volume was subtracted the volume of a single apoBlOO protein molecule, to yield the volume occupied by the phospholipid and cholesterol. From the 1 : 1 molar ratio assumed for these two surface lipids and their partial specific volumes, the density of the surface lipid was determined; then, given the surface lipid volume, its weight was calculated; the weights of phospholipid and cholesterol then followed from their ratio. The weight and volume of the carbohydrate were added to the weight and volume of the remainder of the LDL components before calculating the lipoprotein density. Finally, from the weights of all the components, the percentage of each component was calculated and listed in Table 11. For simplicity of calculation, the core was assumed to contain all of the triglyceride and cholesteryl ester, although it is known that small amounts of the core lipids are dissolved in the surface monolayer, where they represent about 3 mol% of the surface lipids, and a larger fraction, about one ninth of the cholesterol, is dissolved in the core (Miller and Small, 1987).The presence of core lipids in the lipoprotein surface is very important metabolically, for the lipases and transfer proteins have access to these core lipids without having to penetrate the surface monolayer. For the calculation of composition, density, and size, however, the effects of component transfer between surface and core affect these quantities about one part in the fourth significant figure, and have been neglected in Table 11. Once the composition and partial specific volume are known for each lipid, protein, and carbohydrate component of the LDL, then the LDL

TABLE I1 Predicted Physical Properties and Composition of LDL( According to Emulsion Particle ModeP

Composition in weight (percent)

w

0

Radius

Densityb

Molecular weight'

SZ5,,.20d

90.0 91.0 92.0 93.0 94.0 95.0 96.0 97.0 98.0 99.0 100.0 101.0 102.0 103.0 104.0 105.0 106.0 107.0 108.0 109.0 110.0 111.0 112.0

1.0555 1.0527 1.0499 1.0473 1.0447 1& 23 I 1.0399 1.0376 1.0354 1.0333 1.0312 1.0292 1.0273 1.0254 1.0236 1.0219 1.0202 1.0186 1.0170 1.0154 1.0139 1.0124 1.0110

1.96 2.02 2.08 2.14 2.21 2.27 2.34 2.41 2.48 2.55 2.62 2.69 2.77 2.84 2.92 3.00 3.08 3.16 3.25 3.33 3.42 3.51 3.60

-26.42 -27.54 -28.66 -29.79 -30.94 -32.09 -33.25 -34.42 -35.60 -36.79 -38.00 -39.2 1 -40.43 -41.66 -42.91 -44.16 -45.42 -46.70 -47.99 -49.28 -50.59 -51.92 -53.25

s; 1.19 1.68 2.18 2.68 3.17 3.68 4.18 4.69 5.20 5.72 6.24 6.76 7.29 7.82 8.35 8.89 9.43 9.98 10.53 11.08 11.64 12.21 12.78

CE

TG

PL

C

Protein

Carbohydrate/

39.2 39.6 39.9 40.2 40.5 40.8 41.1 41.4 41.6 41.9 42.1 42.3 42.5 42.7 42.9 43.0 43.2 43.3 43.4 43.5 43.6 43.7 43.8

2.1 2.3 2.4 2.6 2.8 3.0 3.2 3.4 3.6 3.8 4.0 4.2 4.5 4.7 5.0 5.2 5.5 5.8 6.0 6.3 6.6 6.9 7.2

20.6 20.9 21.1 21.2 21.4 21.5 21.7 21.8 21.9 22.0 22.1 22.2 22.3 22.4 22.4 22.5 22.5 22.6 22.6 22.6 22.6 22.7 22.7

10.2 10.3 10.4 10.5 10.5 10.6 10.7 10.7 10.8 10.8 10.9 10.9 11.0 11.0 11.0 11.1 11.1 11.1 11.1 11.1 11.2 11.2 11.2

26.2 25.4 24.7 23.9 23.3 22.6 21.9 21.3 20.7 20.1 19.6 19.1 18.5 18.0 17.6 17.1 16.7 16.2 15.8 15.4 15.0 14.6 14.3

1.7 1.7 1.6 1.6 1.5 1.5 1.4 1.4 1.3 1.3 1.3 1.2 1.2 1.2 1.1 1.1 1.1 1.1 1.o 1.o 1.0 1.o 0.9

r4

2

113.0 114.0 115.0 116.0 117.0 118.0 119.0 120.0 121.0 122.0 123.0 124.0 125.0 126.0 127.0 128.0 129.0

1.0096 1.0083 1.0070 1.0057 1.0044 1.0032 1.0020 1.0009 0.9997 0.9986 0.9976 0.9965 0.9955 0.9944 0.9935 0.9925 0.99 15

3.69 3.78 3.88 3.97 4.07 4.17 4.27 4.38 4.48 4.59 4.69 4.80 4.92 5.03 5.14 5.26 5.38

-54.59 -55.95 -57.32 -58.70 -60.09 -61.49 -62.91 -64.34 -65.78 -67.23 -68.70 -70.18 -71.67 -73.18 - 74.69 -76.23 -77.77

13.35 13.93 14.51 15.10 15.69 16.29 16.90 17.51 18.12 18.74 19.37 20.00 20.64 2 1.29 21.94 22.59 23.26

43.8 43.9 43.9 43.9 43.9 43.9 43.9 43.9 43.8 43.8 43.7 43.6 43.5 43.4 43.3 43.2 43.0

7.5 7.8 8.2 8.5 8.8 9.2 9.5 9.9 10.3 10.6 11.0 11.4 11.8 12.2 12.6 13.0 13.5

22.7 22.7 22.7 22.7 22.7 22.7 22.6 22.6 22.6 22.6 22.5 22.5 22.5 22.5 22.4 22.4 22.3

11.2 11.2 11.2 11.2 11.2 11.2 11.2 11.1 11.1 11.1 11.1 11.1 11.1 11.1 11.0 11.0 11.0

13.9 13.6 13.2 12.9 12.6 12.3 12.0 11.7 11.4 11.2 10.9 10.7 10.4 10.2 10.0 9.7 9.5

0.9 0.9 0.9 0.8 0.8 0.8 0.8 0.8 0.7 0.7 0.7 0.7 0.7 0.7

0.6 0.6 0.6

~________

"Shell thickness, 20.2 A. Partial specific volumes of cholesteryl ester (CE), triglyceride (TG), phospholipid (PL), cholesterol (C), protein, and carbohydrate are 1.058, 1.102, 0.984, 1.021, 0.740, and 0.60 g/ml, respectively. A 1 : 1 molar ratio of phospholipid to cholesterol is assumed in the surface. Core triglyceride, as the percentage of total core lipids, is given by core T G = 9.191 - 0.4132R + 0.00408364R2,where R is the lipoprotein radius in angstroms. T h e buoyant densities in this column = (Hc,v,) + 0.0016 g/ml, where the last additive term is to adjust for the differential compressibilities of water and lipoproteins at 52,640 rpm. 'LDL molecular weight X lo6. "Sedimentation coefficient in a KBr solvent with a density of 1.20 glml, in a solvent with the viscosity of 0.8534 centipoise (25°C).T h e negative sign indicates flotation. T h e flotation coefficient in a NaCl solvent with a density of 1.063 g/ml and viscosity of 1.0260 centipoise (26°C). 'The carbohydrate composition was assumed to be 6.5% by weight of the protein concentration.

222


density may be computed. Given the density and radius, the sedimentation coefficient, s, also may be computed from the following expression:

where M is molecular weight, qsOlv is the solvent viscosity, dl, is the lipoprotein density, N is Avogadro's number, dsolvis the solvent density, R is the lipoprotein radius, and f/fi is the translational frictional ratio. For a spherical, anhydrous particle, the frictional ratio is 1.00; however, LDLs are hydrated, and the frictional ratio, determined through diffusion measurements, is 1 . 1 1 (Fisher et al., 1971). This value was used in calculating the sedimentation coefficients listed in Table 11. The lipoprotein density that should be employed in Eq. (2) is the buoyant density, which is the density at which the sedimentation coefficient is equal to zero. T h e buoyant density was obtained experimentally by measuring the sedimentation coefficient at several solvent densities and extrapolating to zero. Kahlon et al. (1982) have shown that the buoyant densities, which they call the u densities, vary with the rotor speed of the centrifuge, reflecting the different compressibilities of water and lipid. In order to convert the lipoprotein density, as determined from its composition, to the buoyant density at a rotor speed of 52,640, the data of Kahlon et al. (1982) were used calculate a correction factor of 0.0016 g/ml, which was added to the compositional densities. T h e values of buoyant densities listed in Table I1 have been calculated by adding 0.0016 g/ml to the density values determined from their compositions. Utilizing the buoyant densities listed in Table I1 and the frictional ratio of 1 . 1 1, values for the sedimentation coefficient have been calculated at two different solvent densities: the standard S,"value routinely used to characterize human serum lipoproteins is defined as value of the flotation coefficient in Svedberg units (the negative sedimentation coefficient x l o p i 3 sec) in an aqueous NaCl solvent with a density of 1.063 g/ml and a viscosity of 1.021 centipoise (the viscosity of a 1.063 g/ml sodium chloride solution at 26°C). These values are listed in Table 11. The S," value is very sensitive to small variations in lipoprotein density because the solvent density is close to the lipoprotein density. T o compare the particle sizes or molecular weights, values of the sedimentation coefficient (s)in a solvent with a density of 1.20 g/ml and the viscosity of KBr at 25°C are preferred, and the computed values are listed in Table 11.

TABLE I11 Experimentul Physical Properties and Composition of Pooled Human LDL" Composition in weight percent Fraction 4 5 6 7 8 9 10 11

12 13 14

Gradient density ( g W

Buoyant densityb ( g W

1.0234 1.0260 1.0286 1.0314 1.0343 1.0372 1.0409 1.0451 I .0502 I .0580 1.OM0

1.0172 1.0196 1.0224 1.0256 1.0289 1.0318 1.0358 1.0404 1.0460 1.0545 1.0632

Sf 9.30 8.50 7.60 6.70 5.90 5.20

CE

TG

PL'

C

Proteind

Carbohydrate'

41.50 41.81 41.61 43.06 43.62 43.43 42.98 4 1.06 39.5 1 40.08 40.95

6.3 1 5.49 4.90 3.77 3.57 3.78 3.89 3.30 3.10 2.27 2.59

22.75 22.64 23.14 22.1 1 22.24 20.88 2 1.35 20.63 22.08 2 1.32 20.57

10.99 11.60 10.71 11.81 11.21 10.93 9.72 9.70 8.77 9.40 9.20

17.33 17.33 18.44 18.08 18.18 19.70 20.71 23.77 24.93 25.28 25.07

1.13 1.13 1.20 1.18 1.18 1.28 1.35 1.54 1.62 1.64 1.63

"Experimental values were taken from Chapman et al., (1988) and adjusted as described in footnotes c and d . 'Buoyant densities were determined from a plot of gradient density versus buoyant densities (udensities), constructed using the appropriate values from Kahlon et al. (1982). 'PL values listed by Chapman et al. (1988)were divided by 0.93 to adjust for the phospholipid not accounted for by the phosphocholine-specific assay used by these workers. dLowry values listed by Chapman et al.(1988)were multiplied by a color correction factor of 0.80 wt apoB/wt BSA, to adjust their Lowry values for the BSA standard. 'The carbohydrate composition was assumed to be 6.5% by weight of the protein.

-2

50 4

+

48-v

v, l-

z

W

46--

0

Z 0

44-

a @a

0 0

42-0

5 W

0

@ a

0

--

40-

0 K

381

r

n *

25 1-

6 IY a L

2

W

O

231

a

a a0

1

ma

21 19/

a

1157 ; Y ( 1.01 0 1.020

1.030

1.040

1.050

1.060

BUOYANT DENSIlY (g/rnl)

FIG.4. A comparison of the observed and calculated sedimentation and compositional were taken from Table 111; the calculated properties of LDLs. The observed properties (0) properties ( 0 )are from Table 11. (A) A comparison of core lipids, defined as the sum of the percentages of cholesteryl esters and triglycerides. (B) A comparison of the percentages of protein. (C) A comparison of the flotation coefficients in a solvent of density 1.063 g/ml (SP values).

225


C

" E I-

z w 0 L LL

w

0 0 Z

0

F

s

1.010

1.020 1.030 BUOYANT DENSITY (g/rnl)

1.040

FIG. 4. (Continued)

How well do the sedimentation coefficients and densities predicted by the model match the values actually observed for LDL? Excellent agreement with the experimental points is shown by the solid curve of Fig. 2, which is a plot of the values for ~ 2 5 , 1 . 2 0given in Table 11. However, this agreement was achieved by selecting a value for the partial specific volume of the cholesteryl esters to make the best fit, yielding the value of 1.058 ml/g for this this quantity. [If a value of 1.044 ml/g were employed for the partial specific volume of the cholesteryl esters, as was used by Sata et al. (1972), the values of ~ 2 5 . 1 . 2 0 listed in Table I1 would have decreased by about 3.5%. The values of Sf in Table I1 would have dropped by 1 to 2 Svedbergs.] How well do the compositional values for the individual lipid components and protein and carbohydrate values predicted by Table I1 agree with experimental values? For this comparison, we have selected the experimental values for fractionated LDL published by Chapman et al. (1988). Before these values could be compared, it was necessary to make three adjustments to the experimental values. These workers employed an enzymatic phospholipid assay (Takayama et al., 1977) specific for phosphocholine-containing lipids, which included phosphatidylcholine, sphingomyelin, and lysophosphatidylcholine, making u p about 93% of LDL lipids (Skipskiet al., 1967). Therefore, the weight of phospholipid has been increased by a divisor of 0.93 in calculating the adjusted experimental values listed in Table 111. Second, Chapman and co-workers

226


(1988) employed the Lowry procedure to determine the apoB content of LDL, using bovine serum albumin as a standard. Such values must be multiplied by a color conversion factor to convert bovine serum albumin weights to apoB weights. Therefore, the weight of protein has been decreased by a factor of 0.80 g apoB polypeptide/l g serum albumin (Fisher and Schumaker, 1986; Margolis and Langdon, 1966b) to adjust the values reported by Chapman et al. (1988) to those reported in Table 111. Finally, Chapman el al. (1988) did not list the carbohydrate content of their LDL; we have assumed that the apoB 100 polypeptide contains 6.5% carbohydrate by weight, the average of the reported values, which vary from about 5 to 8%. The adjusted experimental values, converted to percentages of the total LDL weight, are listed in Table 111. Differences between the adjusted observed values of Table 111 and those calculated from the emulsion particle model in Table I1 fall within the experimental error limits reported by Chapman et al. (1988). This comparison is also provided in graphical form for total core lipids (triglycerides plus cholesteryl esters) (Fig. 4A), for protein (Fig. 4B), and for the values of the flotation coefficient, S,“(Fig. 4C). The agreement is seen to be excellent in all cases. We conclude that the emulsion particle model provides a satisfactory description of the composition, size, density, and hydrodynamic properties of normal human LDL isolated by density gradient centrifugation. IV. STRUCTURAL STUDIES OF APOLIPOPROTEIN B ON LOW-DENSITY LIPOPROTEIN SURFACES What is the configuration of apoB 100 on the surface of the LDL? Is it folded into a single domain, as suggested by the electron microscopy (EM) studies of Lee et al. (1987)? Is it composed of three domains, as suggested by the proteolysis studies of Chen et al. (1989), (Fig. lG); five domains, as suggested by the trypsin releasability studies of Yang el al. (1989) (Fig. 1F);or four domains, as suggested by low-angle X-ray studies (Luzzatti et al., 1979) and freeze-fracture electron microscopy (GulikKryzwicki el al., 1979)? Other EM studies have interpreted the structure of apoB on LDLs to be composed of 20 domains (Pollard et al., 1969) or “strand-like substructures, possibly forming a surface network” (Forte and Nichols, 1972). T o explore this question, three different approaches have been employed to map apoB on the surface of the LDL: (1) lipid extraction after attachment of the LDL to an EM grid, (2) mapping of apoB on the LDL surface with monoclonal antibodies, and (3)determination of how core circumference is related to apoB size during lipoprotein biosynthesis in cell culture.

APOLIPROTEIN B A N D LDL STRUCTURE

227

A . Lipid Extraction after Attachment of Low-Density Lipoproteins to Electron Microscope Grids The first of these techniques (Phillips and Schumaker, 1989) involved adsorbing the LDL to a carbon-coated electron microscope grid, extracting the lipid with 4: 1 ethanokether, which solubilized both polar and nonpolar lipids, and negative staining of the proteinaceous residue, which remained attached to the grid. After negative staining with uranyl acetate, the grid was examined in the electron microscope. Figure 5A shows the intact LDL absorbed to the grid prior to lipid extraction, and Fig. 5B shows the apoB 100 remaining after extraction. The protein (Fig. 5B) is seen to be a long, flexible structure, usually bent into a circular or semicircular configuration, with a diameter approximately equal to that of the LDL (Fig. 5A) from which is was obtained. If the LDLs were first treated with a cross-linking reagent such as glutaraldehyde (Fig. 5C) or water-soluble carbodiimide, then the apoB 100 remaining after extraction appeared as solid circles (Fig. 5D) with diameters equal to those of the LDLs from which they were derived. How were these results to be interpreted in terms of the configuration of apoB 100 on the surface of the LDL? The obvious interpretation was that apoB surrounds the LDL like a belt; moreover, it was also suggested that chemical cross-linking buckles the belt (Fig. 6) (Phillips and Schumaker, 1989). Other interpretations might be advanced; for example, if apoB covered the surface of the LDL like a hair net, then removal of the lipid with the organic solvent might cause a netlike apoB to settle into acircular pool, with much of the mass at the periphery, artifactually creating a circular belt from a structure that was not beltlike at all. Therefore, a second approach was required to test the beltlike model. For this purpose monoclonal antibodies were employed to map apoB 100, in the presence of lipid, on the surface of the LDL.

B . Mapping of Apolipoprotein B on Low-Density Lipoprotein Sulfaces with Monoclonal Antibodies

Electron microscopy of negative-stained native LDL samples does not resolve the distribution of apoB 100 on the lipoprotein surface. However, in electron micrographs of LDL-antibody complexes, antibodies, recognized as small, Y-shaped objects, have been observed protruding from the LDL particles (Chatterton et al., 1991). Therefore, monoclonal antibodies directed against apoB can be used to locate specific regions of apoB on the LDL surface. Because each LDL contains only a single copy

FIG. 5. LDL before and after lipid extraction when bound to the EM grid. After adsorption to the carbon-coated grid, lipid was extracted by brief immersion in ice-cold (4 : 1) ethanol :ether. Negative-stained images include (A) untreated LDL, (B) lipidextracted LDL, (C) glutaraldehyde-treated LDL, (D) lipid-extracted, glutaraldehydetreated LDL, (E) glutaraldehyde-treated LDL, (F) lipid-extracted, glutaraldehyde-treated IDL. Bar = O.1pm for A-F. (Phillips and Schumaker. 1989.)

AFQLIPROTEIN B AND LDL STRUCTURE

229

FIG. 6. ApoB100 is modeled as a belt surroundingthe LDL. The protein is assumed to be embedded in the monolayer, where the inner surface of the belt makes contact with the core lipids and the edges of the belt interact with the phospholipid andlor cholesterol. To fulfill volume requirements, the cross section of the belt would be about 20 x 54 8, and the average length about 585 8,. Cross-linkingwith a chemical reagent “buckles”the belt.

of apoB, and because apoB lacks any repetitive sequences, each LDL should possess only one site complementary to a given monoclonal antibody. As expected, when a single monoclonal antibody was added to LDL, a maximum of one antibody was observed attached to a single LDL. When two different monoclonal antibodies recognizing separate apoB epitopes were simultaneously added to the LDLs, occasionally both types of antibody were seen attached to the same LDL particle (Fig. 7). Visualization of bound antibody was a fortuitous occurrence, requiring that the LDL-antibody complex adsorb to the carbon-coated grid surface with an orientation such that both bound antibodies lay flat on the grid, displacing the thin layer of negative stain (Fig. 8). For a small percentage of the LDLs (1- 10%)this fortuitous orientation did occur, allowing both monoclonals to be visualized and permitting measurement of the angle, at the LDL center, separating the two recognized apoB epitopes (Fig. 7). In a few cases, LDL dimers were seen with two LDLs joined by two different monoclonal antibodies to form a circular complex. Again, the angles could be measured for both members of the pair. Accurate determination of the average angle between a given pair of monoclonal antibodies bound to the same LDL required multiple measurements. In each case, a wide spread of angles was obtained from which an average value could be calculated (Fig. 9). Why was such a large variation in the measured angle observed? Repeating the measurements on the same set of electron micrographs showed that the reading error

FIG. 7. Negative-stained images of monoclonal antibodies binding to apoBlOO on LDL. (A) An equimolar mixture of LDL and each of two anti-apoB100 monoclonal antibodies, MB47 and MB19, were adsorbed to the carbon-coated grid, stained with 1% uranyl acetate, and examined in the electron microscope. The insert shows both open (0) and closed (c) complexes. Bar = 0.1 pm. (B) Complexes formed between LDL and monoclonal antibodies MB47 and MB24. (From Chatterton etal., 1991.)

rI

FIG.8. Illustration of two monoclonal antibodies bound to a flattened LDL. If a sphere of diameter 208 8, was uniformly flattened, holding volume constant, until it had an apparent diameter of 285 8, in projection, it might resemble half of an oblate ellipsoid. A truncated oblate ellipsoid with a major axis of 285 8, and a semiminor axis of 11 1 A would have the same volume as a 208-8, sphere. Diameters of this size or larger were frequently observed. Monoclonal antibody 8, has bound to the LDL above the layer of negative stain, and would not be observed. Monoclonal antibody B has bound to the LDL in the plane of the EM grid, and has displaced the layer of negative stain, SO that it would be observed in the electron micrograph.


23 1

30 20 10

0

60 120 ANGLE (deg)

180

ANGLE (deg)

FIG. 9. Observed angular distributions of pairs of monoclonal antibodies binding to LDL. (A) The distribution of 124 angles measured between 4G3 and B3 yield an average angle of 104". (B) The distribution of 139 angles measured 4G3 and B4 yield an average angle of 60".

was small, about 23". Parallax could not explain this variation because the two antibodies both must have lain in the plane of the grid to be visualized. It was concluded that the spread of measured angles was real, and probably reflected an actual variation in the angle between epitopes caused by flexibility of the apoB embedded in a fluid surface monolayer. To determine the configuration of apoBlOO on the LDL surface, 10 different monoclonal antibodies have been employed (Table IV). In the original model (Chatterton et al., 1991), three antibodies (MBl 1, MB44, and anti-B,,, 16) were placed on a three-dimensional map relative to MB47, MB24, and MB19, which defined the North Pole, the prime meridian, and the handedness, respectively (Fig. 10). In the current model, angles between the first three antibodies (MB19, MB24, and MB 1 1) were used to establish a spherical coordinate system on the LDL surface. To minimize errors inherent in measuring angles between epitopes separated by extremely long distances along the apoB sequence, the next five antibodies (2D8, B4, B3, 4G3, and MB47) were placed on the map by triangulation relative to the three sites that immediately preceded them in the sequence (Chatterton et al., 1993). For example, monoclonal antibody B3 was placed on the map relative to MB 11, 2D8, and B4. The results are shown in Fig. 11. The monoclonal antibodies recognized epitopes that were welldistributed along the first 3500 amino acids of apoBl00. In the threedimensional model (Fig. 1l), the flags, which locate the sites recognized by the eight monoclonal antibodies, are connected by a string in the same order as they appear along the apoB 100 primary sequence. The model was superimposed on a globe and rotated such that antibodies MB19,

232


TABLE IV Location of Epitopes Recognired by Monoclonal Antibodies against Apolipoprotein'

Antibody MB19 MB24 MBI 1 2D8 B4 B3 4G3 MB47 Anti-B,, 16

Sourceb 1 1

1 2 3 3 2 1 2

Binding site location (apoB amino acid residue) 71 399-580 995-1082 1438- 1480 1827- 1943 2330-2376 2980-3084 344 1-3568 4 154-4189

"Determination of epitope positions along the apoB sequence has been reported previously by Knott et al. (1986), Milne et al. (l989), Young and Hub1 (1989), and Pease et al. (1990). bMonoclonalantibodies were obtained from (1) Dr. L. K. Curtiss. Scripps Research Institute, California; (2) Dr. Y. L. Marcel and Dr. R. W.Milne, Clinical Research Institute, Montreal; and (3) Dr. J.-C. Fruchart, The Institute Pasteur, Lille, France.

MB24, and MB 11 occupied approximately the same locations as they did in the original model (Chatterton et al., 1991). Thus, apoB runs from the N terminus near MB19 located on the coast of Colombia, 4" west of Bogota (5"N, 78"W); to MB24 above the Atlantic Ridge, 10" north of the equator (10"N, 36"W); to M B l l off the coast of Uruguay, 6" east of Buenos Aires (35"S, 52"W); to 2D8 off the coast of Antarctica near Enderby Land (62"S, 50"E); to B4 in Queensland, Australia (25"S, 142"E); to B3 in the Indian Ocean off the southern tip of India, near the Maldive Islands (2"N, 75"E); to 4G3 in the Pacific Ocean, near Johnston Island southwest of Hawaii (19"N, 172"W); and to MB47 in the Pacific Ocean, near the Island of Clarion off the western coast of Mexico (2 1"N, 119"W) (J. E. Chatterton, et al. 1994.) Th e most significant difference between this map and the original involves the position of MB47. In the original map, MB47 was placed at the North Pole 80" north of MB19. In the present map, MB47 is only 45.6" from MB19. This difference is believed to be due to flattening of the LDL spheres on the electron microscope grid, leading to an increase

APOLIPROTEIN B A N D LDL STRUCTURE

A

m h l (N. Pole)

B

mAbl

233

Prime Meridian

mAbl

Longitudes

FIG. 10. lnterpretation of mapping measurements. (A) The position where the first monoclonal antibody bound (mAbl) was defined as the North Pole. Therefore, the angle between mAbl and a second monoclonal antibody gave the latitude of the second antibody. (B) The position where mAb2 bound was defined as "Greenwich." Therefore, the great circle that passes through the North Pole (mAbl) and Greenwich (mAb2) was 0" longitude. (C) From the two angles measured between mAbl and mAb2 and any other monoclonal antibody, mAb3, both the latitude and longitude of the other monoclonal antibody were calculated. An ambiguity arose, because the third monoclonal antibody may be placed either in the Eastern or Western Hemisphere. This ambiguity was resolved by arbitrarily placing the third monoclonal antibody in the Western Hemisphere. Thereafter, additional monoclonals were located unambiguously by triangulation. The final map, however, may be a mirror image of the correct map.

in the radius of curvature of the lipid surface, causing the two ends of apoB to drift apart on the surface. This conformational change may reflect a similar change that apoB must undergo in order to exist on the surface of the much larger VLDL particles. In addition, MB44 was

234


FIG.1 1 . Mapping apoB100 on the surface of the LDL. The string between toothpicks connects the epitopes in order as they occur along the primary sequence of apoB (Table IV).

removed from the map due to the extremely broad distributions in measured angle always found with this antibody. Finally, anti-B,,116 has been temporarily removed from the map pending acquisition of additional data. Clearly, the first 3500 amino acids of apoB 100 nearly circumnavigate the globe. T h e gap of 45.6"separating MB47 and MB 19 could easily be spanned by the remaining 1000 amino acids between the site recognized by MB47 and the C terminus of apoB. Making the assumption that apoB is relatively uniform along its length locally, only 400 of the remaining


235

1000 amino acids would be required to complete the circumnavigation, placing amino acid residue 3900 somewhere near the N terminus of apoB. This leaves open the question of where the remaining approximately 600 C-terminal amino acids are located. Does apoB continue from residue 3900 in the same general direction past the N terminus, essentially retracing its own steps? Or does apoB double back, thus placing the C terminus somewhere near the site recognized by MB47 and the LDL receptor? These questions will be resolved as additional monoclonal antibodies between MB47 and the C terminus are placed on the map. Thus, the results of these mapping studies support the cross-linking and lipid extraction studies, and are consistent with a beltlike model for the structure of apoB 100 on the LDL surface. This proposed structure is further supported by a preliminary analysis of LDL in vitreous ice at 34 A resolution by electron microscopy (Atkinson, 1989, 1993)-480 LDL particles were examined to yield five independent sets containing 4- 12 images. “The independent views indicate that LDL is a semi-spherical, 200-220 A diameter particle, with an area of low density (lipid) surrounded by a ring (in projection) of high density believed to represent apolipoprotein B-100. This ring is seen to be composed of four or five (depending on view) regions of high density material that may represent protein superdomains linked by areas of somewhat lower density.” (Atkinson, 1993). Another independent approach to the mapping of apoB on the lipoprotein surface arose unexpectedly during studies on the mechanism of lipoprotein assembly in hepatocyte cell lines, and this will be described next. C . Relating Low-Density Lipoprotein Core Circumferences to Apolipoprotein B Fragment Sizes

A remarkable observation was made by Yao et al. (1991), who expressed a homologous series of C-terminal truncated apoB molecules in stably transfected McArdle 7777 cells, a rat hepatocyte cell line. Plotting the buoyant density of the secreted lipoproteins as a function of the logarithm of the number of amino acid residues, they noticed a linear relationship between these two quantities, implying that the lipoprotein density was inversely proportional to the apoB size. Evidently some fundamental architectural principle was being explored in their studies. A homologous series of C-terminal truncated apoBs in transiently transfected HepG2 cells was prepared by Spring et al. (1992a,b). These data, together with the data taken from Yao et a/. (1991), are presented in Table V, which lists the sizes of the newly expressed apoB molecules using the centile nomenclature of Kane et al. (1980), the observed

236


TABLE V Expression and Secretion of a Series of C-Tenninnlly Truncuted Fragmenfs in Hep-G2 and McArdlc 7777 Cells Cell Type HepG2

McArdle 7777

ApoB size

Densitv (g/ml)

Radiusb(A)

ApoB26 ApoB31 ApoB37 ApoB42 ApoB49 ApoB18 ApoB23 ApoB28 ApoB31 ApoB37 ApoB48 ApoB53 ApoB 100

1.163 1.143 1.136 1.103 1.101 1.23 1.23 1.17 1.17 1.14 1.10 1.06 1.006

43.7 48.7 51.3 57.3 59.5 35.0 37.9 44.2 45.6 50.9 60.0 69.7 104.0

"Experimentally observed density values for HepG2 (Spring et al., 1992b) and McArdle 7777 cells (Yao et al., 1991). "Calculated from the emulsion particle model, given the size of the apoB, the observed density, and the partial specific volumes of the constituent lipid, protein and carbohydrate, a 1 : 1 molar ratio of C to PL,a 25 :6 wt ratio of TG :CE (Thrift et al., 1986), and a shell thickness of 20.2 A.

buoyant density of the secreted lipoproteins, and the predicted lipoprotein radii as calculated assuming the emulsion particle model, as explained in the footnote to Table V. The values listed in Table V are plotted in Fig. 12. The lipoprotein radius is plotted as a function of apoB size, and the points, which include all of the data of Yao et al. (1991) and all of ours, define a single straight line. The existence of this linear relationship strongly suggests that these lipoproteins form a homologous series of emulsion particles; moreover, it provides an additional insight into the nature of the fundamental relationship underlying the structure of these lipoproteins, that is, lipoprotein radius is a linear function of apoB size. Two parameters may be obtained from a straight line, the slope and the intercept. The slope of the line shown in Fig. 12 is 0.166 A/kDa. The intercept is approximately 20 A, which is close to the thickness of the surface monolayer of phospholipid, cholesterol, and protein surrounding the core of the lipoprotein particle. This interesting relationship between lipoprotein radius and apoB size has been confirmed by a study of the sizes of the secreted lipoproteins formed after puromycin treatment of HepG2 cells. This antibiotic causes


237

1

0 0

2E5

4E5

6E5

MOLECULAR MASS OF APOB FRAGMENT FIG. 12. Lipoprotein radii as a function of apoB molecular size. Lipoprotein radii were determined from the measured buoyant densities and apoB fragment molecular masses according to the emulsion particle model (Table V). The straight line through the experimental points has a slope of 0.166hkDa apoB, and a vertical intercept of 19.7 A. A, From Yao el al. (1991); 0,from Spring el al. (1992b).

premature release of the growing polypeptide chain, creating a homologous series of C-terminally truncated apoB polypeptides translated from endogenous message. These truncated apoB polypeptides are used by the HepC2 cells to synthesize small lipoproteins, which they secrete into the medium and which may be isolated from the medium by flotation. Hydrodynamic characterization of these particles yields both flotation coefficients and densities (Table 11), from which the radii may be calculated (Spring et al., 1992a,b). Radii determined in this manner are listed in column 4 of Table VI, and can be seen to increase with the size of the apoB. These radii were obtained without assuming the emulsion particle model. The data of Table VI were plotted to yield the relationship between lipoprotein radius and apoB size (Fig. 13A). Here again, the relationship between the lipoprotein radius and apoB size was linear, with the points defining a straight line intercepting the vertical axis at a radial value of about 20 A, as seen previously for the case of the C-terminally truncated apoBs expressed by the transfected hepatocytes (Fig. 12). The vertical intercept of 20 A (Figs. 12 and 13A) was suggestive of the thickness of the monolayer in the emulsion particle model. In fact, if

238


TABLE Vl Radll of Puromycin-Generated Particles Calculated from Sedimentation Coefficients and Buoyant Densities" ApoB sizeb(centile)

Observed density (g/ml)

80 70 55 51 49 45 42 38 36 33 31 29 25

1.034 1.048 1.058 1.071 1.080 1.09I 1.094 1.104 1.109 1.125 1.144 1.150 1.170

(Svedberg)

5~0,1.2,1

-30.4 -20.1 -14.1 - 12.2 - 10.6 -7.78 - 7.07 -5.82 -5.34 -3.68 -2.51 - 1.83 -0.77

Radius'

(8)

96.1 81.6 70.8 69.1 66.6 60.0 58.0 55.3 54.3 49.7 47.5 42.9 36.0

"Springel al. (1992b). *ApoBsize is expressed in the centile nomenclature,which indicates the apparent molecular weight, as determined by SDS-PAGE as a percentage of the molecular weight of' the full-length apoB100. 'Calculated from the observed buoyant densities and sedimentation coefficients by Eq. (2).

the monolayer thickness, assumed to be 20.2 A, was subtracted from each radial value, the differences became the radii of the lipoprotein cores. In order to replot the data, core radii were multiplied by 27r, to yield core circumferences. It was also convenient to multiply the apoB sizes on the horizontal axis by 513 kDa, the approximate size of the full-length apoB 100 polypeptide. When lipoprotein core circumference was plotted as a function of apoB molecular weight, (Fig. 13B), the points defined a straight line, which now intercepted the vertical axis close to the origin. Th e slope of the line was 1.14 8, of core circumferencelkilodalton of apoB. These data demonstrate that lipoprotein core circumference is directly proportional to apoB molecular weight (Fig. 13B). What is the interpretation of this relationship in terms of molecular structure? Spring et al. (1992a,b) suggested that it is best explained by a beltlike model for apoB, because the circumference of any object is directly proportional to the mass of the belt that surrounds it. Thus, a third, independent approach has yielded a beltlike model for apoB on the surface of small emulsion particles.

A 1 100-

.,.

0

~

20

40

60

80

100

SIZE OF TRUNCATED APOB (PERCENT OF APOBl00)

B

MOLECULAR WEIGHT OF TRUNCATED APO~B(MW x 10-3)

FIG. 13. Lipoprotein size as a function of apoB size. (A) Lipoprotein radius, in di units, is plotted as a function of the length of the apoB polypeptide, in centile units. These data, taken from Table VI, were obtained from the spectra of different sizes of lipoproteins and apoBs secreted by pulse-labeled HepG2 cells after puromycin treatment and analyzed as shown in the previous figure. The straight line through the points has an intercept of 18.8 di and a slope of 0.946 k e n t i l e . (B)Lipoprotein core circumference, in A units, is plotted as a function of the molecular weight of the C-terminally truncated apoB polypeptide. These are the same data as plotted in A, lipoprotein core circumference was calculated as 2a(R - t), where R is lipoprotein radius, and t = 20.2 A, the thickness of the shell.

240

VERNE N. SCHUMAKER ET AL

Why does apo B form a beltlike structure on the surface of these small lipoproteins? What is the functional significance of this architectural design? To answer these questions, it has been suggested that during lipoprotein formation, apoB functions by nucleating and determining the size of the oil droplet that pinches off from the bilayer (Spring et al., 1992b).

V. LIPOPROTEIN ASSEMBLY Cotranslational Lipoprotein Formation in Rough Endoplasmac Reticulum ApoB is synthesized in the rough endoplasmic reticulum (RER), where it was first detected by immunoelectron microscopy (Alexander et al., 1976). In vitro translation experiments utilizing rabbit reticulocytes or wheat germ extracts and dog pancreatic microsomes have shown that the apoB is inserted into the lumen of the ER (Chuck and Lingappa, 1992), and that once the insertion is completed, the apoB appears to be associated with the inner leaflet of the bilayer (Pease et al., 1991). In the in vitro translation systems employed in these experiments, the dog pancreatic microsomes were probably not synthesizing appreciable quantities of phospholipid or triglycerides, and therefore did not incorporate apoB into lipoproteins. In contrast to the in vitro results, Boren et al. (1992) have shown that lipoprotein biosynthesis occurred cotranslationally in HepG2 cells, that is, while the C-terminal portion of apoB was still being synthesized on the ribosome, the N-terminal portion was already incorporated into a small lipoprotein. Figure 14 illustrates the formation of a small lipoprotein particle on the luminal surface of the endoplasmic reticulum. The growing polypeptide chain extends from the ribosome through a membrane channel, and is incorporated into the inner leaflet of the ER membrane. Here, the apoB folds into a continuous strip about 50 8, wide and many hundreds of angstroms long, displacing the phospholipid monolayer to each side. We speculate that the embedded hydrophobic surface of the apoB nucleates the formation of an oil droplet from the supersaturated bilayer, which bulges out into the luminal space because of the presence of the apoB in the inner leaflet of the bilayer. This process is imagined to continue, with more triglyceride added to the growing oil droplet and the apoB growing in length until translation is completed. At this point, the C-terminal end of the apoB polypeptide is released from the ribosome and is free to surround the oil droplet. A.


24 1

FIG. 14. Cotranslational assembly of a lipoprotein from the inner leaflet of the ER bilayer and apoB. In this model, translation of the C-terminal portion of apoB proceeds on a membrane-bound ribosome, while translocation and lipoprotein assembly occur on the luminal side of the ER. The N-terminal portion of apoB is believed to be embedded in the inner leaflet of the bilayer, where it nucleates the formation of an oil droplet from the supersaturated ER membranes. As the hydrophobic inner surface of apoB attempts to surround the oil droplet, it bulges into the lumen, as depicted here. On completion of translation, the two ends of apoB become free to meet, which would automatically result in the detachment of the lipoprotein from the bilayer. (Not drawn to scale.)

242


How is the nascent lipoprotein released from the surface of the ER? This probably would occur spontaneously if the two ends of the apoB were to meet, or come close together. Thus, the phospholipid monolayers that form both hemispheres of the surface coat of the nascent lipoprotein particle are attached by noncovalent forces to the sides of apoB molecule. When the two ends of the apoB touch, continuity be-

SER Lipids

Frc. 15. One version of the two-step model for lipoprotein biosynthesis. This model (Alexander et al., 1976) proposes that VLDL-sized emulsion particles lacking apoB were synthesized in the smooth ER (SER) and that these particles subsequently migrated to the junction between the smooth and the rough ER (RER), where the apoB was incorporated into the surface monolayer of the nascent VLDL. In this review it is suggested that the primary lipoprotein, shown in Fig. 14, is the vehicle that transports the apoB to the emulsion particle and merges with it to complete the assembly of the VLDL. Reproduced from theJouml of Cell Biology (Alexander et a/., 1976), by copyright permission of the Rockefeller University Press.


243

tween the bilayer of the ER and the monolayer surrounding the lipoprotein is automatically broken, and the particle becomes detached from the bilayer. It should be emphasized that according to this model, apoB does not choke the lipoprotein like a noose around the neck of a hanged man, but rather surrounds the lipoprotein along a great circle running from the North Pole to the South Pole. The circle becomes complete when the growing polypeptide is released from the ribosome. Completing the circle automatically separates the lipoprotein from the inner leaflet of the bilayer forming the membrane of the endoplasmic reticulum. B . Two-step Model for Assembly of Triglyceride-Rich Lipoproteins The permanent hepatocyte cell lines that have been studied, HepG2 and McArdle 7777 cells, are defective and do not make the large VLDLs characteristicof normal liver. Instead, they secrete the small, triglyceriderich lipoproteins described in the previous section of this review. We will call these prhury lipoprotein particles. We have suggested (Spring et al., 1992b)that lipoprotein formation is a two-step process, and that the first step is the elaboration of these small primary lipoprotein particles. The second step probably occurs in the smooth ER, where the large, VLDL-sized particle formed does not contain apoB. We will call these the secondar lipoprotein particles. Evidence for the second step was described by Alexander et al. (1976)in an electron microscope study following immunocytochemical staining to localize apoB within various cellular organelles of rat liver hepatocytes. They proposed that VLDL-sized emulsion particles lacking apoB were synthesized in the smooth ER, and that these particles subsequently migrated to the junction between the rough and the smooth ER where they acquired apoB (Fig. 15).A plausible mechanism for the acquisition of apoB would be through coalescence of the primary and secondary lipoproteins to form the nascent VLDL, which would now contain the apoB required for their transfer to the Golgi apparatus and subsequent secretion.

VI. SUMMARY ApoB 100 is a very large glycoprotein essential for triglyceride transport in vertebrates. It plays functional roles in lipoprotein biosynthesis in liver and intestine, and is the ligand recognized by the LDL receptor during receptor-mediated endocytosis. ApoB 100 is encoded by a single gene on chromosome 2, and the message undergoes a unique processing event to form apoB48 message in the human intestine, and, in some

244


species, in liver as well. The primary sequence is relatively unique and appears unrelated to the sequences of other serum apolipoproteins, except for some possible homology with the receptor recognition sequence of apolipoprotein E. From its sequence, structure prediction shows the presence of both sheet and helix scattered along its length, but no transmembrane domains apart from the signal sequence. The multiple carbohydrate attachment sites have been identified, as well as the locations of most of its disulfides. ApoB is the single protein found on LDL. These lipoproteins are emulsion particles, containing a core of nonpolar cholesteryl ester and triglyceride oil, surrounded by an emulsifying agent, a monolayer of phospholipid, cholesterol, and a single molecule of apoBl00. An emulsion particle model is developed to predict accurately the physical and compositional properties of an LDL of any given size. A variety of techniques have been employed to map apoB 100 on the surface of the LDL, and all yield a model in which apoB surrounds the LDL like a belt. Moreover, it is concluded that apoBlOO folds into a long, flexible structure with a cross-section of about 20 X 54 A' and a length of about 585 A. This structure is embedded in the surface coat of the LDL and makes contact with the core. During lipoprotein biosynthesis in tissue culture, truncated fragments of apoBlOO are secreted on lipoproteins. Here, it was found that the lipoprotein core circumference was directly proportional to the apoB fragment size. A cotranslational model has been porposed for the lipoprotein assembly, which includes these structural features, and it is concluded that in permanent hepatocyte cell lines, apoB size determines lipoprotein core circumference. ACKNOWLEDGMENTS Supported by Research Grants GM 13914 and HL 28481 from the National Institutes of Health.

REFERENCES Adams. G. H., and Schumaker, V. N. (1969). Ann. N. Y.Acad. Sci. 164, 130-146. Aggerbeck, L. P., Dezdy, F. J.. and Scanu, A. M. (1976).J. Biol. Chem.251,3823-3830. Alexander, C., Hamilton, R. L., and Havel, R. J. (1976).J. Cell Biol. 69,24 1-263. Atkinson, D. (1989). Bi0phys.J. 55, A208 (Abstr. Tu-AM-D~). Atkinson, D. (1993). Bi0phys.J. 64, A286 (Abstr. W-Pos 202). Baker, M. E. (1988).Bioch.J. 255,1057-1060. Banaszak, L. J., Sharrock, W., and Timmins, P.(1991). Ann. Rev. Biophys. Biophys. Chem. 20, 22 1-246. Blackhart, B. D., Ludwig, E. H., Pierotti, V. R., Caiati, L., Onasch, M. A., Wallis, S . C., Powell, L., Pease, R., Knott, T. J., Chu, M.-L., Mahley, R. W., Scott, J., McCarthy, B. J., and Levy-Wilson, B. (1986).J. Biol. C h . 261, 15364-15367. Boenvinkle, E., and Chan, L. (1989). Nucleic Acid Res. 17,4003.


245

Boerwinkle, E., Xiong, W., Fourest, E., and Chan, L. (1989).Proc. Natl. Acad. Sci. U.S.A. 86,

2 12-2 16. Boren, J., Graham, L., Wettesten, M., Scott,J., White, A., and Olofsson, S.-0. (1992).J . Biol. Chem. 267,9858-9867. Bradley, W. A.. and Gotto, A. M., Jr. (1978).In “Disturbances in Lipid and Lipoprotein Metabolism” (J.M. Dietschy,A. M. Gotto, Jr., and J. A. Ontko, eds.), pp. 1 1 1-137. Am. Physiol. SOC.,Bethesda, MD. Brooks, A. R., and Levy-Wilson, B. (1992).Mol. Cell. Biol. 12, 1134-1 148. Bruns, G. A. P., Karathanasis, D. K., and Breslow, J. L. (1984).Arteriosclerosis (Dallas)4,

97- 102. Butler, R. (1990).In “DNA and Other Polymorphisms in Forensic Science” (H. C. Lee and R. E. Gaenssler, eds.), pp. 135-159. Mosby, St. Louis, MO. Cann, H., and Guyer, M. (1992).Science 258,67-86. Carlsson, P., Darnfors, C.. Olofsson, S O . , and Bjursell, G. (1986).Gene 49,29-51. Chapman, M. J., Goldstein, S., Mills, G. l.,Lagrange, D., and Taylaur, C. E. (1978).Eur. J. Biochem. 87,475-488. Chapman, M. J., Laplaud, P. M., Luc, G., Forgez, P., Bruckert, E., Goulinet, S., and Lagrange, D.(1988).J . Lipd Res. 29,442-458. Chatterton, J. E., Phillips, M. L., Curtiss, L. K.. Milne, R. W., Marcel, Y. L., and Schumaker, V. N. (1991).J.Biol. Chem. 266,5955-5962. Chatterton, J. E. et al. (1994).In preparation. Chen, G. C., Zhu, S., Hardman, D. A., Schilling, J. W., Lau, K., and Kane, J. P. (1989). J. Biol. Chem. 264, 14369-14375. Chen, S.-H., Yang, C. Y.,Chen, P.-F.,Setzer, D.,Tanimura, M., Li, W.-H.,Gotto,A. M., Jr., and Chan, L. (1986).J.Biol. C k m . 261, 12918-12921. Chen, S.-H., Habib, G., Yang, C. Y., Gu, 2.-W., Lee, B. R., Weng, S., Silberman, S. R., Cai, S.-J., Deslypere,J. P., Rosseneu, M., Gotto, A. M., Jr., Li, W.-H., and Chan, L. (1987). Science 238,363-366.

Chuck, S. L., and Lingappa, V. R. (1992).Cell (Cambridge, Mass.) 68,9-21. Cladaras, C., Hadzopoulou-Cladaras, M.. Nolte, R. T., Atkinson, D., and Zannis, V. 1. (1986).E M B O J . 5,3495-3507. Coleman, R. D., Kim, T. W., Gotto, A. M., Jr., and Yang, C.-Y. (1990).Biochim. Biophys. Acta

1037, 129-132. Das, H. K., Leff, T., and Breslow,J. L. (1988).J.Biol. Chem. 263, 11452-11458. Davidson, N. W., Powell, L. M., Wallis, S. C., and Scott, J. (1988).J. Biol. Chem. 263,

13482-13485. Deckelbaum, R.J., Shipley, G. G., and Small, D. M. (1977).J.Eiol. Chon. 252, 744-754. De Loof, H., Rosseneu, M.,Yang, C.-Y., Li, W.-H., Gotto, A. M., Jr., and Chan, L. (1987). J . Lipid Res. 28, 1455-1465. Demmer, L. A., Levin, M. S., Elovson, J., Reuben, M. A., Lusis, A. J., and Gordon, J. I. (1986).Proc. Natl. Acad. Sci. U.S.A. 83, 8102-8106. Dunning, A. M., Renges, H. H., Xu, C. F., Peacock, R., Brasseur, R., Laxer, G., Tikkanen, M. J,, Butler, R., Saha, N., and Hamsten, A. (1992).Am.J. Hum. Genet. 50,208-221. Fisher, W. R., and Schumaker, V. N. (1986).In “Methods in Enzymology” (J. Segrest and J. Albers, eds.). Vol. 128,pp. 247-272. Academic Press, Orlando, FL. Fisher, W. R., Granade, M. E., and Mauldin, J. L. (1971).Eiochonktty 10, 1622-1629. Fisher, W. R., Hammond, M. G., Mengel, M.C., and Warmke, G. L. (1975).Proc. N a d Acad. Sci. U.S.A. 72,2347-235 1. Fojo, S. S., Law, S. W., Brewer, H. B., Jr., Sakaguchi, A. Y., and Naylor, S. L. (1984). Biockm. Biophys. Res. Commun. 112,687-693. Forte, T. M., and Nichols, A. V. (1972).Adu. LipidRes. 10, 1-41.

246


Garcia, Z. C., Paksay, K. S., Bostrom, K., Johnson, D. F., Balestra, M. E., Shechter, I., and Innerarity, T. L. (1992).Arlm’osck?rosisand Thrumb. 12, 172-179. Goldstein, J. L.,and Brown, M. S. (1989).In “The Metabolic Basis of Inherited Disease” (C. R. Scriver, A. L. Beaudet, W. S. Sly, and D. Valle, eds.), 6th ed., Vol. 1, pp. 1215-1250.McGraw-Hill, New York. Goldstein, S., Chapman, M. J., and Mills, G. L. (1977).Atherosclerosis (Shannon,Irel.) 28,

93-100. Gulik-Kryzwicki, T., Yates, M., and Aggerbeck, L. P. (1979).J.Mol. Biol. 131,475-484. Haberland, M. E.,and Reynolds, J. A. (1973).Proc. Natl. Acad. Sci. U.S.A. 70, 2313-

2316. Hauser, H., and Irons, L. (1972).Hoppe-S+r’s 2.Physiol. Chem. 353, 1579-1590. Higuchi, K., Monge, J. C., Lee, N., Law, S. W., and Brewer, H. B., Jr. (1987).Biochem. Biophys. Res. Commun. 144, 1332-1339. Hospattankar, A. V., Higuchi, K., Law, S. W., Meglin, N., and Brewer, H. B. (1987). Biochem. Biophys. Res. Commun. 148,279-285. Huang, L.-S., Miller, P. A., Bruns, G. A. P., and Breslow, J. L. (1986).Proc. Nutl. Acad. Sci. U.S.A. 83,644-648. Huang, L.-S., Ripps, M. E., and Breslow, J. L. (199O).J.LiPidRes. 31,71-77. Ichio, I., Deguchi, K., Kawashima, S., Endo, S., and Ueta, N. (1978).J.Biochem. (Tokyo) 84,

737-749. Jackson, C. L., Bruns, G . A.. and Breslow, J. L. (1984).Proc. Natl. Acad. Sci. U.S.A. 81,

2945-2949. Jeanpierre, M., Weil, D., Hors-Cayla, M. C., Williamson, R., Junien, C., and Humphries, S . E.(1984).Somatic Cell Mol. Genet. 10,645-649. Kahlon, T. S., Adamson, G. L., Shen, M. M. S., and Lindgren, F. T. (1982).Lipids 17,

323-330. Kane, J. P., Hardman, D. A., and Paulus, H. E. (1980).Proc. Natl. Acad. Sci. U.S.A. 77,

2465-2469. Kirchgessner, T. G., Heinzmann, C., Svenson, K. L., Gordon, D. A., Nicosia, M., Lebherz, H. G., Lusis, A. J., and Williams, D. L. (1987).Gene 59,241-251. Knott, T. J., Pease, R. J., Powell, L. M., Wallis, S. C., Rall, S. C., Jr., Innerarity, T. L., Blackhart, B., Taylor, W. H., Marcel, Y.,Milne, R., Johnson, D., Fuller, M., Lusis, A. J., McCarthy, B. J., Mahley, R. W., Levy-Wilson, B., and Scott, J. (1986).Nature (London)

323,734-738. Krauss, R. M., and Burke, D. J. (1982).J.Lipid Res. 23,97-104. Laggner, P., Kostner, G. M., Rakusch, U., and Worchester, D. (1981).]. Biol. C h . 256,

I 1832- 1 1839. Law, A.,andScott, J. (199O).J.LipidRes. 31, 1109-1120. Law, S. E.,Grant, S. M., Higuchi, K., Hospattankar, A.. Lackner, D., Lee, N., and Brewer, H. B., Jr. (1986).Proc. Natl. Acad. Sci. U.S.A. 83,8142-8146. Lee, D. M., Stiers, D. L., and Mok, T. (1987).Biochem. Biophys. Res. Commun. 144,210-216. Lee, P., and Breckenridge, W. C. (1976).Can.J. Biochem. 54,42-49. Levy-Wilson, B., and Fortier, C. (1989).J.Biol. Chem. 264,21196-21204. Levy-Wilson, B., Soria, L., Ludwig, E. H., Argyres, M., Brooks, A. R., Blackhart, B. D., Friedl, W., and McCarthy, B. J. (1991).J.LifndRes. 32, 137-145. Ludwig, E. H., Blackhart, B. D., Pierotti, V. R., Caiati, L., Fortier, C., Knott, T., Scott, J,, Mahley, R. W., Levy-Wilson, B., and McCarthy, B. J. (1987).D N A 6,363-372. Ludwig, E. H., Friedl, W., and McCarthy, B. J. (1989).Am.J. Hum. Genet. 45,458-464. Lund-Katz, S., Ibdah, J. A., Letiza. J. Y., Thomas, T., and Phillips, M. C. (I988).J.Biol.

C h .263,13831-13838.


247

Lund-Katz, S., Innerarity, T. L., Arnold, K. S., Curtiss, L. K., and Phillips, M. C. (1991). J . Biol. Chem. 226,2701-2704. Luzzati, V., Tardieu, A., and Aggerbeck, L. P. (1979).J. Mol. Biol. 131,435-473. Ma, Y., Schumaker, V. N., Butler, R., and Sparkes, R. S. (1987). Artrriosclerosis ( D a l h ) 7, 301-305. Ma, Y., Wang, X., Butler, R., and Schumaker, V. N. (1989). Arteriosclerosis ( D a l h ) 9, 242-246. Maeda, N., Ebert, D. L., Doers, T. M., Newman, M., Hasler-Rapacz, J., Attie, A. D., Rapacz, J., and Smithies, 0. (1988).Gene 70, 213-229. Mahley, R. W., and Holcombe, K. S. (1977).J . LipidRes. 18,314-324. Margolis, S., and Langdon, R. G. (1966a).J.Biol. Chem. 241,469-476. Margolis, S . , and Langdon, R. G. (1966b).J.Biol. Chem. 241,485-493. Marsh, J. B. (1968).Biol. Bull. (Woods Hole, Mass.) 135, 193-199. Matsumoto, A., Hiroyuki, H., Shibasaki, Y., Kodama, T., Fakaku, F., a n d Itakura, H. (1987).Biochem. Biophys. Res. Commun. 144,92-99. Mehrabian, M., Sparkes, R. S., Mohandas, T., Klisak, I. V., Schumaker, V. N., Heinzmann, C., Zollman, S., Ma, Y., and Lusis, A. J. (1986). S m t k Cell Mol. Genet. 12,245-254. Mill, G. L., and Taylaur, C. E. (1978). Protides Biol. Fluidc 25,477-482. Miller, K. W., and Small, D. M. (1987).In “Plasma Lipoproteins” (A. M. Gotto, Jr., ed.), pp. 1-75. Elsevier (Biomed. Div.), New York. Milne, R., Theolis, R., Jr., Maurice, R., Pease, R. J., Weech, P. D., Rassart, E., Fruchart, J.-C., Scott, J., and Marcel, Y. L. (1989).J . Biol. Chem. 264, 19754-19760. OhUigin, C., Chan, L., and Li, W.-H. (1990).Mol. Biol. Euol. 7,327-339. Olofsson, S.-O.,Bjursell, G., Bostrom, K.. Carlsson, P., Elovson, J., Protter, A. A., Reuben, M. A., and Bondjers, G. (1987).Atherosclerosis (Shannon,Irel.) 68, 1-17. Paulweber, B., and Levy-Wilson, B. (1991).J . Biol. Chem. 266,24161-24168. Paulweber, B., Onasch, M. A,, Nagy, B. P., and Levy-Wilson, B. (1991).J.Biol. Chem. 266, 24 149-24 160. Pease, R. J., Milne, R. W.. Jessup, W. K., Law, A., Provost, P., Fruchart, J. C., Dean, R. T., Marcel, Y. L., and Scott, J. (1990).J . Biol. Chem.265,553-568. Pease, R. J., Harrison, G. B., and Scott, J. (1991). Nature (London)353,448-450. Perez, L. E., Fenton, M. J., and Callard, I. P. (1991). Comp. Biochem. Physiol. B 100B, 82 1-826. Perutz, M. F., Kendrew, J. C., and Watson, H. C. (1965).J. Mol. Biol. 13,669-678. Phillips, M. L., and Schumaker, V. N. (1989).J.LipzdRes. 30,415-422. Pollard, H., Scanu, A. M., and Taylor, E. W. (1969). Proc. Natl. Acad. Sci. U.S.A. 64, 304-310. Powell, L. M., Wallis, S. C., Pease, R. J., Edwards, Y. H., Knott, T.J., and Scott,J. (1987).Cell (Cambridge, Mass.) 50,831-840. Raag, R., Appelt, K., Xuong, N. H., and Banaszak, L. J. (1988).J. Mol. Biol. 400, 553569. Rapacz, J., Chen, L., Butler-Brunner, E., Wu, M.-J., Hasler-Rapacz, J. O., Butler, R., and Schumaker, V. N. (1991). Proc. Natl. Acad. Sci. U.S.A. 68, 1403-1406. Reuben, M. A., Svenson, K. L., Doolittle, M. H., Johnson, D. F., Lusis, A. J., a n d Elovson, J. (1988).J.LipdRes. 29, 1337-1347. Sasak, W. V.,Lown, J. S., and Colburn, K. A. (1991).Bi0chem.J. 474, 159-165. Sata, T., Havel, R. J., and Jones, A. L. (1972).J. L i e Res. 13,757-768. Schumaker, V. N. (1973). Acc. Chem. Res. 6,398-403. Schumaker, V. N., and Puppione, D. L. (1986).In “Methods in Enzymology” (J. Segrest and J. Albers, eds.), Vol. 128, pp. 155-170. Academic Press, Orlando, FL.

248


Segrest, J. P., Jones, M. K., De Loof, H., Brouillette, C. G., Venkatachalapathi, Y. V., and Anantharamaiah, G . M. (1992).J. LipidRes. 33, 141-166. Shen, B. W., Scanu, A. M., and Kezdy, F. J. (1977).Proc. Natl. Acad. Sci. U.S.A. 74,837-841. Shireman, R. B., and Fisher, W. R. (1979). Biochim. Biophys. Acta 572, 532-540. Skipski, V. P., Barclay, M., Barclay, R. K., Fetzer, V. A., Good, J. J., and Archibald, F. M. (1967). Biochem.J. 104, 340-352. Small, D. M. (1986). Handb. LipldRes. 4,368, 512. Soria, L. F., Ludwig, E. H., Clarke, H. R. G.,Vega, G . L., Grundy, S. M., and McCarthy, B. J. (1989). Proc. Natl. Acad. Sci. U.S.A. 86,587-591. Spring, D. J., Lee, S.-M., Puppione, D. L., Phillips, M. L., Elovson, J., and Schumaker, V. N. (1992a).J.Lipid Res. 33,233-240. Spring, D. J., Chen-Liu, L. W., Chatterton,J. E., Elovson, J., and Schumaker, V. N. (1992b). J . Biol. Chem. 267, 14839-14845. Takayama, M., Itoh, S., Nagasaki, T., and Tanimizu, I. (1977).Clin. Chim. Acta 79,93-98. Taniguchi, T., Ishikawa, Y., Tsunemitsu, M., and Fukuzaki, H. (1989). Arch. Biochem. Biophys. 273, 197-205. Thrift, R. N., Forre, T. M., Cahoon, B. E., and Shore, V. G . (1986).J.LipidRes. 27,236-250. Thrift, R. N., Drisko, J., Dueland, S., Trawick, J. D., and Davis, R. A. (1992).Proc. Natl. Acad. Sci. U.S.A. 89,9161-9165. Timmins, P. A., Poliks, B., and Banaszak, L. J. (1992). Science 257,652-655. Triplett, R. B., and Fisher, W. R. (197Q.J. Lipid Res. 19,478-488. Vauhkonen, M. (1986). Glycoconjugate 3 , 3 5 4 3 . Wagener, R., Pfitzner, R., and Stoffel, W. (1987).Biol. Chem. Hoppe-Seyler 368,419-425. Wang, X., Schlapfer, P., Ma, Y., Butler, R., Elovson, J., and Schumaker, V. N. (1988). Arteriosclerosis (Dallas) 8,429-435. Wilson, C., Wardell, M. R., Weisgraber, K. H., Mahley, R. W., and Agard, D. A. (1991). Science 252, 1817-1822. WU. M.-J., Butler, E., Butler, R., and Schumaker, V. N. (1991). Arteriosclerosis Thromb. 11, 379-384. Xu, C., Nanjee, N., Tikkanen, M. J., Huttunen, J. K., Pirjo, P., Butler, R., Algelico, F., Del Ben, M., Mazzarella, B., Antonio, R., Miller, N. G., Humphries, S., and Talmud, P. J. (1989).Hum. Genet. 82,322-326. Yang, C.-Y., Chen, S.-H., Gianturco, S. H., Bradley, W. A., Sparrow, J. T., Tanimura, M., Li, W.-H., Sparrow, D. A., DeLoof, H., Rosseneu, M., Lee, F A . , Gu, 2.-W., Gotto, A. M., Jr., and Chan, L. (1986). Nature (London) 323,738-742. Yang, C.-Y., Gu, 2.-W., Weng, S.-A., Kim, T. W., Chen, S.-H., Pownall, H. J., Sharp, P. M., Liu, S.-W., Li, W.-H., Gotto, A. M., Jr., and Chan, L. (1989).Arteriosclerosis (Dallas) 9, 96- 108. Yang, C.-Y., Kim, T. W., Weng, S.-A., Lee, B., Yang, M, and Gotto, A. M., Jr. (1990). Proc. Natl. Acad. Sci. U.S.A. 87,5523-5527. Yao, Z., Blackhart, B. D., Linton, M. F., Taylor, S. M., Young, S. G., and McCarthy, B. J. (1991).J. Biol. Chem. 266,3300-3308. Yeagle, P. L., Martin, R. B., Pottenger, L., a n d Langdon, R. G . (1978). Biochemistry 17, 2707-27 10. Young, S. G., and Hubl, S. T. (1989).J. Lipid Res. 30, 443-449. Young, S. G., Bertics, S. J., Scott, T. M., Dubois, B. W., Curtiss, L. K., a n d Witztum, J. L. (1986).J.Biol. Chem. 261,2995-2998. Zhuang, H., Chuang. S. S., and Das, H. K. (1992).Mol. Cell. Biol. 12, 3183-3191.

APOLIPOPROTEIN E: STRUCTURE-FUNCTION RELATIONSHIPS

.

By KARL H WEISGRABER Gladntono Inntltute of Cardiovascular Dlneane. Son Francinco. California 94141-9100

1. Introduction . . . . . . . . . . . ........................................ I1. Apolipoprotein E . . . . . . . . . . . . . . . . ............................ A . Discovery ...................................................... B . Primary Structure .............................................. C. Heterogeneity .................................................. D. Physical Properties and Domain Structure ......................... Ill . Function .................... ................................ A . Component of Plasma Lipoproteins .............................. B. Interaction with Lipoprotein Receptors ........................... C. Mediator of Lipoprotein Metabolism ............................. D. Receptor-Binding Region ....................................... E. Interaction with Heparin ........................................ 1V. Impact of Structure on Function ..................................... A . Relationship of Apolipoprotein E Varlants to Type 111 Hyperlipoproteinemia .......................................... B. Effect of Apolipoprotein E Heterogeneity on Plasma Lipid Concentrations ................................................. C . Domain Interactions ............................................ D. Effect of Cysteinyl Residues on Function .......................... V . Three-Dimensional Structure of Apolipoprotein E 22-kDa Fragments ... A . Apolipoprotein E3 .............................................. B . Comparison of Apolipoprotein E2, E4, and E3 Structures .......... V I . Lipid Binding ..................................................... A . Effect of Lipid on Structure ..................................... B. Carboxyl-Terminal Lipid-Binding Regions ........................ VII . Future Directions .................................................. References ........................................................

249 251 251 251 255 257 260 260 262 265 267 274 277 277 281 282 284 285 285 288 290 290 292 294 295

I . INTRODUCTION Of the approximately 14 plasma apolipoproteins that have been described to date. apolipoprotein E (apoE) is one of the best characterized in terms of its structural and functional properties . In general. plasma apolipoproteins serve to regulate lipoprotein metabolism and to control the transport and redistribution of lipids among tissues and cells. Specifically. apolipoproteins can perform one of three major roles . First. because of their ability to bind lipid. a property they all share. apolipoproteins stabilize the pseudomicellar structure of lipoprotein partiADVANCES IN PROTEIN CHEMISTRY Vul. 45

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CopyrighlO1994 hy Academic Press. Inc . All righls of reproduction in any form reserved.

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KARL H. WEISGRABER

cles. Second, apolipoproteins can act as cofactors or activators of various enzymes or lipid transfer proteins that participate in the metabolism of “remodeling” of lipoproteins as they circulate in plasma. A third function of plasma apolipoproteins-one that is restricted to apoBlOO and ApoE-is to serve as a ligand for cell surface lipoprotein receptors. Because of this special function, apoBlOO and apoE direct the delivery and redistribution of lipids to cells expressing receptors. Receptor-mediated pathways are key components of lipoprotein metabolism and cholesterol homeostasis. [For a review of the low-density lipoprotein (LDL) receptor pathway, see Brown and Goldstein (1986) and Myant, (1990); for a discussion of the role of apoE in the redistribution of cholesterol in an autocrine or paracrine manner among cells and tissues, see Mahley (1988).] In discussing the structure and function of apoE, this review focuses on the role of apoE in lipoprotein metabolism, the most completely described function of the protein. The discussion is limited to cases in which the structure of apoE is known to influence its functional properties. Other functions and properties ascribed to apoE that will not be discussed herein include its roles in immunoregulation (Hui et al., 1980; Avila et al., 1982; Pepe and Curtiss, 1986), nerve regeneration (Ignatius et al., 1986, Snipes 1987; el al., 1986; Boyles et al., 1989; Handelmann et al., 1992), and modulation of intracellular cholesterol utilization and steroidogenesis in adrenal cells (Reyland et al., 1991; Reyland and Williams, 1991), and as an activator or modulator of hepatic lipase (HL) (Ehnholm et al., 1984; Landis et al., 1987; Thuren et al., 1991, 1992), lipoprotein lipase (LPL) (Quarfordt et al., 1977; Yamada and Murase, 1980; Ehnholm et al., 1984; Clark and Quarfordt, 1985), and lecithincholesterol acyltransferase (LCAT) (Zorich et al., 1985; Steinmetz et al., 1985; Chen and Albers, 1985). One of the more provocative and newer functions suggested for apoE is based on the observations that apoE is found in amyloid plaques associated with Alzheimer’s and CreutzfeldtJakob diseases (Namba et al., 1991), as well as in a variety of types of cerebral and systemic amyloidoses (Wisneiwski and Frangione, 1992). As a result of this widespread presence in amyloid plaques, it has been suggested that apoE may function as a pathological chaperone protein-that is, one that induces P-pleated conformation in amyloidogenic polypeptides (Wisniewski and Frangione, 1992). Along these lines, one of the most intriguing observations related to Alzheimer’s disease is that the frequency of the apoE4 allele (one of the three major isoforms of the protein; see Section II,B) is three times higher in unrelated subjects with familial Alzheimer’s disease than in the general population (Strittmatter et al., 1993), suggesting an association of one of the apoE variants with this disorder.

APOLIPOPROTEIN E STRUCTURE-FUNCTION

25 1

11. APOLIPOPROTEIN E

A . Discovely The initial description of apoE was published in 1973 by Shore and Shore. It was identified as a component of triglyceride-rich very lowdensity lipoproteins (VLDL) and was referred to as the “arginine-rich” protein (ARP), because it contained a relatively high content of arginine compared to the other apolipoproteins whose amino acid compositions were known at that time. As determined by sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE), the protein migrated as a broad band with molecular weights reported to be between 33,000 and 39,000 (Shelburne and Quarfordt, 1974; Utermann, 1975). In 1975, Utermann suggested the designation “apoE” for this protein, consistent with the alphabetical nomenclature that was becoming commonly used in this field. However, this designation was not universally adopted for several years. As a result, during the late 1970s the protein was referred to both as ARP and as apoE.

B . Prima? Structure The first apoE sequence to be determined was the amino acid sequence of human apoE, elucidated in 1982 (Rall et al., 1982a). T h e structure was determined by direct protein sequencing of 299 amino acids ( M I 34,200) (Fig. 1). Later, the protein sequence was confirmed by nucleotide sequencing of a full-length cDNA of apoE mRNA (McLean et al., 1984). The apoE gene is located on chromosome 19 (Olaisen et al., 1982; Das et al., 1985). The complete gene has been sequenced (Paik et al., 1985) and found to be 3597 nucleotides in length and to contain four exons and three introns (Paik et al., 1985). Other apolipoprotein genes share a similar structure (Li et al., 1988), suggesting that this class of proteins belongs to a multigene family. T h e mRNA codes for an 18-residue signal peptide that is removed cotranslationally (McLean et al., 1984; Zannis et al., 1984). Th e primary structures of apoE from 10 species have been determined (Fig. 1). They range in length from 279 residues to 310 residues for guinea pig and sea lion apoE, respectively. Overall, there is a high degree of sequence conservation across species, with notable exceptions at the amino and carboxyl termini. The lack of sequence conservation at the amino terminus was recognized in several species in 1980 by amino acid sequencing of the amino termini of the intact proteins (Weisgraber et al., 1980). It is interesting to note that the aminoterminal extension present in the dog and sea lion sequences (Fig. 1) is

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FIG. 1. Comparison of amino acid sequences of apolipoprotein E from 10 species. Sequences are aligned against human apoE4. Hu, Human (Rall et al., 1982a); Ba, baboon (Hixson et al., 1988); CynM, cynomolgus monkey (Marotti et al., 1989); Rt, rat (McLean el al., 1983); Mo, mouse (Rajavashisth et al., 1985); GP, guinea pig (Matsushima el al., 1990); Rb, rabbit (Lee et al., 1991); cow (Chan and Li, 1991); dog (Luo et al., 1989; Weisgraber et al., 1980); SeaL, sea lion (Davis et al., 1991). Blanks indicate identity to human sequence; dashes (-) indicate deletions inserted to maximize homology with the human sequence. One-letter amino acid designations are used: A, alanine; C, cysteine; D, aspartic acid; E, glutamic acid; F, phenylalanine; C, glycine; H, histidine; I, isoleucine; K, lysine; L,


253

shared by the walrus and dolphin (Davis et al., 1991). As indicated in Fig. 1, homology begins in the vicinity of residue 26 in the human sequence and continues to approximately residue 288. It has been noted that the most conserved region of the protein is a block of 33 amino acids (residues 29-61) (Chan and Li, 1991), suggesting that this region is important in terms of structure and/or function, although a specific function has not been assigned at this time. Another highly conserved region is the receptor-binding region of apoE. Studies in the late 1970s using selective chemical modification of specific amino acids suggested that arginyl and lysyl residues were important in the interaction of both apoB and apoE with the LDL receptor (Mahley et al., 1977a; Weisgraber et al., 1978). As is discussed in Section III,D, this has been substantiated for apoE. The region of the protein involved in receptor interaction, shown to be in the vicinity of residues 136-158, is enriched in basic amino acids, with 10 residues being either lysine or arginine. It has been postulated that residues 136-150 interact directly with the receptor (see Section II1,D for discussion). Basic residues are conserved in the 136-158 region of the apoE across species with only two exceptions: the dog (arginine is substituted for lysine at position 157) and the cow (proline is substituted for arginine at 145). Based on the Chou-Fasman algorithm (Chou and Fasman, 1974) for prediction of secondary structure, human apoE is predicted to be highly helical (Rall et al., 1982a). As shown in Fig. 2, the predicted structure of apoE segregates into two ordered segments, one in the amino- and one in the carboxyl-terminal region of the protein. The two ordered regions are connected by a segment whose structure is predicted to be random (residues 165-200) (Rall et al., 1982a). T h e protein is predicted to contain multiple a helices of various length and character (Rall et al., 1982a). It is of interest that only five of these helices contain elements that satisfy some or all of the properties thought to be important in the association of the amphipathic helices of apolipoproteins with lipid. The carboxyl-terminal region contains three of these helical stretches-residues 203-22 1, 226-243, and 245-266, which have all the characteristics typical of apolipoprotein helices. On this basis the carboxyl terminus was predicted to be a major lipidbinding region of the protein (Rall et al., 1982a). (For a more complete leucine; M , methionine; N. asparagine; P, proline; Q, glutamine; S, serine; V, valine; W, tryptophan; Y, tyrosine. *, Dog sequence contains amino-terminal extension: DVQPEPELERELEP; t, SeaL sequence contains amino-terminal extension: DVEPESPLEENLEPEL + EPKR.

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KARL H. WEISCRABER

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FIG. 2. Predicted secondary structure of human apolipoprotein E3. One-letter amino acid designations as in Fig. 1. The predicted secondary structure was determined by applying the Chou-Fasman algorithm (Chou and Fasman, 1974) and is predicted to contain two ordered segments, one in the amino- and one in the carboxyl-terminal region of the protein (residues 1-164 and 200-290, respectively).


255

description of the characteristics of the helices in apoE and the importance of the amphipathic a helix of apolipoproteins in lipid binding, see Segrest et al., this volume.)

C. Heterogeneity T h e heterogeneity of apoE was recognized by Shore and Shore (1973)in the initial description of the protein. They observed that the protein eluted in multiple peaks on DEAE ion-exchange chromatography and that the proteins contained in these peaks appeared similar in electrophoretic mobility and amino acid composition. Use of isoelectric focusing on polyacrylamide gels to demonstrate heterogeneity, introduced by Utermann in 1975,showed the protein to focus in multiple bands, with pZ values ranging from 5 to 6 (Utermann, 1975).Today, this technique remains as standard for examining apoE polymorphism [for a review of variations of this technique as applied to apoE, see Davignon et al. (1988)l.Utermann, the first investigator to recognize that apoE polymorphism was genetically determined, proposed a twoallele model to explain the different isoelectric focusing patterns observed in family studies (Utermann et al., 1977). This model was later modified to accommodate the presence, in some subjects, of an additional band revealed in more extensive screening studies (Utermann et al., 1980). However, the model remained a two-allele model, with the presence or absence of the additional band indicated by plus (+) or minus (-). Based on two-dimensional gel analysis, a major advance in the understanding of the genetic basis of apoE polymorphism occurred in 1981 when Zannis and Breslow proposed a three-allele model to account for the multiple-banded patterns. T h e three alleles, designated 11, 111, and IV, were shown to differ progressively by a single charge unit. In addition, a key feature of the model was that posttranslational glycosylation also contributed to the observed charge and molecular weight heterogeneity. Treatment with neuraminidase demonstrated that the minor, more acidic bands in each pattern were sialylated derivatives of a major band (Zannis and Breslow, 1981).Thus, the apoE polymorphism, as it was then understood, could be explained by the presence of three alleles at a single gene locus in combination with the addition of one or more sialic acid residues to the product of each allele (Zannis and Breslow, 1981).This three-allele model predicted the presence of three homozygous and three heterozygous apoE phenotypes, each with a unique isoelectric focusing pattern. This model fit the data from screening and family studies (Zannis and Breslow, 1981).

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KARL H. WEISCRABER

In 1982, a standard nomenclature was proposed that has been generally adopted. The three allele products were designated apoE2, apoE3, and apoE4. The numerical designation corresponded to the relative pt of each protein on isoelectric focusing gels, with apoE2 being the most acidic isoform (pt -5.7) and apoE4 the most basic (pl -6.1); the apoEl position was reserved for sialylated isoforms (Zannis et al., 1982). The three homozygous phenotypes were designated E2/2, E313, and E4/4; the heterozygous phenotypes were designated E413, E4/2, and E3/2. A schematic representation of the isoelectric focusing patterns of the six common phenotypes is presented in Figs. 3A and B, including the effect of posttranslational addition of sialic acid on the patterns. As determined in a number of populations, the average apoE allelic frequencies are ~2 0.073, ~3 0.783, and ~4 0.143. In general, across all

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FIG. 3. Diagram of the isoelectric focusing patterns of the six common apolipoprotein E phenotypes. E, Protein; E , apoE allele. Top: Focusing position of the three common apoE isoforms, designated E2,E3, and E4, as they appear in the three hornozygous and three heterozygous phenotypes. Bottom: Effect of posttranslational sialylation on the isoform patterns. Sialylated derivatives are indicated as E,.


257

populations examined, the allelic frequencies are similar, with apoE3 being the most common form (Hallman et al., 1991; for review, see Davignon et al., 1988). T h e molecular basis for the apoE polymorphism was determined in 1981 to result from cysteine-arginine interchanges at two positions in the protein (Weisgraber et al., 1981), later identified as residues 112 and 158 (Rall et al., 1982a). These single amino acid substitutions account for the known charge differences among the major isoforms, supporting the three-allele model of Zannis and Breslow (1981). T h e most common isoform, apoE3, contains a single cysteine in its structure at residue 112; it contains arginine at 158. Both positions have cysteine in the apoE2 isoform and arginine in apoE4. As is discussed in Section III,D, a number of additional rare apoE variants have since been described. Functional consequences of these amino acid substitutions are discussed in Sections III,D and IYA-IV,D. It is interesting to note that cysteine is found only in human apoE2 and apoE3 and rabbit and bovine apoE (Fig. 1). The impact of cysteinyl residues on properties of apoE is discussed in Section IV,C.

D . Physical Properties and Domain Structure In common with other apolipoproteins, apoE was demonstrated to self-associate in the absence of lipid (Yokoyama et al., 1985; Aggerbeck et al., 1988a). However, apoE differed in one interesting aspect: it remained associated as a tetramer over a wide range of concentrations (50 pg/ml to 15 mg/ml) (Yokoyama et al., 1985; Aggerbeck et al., 1988a). All other soluble apolipoproteins are monomeric at low protein concentrations and self-associate as the concentration increases, forming a progression of high molecular weight oligomers in a concentration-dependent manner. In addition, apoE was shown to differ in another important aspect. As illustrated in Fig. 4, when apoE is denatured by treatment with guanidine, and protein unfolding is monitored by circular dichroism, apoE displays two transitions (Wetterau et al., 1988). The first transition midpoint occurs at a guanidine concentration of 0.7 M ,a point at which all other soluble apolipoproteins are completely unfolded. The second midpoint occurs at a much higher guanidine concentration (2.4 M), which translates to a free energy of stabilization of 12 kcal/mol. These results indicated that apoE contained two regions that were independently folded and that differed markedly in their stabilities. Furthermore, these two regions apparently represent two independent structural domains in apoE. The stability of the domain, corresponding to

-

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KARL H. WEISGRABER

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the second transition, is in the range of stable, soluble globular proteins. The indication of two structural domains, particularly one with such a high stability, is unique among plasma apolipoproteins. Additional evidence supporting the existence of two domains in apoE was obtained when apoE was subjected to limited proteolysis with a battery of five proteases having widely varying specificities (Wetterau et al., 1988). The results, summarized in Fig. 5, demonstrate that there are two protease-resistant regions in the amino-terminal (residues 20165) and carboxyl-terminal (residues 225-299) portions of the protein, suggesting that these regions constitute the structural domains. The central portion of the protein, which is highly susceptible to proteolysis (residues 165-210), is the portion of apoE that is predicted to exist as a random structure (Fig. 2). Thrombin cleaves apoE at residues 191 and 215 (Bradley et al., 1982; Innerarity et ad., 1983), generating a 22-kDa amino-terminal fragment and a 10-kDa carboxyl-terminal fragment. Because the two thrombolytic fragments closely approximated the two protease-resistant domains (Fig. 5 ) , these fragments were used to model the two domains.

259


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FIG. 5. Proteolytic cleavage sites in apolipoprotein E. Top: Linear representation of the structure of human apoE demonstrating cleavage sites when a p E 3 was subjected to limited proteolysis with six enzymes: 0,trypsin; 0,elastase; U, chymotrypsin; 0, subtilisin; Stufihylococcw a u r m V8; 0 , thrombin. Bottom: Linear representation of the two thrombolytic fragments of apoE, the 22- and 10-kDa fragments.

+,

When each fragment was denatured with guanidine, the denaturation curve of the amino-terminal fragment closely approximated the second, highly stable domain, and the carboxyl-terminal fragment closely approximated the first, less stable domain (Wetterau et al., 1988). These results demonstrated the usefulness of the two thrombolytic fragments as models for the two domains of apoE. Sedimentation equilibrium ultracentrifugation of the fragments and proteolysis of the intact protein followed by HPLC chromatography revealed that the 22-kDa fragment was monomeric and the 10-kDa fragment was tetrameric (Aggerbeck et al., 1988a). These results suggested that the tetramerization of the intact protein was mediated by the carboxyl-terminal domain and also that the domains did not interact with each other when the protein was free in solution. Characterization of other hydrodynamic properties indicated that the 22-kDa fragment was compact and globular and that the 10-kDa fragment was elongated in shape (Aggerbeck et al., 1988a). Based on the physical-chemical studies, a two-domain model of apoE was suggested, having a stable, globular amino-terminal domain and an elongated carboxyl-terminal domain. The domains are connected by a “hinge region”-the protease-susceptible region. As will be discussed, the two domains have different functional properties: the aminoterminal domain contains the receptor-binding function and the carboxyl-terminal domain contains the lipid-binding function. As assessed by circular dichroism, the intact protein was shown to be helical in form (Shore and Shore, 1974; Roth et al., 1977; Cassel et al., 1984; Chen et al., 1984; Yokoyarna et al., 1985; Aggerbeck et al., 1988a; Mims et al., 1990). Both domains also were highly helical (approximately 54%) (Aggerbeck et al., 1988a). in good agreement with the Chou-Fasman prediction (Fig. 2). Based on the a-helical nature of the carboxyl-terminal fragment and the fact that these helices are amphipathic (Rall et al., 1982a), it was suggested that the carboxyl-terminal do-

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main self-associates by forming a tetrameric a-helical bundle (Aggerbeck et al., 1988a). In this situation the hydrophobic surfaces of the amphipathic helices would be sequestered within the interior of the bundle and would be shielded from the aqueous environment. The bundle could form by arrangement of the helices in either a parallel or an antiparallel manner. However, an antiparallel arrangement would provide additional stability from helix-dipole interactions. 111. FUNCTION A . Component of Plasma Lipoproteins Although apoE was recognized first as a component of VLDLs (Shore and Shore, 1973; Shelburne and Quarfordt, 1974; Utermann, 1975; Kane et al., 1975), it has been demonstrated to be present in most other lipoprotein classes as well. In addition to occurring in the other triglyceride-rich lipoproteins, chylomicrons and their remnants, and the intermediate-density lipoproteins (IDLs), apoE is present in a subclass of the cholesterol-rich high-density lipoproteins (HDLs), referred to as “HDL-with apoE” (Mahley, 1978). By use of SDS-PAGE (Fig. 6), the M , 34,200 apoE is easily distinguished from the other common apolipoproteins that also are present in the various human lipoprotein classes. Shortly after apoE was initially described, some unusual characteristics were noted that provided the first clues to the functional importance of the protein in lipoprotein metabolism. Havel and Kane observed that apoE was present in elevated amounts in the cholesteryl ester-rich lipoproteins, referred to as P-VLDLs, which accumulate in plasma and characterize subjects with the lipoprotein disorder type I1 I hyperlipoproteinemia, or familial dysbetalipoproteinemia (Havel and Kane, 1973). Shore and Shore also observed that apoE accumulated in the cholesteryl ester-rich lipoproteins from type I I I hyperlipoproteinemic subjects, as well as in similar lipoproteins that accumulate in the plasma of rabbits fed a diet enriched in cholesterol and fat (Shore and Shore, 1974). These early observations led to the suggestion that apoE probably was involved in the transport and metabolism of cholesteryl esters and/or unesterified cholesterol. Beginning in 1974, a series of studies characterizing the lipoproteins in a number of choIesteroVfat-fed animal models, including dogs (Mahley et al., 1974; Mahley and Weisgraber, 1983), pigs (Mahley et al., 1975), rats (Mahley and Holcombe, 1977; Weisgraber and Mahley, 1983), rabbits (Shore et


Chylomicrons

VLDL

LDL

26 1

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FIG. 6. Sodium dodecyl sulfate polyacrylamide gel electrophoresis of various human lipoprotein classes. The apolipoprotein components of the major lipoprotein classes were separated by SDS-PAGE on a 10% acrylamide gel in the absence of reducing agents. From Mahley and Innerarity (1983), with permission from Elsevier Science Publishers BV.

al., 1974), and patas monkeys (Mahley et al., 1976), demonstrated that one of the common responses in these models was the accumulation of cholesteryl ester-rich lipoproteins, which closely resembled the flVLDLs of type 111 hyperlipoproteinemia and which contained high concentrations of apoE. These results supported the suggestion that apoE was an important component of cholesterol metabolism. Utermann was the first to recognize that a particular isoform pattern is associated with type 111 hyperlipoproteinemia (Utermann et al., 1975), indicating that apoE polymorphism was associated with this disorder. Later, with the introduction of the three-allele model by Zannis and Breslow (1980), it was shown that type 111 hyperlipoproteinemia was associated with the E2/2 phenotype. The underlying basis for this association is discussed in Section IYA. An additional feature of cholesteroVfat-fed animal models is the accumulation of HDL-with apoE. In several species, including dogs (Mahley et al., 1974; Mahley and Weisgraber, 1983), rats (Mahley and Holcombe, 1977; Weisgraber and Mahley, 1983), pigs (Mahley et d., 1975), and mice (de Silva et al., 1992), the HDL-with apoE is a prominent lipoprotein class that extends into the lower density ranges overlapping in its density distribution with LDLs. The HDL-with apoE in these spe-

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KARL H. WEISGRABER

cies also has been referred to as HDL, or HDL, when the animals are cholesterol/fat fed (for review, see Mahley, 1983, 1985). In humans, the HDL-with apoE is less prominent, transporting approximately 20% of plasma cholesterol (Weisgraber and Mahley, 1980). T h e prominence of the HDL-with apoE in a given species is associated with cholesteryl ester transfer protein (CETP) activity in that particular species (Tall, 1986). The CETP transfers cholesteryl esters from HDLs to VLDLs, IDLs, and LDLs, in exchange for triglyceride. Species with low or no transfer activity (rats, mice, dogs, and pigs) transport a majority of their plasma cholesterol in HDLs, of which the HDL-with apoE (HDLI) is a prominent subclass. In species with moderate to high CETP activity (humans, rabbits, and primates), the HDLs are less prominent and the majority of plasma cholesterol is transported in LDLs. This low HDL/high LDL pattern is reversed in humans with CETP deficiency (Koizumi et al., 1985; Brown et al., 1989; Yamashita et al., 1990a). In addition, the HDL-with apoE is a major cholesterol transporter in these subjects (Yamashita et al., 1990b). In the various animal models, as well as in humans, the HDL-with apoE increases with cholesterol feeding (for review, see Mahley, 1983, 1985). As is discussed in Section III,B, there is a particular HDL, subclass, which can be isolated from the plasma of cholesterol/fat-fed dogs, that is unique in that apoE is the only apolipoprotein on the particle (Mahley et al., 1977b). This apoE HDL, has proved to be very useful in studying the interaction of apoE with the LDL receptor.

B . Interaction with Lipoprotein Receptors In the initial studies on the LDL receptor, it was thought that the apoBlOO of LDLs was the only apolipoprotein capable of binding to the receptor, although there were some suggestions that HDLs also could bind (Carew et al., 1976; Miller et al., 1977). In 1976, it was demonstrated that swine HDL,, which contained significant amounts of apoE in addition to apoA-I, bound to the LDL receptor, competed with LDL for binding, and delivered cholesterol to cells (Bersot et al., 1976). It was shown that binding to the LDL receptor was proportional to the apoE content of the HDL-with apoE and that HDLs from which the apoEcontaining subclasses were removed did not bind (Innerarity and Mahley, 1978; Mahley and Innerarity, 1977). Selective chemical modification studies demonstrated that lysyl and arginyl residues were important in the interaction of apoE, as well as of apoBl00, with the LDL receptor (Mahley et al., 1977a; Weisgraber et al., 1978). Thus, there are close parallels in the binding of both apoli-


263

poproteins to the LDL receptor, which is sometimes referred to as the apo-B,E receptor. However, one very important difference was established with apoE HDL,, the HDL-with apoE subclass, obtained from cholesteroVfat-fed dogs, that only contains apoE and no other apolipoproteins: the apoE HDL,s bound to the LDL receptors with a 20- to 25-fold higher affinity than did LDLs (Pitas et al., 1979). In competition studies the apoE HDL,s were 80- to 100-fold more effective than LDLs in competing with ‘251-labeledLDL for receptor sites (Pitas et al., 1979). In terms of size of the respective lipoprotein particles and their chemical composition, both lipoproteins were similar. Therefore, the difference in binding affinity was the direct result of the presence of apoE. Detailed kinetic analysis revealed that the Kd values for LDL and apoE HDL, were approximately 2.8 X lo-’ and 1.0 X lO-’OM, respectively (Pitas et al., 1979). In addition, saturation of the available receptor sites was shown to require approximately four times as many LDL particles as HDL, particles. Based on these results, it was proposed that the HDL,s bound to multiple (up to four) sites on LDL receptors, whereas LDLs bound to one site (Pitas et al., 1979). Thus, according to this model, the increased binding affinity of the apoE HDL, resulted from multiple interactions with the LDL receptors, which are present as dimers on the cell surface. When it was determined several years later that one LDL particle bound to a single receptor molecule (van Driel et al., 1989), the model was modified so that apoE HDL, bound to multiple receptors-four receptors o r two dimers. This revised model is still consistent with the higher affinity of HDL, and the 4: 1 binding ratio of LDL:apoE HDL,. A property that all the soluble apolipoproteins share is the ability to combine with phospholipid vesicles to form discoidal-shaped particles. The phospholipid in these artificial lipoprotein complexes is organized in a bilayer, with the apolipoprotein oriented on the periphery of the “disc.” When apoE is complexed with dimyristoylphosphatidylcholine (DMPC) at a weight ratio of 3.75: 1 (phospholipid : protein), discoidal particles containing four apoE molecules are formed (Innerarity et al., 1979; Pitas et al., 1980). Using these discoidal particles with various ratios of receptor-inactive and receptor-active (chemically modified to abolish receptor activity) apoE, it was determined that a single apoE bound to the LDL receptor with an affinity similar to that of LDL (Kd 2.6 x lO-’M) (Pitas et al., 1980). In addition, it was determined that the Kd for the apoE phospholipid discs, which contained four receptoractive molecules, was similar to the Kd for apoE HDL, (Pitas et ad., 1979). Taken together, these results support the model that the higher affinity of apoE-containing lipoproteins, compared to that of LDLs, re-

-

264

KARL H. WEISGRABER

sulted from the interaction of four apoE molecules with four LDL receptors. Model studies in which the number of apoE molecules is varied on microemulsion particles support the multireceptor model and demonstrate a marked increase in receptor activity as the number of apoE molecules on the emulsion particles is increased from one to four (Funahashi et d., 1989). Addition of more than four apoE molecules to the particles does not further increase binding activity (Funahashi et al., 1989). Although intact apoE forms a stable tetramer in solution (Yokoyama et al., 1985; Aggerbeck el al., 1988a), tetramerization does not appear to be required for lipid binding and the protein does not appear to self-associate on lipid surfaces (Yokoyama, 1990). The multireceptor binding ability of apoE has important consequences in lipoprotein metabolism. Lipoprotein particles with one apoE per particle will compete on a more or less equal basis with LDL for LDL receptors and would be expected to be cleared at approximately equal rates. However, when the apoE content increases to four or more molecules per particle, the binding affinity increases dramatically through multireceptor interaction, with a corresponding increase in removal rate. In addition to binding to the LDL receptor, apoE also binds to the LDL receptor-related protein (LRP), another receptor molecule that is potentially important in lipoprotein metabolism. As the name implies, the LRP bears a strong resemblance to the LDL receptor (Herz et al., 1988; Brown et al., 1991). It contains multiple copies of two types of cysteine-rich repeats that comprise either the ligand-binding domain or the epidermal growth factor precursor region of the LDL receptor. However, in contrast to the seven cysteine repeats found in the ligandbinding domain of the LDL receptor, the LRP contains 3 1 such repeats. Apparently unrelated to its role in lipoprotein metabolism, the LRP also serves as the receptor for a2-macroglobulin (Strickland et al., 1990) [for a review of LRP, see Brown et al., (1991)l. Although its role in lipoprotein metabolism has not been delineated fully, recent studies clearly imply that LRP is responsible for the clearance of a significant percentage of chylomicron remnants (Hussain et al., 1991; Mahley and Hussain, 1991). T h e interaction of apoE with the LRP has been most extensively studied using rabbit P-VLDLs (Kowal et al., 1989, 1990). These cholesterol-enriched lipoproteins represent chylomicron remnants derived from the intestine and VLDL remnants from the liver. They contain multiple apoE molecules in addition to apoB and the low molecular weight apoC molecules. Interestingly, for rabbit /3-VLDLs to interact effectively with the LRP receptor, the apoE content of these lipoproteins must be first enriched by incuba-

265


tion with exogenous apoE (Kowal et al., 1989, 1990). T h e apoEdependent binding is inhibited by a mixture of the apoC proteins (Kowal et al., 1990). It was demonstrated that apoC-I was the most effective inhibitor and that the decrease in binding to the LRP correlated with the decrease in apoE content, suggesting that the inhibition by apoC-I was the result of displacement of apoE below a critical level required for interaction with the LRP receptor (Weisgraber et al., 1990). This level appeared to be approximately 50% of the apoE content of the apoE-enriched P-VLDLs. C . Mediator of Lipoprotein Metabolism Chylomicrons are synthesized in the intestine and transport dietary triglycerides and cholesterol. While circulating, the core triglycerides in these particles are hydrolyzed by lipoprotein lipase, which results in the production of a cholesterol-enriched remnant particle. When synthesized and initially released by the intestine, chylomicrons contain essentially no apoE, but as they circulate and are processed to remnants, the particles acquire apoE from other lipoprotein classes. This results in a shift of the distribution of apoE in plasma to the triglyceride-rich remnants in the absorptive state (Blum, 1982). A very simplified scheme of lipoprotein metabolism is shown in Fig. 7. I t is intended only to highlight the role of apoE in major metabolic processes and the central importance of the liver in lipoprotein metabolism [for a general review of lipoprotein metabolism, see Have1 and

I

LDL

VLDL Chylomicron Remnants

/

-

IDL

LDL

FIG. 7. Role of hepatic lipoprotein receptors in lipoprotein metabolism. The central role of hepatic receptors and the importance of apoE in the clearance of chylomicron remnants (remnant receptor), VLDL (LDL receptors), IDL (LDL receptors), and HDLwith apoE (LDL receptors) are indicated. In addition, the suggested role of apoE and hepatic lipase (HL) in the conversion of IDL to LDL is shown.

266

KARL H. WEISGRABER

Kane (1989)l. Chylomicron remnants are rapidly removed from plasma in a process known to be mediated by apoE (Shelburne et al., 1980; Sherrill et al., 1980; Windler et al., 1980). T h e full details of this uptake process have not been completely defined. It has been postulated that the LRP receptor functions as the so-called remnant receptor (Kowal el al., 1989, 1990).In vivo evidence indicates that LRP is involved in uptake of chylomicron remnants (Hussain et al., 1991; Mahley and Hussain, 1991). In addition, LDL receptors also appear to play a role in uptake (Choi et al., 1991). Thus, at this point it appears that remnants may be cleared by t w o receptor systems. Therefore, in this regard, the scheme depicted in Fig. 7 is oversimplified. However, what clearly has been established is that apoE is a critical component of the chylomicron clearance process regardless of the receptor or receptors that are involved. Very low-density lipoproteins also are triglyceride-rich lipoproteins that are synthesized by the liver. These lipoproteins contain apoE and apoBl00. In a manner similar to that of chylomicrons, VLDL particles pass through a lipolytic cascade as these particles are acted on by lipoprotein lipase (LPL). A spectrum of particles of progressively decreasing size is produced, including VLDL remnants and intermediate density lipoproteins. The cholesterol-rich LDLs represent the final stage of this process (Fig. 7). Although both VLDLs and IDLs contain apoE and apoB 100, these particles are cleared through apoE interaction with the LDL receptor. As indicated in Fig. 7, not all IDLs are cleared by the liver. In humans, a major portion of the IDL is converted to LDL, a process that involves a second lipase, hepatic lipase (HL). In addition, it has been suggested that apoE is involved in this process by serving as an activator of HL (Thuren et al., 1991, 1992). Once LDL particles are produced, the apoE has been lost from the surface, and the apoBlOO remains as the sole apolipoprotein component. T h e clearance of the LDL is then via apoBlOO through the LDL receptor. Thus, a progression from apoE- to apoB-mediated clearance occurs as VLDL particles transverse the lipolytic cascade (Bradley el al., 1984). In addition, in species with high concentrations of HDL-with apoE, it has been postulated that this lipoprotein class serves to transport excess cholesterol from peripheral cells to the liver for elimination from the body (Mahley et al., 1980). This transport process is referred to as the reverse cholesterol transport process (Glomset, 1968). In contrast, in species with high CETP activity, the excess cholesterol from the periphery is transferred from the typical non-apoE-containing HDL to the lower density lipoprotein classes (VLDL, IDL, and LDL) for clearance by the liver.


267

It has been shown that infusion of apoE into cholesteroVfat-fed rabbits, effectively raising plasma apoE levels, results in lowering of plasma cholesterol concentrations (Mahley et al., 1989; Yamada et al., 1989). Based on these results, it was suggested that the availability or concentrations of apoE in plasma may be rate limiting for lipoprotein clearance (Mahley et al., 1989). Consistent with this suggestion are the recent studies in which apoE has been overexpressed in transgenic mice, resulting in decreased plasma cholesterol levels in chow-fed animals (Shimano et a/., 1991) and a resistance to hypercholesterolemia when overexpressors were fed a high-cholesteroVfat diet (de Silva et al., 1992; Shimano et al., 1992). The importance of apoE in lipoprotein metabolism also is emphasized in apoE-deficient subjects, who display features of type I11 hyperlipoproteinemia (Schaefer et al., 1986). Mice in which the apoE has been inactivated by gene targeting also exhibit the massive accumulation of remnant lipoproteins in their plasma (Zhang et al., 1992; Plump et al., 1992). D. Receptor-Binding Region

Because of the importance of apoE interaction with the LDL receptor in lipoprotein metabolism, considerable effort has been directed zt determining the region of apoE that binds to the receptor. Attention was drawn first to a region in the center of the protein (residues 136-158, Fig. 2) that was enriched in basic amino acid residues (Rall et al., 1982a). This region attracted attention because of the earlier chemical modification studies that implicated arginyl and lysyl residues in the binding process (Mahley et al., 1977a; Weisgraber et al., 1978). The first indication that this region was involved in receptor binding became apparent when the structural basis for the three major isoforms was determined (Weisgraber et al., 1981), and the three isoforms were tested for receptor-binding activity (Weisgraber et al., 1982). To test the receptor-binding activity, apoE is complexed with phospholipid, because the protein is essentially inactive in the lipid-free state (Innerarity et al., 1979). As shown in Fig. 8, apoE4 and apoE3 phospholipid complexes were equally effective in competing with '251-labeled LDL for binding to LDL receptors on cultured human fibroblasts (Weisgraber et al., 1982). This finding indicated that either cysteine or arginine is tolerated at position l 12 without affecting receptor-binding activity. However, the apoE2 (with cysteine substituted for arginine at position 158) possessed only 1% of the binding activity of apoE3 and E4. The relevance of the relationship between the defective binding of the apoE2 and type 111 hyperlipoproteinemia is discussed in Section IV,A.

268

KARL H. WEISGRABER

3

z

40-

i I =

20-

ApoE3 and ApoW m

FIG. 8. Ability of apolipoprotein E*dimyristoylphosphatidylcholine (DMPC) complexes of the three major isoforms to compete with human 1251-labeledLDL for binding to normal human fibroblasts. Cells incubated in medium containing 10% human lipoprotein-deficient serum received 1 ml of the same medium with 2 pg/ml of 1251-labeled LDL and the indicated concentrations of apo-E.DMPC complexes. After a 2-hr incubation on ice at 4"C, the cells in 35-mm petri dishes were extensively washed and the '251labeled LDL bound to the cells was determined. From Weisgraber el a/. (1982).

In terms of the present discussion of the receptor-binding region of apoE, the effect of the loss of Arg-158 on receptor interaction was consistent with the chemical modification studies (Mahley et al., 1977a; Weisgraber et al., 1978) and with the fact that Arg-158 was within the basic region of apoE noted above. Three independent approaches were used to define the receptorbinding region of apoE. T h e first involved screening human populations, primarily type 111 subjects, to look for other mutation sites in the molecule that resulted in defective binding to the LDL receptor. These studies uncovered a number of additional mutations that helped define the binding region. These variants are listed in Table I along with other known variants that have been described. In addition to relying solely on natural mutations for insight, site-directed mutagenesis also was used to probe this structure-function question (Lalazar et al., 1988). In the second approach to define the binding region, fragments of apoE were generated by chemical o r enzymatic digestion and tested for bind-

TABLE I Genetic Variants of Apolipoprotcin E

Mutation

I soelectric focusing position

Receptor-binding activity

Associated with type I11 hyperlipoproteinemia/mode of inheritance

E4 E2 El

Normal Defective Defective

No/NA Yeslrecessive Yedrecessive

E2

Defective

Yes/un known

E3 E3 E2 E4 E2 El

Defective Defective Defective Unknown Defective Defective

NolNA Yesldominant Yeslunknown Yedun known Yesldominant Yeddominant

C Y S ' ' -+ ~ Arg, seven-amino acid tandem insertion (residues 12 1-127) GluS + Lys

E3

Defective

Yeddominant

E5

Above normal

NolNA

Glu'3 + Lys Proa4+ Arg Arg*' + Cys GlU'* + Lys, GIU245-+ Lys

E5 E5 E2 E7

Unknown Normal Normal Defective

NolNA NolNA NolNA NolNA

Unknown E2

Unknown Unknown

Yeslunknown Yeslunknown

Cys"' + Arg Arg'58 + Cys Arg'58 + Cys, Glyl*' Arg'"

+ Ser

ArgIS6 + His ArgI4' + Cys, Cys"' Arg'45 4Cys Arg'45 + Cys, Glu'' Lys146+ Gln Lys'4" + Glu

209 truncation ArgIS4 + Cys a

+ Asp

+ Arg + Lys

NA, Not applicable. A. Minnich, K. H. Weisgraber, Y.Newhouse, and J. Davignon, unpublished observation. K. H.Weisgraber, Y. Newhouse, and R. Illingworth, unpublished observation.

Ref, Weisgraber et al. ( 1981, 1982) Weisgraber et al. (1981, 1982) Weisgraber et al. (1984), Steinmetz et al. (1990) Wardell et al. (1987). Lalazar et al. ( 1988) -b Have1 et al. (1983). Rall et al. (1989) Rall p t al. (1982b), Emi el al. (1988) Lohse et al. (1991) Rall et al. (1983b), Smit et al. (1990) Mann et al. (1989).Moriyama et al. ( 1992) Wardell et al. (1989) Yamamura et al. (1984a,b), Dong et al. (1990). Wardell et al. (1991) Mailly et al. (1991) Wardell et al. (1991) Wardell et al. (1990) Yamamura et al. (1984b), Maeda et al. (1989), Tajima et al. (1989) Lohse el al. ( 1992)

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270

KARL H. WEISGRABER

ing activity (Innerarity et al., 1983). The third approach involved screening monoclonal antibodies for their ability to block receptor interaction and then defining the epitope of the antibodies (Weisgraber el al., 1983). The results of the three approaches can be summarized with the aid of Fig. 9. The positions at which natural mutants were identified are indicated above the linear representation of the apoE molecule. As discussed above, the cysteine-arginine interchange that distinguishes apoE3 and E4 was without effect, and the cysteine-for-arginine substitution that defines apoE2 resulted in a dramatic reduction in binding activity. Substitution of either arginine or lysine at positions 136, 142, 145, and 146 by neutral residues all resulted in defective binding (from 20 to 45% of normal apoE3 binding activity), whereas cysteine substitution for Arg-228 was without effect. These results indicated that the

Natural Mutants:

145 146 '42,

I 9 COOH

Site Directed: 143

Fragment 1 Binding:

22-kDa

191 126

mAb

inhibition:

7m

CNBr

2U@

0-

m

218

1D7

FIG. 9. Determination of the receptor-binding site on human apolipoprotein E. Summary of the various lines of evidence that indicate that the central region of apoE (residues 136- 158, solid area) contains the LDL receptor-binding site. Top: Linear representation of the linear structure of apoE, above which are indicated the positions of naturally occurring mutants of apoE; the solid numerals indicate mutation sites that result in defective binding to LDL receptors and the open numerals indicate mutation sites that have no effect on receptor binding. Below the linear sequences are displayed the sites that have been mutated by site-specific mutagenesis. Middle (fragment binding): The two thrombolytic fragments of apoE (22 and 10 kDa) and the largest cyanogen bromide fragment (CNBr) are displayed; the solid line indicates that the fragment possessed receptorbinding activity and the crosshatched line indicates that the fragment lacked activity. Bottom (mAb inhibition): Epitopes of three monoclonal antibodies (7C9, 1D7, and 3H1) to apoE are shown; the solid numeralslletters indicate antibody effectively blocked binding of apoE to the LDL receptor and the open numerals/letters indicate antibody did not block activity.


27 1

central region (highlighted by the solid bar in Fig. 9) was critical for receptor interaction. Site-directed mutagenesis within this region (shown below the bar in Fig. 9) provided additional evidence that basic amino acids were involved in receptor interaction (Lalazar et al., 1988). When the two fragments that are generated by thrombin digestion were recombined with phospholipid and tested for binding activity, only the 22-kDa fragment (residues 1-191) was active (as active as the intact protein); the 10-kDa fragment was completely inactive (Innerarity et al., 1983) (Fig. 9). Of the four cyanogen bromide fragments of apoE that were tested, only the fragment encompassing residues 126218 possessed binding activity (Innerarity et al., 1983) (Fig. 9). Taken together, the fragment results support the conclusion that the receptorbinding region of the molecule is located in the center of the protein. In the monoclonal antibody approach, only one antibody, 1D7, was found to inhibit receptor binding (Weisgraber et al., 1983) (Fig. 9). Its epitope was determined to encompass residues 136-150 (Weisgraber et al., 1983). Thus, the evidence from all three independent approaches supports the conclusion that the central portion of apoE in the vicinity of residues 136-160 is involved in binding to the LDL receptor. In addition, basic amino acids in this region are key to this interaction. Because the cysteine-rich repeats that comprise the ligand-binding domain of the LDL receptor are enriched in acidic residues (Yamamoto et al., 1984), it was suggested that the basic receptor-binding region of apoE binds to the LDL receptor via an ionic interaction (Mahley et al., 1986). Because no single substitution of a basic residue within the receptor-binding region of apoE completely disrupts binding to the LDL receptor, it seems likely that no one residue is critical for interaction and that the basic residues in this region are acting in a cooperative manner. It is now thought that Arg-158 does not interact directly with the receptor but that charge at this position helps to maintain the receptorbinding region in a conformation that can effectively interact with the LDL receptor (Innerarity et al., 1984). Only the basic residues between residues 136 and 150 are believed to interact directly with the receptor (Mahley et al., 1990). The evidence for an indirect role for Arg-158 was suggested when the binding activity of the 22-kDa fragment of this variant was compared to that of the intact protein (Innerarity et al., 1984). It was found that simply removing the carboxyl terminus (residues 192-299) by digestion with thrombin increased binding approximately 10-fold (from 1 to 10% binding activity). In addition, when the intact variant protein was charge modified with cysteamine (a reagent that converts cysteinyl side chains to charged lysinelike derivatives), the

272

KARL H. WEISCRABER

binding activity also increased approximately 1O-fold (Weisgraber et al., 1982; Innerarity et al., 1984). Although the apoE2 contains two cysteinyl residues (positions 112 and 158) that can be charge modified by cysteamine, this increase in binding activity is most probably due to charge modification of Cys- 158 because the presence of positive charge at position 112 is without effect, i.e., binding activities of apoE3 and apoE4 are equal (Fig. 8) (Weisgraber et al., 1982). However, when the apoE2 22-kDa fragment was modified with cysteamine, the binding activity increased 10-fold from 10% to normal (100% of apoE3) (Innerarity et al., 1984). Furthermore, when the cysteamine modification was reversed by reduction of the apoE2 22-kDa phospholipid complexes with 2-mercaptoethanol, if the receptor activity was tested immediately, it was found to be normal (Innerarity et al., 1984). However, with time, the binding activity decreased to approximately lo%, similar to that of the unmodified 22-kDa apoE2 fragment. Thus, these results suggest that charge at position 158 is important for receptor activity but that the role of charge at this position is to help induce the proper conformation when the protein binds to lipid. This conformation in the lipid-bound state can be maintained for a short time after the charge is removed from position 158. Crystallographic results (see Section V,B) provide a potential explanation for the indirect effect of position 158 on receptor-binding activity. Results from a number of other studies are consistent with the concept that the conformation of apoE on the surface of a lipoprotein particle can be modulated by a number of factors, including lipid composition. It has been demonstrated that apoE on the surface of large VLDLs isolated from hypertriglyceridemic patients binds with a higher affinity to LDL receptors than to smaller hypertriglyceridemic VLDL subclasses (Gianturco el al., 1983). In addition, a greater susceptibility to cleavage with thrombin was found for the apoE on the large hypertriglyceridemic VLDL than on smaller particles, and thrombin digestion resulted in the loss of receptor-binding activity (Gianturco et al., 1983). In a separate study, the conformation of apoE in the vicinity of the receptor-binding region was examined on several different lipoprotein classes with a panel of apoE monoclonal antibodies, and the LDL receptor-binding activity of these lipoproteins was determined (Krul et al., 1988). T h e results demonstrated that there is considerable heterogeneity in the expression of apoE epitopes in these various lipoprotein classes and that this heterogeneity was responsible, in part, for the differences observed in the receptor-binding activities within a given lipoprotein class. In addition, lipolysis of VLDL triglycerides from normal subjects also affects apoE-mediated binding to LDL receptors. Following lipolysis,


273

there is a marked increase (2- to 20-fold) in the ability of the lipolyzed products to interact with receptors and deliver cholesterol to cells (Sehayek et al., 1991), indicating that the apoE on these particles is changing from a receptor-inactive to a receptor-active form during hydrolysis of core triglycerides. Although these results can be interpreted as arising from a conformational change in apoE with remodeling of the lipid content of the VLDLs, other factors also could contribute, such as protein-protein interactions or the loss of the C apolipoproteins during lipolysis. Along similar lines, manipulation of the lipid composition of VLDLs by lipolysis or with lipid transfer proteins demonstrated that the VLDL lipid composition is a major factor in the ability of apoE to bind to the lipoprotein particle (Ishikawa et al., 1988). A major effect of cholesterol on the conformation of apoE was revealed by comparing the conformation on DMPC discs, on HDL,, and on spherical artificial microemulsion particles by circular dichroism (Mims et al., 1990). Conformational differences of apoE on different types of particles also were demonstrated using 13C NMR to probe lysyl microenvironments. When the apoE lysyl residues were labeled by reductive methylation with ["C]formaldehyde to allow detection, the lysyl microenvironments manifested dramatic differences on a discoidal particle compared to spherical particles (S. Lund-Katz et al., 1993). On spherical particles, two lysine microenvironments were observed, but on discoidal particles eight peaks were observed (apoE has 12 lysyl residues). These results indicate that apoE structure differs significantly on the two lipid surfaces. In a systematic study of the effect of the particle lipid composition on the conformation of apoE, conformation was shown to be affected by a number of parameters (Mims et al., 1990). The a-helical content was lower when apoE was bound to a spherical particle compared to a discoidal particle. It was concluded that this probably reflects the different ways in which the amphipathic helices interact with phospholipid on the two particles. With discoidal particles the interaction is primarily with phospholipid acyl side chains, whereas with spherical particles the interaction is with polar phospholipid head groups. In addition, the conformation of apoE was influenced by the diameter of the microemulsion particle and possibly by the order/ disorder of the lipid components. As discussed above, the 22-kDa fragment (residues 1-191) retains full receptor-binding activity. To define further the minimal amount of sequence necessary for maintaining full activity, a series of carboxylterminal truncations was produced by site-directed mutagenesis and expressed in Escherichiu coli (Lalazar et al., 1989). The truncations comprised residues 1-166, 1-170, 1-174, and 1-183. When the truncated proteins were complexed with DMPC and tested for receptor activity,

274

KARL H. WEISGRABER

the two shortest truncated proteins were found to be essentially inactive (-l%), whereas the 1-174 and the 1-183 variants possessed 19% and 85% normal binding activity, respectively. It was concluded that the 17 1-183 region contained sequences critical to receptor binding and that residues beyond 183 were not required. Either residues 171-183 contain one or more residues that interact directly with the receptor, or this region contains critical elements that help maintain the receptorbinding region (residues 136- 150) in an active binding conformation. Based on the apoE2 mutant discussed above, the latter alternative seems more probable. Because amino-terminal deletions have not been studied as systematically as carboxyl-terminal truncations, it is not clear exactly how much of the amino terminus is required to maintain receptor activity. Receptor-binding studies performed with the large cyanogen bromide fragment (residues 126-218; Fig. 9) and a synthetic peptide fragment spanning residues 129- 169 provide some insight (Innerarity et al., 1983). Although both fragments formed stable complexes with DMPC, only the cyanogen fragment displayed significant and reproducible binding activity (-10%) at the concentrations that were examined. In more recent studies performed with lipid-free synthetic peptide fragments, comprising residues 141-155 and a dimeric peptide of this sequence, no activity was detected with the monomer, but interestingly, very low levels were observed with the dimer (-1% of LDL activity) (Dyer et al., 1991). Together, these results can be interpreted as indicating that more than the immediate region around residues 136-150 is required for high-affinity binding to the LDL receptor and that critical residues amino terminal to residue 126 are required for full activity. Th e supposition is that a critical length of sequence within residues 1-135 also is required to maintain residues 136-150 in a proper conformation for high-affinity interaction with the LDL receptor when apoE is bound to lipid. This requirement is similar to the role suggested for residues 17 1- 183, discussed above.

E . Interaction with Heparin

Another property of potential metabolic importance that apoE shares with apoB is the ability to bind to various glycosylaminoglycans, including heparin (Mahley et al., 1979; Cardin et al., 1986; Weisgraber et al., 1986). Advantage has been taken of the apoE-heparin interaction to subfractionate apoE-containing from non-apoE-containing lipoproteins (Shelburne and Quarfordt, 1977; Weisgraber and Mahley, 1980). Gly-


275

cosylaminoglycans are components of proteoglycans, which are found in the extracellular matrix of tissues and organs. It has been suggested that the proteoglycans of the arterial wall interact with lipoproteins and, through this interaction, participate in the deposition of cholesterol associated with atherosclerosis (Srinivasan et al., 1972). In addition, lipoprotein lipase is present on the endothelial cell surfaces lining capillary beds in association with heparin-like proteoglycans, where it acts to hydrolyze the triglyceride-rich lipoproteins. It has been suggested that triglyceride-rich lipoprotein particles are anchored to the proteoglycans, allowing access of LPL to the triglyceride core (Landis et al., 1987). Consistent with the suggestion of a role for apoE in the anchoring process are studies demonstrating that addition of apoE to triglyceride emulsion particles resulted in the enhanced hydrolysis of these emulsions by LPL that was bound to a heparin-Sepharose matrix (Landis et al., 1987). It has been postulated that heparan sulfate proteoglycans on the surface of hepatocytes are responsible for the initial sequestration and binding of chylomicron remnants in the space of Disse (Mahley and Hussain, 1991). Because all of the major proteins involved in chylomicron metabolism (apoE, LPL, and HL) bind to heparan sulfate proteoglycans and have been demonstrated in the space of Disse (Clarke et al., 1983; Doolittle et al., 1987; Landis et al., 1987; Hamilton et al., 1990) along with heparan sulfate proteoglycans (Stow et al., 1985), the potential exists for concentrating key components in this compartment, where chylomicrons could be sequestered, processed, and taken up by hepatic receptors. In addition, LPL has been shown to increase the binding of remnants to LRP (Beisiegel et al., 1991). In support of the sequestration model, it has been demonstrated that the increased binding and uptake of apoE-enriched P-VLDL exhibited by a variety of cell types results from binding to cell surface heparan sulfate proteoglycans (Ji et al., 1993). These results are consistent with a role for heparan sulfate proteoglycans in the rapid clearance of chylomicron remnants from circulation, although it is not clear at present if heparan sulfate proteoglycans function alone or in combination with lipoprotein receptors in uptake by the liver. Using heparin as a model compound for heparan sulfate proteoglycan interactions, the region of apoE that binds to heparin was determined. It was known from earlier studies that the interaction of apoB and apoE with heparin involved arginyl and lysyl groups, probably in an ionic interaction with the negatively charged sulfate and carboxylate groups present in the carbohydrate structure of heparin (Mahley et al., 1979). Using an approach similar to that employed to map the region of

276

KARL H. WEISGRABER

apoE that bound to the LDL receptor, apoE was found to possess two heparin-binding sites (Cardin et al., 1986; Weisgraber et al., 1986). T h e first binding site, in the vicinity of residues 142-147, corresponds almost exactly to the region that binds the LDL receptor. Heparin binding to this site, which coincides with the epitope of the monoclonal antibody 1D7, is inhibited by this antibody (Weigraber et al., 1986) (Fig. 10). T h e location of the second heparin-binding site is less clearly defined. This site is partially blocked by a monoclonal antibody, 3H1, whose epitope is located between residues 243 and 272 (Weisgraber et al., 1986). However, studies with synthetic peptides of sequences in the carboxyl terminus of apoE indicate that there is a heparin-binding site between residues 202 and 243 (Cardin et al., 1986). The partial inhibition by the 3H 1 antibody could result from incomplete steric interference of the 202-243 site or from the antibody blocking a third binding site

1107

1D7

IU D I 0 Heparin Binding Antibody Epitope FIG. 10. Schematic model depicting the location of the two apo-E heparin-binding sites and their spatial relationship to the 1D7 and 7C9/6C5 epitopes. Left: I t is proposed that one heparin-binding site is located in the center of the apo-E molecule and that this binding site coincides with the 1D7 epitope. A second heparin-binding site is located in the carboxyl-terminal region of the protein. In the free protein, it is proposed that the second heparin-binding site is in a close spatial relationship with the 6C5 epitope such that antibody interaction with this epitope inhibits heparin binding to the second site. The epitope for 7C9 is also located at the amino terminus of apoE; however, it differs from the 6C5 epitope in that antibody interaction with this site does not interfere with heparin binding at the carboxyl-terminal heparin-binding site. Rzghf: When apo-E is complexed with DMPC, a recombinant is produced in which the first heparin-binding site and the 6C5 and 7C9 epitopes are available for heparin binding or antibody interaction, respectively, but the second heparin-binding site is “masked” by the DMPC and does not bind to heparin. From Weisgraber el al. (1986).


277

within its epitope and allowing the 202-243 site to interact. In either case, the carboxyl-terminal site(s) is only available for binding when apoE is in the lipid-free state; this site is masked in the presence of lipid (Weigraber et al., 1986).

IV. IMPACTOF STRUCTURE ON FUNCTION A. Relationship of Apoliprotein E Variants to Type III Hyperlipo@oteinemia

The early studies of Utermann (Utermann et al., 1975) and Zannis and Breslow (1980) recognized that a particular apoE phenotype, later designated as the E2/2 phenotype, was associated with type 111 hyperlipoproteinemia. However, only 1-2% of E2/2 subjects develop clinical features of the disorder [for a review of this disorder, see Mahley and Rall (1989)l. T o explain this discordance, it was suggested that for expression of the disorder, a second factor (either genetic or environmental) was required in addition to the E2/2 phenotype (Utermann et al., 1979). Thus, the primary defect in type 111 hyperlipoproteinemia is the presence of a defective form of apoE, i.e., apoE2, but this presence is not sufficient in itself to cause the disorder. It was demonstrated that the structures of apoE2 from normal as well as hypocholesterolemic subjects were identical, with cysteine at position 158 (Rall et al., 1983a). Also, regardless of the source, all apoE2s were equally defective. Thus, a structural heterogeneity in the apoE2 was not the basis for the fact that not all subjects with the E2/2 phenotype develop type 111 hyperlipoproteinemia. However, heterogeneity has been observed in the receptor-binding activity (1-60% of normal) of apoE from a number of type I1 I hyperlipoproteinemic subjects, suggesting that structural heterogeneity does occur (Schneider et al., 1981). This suggestion was confirmed in 1982 when differences in the receptor-binding activities of apoE2 were demonstrated to result from structural heterogeneity (Rall et al., 1982b). Since then, a number of other apoE variants have been identified that focus in several different positions (see Table I). Thus, with structural heterogeneity at the major isoelectric focusing positions, it is not sufficient to designate apoE isoforms solely on the basis of isoelectric focusing position, i e . , E2, E3, or E4. The convention used in this review is to relate variant structures to the common apoE3 structure, often referred to as the parent form of the protein (Zannis et al., 1982), i.e., apoE2(Arg15*-3 Cys) or apoE4(Cys1l 2 --* Arg). However, as a matter of convenience, the two common apoE variants, apoE2(Arg15' + Cys) and apoE4(Cys'l2 + Arg), are referred to simply as apoE2 and

278

KARL H. WEISGRABER

apoE4, respectively, and the other variants are referred to by the more descriptive designations. Another interesting feature of type 111 hyperlipoproteinemia has been recognized. Although the common apoE2 isoform requires the homozygous state plus an additional factor for expression of the disorder (recessive transmission with low penetrance), with several less common variants the disorder is expressed in the heterozygous state, and most, if not all, carriers display clinical features (dominant transmission with high penetrance) (Rall and Mahley, 1992) (Table I). T h e basis for this distinction in expression is not clear at present. The position of the substitution has been suggested as critical (Rall and Mahley, 1992), ie., substitutions within the receptor-binding region lead to a dominant variant, whereas the 158 substitution is recessive and exhibits a very low degree of penetrance. In addition, one difference between dominant and recessive variants also appears to be related to whether the receptor-binding activity of the variant can be modulated. It was suggested that with dominant variants, receptor activity is permanently defective (Rall and Mahley, 1992). Therefore, even in the presence of a receptor-competent form, i.e., the heterozygous condition, the lipoprotein particles with mixtures of the two forms would be defective because multiple apoE molecules participate in the interaction with receptors in the clearance of these particles. In contrast, the binding of the apoE2 is thought to be variable, ranging from severely defective to near normal. As discussed earlier, the binding activity of apoE2 can be modulated by charge modification with cysteamine o r by the removal of the carboxyl terminus of the protein, whereas charge modification of the apoE3(Arg112+ Cys, ArgI4' + Cys) has no effect on binding activity (Rall et al., 1989). This indicates that the arginine side chain at position 142 is an absolute requirement for receptor interaction and that the presence of positive charge in itself is not sufficient. The requirement for the arginyl side chain at position 142 also argues for the direct involvement of this residue in binding to the receptor. In addition, the receptor-binding activity of the apoE2 can be modulated by the lipid composition of lipoprotein particles (Innerarity et al., 1986). The binding activity of the d < 1.006 lipoproteins (P-VLDLs) of a type 111 subject with this variant was tested before and after dietary intervention. Before intervention, the subject expressed the clinical features of type 111 hyperlipoproteinemia: plasma triglycerides and cholesterol were elevated (670 and 725 mg/dl, respectively). After dietary intervention (a common treatment for this disorder, to which patients often respond satisfactorily), the subject lost 34 Ibs. and his


279

plasma lipid levels were reduced dramatically (final concentration of 77 and 92 mg/ml for plasma triglyceride and cholesterol levels, respectively). The pre- and postdiet P-VLDLs differed markedly in their lipid composition, with the postdiet P-VLDLs being less cholesterol rich (a threefold reduction in the cholesterol: triglyceride ratio). T h e prediet P-VLDLs bound to LDL receptors with a relatively low affinity and were significantly activated by cysteamine treatment. In marked contrast, the postdiet P-VLDLs bound with high affinity to LDL receptors (30-fold better) and these lipoproteins were not significantly activated by cysteamine modification. The binding activities of both P-VLDLs were shown to be mediated through apoE with the monoclonal antibody 1D7. These results clearly indicate that receptor-binding activity of the 158 variant can be influenced by the lipid composition of the lipoprotein particle to which it is bound. This modulation by lipid composition is consistent with the fact that expression of type I11 hyperlipoproteinemia in E2/2 subjects requires a second factor, which in many instances is a familial lipid disorder (Utermann et al., 1979), and that subjects with normal or below-normal lipid levels do not express the disorder. In contrast to apoE2, apoE3-Leiden is another example of a dominant variant (Table I). It does not satisfy the criterion that the binding activity of a dominant variant is not modulated. This variant is unlike all other variants of apoE in that it does not arise as the result of a point mutation. This variant has a seven-amino acid insertion, a direct repeat of residues 121 to 127 (Wardell et al., 1989). The binding activity is approximately 25% of normal; however, when the carboxyl-terminal domain is removed by thrombin digestion, the binding activity of the 22-kDa domain is near normal (Wardell et al., 1989). This result indicates that conformational constraint on the receptor-binding region is imparted by the insertion that is relieved when the carboxyl terminus is removed, similar to the case of the apoE2 variant. Although this might be viewed as a form of modulation, in the intact molecule, the binding defect of apoE3-Leiden has been suggested as permanent in the sense that its activity cannot be modulated on lipoprotein particles. In addition, apoE3-Leiden has recently been shown to have an enhanced preference for VLDL particles, a property that results in higher levels of the apoE-Leiden on these particles relative to the LDL receptoractive product of the second allele in the heterozygous state. The higher proportion of a defective form of apoE relative to normal forms on remnant particles probably further contributes to the delayed clearance of these particles (Fazio et al., 1993). This differential distribution of apoE variants to triglyceride-rich lipoproteins, including rem-

280

KARL H. WEISGRABER

nants, has been suggested to be a second potential mechanism of dominant expression of type 111 hyperlipoproteinemia (Fazio et al., 1993). As discussed above, the apoE3(ArglL2--* Cys, Arg'42 --* Cys) variant also is associated with dominant expression of the type 111 disorder, and the receptor-binding activity of the protein cannot be modulated by cleavage (Horie et al., 1992) or charge modification (Rall et al., 1989). However, additional characteristics of this variant may be important for dominant expression. Because of the presence of arginine at 112, as in apoE4, this variant displays a preference for the triglyceride-rich lipoproteins (Horie et al., 1992) (see Sections IV,C and D). This preference means that in affected heterozygotes with a normal apoE3 allele, as is the case in the family in which this mutation occurs, the defective variant is enriched relative to normal apoE3 on the triglyceride-rich particles, similar to the situation with apoE3-Leiden. In addition, the 142 substitution has been shown to reduce the affinity of this variant for heparin (Horie et al., 1992). If binding to heparin-like structures mediated by apoE is important in the lipolytic processing of remnant particles or in the clearance of these particles by the liver, as recently suggested (Mahley and Hussain 1991; Ji et al., 1993) and discussed in Section III,E, then the reduced affinity for heparin displayed by this variant also might contribute to the accumulation of remnant lipoproteins in this family. Thus, a reduced affinity for heparan sulfate proteoglycans represents a potential third mechanism for dominant expression of type 111 hyperlipoproteinemia (Horie et al., 1992).

FIG. 11. Effect of receptor-binding-defectiveforms of apolipoprotein E on the hepatic clearance of plasma lipoproteins. Defective forms of apoE result in reduced clearance of chylomicron remnants, VLDLs, and IDLs and their accumulation in plasma. The chylomicron remnants and VLDLs are enriched in cholesteryl esters as the result of CETP activity, and together they constitute the P-VLDLs, which are a hallmark of type Ill hyperlipoproteinemia. A block in the conversion of IDLs to LDLs by hepatic lipase by the presence of an abnormal apoE form also is indicated.


28 1

The effect of a defective form of apoE on lipoprotein clearance is summarized in Fig. 11. Chylomicron and VLDL remnants accumulate in the plasma as the direct result of a decrease in the normal uptake by hepatic receptors caused by the presence of the defective apoE on these particles. As these particles circulate for prolonged periods, they become cholesterol enriched by the action of transfer of cholesteryl ester from HDL in exchange for triglyceride by CETP. Together, the cholesteryl ester-enriched chylomicron and VLDL remnants make up the P-VLDLs, which are a hallmark of type 111 hyperlipoproteinemia. In addition, IDLs also accumulate because of a decrease in apoE-mediated clearance. Finally, it appears that conversion of IDLs to LDLs also is impaired in the presence of apoE2. In vitro studies demonstrate that apoE2 is less effective than apoE3 in promoting the lipolysis of P-VLDLs (Ehnholm et al., 1984). Similarly, this variant is also less effective than apoE3 in activating hepatic lipase in model monolayer systems (Thuren et al., 1992). B . Effect of Apolipopotein E Heterogeneity on P h m a Lipid Concentrations

It has been established in a number of populations that the three major isoforms of apoE have a significant impact on interindividual plasma cholesterol and LDL concentrations [for a review of this topic, see Davignon et al. (1988) and Hallman et al. (1991)l. For the purposes of the discussion on apoE structure and function, the results can be summarized as follows: It has been estimated that 60% of the variation in plasma cholesterol levels is genetically determined and that approximately 14% of that variation is the result of apoE heterogeneity (Davignon et al., 1988). The ~4 allele is associated with the highest plasma cholesterol levels and the 82 allele with the lowest levels, with the 83 allele being intermediate. Although apoE2 is associated with type 111 hyperlipoproteinemia, clinical expression of the disorder with elevation of plasma lipid concentrations requires a second genetic or environmental factor, as discussed above. The majority of subjects with the E2/2 phenotype actually have low plasma cholesterol concentrations. With what is known regarding the properties of the three common isoforms, we can begin to understand the basis for the differences in cholesterol levels among the three alleles. In the case of apoE2, remnant lipoproteins are cleared from circulation at a slower rate than normal and the conversion of VLDLs to LDLs appears to be retarded. This defect in remnant clearance leads to the up-regulation of hepatic LDL receptors, which contribute to a further lowering of plasma LDL concentrations (Davignon et al., 1988). It is known that E2/2 subjects with

282

KARL H. WEISGRABER

normal or subnormal plasma lipid concentrations accumulate j3-VLDLlike lipoproteins and have elevated plasma apoE concentrations (Utermann, 1985). Although apoE3 and apoE4 have identical affinities for the LDL receptor (Weisgraber et al., 1982), it is known that remnants are cleared more efficiently in subjects with apoE4 (Weintraub et al., 1987). T h e basis for the more rapid clearance appears to be associated with the property that, relative to apoE3 and E2, apoE4 displays a preferential distribution to the triglyceride-rich lipoproteins in plasma (Gregg et al., 1986; Steinmetz et al., 1989; Weisgraber, 1990). T h e resulting higher concentration of apoE on these particles would account for their efficient clearance. The structural basis for this differential distribution is discussed in detail in Sections IV,C and IV,D. It has been suggested that this effective clearance of remnant lipoprotein particles leads to a down-regulation of the hepatic LDL receptors, resulting in elevation of plasma LDL concentrations (Gregg et al., 1986; Weintraub et al., 1987; Davignon et al., 1988).

C . Domain Interactions Although the amino- and carboxyl-terminal structural domains of apoE appear to be independently folded in the lipid-free state, as discussed in Section II,C, there are several examples wherein amino acid substitutions in one domain appear to affect the properties of the other domain. These effects fall into one of two general categories: (1) the effect on the distribution of apoE among the various lipoprotein classes and (2) the effect on receptor-binding activity. Domain interactions were first observed when ultracentrifugation was used to examine the lipoprotein profile of subjects homozygous for apoE3 or apoE4. It was observed that apoE3 and apoE4 displayed differences in their distribution among lipoprotein classes, with apoE3 showing a preference for HDL and apoE4 a preference for VLDL (Gregg et al., 1986). This differential distribution has been confirmed in heterozygous E4/3 subjects, using agarose column chromatography to separate lipoprotein classes (Steinmetz et al., 1989; Weisgraber, 1990). This phenomenon was studied further by incubating '251-labeled isoforms and fragments with plasma and then separating the lipoproteins by agarose column chromatography (Weisgraber, 1990). As shown in Fig. 12, when incubated in plasma from an E3/3 subject, apoE4 displayed a preference for VLDL, and apoE3 a preference for HDL; apoE2 also displayed a preference for HDL, indicating that the cysteine-arginine interchange at position 158 has little, if any, effect on lipoprotein distribution (Weisgraber, 1990).


283

Fraction Number FIG. 12. Distribution of 1251-labeledapoE4, apoE3, and apoE2 among plasma lipoproteins. Iodinated apoE was incubated with plasma for 2 hr at 37°C. The plasma was then subjected to agarose chromatography on Bio-Gel A-5m. Arrows indicate the elution positions of VLDL, LDL, and HDL. From Weisgraber et al. (1990).

Because the apoE3 and apoE4 variants differ only by the cysteinearginine interchange at position 112, this substitution must be involved in the differences in lipoprotein distribution. The distribution of cysteamine-modified apoE3 was shown to be indistinguishable from the distribution of apoE4, indicating that charge at 112 was critical for the effect and that arginine specifically was not required (Weisgraber, 1990). The requirement for positive charge at 112 suggests that salt bridges may be involved. When the distributions of the apoE3 and apoE4 22-kDa fragments were determined, both were found to distribute primarily in the lipoprotein-poor region of the column, i.e., they did not associate with any of the major lipoprotein classes (Weisgraber, 1990). The 10-kDa fragment, which has been suggested to represent the major lipid-binding region of the protein, displayed a unique lipoprotein distribution of its own, in that it did not resemble that of either apoE3 or apoE4 (Weisgraber, 1990); it distributed primarily with LDL and small HDL. Based on these results, it was suggested that the amino acid at position 112, within the amino-terminal domain, influences the

284

KARL H. WEISGRABER

lipid-binding properties of the carboxyl-terminal domain, indicating an interaction or communication between the domains (Weisgraber, 1990). As discussed in Section V,B, the X-ray crystal structures of apoE3 and apoE4 provide clues as to how this domain interaction might occur. The other examples of domain interaction involve binding to the LDL receptor. Two of these have been considered in another context in Section IV,A. As discussed, the binding activity of both the apoE2 and the apoE3-Leiden variants increases when the carboxyl-terminal domain is removed by thrombin digestion (Innerarity et al., 1984; Wardell et al., 1989). These results have been interpreted as indicating that, with these mutations, the carboxyl-terminal domain is capable of modulating the receptor-binding activity of the amino-terminal domain, probably by influencing the conformation of the receptor-binding region. In addition, the apoE5(Glu3+ Lys) displays the interesting property of having binding activity that is increased significantly over normal levels (Dong et al., 1990; Wardell et al., 1991), indicating that the amino terminus also is capable of affecting receptor-binding activity. D. Effect of Cysteinyl Residues on Function The cysteine at position 112 in apoE3 has been shown to be capable of forming either homodiiners (Weisgraber and Shinto, 1991) or heterodimers with apoA-I1 (Weisgraber and Mahley, 1978) (Fig. 13). It has been demonstrated that both forms display a marked preference for HDL, indicating that the disulfide-linked dimers also contribute to the preference of apoE3 for HDL (Weisgraber, 1990; Weisgraber and Shinto, 1991). The homodimer of apoE3 is readily removed from HDL by ultracentrifugation and is observed in HDL only when column chromatography is used to fractionate plasma (Weisgraber and Shinto, 1991). The disulfide-linked forms represent a substantial proportion of the apoE3 in plasma, with the apoE3-A-I1 heterodimer and the homodimer accounting for -26% and -28%, respectively, of total apoE3 (Weisgraber and Shinto, 1991). It is not clear how o r where the dimerization takes place. Studies with liver-derived cells in culture suggest that the dimerization takes place extracellularly (Weisgraber and Shinto, 1991). Possibly this could occur on the surface of lipoprotein particles, where the monomeric units could be brought together in close spatial contact. Both dimers reduce the ability of apoE3 to bind to the LDL receptor. The binding activity of apoE3-A-I1 and the homodimer is 30% (Innerarity et al., 1978) and 20% (Weisgraber and Shinto, 1991), respectively, of the activity of the monomer. It has been suggested that the forma-


285

FIG.13. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis and immunoblotting of human plasma. Plasma (1 PI) from subjects with the E3/3, E4/4, and E4/4 phenotypes was subjected to SDS-PAGE followed by transfer to a nitrocellulose filter and detection with '251-labeledaffinity-purified antihuman apoE immunoglobulin G. One sample included 2-mercaptoethanol (tBME)as a disulfide reducing agent. From Weisgraber and Shinto (1991).

tion of the disulfide-linked dimers modulates the in vzvo binding activity of apoE3 (Innerarity et al., 1978; Weisgraber and Shinto, 1991). In addition, it has been postulated that the formation of receptor-bindingdefective dimers contributes, in part, to the postulated deficiency of plasma apoE3 that is available for the clearance of lipoproteins (Weisgraber and Shinto, 1991), which was discussed in Section II,D.

v.

THREE-DIMENSIONAL STRUCTURE OF APOLIPOPROTEIN E 22-kDa FRAGMENTS

A . Apolipopfotein E 3 Because the 22-kDa fragment of apoE3 is a stable, monomeric structure and contains the receptor-binding function of the protein, it was an attractive first target for crystallographic efforts. The first crystals suitable for X-ray diffraction studies were obtained using polyethylene glycol (PEG) 8000 and n-octyl-a-D-glucopyranosideand belonged to the orthorhombic space group P21212, (Aggerbeck et aL, 1988b). For the final structure determination, the conditions were modified to use 15% PEG 400 and n-octyl-a-D-glucopyranoside (Wilson et al., 1991). The crystals diffract to 2.5 A and had the following unit cell dimensions: a =

286

KARL H. WEISGRABER

-

41.3 A, b = 54.5 A, c = 87.0 A, a = p = y 90". The final model contains residues 23-165 but lacks residues 1-22 and 166-191, which are believed to be disordered in the crystal (Fig. 14) (Wilson et al., 1991). The structure contains five helices, representing more than 80% of the residues present in the 22-kDa fragment. Four of the helices are arranged in an antiparallel four-helix bundle, which is a common folding motif of a-helical proteins (Wilson et al., 1991). As indicated in Fig. 14, helix 1 begins at residue 23 and extends to residue 42. Helix 1 is connected to helix 2 by a short connecting helix, encompassing residues 44-53. Helix 2 begins with residue 54 and extends to residue 81. The chain is reversed with a turn defined by residues 82-86. Helic 3 extends from residues 87 to 122. The chain reverses again with a turn (residues 123-129) and helix 4 extends from residue 130 to 165. It is within helix 4 that the receptor-binding region of the protein is located (residues

FIG. 14. Ribbon model of the structure of the 22-kDa fragment of human apolipoprotein E3. Four of the five helices (helices 1-4) are arranged in an antiparallel fourhelix bundle. The four-helix bundle can be viewed as a rectangle, with approximate dimensions of 20 X 20 x 65 A. The receptor-binding region of apoE (-residues 130150) is indicated on helix 4. The residue numbers at the start and end of each helix are indicated.


287

136-150). The lengths of the four helices in the four-helix bundle are 19, 28, 36, and 35 amino acids, respectively. This structure is unusual among four-helix bundle proteins in that three of the four helices are much longer than the average helical length of 18 residues in the other proteins with this folding motif. It is interesting to note the similarities of the apoE3 22-kDa fragment structure and apolipophorin 111 (Breiter et al., 1991), an apolipoprotein isolated from the hemolymph of miqratory locusts (see Soulages and Wells, this volume). Both contain elongated helical bundles-five extended helices in the case of apolipophorin I11 (Breiter et al., 1991). It will be of interest in the future, as structures of other apolipoproteins are solved, to determine whether the elongated helical bundle is a common structural element of other plasma apolipoproteins. The helices in the apoE3 22-kDa fragment are amphipathic in nature, although their properties, hydrophobic moments, and charge distributions differ from those of the typical apolipoprotein helix (Segrest et al., 1992). The helices in the 22-kDa fragment are classified as G* according to the nomenclature of Segrest et al. (1992). According to this classification, helices with the G* designation share some properties in common with both typical apolipoprotein helices and helices from stable, globular proteins (for more complete discussion, see Segrest et al., this volume). The hydrophobic side chains are sequestered in the interior of the bundle and the hydrophilic side chains are solvent exposed on the surface. This packing of the hydrophobic residues in the interior of the bundle probably contributes to the stability of the structure. Leucine side chains occur approximately every seven residues, forming a leucine zipperlike structure originally proposed by Landschulz et al. for the dimerization of the C/EBP-type transcription factors (Landschulz et al., 1988). This arrangement of leucine side chains in the bundle appears to stabilize the interfaces between helix 1 and helix 4 and between helix 2 and helix 3. The 22-kDa structure includes 24 acidic and 24 basic residues, most of which are involved in intra- and interhelical salt bridges (Table 11). It is likely that the interhelical salt bridges also contribute, along with the hydrophobic packing, to the unusual stability of this fragment compared to other apolipoproteins. It is interesting to note that, with the exception of Arg-147 and Arg-150, the basic residues within the receptor-binding region (residues 136- 150) are not involved in salt bridges and are solvent exposed. Exposure of these basic amino acids results in a large area of positive electrostatic potential over the receptor-binding region in helix 4 (Wilson et al., 1991). In addition, because most of the residues within this region are not salt bridged,

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KARL H. WEISGRABER

TABLE I1 Salt Bridges in Four-Helix Bundle Structures of Apolipoprotein E3 22-kDa Fragments Intrahelical

Helix

Interhelical

Helices

Asp-35 - Arg-38 Arg-6 1 - Asp-65 Glu-66 - LYS-69 Arg-92 -Glu-96 Arg-114 - Glu-12 1 Glu- 13 1 - Arg- 134 Arg-147 - Asp-151 Arg-150 - Asp- 151 ASP-153-LYS-157 ASP-154-LYS-157 Asp- 154 - Arg- 158

1 2 2 3 3 4 4 4 4 4

Arg-32 - Glu-66 Arg-25 - Glu-70 Asp-107 - Arg-147 Asp-1 10 - Arg- 147 Arg- 103 - Asp-15 1 Arg-103 - Asp-154 Glu-96 - Arg-158 -

1 and 2 1 and 2 3 and 4 3 and 4 3 and 4 3 and 4 3 and 4

4

-

-

-

they would be free to interact with the LDL receptor, as suggested in Section III,D. B . Comparison of Apolipoprotein E2, E 4 , and E3 Structures The structures of the apoE2 (Wilson et al., unpublished observation) and apoE4 (Wardell et al., 1993; M. R. Wardell et al., unpublished observation) 22-kDa fragments have recently been determined. In general, the apoE2 structure resembles the apoE3 structure in that the four-helix bundle folding motif is retained; however, there are significant changes in the vicinity of the residue 158 substitution site (Wilson et al., 1993). In the apoE3 structure, the Arg-158 (helix 4) forms a salt bridge with Asp-154 (helix 4) and Glu-109 (helix 3) (Table 11) (Fig. 15). In addition, salt bridges are formed between Arg-92 and Glu-96, Asp151 and Arg-103, Asp-151 and Arg-150, and Asp-154 and Arg-103 (Table 11). Several of these salt bridges link helix 3 with helix 4. With the neutral cysteine at position 158 in apoE2, this pattern of salt bridges is eliminated with the exception of the Arg-92 and Glu-96 pair. A new salt bridge between Arg-150 and Asp-154 is added (Fig. 15) (Wilson et al., 1993). These salt bridge rearrangements produce significant distortions in the backbone of helix 3. The slight kink in the apoE3 structure at Gly-105 becomes more pronounced in apoE2, with the aminoterminal half of this helix being displaced away from the bundle (Wilson et d., 1993). As discussed in Section IILD, several lines of evidence indicate that Arg-158 does not interact directly with the LDL receptor, but that this residue has an indirect effect on the receptor-binding region (residues


289

APOm

ApoE3

FIG. 15. Salt bridges in the 22-kDa fragments of apolipoprotein E3 and E2. In apoES, Arg-158 forms a salt bridge (---) with Glu-96 and Asp-154. In apoE2 the loss of Arg-158 results in the formation of a salt bridge between Asp-154 and Arg-150. Arg-150 swings out of the basic cluster of side chains in the receptor-binding domain to form this bond.

136-150). The crystal structure of apoE2 provides a plausible explanation for the indirect effect. i n the apoE3 structure, Arg-150 is exposed to solvent and lies within the region of positive electrostatic potential (Fig. 15). However, in apoE2, with Arg-150 paired to Asp154, the Arg-150 side chain swings out of this region of positive potential, reducing the area of positive potential over residues 136-145 (Wilson et d.,1993).Thus, the removal of the Arg-150 side chain from the receptor-binding region could provide the mechanism by which the substitution of Arg- 158 by cysteine indirectly reduces the receptorbinding activity of the apoE2 variant.

GlylOS Arg 61 Arb112

FIG. 16. Salt bridges in the 22-kDa fragment of apolipoprotein E3 and E4. Substitutions of arginine at position 112 in apoE4 results in the formation of a new salt bridge between Arg-112 and Glu-109. The Arg-61 side chain assumes a new position in the apoE4 structure compared to that in apoE3.

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KARL H. WEISCRABER

The structural changes in the apoE4 22-kDa fragment are more subtle. T h e four-helix bundle is indistinguishable from the apoE3 structure (Wardell et al., 1993; M. R. Wardell el al., unpublished observation). The only changes are the reorientation of two side chains at positions 109 and 61. In apoE4, the Glu-109 swings down to form a salt bridge with Arg-112. T o accommodate this shift, Arg-61, which normally occupies the space above Cys-112, swings out of the way (Fig. 16). Because this substitution appears to have an effect on the lipid-binding properties of the carboxyl-terminal domain, it is likely that this substitution is acting in an indirect manner, possibly altering salt bridge interactions between the two domains. T h e answer to this question awaits the crystal structure of the intact protein. VI.

LIPIDBINDING

A. Effect of Lipid on Structure

Determination of the three-dimensional structure of the 22-kDa fragment of apoE represents a major advance in the understanding of the structure-function of this plasma apolipoprotein. However, a complete correlation of the structure and the function of apoE will require an equally comprehensive understanding of the relationship between the lipid-free and lipid-associated forms, as well as of the effect of lipid on conformation. This point is underscored by the fact that lipid association is required for high-affinity binding to the LDL receptor (Innerarity et al., 1979). The hydrophobic side chains of the 22-kDa four-helix bundle are directed toward the interior of the bundle. Because the hydrophobic faces of apolipoprotein amphipathic helices are thought to interact with the hydrophobic acyl chains of phospholipids, the working hypothesis is that the four-helix bundle undergoes a conformational change when associated with lipid, such that the bundle opens without a major disruption of the a-helical structure (Fig. 17). This “opened” structure would have a broad hydrophilic face and a hydrophobic face. The hydrophobic face would then be available to interact with lipid. T o test this model, the surface properties of the 22-kDa fragment at an air-water interface have been examined. The air-water interface system has been used extensively to model the interaction of apolipoproteins with lipid (Phillips and Sparks, 1980; Shen and Scanu, 1980; Camejo and MUAOZ,1981; Phillips and Krebs, 1986). When the 22-kDa fragment was spread as monomolecular film in a Langmuir trough, the surface pressure-molecular area isotherm was calculated to be 16 A2/

-

APOLJPOPROTEJN E STRUCTURE-FUNCTION

29 1

Hydrophlllc Faces Unfolding Hydrophoblc Faces

4-helix bundle

FIG. 17. Model of the interaction o f the four-helix bundle structure of the apoE3 22-kDa fragment with lipid. The four-helix bundle structure as it exists in solution is shown on the left. On the right, in the presence of lipid, the bundle “opens” without disrupting a helices, exposing the hydrophobic core of the bundle and making it available to interact with lipid (Weisgraber el al., 1992).

residue (Weisgraber et al., 1992), comparable to data for other apolipoproteins (Shen and Scanu, 1980; Yokoyama et al., 1985; Krebs et al., 1988) and in the range (13- 19 A’lresidue) for a-helical homopolypeptides, which lie coplanar with the surface (Malcolm, 1973). An apoA-I control gave a similar value (Weisgraber et al., 1992). This agreement of the limiting molecular areas of the 22-kDa fragment and apoA-I indicates that both adopt a similar conformation at the air-water interface. Agreement with the model peptide values indicates that the helices of the 22-kDa fragment lie coplanar with the surface. The molecular area of -16 A2/residue implies that one molecule of the 22-kDa fragment should occupy an area of -3000 A2. As shown in Fig. 14, the four-helix bundle can be viewed as a rectangular “box” with approximate dimensions of 65 X 20 X 20 A, with each helix forming an edge of the box. Shown in Fig. 18 are three possible orientations of the bundle at the air-water interface with the calculated areas that each orientation would occupy at the interface. In the first possibility, the bundle is oriented so only one of the minor faces of the bundle is at the interface. In the second, one of the major faces is aligned with the interface. With these two possibilities there is no rearrangement of the structure and the two orientations would occupy -400 and 1300 A2, respectively. However, if the bundle undergoes a conformational change by interacting with the interface such that the

-

292

KARL H. WEISCRABER

FIG. 18. Three possibilities for the orientation of the four-helix bundle structure of the apoE3 22-kDa fragment at an air-water interface. The calculated molecular area of each orientation based on the bundle dimension in Fig. 16 is indicated above each structure.

bundle “opened up” or unfolded while maintaining the helical structure, it would be predicted to occupy -2600 (third orientation in Fig. 18). This value is in good agreement with the value obtained from the surface studies. Thus, the conclusion was reached that the four-helix bundle undergoes a structural reorganization at the interface surface (Weisgraber et al., 1992). In this reorganization, the hydrophobic faces of helices, which are shielded from solvent in the interior of the bundle, would now be directed toward air, maintaining a hydrophobic environment. By extrapolation to a lipid surface, the four-helix bundle would be predicted to undergo a similar structural reorganization. This conformational reorganization is similar to what has been suggested for apolipophorin I11 when the circulating lipid-free form of the protein becomes associated with the high-density lipoprotein particle present in hemolymph (see Soulages and Wells, this volume). B . Carboxyl-Terminal Lip&-Binding Repons Across species the carboxyl-terminal region of apoE up to approximately residue 288 in the human sequence is highly conserved (Fig. 1). Studies to determine the carboxyl-terminal regions of apoE responsible for lipid binding and tetramer formation have been performed using three carboxyl-terminal truncations (Westerlund and Weisgraber, 1993). As shown in Fig. 19, the carboxyl terminus beyond position 191 contains three predicted a-helical regions. Two of the helices, residues

293


Class A amphipathic helices

Class 0' helix

-.

Characteristics binds to VLDL, IDL, HDL exists as tetramer binds to VLDL HDL 25% lipoproteh free monomerk

COOH 223

FIG. 19. Carboxyl-terminal truncations of apolipoprotein E3. The predicted secondary structure of the carboxyl terminus of each of the four carboxyl-terminal truncated proteins is compared with that of intact apoE (top) and the 22-kDa fragment (bottom). Shown on the right are the characteristics of each protein with respect to lipoprotein binding and the ability of the lipid-free form to tetramerize.

203-223 and 225-266, are classified as the class A type (Segrest et al., 1992; also see Segrest et al., this volume), whereas the third, residues 268-289, is a G' helix. The structure beyond residue 289 is predicted to be random. The first variant was truncated at residue 266, eliminating the G' helix and the unstructured region (Fig. 19). T h e second and third variants were truncated at residue 244 or 223, respectively. These truncations eliminated, respectively, half or all of residues 225-266, Class A (Fig. 19). Using intact apoE and the 22-kDa fragment as controls, iodinated truncated variants were assessed for their ability to associate with lipoproteins after incubation with plasma followed by gel filtration to separate the VLDL, IDL, and HDL classes. In addition, the state of association of the various fragments in the lipid-free state was assessed by gel filtration and sedimentation equilibrium centrifugation (Westerlund and Weisgraber, 1993) The results are summarized in Fig. 19. Removal of the terminal 33 residues resulted in a partial loss in lipoprotein binding, particularly in HDL; -25% of the 266 variant eluted in the lipoprotein-poor region of the column. Agarose gel electrophoresis indicated that this fraction was not associated with a lipoprotein particle (Westerlund and Weisgraber, 1993). Also, the 266 variant existed as a monomer in the lipid-free state. The results suggest that the G' helix

294

KARL H. WEISGRABER

plays a role in lipoprotein association and is critical for tetramer formation. T h e unstructured region beyond residue 289 is probably not important in either process because it is not well conserved across species. With removal of residues 245-266, the binding to VLDL and most of the remainder of binding to HDL was lost, and >65% of the protein was eluted in the lipoprotein-free fraction. T h e 223 variant exhibited similar characteristics, as did the 244 variant, including being monomeric in solution. It is interesting to note that, like the 22-kDa fragment (residues 1-191), the 244 and 223 variants still retained the ability to bind to phospholipid (DMPC) and form discoidal particles (Westerlund and Weisgraber, 1993). The results of this analysis indicate that major determinants for lipoprotein association are located at the extreme carboxyl terminus of apoE, the G* helix, and the carboxyl-terminal half of the Class A helix spanning residues 225-266. Elements that contribute to VLDL association appear to reside in the 245-266 region. I t is interesting that although the 244 and 223 variants contain long stretches of putative a-helical structure with class A potential, these fragments do not associate significantly with lipoprotein particles. Results from lipid-binding studies with synthetic peptide fragments of sequences within the carboxyl terminus of apoE are consistent with the truncation results. Four peptides (residues 202-243, 2 1 1-243, 263-286, and 267-286) were examined for their ability to bind to DMPC (Sparrow et al., 1992). It was interesting that only the 263-286 peptide formed a stable complex with DMPC, with -80% helical content, indicating the importance of the G* helix in this region for lipid binding. It was concluded that the two class A helices between residues 202 and 243 lacked a high enough hydrophobic moment to form stable complexes, whereas the 263-286 region could form a complex because of a higher hydrophobic moment (Sparrow et al., 1992).

FUTUREDIRECTIONS T h e determination of the three-dimensional crystal structure of the 22-kDa fragment of apoE in 1991 represented a major milestone in the studies of the structure and function of apoE. With this structure, it is now possible to understand and interpret much of what was known previously about the protein. In addition, identification of the structures of the apoE2 and apoE4 variants provides new insight into how apoE interacts with the LDL receptor and how the preference for different lipoprotein classes might be influenced by structure. These structures represent the beginning of the next level of understanding of how VII.


295

this protein functions in its various roles in lipoprotein metabolism. However, much remains to be learned about both structure and function. Some of the challenges for the future in terms of structure include determining the structure of the intact protein, delineating the details of the interaction between the amino- and carboxyl-terminal domains, determining the structure of the protein when it is associated with lipid, and determining the interaction of the receptor-binding region of apoE with the LDL receptor at the molecular level. With regard to function, a major unresolved issue is the basis for dominant expression of type 111 hyperlipoproteinemia. Other areas of interest related to lipid metabolism include a further definition of the role of apoE and heparan sulfate proteoglycans in chylomicron remnant metabolism, the role of apoE as a cofactor in lipolytic processing of triglyceride-rich lipoproteins, and the role of apoE in the reverse cholesterol transport process. Finally, as the studies implicating apoE in Alzheimer’s disease, nerve regeneration, and immunoregulation indicate, additional roles for this protein may yet remain to be discovered.

ACKNOWLEDGMENTS I wish to acknowledge and express gratitude to my colleagues who have shared my interest in apoE and who have made significant contributions to the current understanding of this protein: R. W. Mahley, S. C. Rall, Jr.. T. L. Innerarity, R. E. Pitas, M. R. Wardell, J. R. Wetterau, L. P. Aggerbeck, C. Wilson, and D. A. Agard. 1 am grateful to S. C. Rall, Jr., R. W. Mahley, and T. L. lnnerarity for critical reading of this article and for helpful suggestions, to K. Humphrey and S. Richmond for manuscript preparation, to D. Read for editorial assistance, and to L. Jach for graphics preparation. This work was supported in part by the National Institutes of Health Program Project Grant HL41633.

REFERENCES Aggerbeck, L. P., Wetterau, J. R., Weisgraber, K. H., Wu, C.-S. C., and Lindgren, F. T. (1988a).J. B i d . Chem. 263,6249-6258. Aggerbeck, L. P., Wetterau, J. R., Weisgraber, K. H., Mahley, R. W., and Agard, D. A. (1988b).J. Mol. Biol. 202, 179-181. Avila, E. M., Holdsworth, G., Sasaki, N., Jackson, R. L., and Harmony, J. A. K. (1982). J . B i d . Chem. 257, 5900-5909. Beisiegel, U., Weber, W., and Bengtsson-Olivecrona, G. (1991). Proc. Natl. Acad. Sci. U.S.A. 88-8342-8346. Bersot, T. P., Mahley, R. W., Brown, M. S., and Goldstein, J. L. (1976).J. Eiol. Chem. 251, 2395-2398. Blum, C . B. (1982).J. LipidRex 23, 1308-1316. Boyles, J. K., Zoellner, C. D., Anderson, L. J., Kosik, L. M., Pitas, R. E.,Weisgraber, K. H., Hui, D. Y., Mahley, R. W.,Gebicke-Haerter, P. J.. Ignatius, M. J , , and Shooter, E. M. (1989).J. Clzn. Invesf. 83, 1015-1031.

296

KARL H. WEISGRABER

Bradley, W. A., Gilliam, E. B., Gotto, A. M., Jr., and Gianturco, S. H. (1982). Biochem. Biophys. Res. C m m u n . 109, 1360-1367. Bradley, W. A., Hwang, S.-L. C., Karlin, J. B., Lin, A. H. Y., Prasad, S. C., Gotto, A. M., Jr., and Gianturco, S. H. (1984).J. Biol. Chem. 259, 14728-14735. Breiter, D. R., Kanost, M. R., Benning, M. M., Wesenberg, G., Law, J. H., Wells, M. A., Rayment, I., and Holden, H. M. (1991). Biochemistry 30,603-608. Brown, M.L., Inazu, A., Hesler, C. B., Agellon, L. B., Mann, C., Whitlock, M. E., Marcel, Y. L., Milne, R. W., Kolzumi, J., Mabuchi, H., Takeda, R., and Tall, A. R. (1989). Nature (London) 342,448-45 1. Brown, M.S., and Goldstein, J. L. (1986). Science 232,34-47. Brown, M. S., Herz, J., Kowal, R. C., and Goldstein, J. L. (1991). Curr. Opin. Lipidol. 2, 65-72. Camejo, G., and Munoz, V. (1981). I n “High-Density Lipoproteins” (C. E. Day, ed.), pp. 131-147. Dekker, New York. Cardin, A. D., Hirose. N., Blankenship, D. T., Jackson, R. L., and Harmony, J. A. K. (1986).Bwchem. Biophys. Res. Commun. 134,783-789. Carew,T. E., Koschinsky,T., Hayes, S. B.,and Steinberg, D. (1976).Lancet1, 1315-1317. Cassel, D. L., Phillips, M. C., Rostron, P., Rothblat, G. H., and Utermann, G. (1984). Atherosclerosis (Shannon, Irel.) 52,203-2 18. Chan, L., and LI, W.-H. (1991). Curr. Opin. Lipulol. 2,96-103. Chen, C. H., and Albers, J. J. (1985). Biochim. Biophys. Acta 836,279-285. Chen, G. C., Guo, L. S . S., Hamilton, R. L., Gordon, V., Richards, E. G.. and Kane, J. P. (1984). Biochemistry 23,6530-6538. Choi, S. Y., Fong, L. G., Kirven, M. J., and Cooper, A. D. (1991).J. Clzn. Invest. 88, 1173-1181. Chou, P. Y., and Fasman, C. D. (1974). Biochemistry 13,222-245. Clark, A. B., and Quarfordt, S. H. (1985).J.Biol. Chem. 260,4778-4783. Clarke, A. R., Luscombe, M., and Holbrook, J. J. (1983). Biochim. Biophys. Acta 747, 130137. Das, H. K., McPherson, J., Bruns, G. A. P., Karathanasis, S. K., and Breslow, J. L. (1985). J . Biol. Chem. 260,6240-6247. Davignon, J., Gregg, R. E., and Sing, C. F. (1988). Arteriosclerosis (Dallas) 8, 1-21. Davis, R. W., Pierotti, V. R., Lauer, S . J., Hubl, S. T., McLean, J. W., Witztum, J. L., and Young, S. G. (1991).J.L i p d Res. 32, 1013-1023. de Silva, H. V., Lauer, S. J., Wang, J.. Newhouse, Y., Shinto, L., Mahley, R. W., Weisgraber, K. H., and Taylor, J. M. (1 992). Circulation 86,l-47 1. Dong, L.-M., Yamamura, T.. and Yamamoto, A. (1990). Biochem. Biophys. Res. C m m u n . 168,409-414. Doolittle, M. H., Wong, H., Davis, R. C., and Schotz, M. C. (1987). J . Lipid Res. 28, 1326-1334. Dyer, C. A., Smith, R. S., and Curtiss, L. K. (1991).]. Biol. Chem. 266, 15009-15015. Ehnholm, C., Mahley, R. W., Chappell, D. A., Weisgraber, K. H., Ludwig, E., and Witztum, J. L. (1984).Proc. Natl. Acad. Sci. U.S.A. 81,5566-5570. Emi, M., Wu, L. L., Robertson, M. A., Myers, R. L., Hegele, R. A., Williams, R. R., White, R., and Lalouel, J.-M. (1988). Genomics 3, 373-379. Fazio, S., Horie, Y.,Weisgraber, K. H., Havekes, L. M., and Rall, S. C., Jr. (1993).J.Lipzd Res. 34,447-453. Funahashi, T., Yokoyama, S., and Yamamoto, A. (1989).J.Biochem. (Tokyo) 105,582-587. Gianturco, S. H.. Gotto, A. M., Jr., Hwang, S.-L. C., Karlin, J. B., Lin, A. H. Y., Prasad, S. C., and Bradley, W. A. (1983).J. Biol. Chem. 258,4526-4533. Glomset, J. A. (1968).J. Lipul Res. 9, 155-167.


697

Gregg, R. E., Zech, L. A., Schaefer, E. J., Stark, D., Wilson, D., and Brewer, H. B., Jr. (1986).J. Clin. Invest. 78,815-821. Hallman, D. M., Boerwinkle, E., Saha, N., Sandholzer, C., Menzel, H. J., CsPzPr, A., and Utermann, G. (1991). A m . ] . Hum. Genet. 49,338-349. Hamilton, R. L., Wong, J. S., Guo, L. S. S., Krisans, S., and Havel, R. J. (1990). J . Lipid Res. 31,1589-1603. Handelmann, G. E., Boyles, J. K., Weisgraber, K. H., Mahley, R. W., and Pitas, R. E. (1992).J. L i p d R e s . 33, 1677-1688. Havel, R. J., and Kane, J. P. (1973). Proc. Natl. Acad. Sci. U.S.A. 70,2015-2019. Havel, R. J., and Kane, J. P. (1989). In “The Metabolic Basis of Inherited Disease” (C. R. Scriver, A. L. Beaudet, W. S. Sly, and D. Valle, eds.), 6th ed., Vol. 1, pp. 1129-1 138. McGraw-Hill, New York. Havel, R. J., Kotite, L., Kane, J. P., Tun, P., and Bersot, T. (1983). J . Clin. Invest. 72, 379-387. Herz, J., Hamann, U., Rogne, S., Myklebost, O., Gausepohl, H., and Stanley, K. K. (1988). E M B O J . 7,4119-4127. Hixson, J. E., Cox, L. A., and Borenstein, S. (1988). Genomics 2,315-323. Horie, Y., Fazio, S., Westerlund, J. R., Weisgraber, K. H., and Rall, S. C., Jr. (1992). J . Biol. Chem. 267, 1962-1968. Hui, D. Y., Harmony, J. A. K., Innerarity, T. L., and Mahley, R. W. (1980).J. Biol. Chem. 255,11775-11781. Hussain, M. M., Maxfield, F. R., MPs-Oliva, J., Tabas, I., Ji. Z.-S., Innerarity, T. L., and Mahley, R. W. (1991).J. Biol. Chem. 266, 13936-13940. Ignatius, M. J., Gebicke-Harter, P. J., Skene, J. H. P., Schilling,J. W., Weisgraber, K. H., Mahley, R. W., and Shooter, E. M. (1986). Proc. Natl. Acad. Sn’. U.S.A. 83, 1125-1 129. Ignatius, M. J., Shooter, E. M., Pitas, R. E., and Mahley, R. W. (1987). Science 236, 959962. lnnerarity, T. L., and Mahley, R. W. (1978). Eiochemlstry 17,1440-1447. lnnerarity, T. L., Mahley, R. W., Weisgraber, K. H., and Bersot, T. P. (1978). J. Biol. Chem. 253,6289-6295. Innerarity, T. L., Pitas, R. E., and Mahley, R. W. (1979).J. Biol. Chem. 254,4186-4190. Innerarity, T. L., Friedlander, E. J.. Rall, S. C., Jr., Weisgraber, K. H., and Mahley, R. W. (1983).J. Eiol. Chem. 258, 12341-12347. Innerarity, T. L., Weisgraber, K. H., Arnold, K. S., Rall, S. C., Jr., and Mahley, R. W. (1984).J. Biol. Chem. 259,7261-7267. lnnerarity, T. L., Hui, D. Y., Bersot, T. P., and Mahley, R. W . (1986). In “Lipoprotein Deficiency Syndromes” (A. Angel and J. Frohlich, eds.), pp. 273-288. Plenum, New York. Ishikawa, Y., Fielding, C. J.. and Fielding, P. E. (1988).J. Bwl. Chem. 263,2744-2749. Ji, Z.-S., Brecht, W. J., Miranda, R. D., Hussain, M. M., Innerarity, T. L., and Mahley, R. W. (1993).J. Eiol. Chem. 268,10160-10167. Kane, J. P., Sata, T., Hamilton, R. L., and Havel, R. J. (1975).J. Clan. Invest. 56, 16221634. Koizumi, J., Mabuchi, H., Yoshimura, A., Michishita, I., Takeda, M., Itoh, H., Sakai, Y., Sakai, T., Ueda, K., and Takeda, R. (1985). Anthosclerosis (Shannon, Ire/.) 58, 175186. Kowal, R. C., Herz, J., Goldstein, J. L., Esser, V., and Brown, M. S. (1989). Proc. Natl. Acad. SCZ’.U.S.A. 86,5810-5814. Kowal, R. C., Herz, J., Weisgraber, K. H., Mahley, R. W., Brown, M. S., and Goldstein, J. L. (199O).J. Eiol. C h . 265, 10771-10779. Krebs, K. E., lbdah, J. A., and Phillips, M. C. (1988). Biochim. BiOphys. Acta 959,229-237.

298

KARL H. WEISCRABER

Krul, E.S., Tikkanen, M. J.. and Schonfeld, G. (1988).J. Lipid Res. 29, 1309-1325. Lalazar, A., Weisgraber. K. H., Rall. S. C., Jr., Giladi, H., Innerarity, T. L., Levanon, A. Z., Boyles, J. K., Amit, B., Gorecki, M., Mahley, R. W., and Vogel, T. (1988). J. Biol. Chem. 263,3542-3545. Lalazar, A., Ou, S.-H., and Mahley, R. W. (1989).J. Biol. Chem. 264,8447-8450. Landis, B. A., Rotolo, F. S., Meyers, W. C., Clark, A. B.,and Quarfoldt, S. H. (1987). Am. J. Physiol. 452, G805-CS10. Landschulz, W. H., Johnson, P. F., and McKnight, S. L. (1988). Science 240, 1759-1764. Lee, B., Miller, J. M., Yang, C.-Y., Ramdas, L., Yang, M., Morrisett, J. D., and Mims. M. P. (1991).J.LipidRes. 32, 165-171. Li, W.-H., Tanimura, M., Luo, C.-C.. Datta, S., and Chan, L. (1988). J. L i p d Res. 29, 245-27 1. Lohse, P., Mann, W. A., Stein, E. A., and Brewer, H. B., Jr. (1991).J. Biol. Chem. 266, 10479- 10484. Lohse, P., Brewer, H. B., I l l , Meng, M. S., Skarlatos, S. I., LaRosa, J. C., and Brewer, H. B., Jr. (1992).J. LipulRes. 33, 1583-1590. Lund-Katz, S., Weisgraber, K. H., Mahley, R. W., and Phillips, M. C. (1993).]. Biol. Chem. in press. Luo, C.-C., Li, W.-H., and Chan, L. (1989).J. Lipid Res. 30, 1735-1746. Maeda, H., Nakamura, H., Kobori, S., Okada, M., Mori, H., Niki, H.. Ogura, T., and Hiraga, S. (1989).]. Biochem. (Tokyo) 105, 51-54. Mahley, R. W. (1978). In “Disturbances in Lipid and Lipoprotein Metabolism” (J. M. Dietschy, A. M. Gotto, Jr., and J. A. Ontko, eds.), pp. 181-197. Am. Physiol. SOC., Bethesda, MD. Mahley, R. W. (1983). Arch. Pathol. Lab. Med. 107, 393-398. Mahley, R. W. (1985). Circulation 72,943-948. Mahley, R. W. (1988). Science 240, 622-630. Mahley, R. W., and Holcombe, K. S. (1977).J. LzPldRes. 18,314-324. Mahley, R. W., and Hussain, M. M. (1991). Curr. Opin. Lipidol. 2, 170-176. Mahley, R. W., and Innerarity, T . L. (1977).J. Biol. Chem. 252,3980-3986. Mahley, R. W., and Innerarity, T. L. (1983). Biochim. Biophys. Acta. 737, 197-222. Mahley, R. W., and Rall, S. C., Jr. (1989). In “The Metabolic Basis of Inherited Disease” (C. R. Scriver, A. L. Beaudet, W. S . Sly, and D. Valle, eds.), 6th ed., Vol. 1 , pp. 1195-1213. McCraw-Hill, New York. Mahley, R. W., and Weisgraber, K. H. (1983). In “CRC Handbook of Electrophoresis. Lipoproteins of Nonhuman Species” (L. A. Lewis and H. K. Naito, eds.), Vol. 4. pp. 151-173. CRC Press, Boca Raton, FL. Mahley, R. W., Weisgraber, K. H., and Innerarity, T. L. (1974). Circ. Res. 35, 722-733. Mahley, R. W., Weisgraber, K. H.,Innerarity, T., Brewer, H. B., Jr.. and Assmann, G. (1975). Biochemistty 14,2817-2823. Mahley, R. W., Weisgraber, K. H., and Innerarity, T. (1976). Biochemistry 15, 29792985. Mahley, R. W., Innerarity, T. L., Pitas, R. E.,Weisgraber, K. H., Brown, J. H., and Gross, E. (1977a). J. Biol. Chem. 252,7279-7287. Mahley, R. W., Innerarity, T. L., Weisgraber, K. H., and Fry, D. L. (1977b). A m . ] . Pathol. 87,205-226. Mahley, R. W., Weisgraber, K. H., and Innerarity, T. L. (1979). Biochim. Biophys. Acta 575, 81-91. Mahley, R. W., lnnerarity, T. L., and Weisgraber, K. H. (1980). Ann. N. Y.Acad. Sci. 348, 265-280.


299

Mahley, R. W.. lnnerarity, T . L., Weisgraber, K. H., Rall, S. C., Jr., Hui, D. Y., Lalazar, A., Boyles, J. K., Taylor, J. M., and Levy-Wilson, B. (1986).Cold Sprtng Harbor Symp. Quant. Biol. 51, 821-828. Mahley, R. W., Weisgraber, K. H., Hussain, M. M., Greenman, B., Fisher, M., Vogel, T., and Gorecki, M. (1989).J. Clin. Invest. 83,2125-2130. Mahley, R. W., and Innerarity, T. L. (1983).Biochim. Biophys. Acfa. 737, 197-222. Mahley, R. W., and Rall, S. C., Jr. (1989).In “The Metabolic Basis of Inherited Disease” (C. R. Scriver, A. L. Beaudet, W. S. Sly, and D. Valle, eds.), 6th ed., Vol. 1, pp. 1195-1213. McCraw-Hill, New York. Malcolm, B. R. (1973).Prog. Sut$ Membr. Sci. 7, 183-229. Mann, W. A., Gregg, R. E., Sprecher, D. L., and Brewer, H. B., Jr. (1989).Biochim. Biophys. Acta 1005,239-244. Marotti, K. R., Whitted, B. E., Castle, C. K., Pokes, H. C., and Melchior, G. W. (1989). Nucleic Acids Res. 17, 1778. Matsushima, T., Getz, C. S., and Meredith, S. C. (1990).Nucleic Actdc Res. 18,202. McLean, J. W., Fukazawa, C., and Taylor, J. M. (1983).J . Biol. Chem. 258, 8993-

9000. McLean, J. W., Elshourbagy, N. A., Chang, D. J., Mahley, R. W., and Taylor, J. M. (1984). J . Biol. Chem. 259,6498-6504. Miller, N. E.,Weinstein, D. B., Carew, T. E., Koschinsky, T., and Steinberg, D. (1977). J . Clin. Invest. 60,78-88. Mims, M. P., Soma, M. R., and Morrisett, J. D. (1990).Biochemistry 29,6639-6647. Moriyama, K., Sasaki, J., Matsunaga, A.. Arakawa, F., Takada, Y., Araki, K., Kaneko. S., and Arakawa, K. (1992).Biochim. Biophys. Acfa 1128, 58-64. Myant, N. B. (1990).“Cholesterol Metabolism, LDL, and the LDL Receptor.” Academic Press, San Diego. Namba, Y., Tomonaga, M., Kawasaki, H., Otomo, E., a n d Ikeda, K. (1991.Bruin Res. 541,

163- 166. Olaisen, B., Teisberg, P., and Gedde-Dahl, T., Jr. (1982).Hum. Genet. 62,233-236. Paik, Y.-K., Chang, D. J., Reardon, C. A., Davies, C. E., Mahley, R. W., a n d Taylor, J. M. (1985).Proc. Natl. Acad. Sci. U.S.A. 82,3445-3449. Pepe. M. G., and Curtiss, L. K. (1986).J. Immunol. 136,3716-3723. Phillips, M. C., and Krebs, K. E. (1986).In “Methods in Enzymology” (J. Segrest and J. Albers, eds.), vol. 128,pp. 387-403. Academic Press, Orlando, FL. Phillips, M. C., and Sparks, C. E. (1980).Ann. N . Y. Acad. Sci. 348, 122-137. Pitas, R. E.,Innerarity, T . L., Arnold, K. S., and Mahley, R. W. (1979).Proc. Null. Acad. Sci. U.S.A. 76,231 1-2315. Pitas, R. E., Innerarity, T. L., and Mahley, R. W. (1980).J. Biol. Chem. 255, 5454-

5460. Plump, A. S., Smith, J. D., Hayek, T., Aalto-Setala, K., Walsh, A., Verstuyft, J. G., Rubin, E. M., and Breslow, J. L. (1992).Cell (Cambridge, Marc.) 71,343-353. Quarfordt, S. H., Hilderman, H., Greenfield, M.R., and Shelburne, F. A. (1977).Biochem. Biophys. Res. Commun. 78,302-308. Rajavashisth, T. B., Kaptein, J. D., Reue, K. L., and Lusis, A. J . (1985).Proc. Natl. Acad. Sci. U.S.A. 82,8085-8089. Rall, S. C., Jr., and Mahley. R. W. (1992).J . I n t m . Med. 231,653-659. Rall, S. C., Jr., Weisgraber, K. H., and Mahley, R. W. (1982a).J.Biol. Chem. 257, 4171-

4 178. Rall, S. C.,Jr., Weisgraber, K. H.. Innerarity, T. L., and Mahley, R. W. (1982b).Proc. Natl. Acad. Sci. U.S.A. 79,4696-4700.

300

KARL H. WEISGRABER

Rall, S. C., Jr., Weisgraber, K. H., Innerarity, T. L., Mahley, R. W., and Assmann, G. (1983a).J.Clin. Invest. 71, 1023-1031. Rall, S. C., Jr., Weisgraber, K. H., Innerarity, T. L., Bersot, T. P., Mahley, R. W., and Blum, C.B. (1983b).J. Clin. Invest. 72, 1288-1297. Rall, S. C., Jr., Newhouse, Y. M., Clarke, H. R. G., Weisgraber, K. H., McCarthy, B. J., Mahley, R. W., and Bersot, T. P. (1989).J.Clin. Invest. 83, 1095-1101. Reyland, M. E., a n d Williams, D. L. (1991).J.Biol. Chem. 266,21099-21104. Reyland, M. E., Gwynne, J. T., Forgez, P., Prack, M. M., and Williams, D. L. (1991).Proc. Nafl. Acad. Sci. U.S.A. 88,2375-2379. Roth, R. I., Jackson, R. L., Pownall, H. J., and Gotto, A. M., Jr. (1977).Biochemisfty 16, 5030-5036. Schaefer, E. J., Gregg, R. E., Ghiselli, G., Forte, T. M., Ordovas, J. M., Zech, L. A., and Brewer, H. B., Jr. ( 1986).J . Clin. Invest. 78, 1206- I219. Schneider, W. J., Kovanen, P. T., Brown, M. S., Goldstein, J. L., Utermann, G., Weber, W., Havel, R. J., Kotite, L., Kane, J. P., Innerarity, T. L., a n d Mahley, R. W. (1981). J . Clin. Invest. 68, 1075-1085. Segrest, J. P., Jones, M. K., De Loof, H., Brouillette, C. G., Venkatachalapathi, Y. V., and Anantharamaiah, G. M. (1992).J.Lipid Res. 33, 141-166. Sehayek. E., Lewin-Velvert, U., Chajek-Shaul, T., and Eisenberg, S. (1991).J.Clin. Invesf. 88,553-560. Shelburne, F. A., a n d Quarfordt, S. H. (1974).J.Biol. Chem. 249, 1428-1433. Shelburne, F. A., and Quarfordt, S. H. (1977).J.Clin. Invest. 60,944-950. Shelburne, F. A., Hanks, J., Meyers, W., and Quarfordt, S. H. (1980).J.Clin. Invest. 65, 652-658. Shen, B. W., and Scanu, A. M. (1980).Biochemisfry 19,3643-3650. Sherrill, B. C.,Innerarity, T. L., and Mahley, R. W. (1980).J. Biol. Chem. 255, 1804-1807. Shimano, H., Yamada, N., Shimada, M., Ohsawa, N., Fukazawa, C., Yazaki, Y.. Takaku, F., a n d Katsuki, M. (1991).Biochim. Biophys. Acfa 1090,91-94. Shimano, H., Yamada. N., Katsuki, M., Yamarnoto, K., Gotoda, T., Harada, K., Shimada, Clin. Invest. 90,2084-2091. M., and Yazaki, Y. (1992).J. Shore, B., and Shore, V. (1974).Biochem. Biophys. Res. Commun. 58, 1-7. Shore, V. G., and Shore, B. (1973).Biochemisfry 12,502-507. Shore, V. G., Shore, B., and Hart, R. G. (1974).Biochemisfly 13, 1579-1585. Smit, M., de Knijff, P., van der Kooij-Meijs, E., Groenendijk, C., van den Maagdenberg, A. M. J. M., Gevers Leuven, J. A., Stalenhoef, A. F. H., Stuyt, P. M. J., and Frants, R. R., and Havekes, L. M. (199O).J.Lipid Res. 31,45-53. Snipes, G . J., McGuire, C. B., Norden, J. J., and Freeman, J. A. (1986).Proc. Nafl. Acad. Sca. U.S.A. 83, 1130-1 134. Sparrow, J. T., Sparrow, D. A., Fernando, G., Culwell, A. R., Kovar, M., and Gotto, A. M., Jr. (1992).Baochemisfry31, 1065-1068. Srinivasan, S. R., Dolan, P., Radhakrishnamurthy, B., a n d Berenson. G. S. (1972).Afherosclerosis (Shannon, I d ) 16,95-104. Steinmetz, A., Kaffarnik, H., and Utermann, G . (1985).Eur. J. Biochem. 152,747-751. Steinmetz, A., Jakobs, C., Motzny, S., and Kaffarnik, H. (1989).Arteriosclerosis (Dallas) 9, 405-4 1 1. Steinrnetz, A., Assefbarkhi, N., Eltze, C., Ehlenz, K., Funke, H., Pies, A., Assmann, G., and Kaffarnik, H. (199O).J.LipidRes. 31, 1005-1013. Stow, J. L., Kjellen, L., Unger, E., Hook, M., and Farquhar, M. G . (1985).J. Cell Biol. 100, 975-980. Strickland, D. K., Ashcorn, J. D., Williams, S., Burgess, W. H., Migliorini, M.. and Argraves, W. S. (199O).j.Biol. c h . 265, 17401-17404.


30 1

Strittmatter, W. J., Saunders, A. M., Schmechel, D., Pericak-Vance, M., Enghild, J., Salvesen, G. S., and Roses, A. D. (1993).PYOC. Natl. Acad. Sci. U.S.A. 90, 1977-1981. Tajima, S., Yamamura, T.. Menju, M., and Yamamoto, A. (1989).J. Biochem. (Tokyo) 105, 249-253. Tall, A. R. (1986).J.Lipid Res. 27,361-367. Thuren, T., Wilcox, R. W., Sisson, P., and Waite, M. (1991).J. B i d . Chem. 266, 48534861. Thuren, T., Weisgraber, K. H., Sisson, P., and Waite, M. (1992). B i o c h i r t t y 31, 23322338. Utermann, G. (1975). Hoppe-Seyler’s Z. Physiol. Chem. 356, 1 1 13-1 121. Utermann, G. ( 1985). In “Diabetes, Obesity and Hyperlipidemias-111” (C. Crepaldi, A. Tiengo, and G. Baggio, eds.), pp. 1-28. Elsevier, Amsterdam. Utermann, G., Jaeschke, M., and Menzel, J. (1975). FEBS Lett. 56,352-355. Utermann, G., Hees, M., and Steinmetz, A. (1977).Nature (London) 269,604-607. Utermann, G.. Vogelberg, K. H., Steinmetz, A., Schoenborn, W., Pruin, N., Jaeschke, M., Hees, M., and Canzler, H. (1979). Clin. Genet. 15,37-62. Utermann, G., Langenbeck, U., Beisiegel, U., and Weber, W. (1980).Am. J. Hum. Genet. 32,339-347. van Driel, I. R., Brown. M. S., and Goldstein, J. L. (1989).J.Biol. Chem. 264,9533-9538. Wardell, M. R., Brennan, S. O., Janus, E. D., Fraser, R., and Carrell, R. W. (l987).J.Clin. Invest. 80,483-490. Wardell, M. R., Weisgraber, K. H., Havekes, L. M., and Rall, S. C., Jr. (1989). J . B i d . Chem. 264,21205-21210. Wardell, M. R., Rall, S. C., Jr., Brennan, S. O., Nye, E. R., George, P. M., Janus, E. D., and Weisgraber, K. H. (1990).J.LipidRes. 31,535-543. Wardell, M. R., Rall, S. C., Jr., Schaefer, E. J., Kane, J. P., and Weisgraber, K. H. (1991). J . Lipid Res. 32, 52 1-528. Wardell, M. R., Wilson, C., Agard, D. A., Mahley, R. W., and Weisgraber, K. H. (1993). In “NATO Advanced Research Workshop. Human Apolipoprotein Mutants. 3. Apolipoproteins in the Diagnosis and Treatment of Disease” (C. Sirtori and C. Franceschini, eds.), pp. 81-96. Springer-Verlag, Berlin. Weintraub, M. S., Eisenberg, S., and Breslow, J. L. (1987).J.Clin. Invest. 80, 1571-1577. Weisgraber, K. H. (1990).J.LipdRes. 31, 1503-1511. Weisgraber, K. H., and Mahley, R. W. (1978).J.Biol. Chem. 253,6281-6288. Weisgraber, K. H., and Mahley, R. W. (1980).J.LipidRes. 21,316-325. Weisgraber, K. H., and Mahley, R. W. (1983). In “CRC Handbook of Electrophoresis. Lipoprotein Studies of Nonhuman Species” (L. A. Lewis and H. K. Naito, eds.), Vol. 4, pp. 103-132. CRC Press, Boca Raton, FL. Weisgraber, K. H., and Shinto, L. H. (1991).J.Biol. Chem. 266, 12029-12034. Weisgraber, K. H., Innerarity. T. L., and Mahley, R. W. (1978).J.Biol. Chem.253,90539062. Weisgraber, K. H.. Troxler. R. F., Rall, S. C., Jr., and Mahley, R. W. (1980). Biochem. Biophys. Res. Commun. 95, 374-380. Weisgraber, K. H., Rall, S. C., Jr., and Mahley, R. W. (1981).J. Biol. Chem. 256, 90779083. Weisgraber, K. H., Innerarity, T. L.. and Mahley. R. W. (1982).J.Biol. Chem. 257, 2518252 I . Weisgraber, K. H., Innerarity, T. L., Harder, K. J., Mahley, R. W., Milne, R. W., Marcel, Y. L., and Sparrow, J. T. (1983).J.Biol. Chem. 258, 12348-12354. Weisgraber, K. H., Rall, S. C., Jr., Innerarity, T. L., Mahley, R. W., Kuusi, T., and Ehnholm, C. (1984).J.Clin. Invest. 73, 1024-1033.

302

KARL H. WEISGRABER

Weisgraber, K. H., Rall, S. C., Jr., Mahley, R. W., Milne, R. W., Marcel, Y. L., and Sparrow, J. T. (1986).J.Biol. Chem. 261,2068-2076. Weisgraber, K. H., Mahley, R. W., Kowal, R. C., Herz, J., Goldstein, J. L., and Brown, M. S. (199O).J. Biol. Chem. 265,22453-22459. Weisgraber, K. H., Lund-Katz, S., and Phillips, M. C. (1992).I n “High Density Lipoproteins and Atherosclerosis 111” (N. E. Miller and A. R. Tall, eds.), pp. 175-181. Elsevier, Amsterdam. Biol. Chem. 268, 15745-15750. Westerlund, J. A., and Weisgraber, K. H. (1993).J. Wetterau, J. R., Aggerbeck, L. P., Rall, S. C., Jr., and Weisgraber, K. H. (1988).J.Biol. Chent. 263,6240-6248. Wilson, C.,Wardell, M. R., Weisgraber, K. H., Mahley, R. W., and Agard, D. A. (1991). Science 252, 1817-1822. Windler, E., Chao, Y.-S., and Havel, R.J. (1980).J.Biol. Chem. 255,8303-8307. Wisniewski, T., and Frangione, B. (1992).Neurosci. Leu. 135,235-238. Yamada, N., and Murase, T. (1980).Biochem. Biophys. Res. Cummun. 94,710-715. Yamada, N.,Shimano, H., Mokuno, H., Ishibashi, S., Gotohda, T., Kawakami, M., Watanabe. Y., Akanuma, Y., Murase, T., and Takaku, F. (1989).Proc. Natl. Acad. Sci. V.S.A. 86,665-669. Yamamoto, T., Davis, C. G., Brown, M. S., Schneider, W. J., Casey, M. L., Goldstein, J . L., and Russell, D. W. (1984).Cell (Cambrulge, Mas.) 39,27-38. Yamamura, T., Yamamoto, A., Hiramori, K., and Nambu, S. (1984a).Afherosclerosis (Shunnon, Irel.) 50, 159-172. Yamamura, T., Yamamoto, A., Sumiyoshi, T., Hiramori, K., Nishioeda, Y., and Nambu, S. (1984b).J. Clin. Invesf. 74, 1229-1237. Yamashita, S., Hui, D. Y., Sprecher, D. L., Matsuzawa, Y., Sakai, N., Tarui, S., Kaplan, D., Wetterau, J. R., and Harmony, J. A. K. (1990a).B i o c h . Biophys. Res. Commun. 170,

1346- 1351.

s., Sprecher, D. L., Sakai, N., Matsuzawa, Y., Tarui, S., and Hui, D. Y. (1990b).J.Clin. Invest. 86,688-695. Yokoyama, S. (1990).Biochim. Biophys. Acta 1047,99-101. Yokoyama, S., Kawai, Y., Tajima, S., and Yamamoto, A. (1985).J. Biol. Chem. 260, 16375- 16382. Zannis, V. I., and Breslow, J. L. (198O).J.Biol. C h . 255, 1759-1762. Zannis, V. I., and Breslow, J. L. (1981).Biochemistsy 20, 1033-1041. Zannis, v. I., Breslow, J. L., Utermann, G., Mahley, R. W., Weisgraber, K. H., Havel, R. J., Goldstein, J. L., Brown, M. S., Schonfeld, G., Hazzard, W. R., and Blum, C. (1982).J. LipulRes. 23,911-914. Zannis. v. I., McPherson, J., Goldberger, G., Karathanasis, S., and Breslow, J. L. (1984). J. Biol. Chem. 259,5495-5499. Zhang, S. H., Reddick, R. L., Piedrahita, J. A., and Maeda, N. (1992).Science 258, 468471. Zorich, N.,Jonas, A., and Pownall, H. J. (1985).J.Biol. Chem. 260,8831-8837. Yamashita,

THE AMPHIPATHE a HELIX: A MULTIFUNCTIONAL STRUCTURAL MOTIF IN PLASMA APOLIPOPROTEINS By JERE P. SEGREST,’ DAVID W. QARBER,‘ CHRISTIE 0. BROUILLETTE,t STEPHEN C. HARVEY,* and 0. M. ANANTHARAMAIAH’ Departments of Medklne and Blochomlatry, Atherorclerorla Research Unit, Unlvoralty of Alabama at Blrmlngham M d k a l Center, Blrmlngham,Alabama 35194; t Southern Rosearch Institute, Blrmlngham, Alabama 35205; Department of Blochemlatry,Unlveralty of Alabama at Blrmlngham M d k a l Center, Blrmlngham. Alabama 35294

*

I. Plasma Lipoproteins and Apolipoproteins ............................. 11. The Amphipathic a Helix ........................................... A. Methods for Characterization of Amphipathic Helices ..............

B. Amphipathic Helix Classes ...................................... C. Amphipathic a Helices in Exchangeable Apolipoproteins ........... D. Structure-Function Studies of Amphipathic Helices in Apolipoproteins ................................................ 111. Conclusions ........................................................ References .........................................................

303 309 310 313 322 346 363 363

I. PLASMA LIPOPROTEINS AND APOLIPOPROTEINS Lipoproteins, as the name implies, are complexes of lipids and proteins. The general structure of lipoproteins, as shown in Fig. 1, is an oil droplet consisting of a outer unilamellar membrane of phospholipids, unesterified cholesterol, and proteins, with a core of neutral lipids, predominantly cholesterol ester and triglycerides. The main function of lipoproteins is to transport lipids and lipid-soluble material throughout the body. As shown in Table I, lipoproteins are commonly classified by their density. Although the structures of these lipoprotein classes are similar, they differ in relative proportion of lipids, in the apolipoprotein: lipid ratio, and in the apolipoprotein species present (Table I). In much of the following discussion, lipoprotein classes are considered as homogeneous. However, this is incorrect, in that each lipoprotein class can be subdivided both structurally and metabolically, into a number of subclasses, such as HDL2 and HDLs (Fig. 2). Abbreviations of lipoprotein names are given in Table I. Proteins associated with lipoproteins are referred to as apolipoproteins (abbreviated “apo”). Apolipoproteins are amphipathic in nature in that they have both hydrophobic and hydrophilic regions, and can therefore interact both with the lipids of the lipoprotein and with the aqueous environment. Because of the nature of these amphipathic regions, termed amphipathic a helices, they act as protein detergents and have a ADVANCES I N PROTEIN CHEMISTRY, Vol. 45

303

Copyright Q 1994 by Academic Press, Inc. All rights of reproduction in any form reserved.

304

JERE P. SEGREST E T AL.

i

C:ll

.C.lll

A-I

A-ll

mud FIG. 1. General oil-droplet model of lipoproteins is presented for chylomicron, very low-density lipoprotein (VLDL), low-density lipoprotein (LDL), and high-density lipoprotein (HDL) structures. Apolipoproteins in the outer phospholipid membrane, designated by letters, are defined in Table 11. The major differences between the lipoproteins are the size of the neutral lipid (triglyceride and esterified cholesterol) core, liquid composition in the core, and apolipoprotein composition. (E) Triglycerides, ( 8 ) phospholipids, and ( ) esterified cholesterol are shown. Although not shown, unesterified cholesterol is found predominantly in the phospholipid monolayer.

TABLE I Chsifcation and Composition of Plasma Lipoproteins Abbreviation

Diameter (nm)

Density

Chylomicrons

Chylo. CM

75- 1200

0.93

TG

Very low density Intermediate density L o w density High density

VLDL

30-80

0.93-1.006

TG

848. E. C. A-1, A-11. A-IV B 100. E, C

IDL

25-35

1.006-1.019

TG

8100, E, C

LDL HDL

18-25 5- 12

1.019-1.063 1.063-1.21

CE Cholesterol, CE, phospholipid

8100

Lipoprotein

TG, Triglyceride; CE, cholesteryl ester.

Major lipids"

Major apolipoproteins

A-I, A-11, C

THE AMPHlPATHlC 0-HELIX MOTIF

305

FIG. 2. Size-density distribution of lipoproteins. Lp(a), Lipoprotein (a); other lipoprotein abbreviations are as given in Table I.

major role in determining and stabilizing the size and structure of the lipoprotein particle. In addition, apolipoproteins act as mediators of metabolism, either as ligands for cellular receptors or as cofactors for enzymes involved in lipoprotein metabolism. Species of apolipoproteins as well as some putative functions are summarized in Table 11. The metabolism of lipoproteins is complex and will be briefly summarized here (Fig. 3). Triglyceride-rich lipoproteins (chylomicrons, very low-density lipoproteins, and intermediate-density lipoproteins) follow two parallel metabolic courses, the exogenous and endogenous pathways. The exogenous pathway deals with dietary lipids and begins with intestinal absorption of those lipids. T h e lipids are secreted into the blood by the intestine as chylomicrons, which are large, buoyant lipoproteins containing a high proportion of triglyceride. Once in the blood, chylomicrons are acted on by the enzyme lipoprotein lipase, which catabolizes triglycerides to free fatty acids and glycerol; the resulting particle is referred to as the chylomicron remnant. Lipoprotein lipase is a triglyceridase present on the endothelial surface of most vascular beds, and requires apoC-I1 as an activator (Havel et al., 1970). The chylomicron particle diameter decreases as a result of triglyceride depletion through the action of lipoprotein lipase. Excess surface materials, such as apolipoproteins and lipids, leave the particle and enter the high-density lipoprotein (HDL) density class (Havel el al., 1973). The chylomicron remnant is rapidly tleared from the blood through receptor-mediated uptake

306

JERE P. SEGREST ET AL.

TABLE I1 Molecular Weights and Functions of Major Apolipoproleins Apolipoprotein

Molecular weight"

ApoBlOO ApoB48 APE APOC-I ApoC-I1 APOC-II1

550,000 264,000 33,000 6630 8900 8800

ApoA-1 ApoA-I1 ApoA-IV

28,000 17,400 44,500

Putative function Structural: ligand for LDL (B/E) receptor Structural Ligand for various receptors Inhibition of interaction with hepatic receptors Activation of lipoprotein lipase Inhibition of interaction with hepatic receptors; inhibition of lipoprotein lipase Activation of lecithin-cholesterol acyltranferase Structural Surface activity buffer

" Molecular weights shown are for human apolipoproteins; the weight for apoA-I1 is for the dimer.

FIG. 3. Metabolic interrelationships of lipoproteins (lipoprotein abbreviations are as given in Table 1). LpL, Lipoprotein lipase; LCAT, lecithin-cholesterol acyltransferase; HL, hepatic lipase; CETP, cholesteryl ester transfer protein. Solid lines represent interconversion of particles; regular dashed lines represent movement of cholesterol: irregular dashed lines represent transfer of lipids mediated by CETP.

THE AMPHIPATHIC a-HELIX MOTIF

307

)y the liver. The receptor responsible for this clearance has been called *he remnant receptor; apoE acts as the ligand for this receptor (Sherrill et al., 1980). The endogenous pathway is similar to the exogenous pathway summarized above, except that it deals with lipids already present in the body. The liver synthesizes and secretes very low-density lipoproteins (VLDLs), which, like chylomicrons, are triglyceride-rich lipoproteins. Again, lipoprotein lipase catabolizes the triglycerides in VLDLs to produce VLDL remnants called intermediate-density lipoproteins (IDLs), with production of excess surface material; these surface remnants are taken up by HDLs. In humans, 20-60% of VLDL is ultimately converted to LDL, with the rest being cleared from the blood Uanus et al., 1980; Parhofer et al., 1991). Hepatic lipase is a triglyceridase on the endothelium of the liver vasculature and differs from lipoprotein lipase in that apoC-I1 is not a cofactor for its activation (Nilsson-Ehle et al., 1980). The catabolic cascade depletes most of the triglyceride from the particle; thus, LDL is a cholesterol-rich particle. A major function of LDL is to provide cholesterol to peripheral (nonhepatic) tissues. LDL is taken up by these tissues through the LDL receptor, which recognizes apoBlOO and apoE as its ligands (Jones et ad., 1984; Goldstein et al., 1983). Epidemiological evidence suggests that the level of LDL cholesterol is directly correlated with the incidence of atherosclerosis (Kannel et al., 1971). Trigiyceride levels and IDL cholesterol are also now considered to be independent risk factors for atherosclerosis (Austin, 1989; Tatami et al., 1981). HDL, like LDL, is a cholesterol-rich particle, and is distinct from the other lipoprotein classes in that it does not contain apoB. HDL levels are inversely correlated with risk for atherosclerosis (Wilson et al., 1988). Nascent HDL particles are produced by direct synthesis (Hamilton, 1984), and excess surface remnants from chylomicrons and VLDL produced during the action of lipoprotein lipase (as noted above) enter the HDL density class. HDL appears to be involved in delivery of cholesterol to steroidogenic tissues as well as the removal of excess cholesterol from peripheral tissues and excretion from the system. This HDL-mediated removal of cholesterol has been termed reverse cholesterol transport (Glomset, 1968). Although apolipoproteins present in HDLs are cleared by the liver, the reverse cholesterol transport pathway has never been directly demonstrated. HDL can remove cholesterol from tissues, a process that may be partially mediated by interaction with a putative HDL receptor, with apoA-I as the ligand for that receptor (Oram et al., 1983). The existence of an HDL receptor remains controversial; saturable HDL binding may not be mediated by a specific apolipoprotein ligand and may not even be required for transfer of cholesterol from cells to

308


HDL (Johnsonet al., 1988; Slotte et al., 1987). However, binding of HDL to the putative HDL receptor may induce movement of cholesterol from intracellular pools to the cell surface, where it is available for transfer to HDL (Slotte et al., 1987). After HDL has taken up cholesterol, the enzyme lecithin-cholesterol acyltransferase (LCAT) mediates the conversion of free cholesterol to cholesterol ester (Glomset, 1968); as a neutral lipid, the cholesterol ester partitions into the core of the particle. LCAT is primarily activated by apoA-I (Kottke et al., 1986). In humans and a number of animal species, the next step in the removal of cholesterol ester involves the enzyme cholesterol ester transfer protein (CETP), which transfers neutral lipids between lipoproteins (Morton and Zilversmit, 1982). CETP may be in part responsible for the relatively high levels of LDLs in humans; species that lack CETP activity, such as rats and mice, have much lower LDL : HDL cholesterol ratios than do those with CETP activity Uiao et al., 1991; Groener et al., 1989). Humans who have genetically low or no CETP activity also have elevated HDL levels, as well as family histories of longevity (Koizumi et al., 1991). CETP also transfers cholesterol from HDLs to triglyceride-rich lipoproteins, and may provide the route by which HDLs mediate removal of cholesterol from the system through hepatic uptake of these rapidly removed particles (Tall et al., 1984). CETP has no known requirement for apolipoprotein cofactors, although the apolipoprotein content of particles may affect their donor activity, perhaps by modifying particle structure (Rye et al., 1992). HDLs in humans are a heterogeneous population of particle subclasses, with apoA-I as the major apolipoprotein species in all subclasses. Based on size, two major subclasses are present, HDL2 and HDL3. However, separation done by nondenaturing gradient gel electrophoresis demonstrates up to 11 subclasses (Cheung et al., 1987). In addition, HDL subclasses can be separated, based on apolipoprotein composition, into particles containing apoA-I and apoA-11, and those containing apoA-I without apoA-I1 (Cheung et al., 1987); other apolipoproteins are present on both of these particles. Subclass particle size, structure, and function may be due to changes in the conformation of apoA-I on the particle surface, due in part to a “hinge domain” region of apoA-I, first proposed by Brouillette et ad., (1984), which will be discussed in detail later. Interactions between apolipoproteins may also affect subclass particle structure. An understanding of structural motifs of apolipoproteins is necessary to determine how apolipoprotein content and interactions modify lipoprotein structure and function. Extensive literature exists concerning the role of HDLs in the reduction of risk for atherosclerosis. An enlarged body of evidence suggests


309

that HDLs and their major apolipoprotein, apoA-I, are also involved in many other beneficial effects. ApoA-I has been shown to interact with many cellular systems. It interacts with neutrophils to diminish neutrophi1 activation by various activators, and thus suppresses the secretion of cellular contents, suggesting apoA-I has an anti-inflammatory action (Blackburn et al., 1991). ApoA-I also inhibits cell fusion mediated by enveloped viruses [such as human herpes simplex virus (HSV) and human immunodeficiency virus (HIV) ] and thus acts as an antiviral agent (Owens et al., 1990; R. V. Srinivas et al., 1990). ApoA-I inhibits hemolysis of erythrocytes induced by toxins such as mastoparan o r comlement proteins (unpublished data from this laboratory 1992). An increase in apoA-I and HDL levels during pregnancy has been correlated with an increase in the levels of human placental lactogen (Handwerger et al., 1987); apoA-I induces the secretion of human placental lactogen in vitro (Handwerger et al., 1987; Jorgensen et al., 1989). Apolipoproteins can be grouped into two general classes, the nonexchangeable apolipoproteins (apoB 100 and apoB48) and the exchangeable apolipoproteins (all other apolipoproteins). The B apolipoproteins, present in chylomicrons, VLDLs, IDLs, LDLs, and Lp(a), are highly insoluble in aqueous solutions and thus remain with the lipoprotein particle throughout its metabolism. Because of their size and insoluble nature, it has been difficult to deduce the structural motif(s) responsible for the lipid-associating properties of B apolipoproteins. On the other hand, the exchangeable apolipoproteins are soluble in water and have been extensively studied to determine the structural motif responsible for their lipid association. Since the suggestion of a structural motif for lipid association of exchangeable apolipoproteins in 1974 from our laboratory (Segrest et al., 1974), research in this area has rapidly expanded. The common structural motif in exchangeable apolipoproteins that provides amphipathicity is the amphipathic a helix. The presence of this structural motif has been confirmed by many lines of investigation, including studies using native protein fragments and synthetic peptides. The amphipathic a helix will be discussed in detail in this review. As will be seen, this motif is not only responsible for lipid association, but is also an innate part of many biological functions mediated by apolipoproteins. 11. THEAMPHIPATHIC a HELIX

The amphipathic a helix, defined as an a helix with opposing polar and nonpolar faces oriented along its long axis, is a common secondary structural motif in biologically active peptides and proteins. T h e discovery of this structural motif was made by studying space-filling models of

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the sequences of exchangeable apolipoproteins. This was described by Segrest et al. (1974) as a unique structure-function motif involved in lipid interaction of exchangeable apolipoproteins. Prior to this, Perutz et al. (1965) had noted that a helices in globular proteins often have narrow nonpolar edges (parallel to the long axis of the helix) that face the nonpolar interior of the protein. T h e present review will deal with amphipathic a helices of the exchangeable apolipoproteins. It has been known for some time that the amphipathic a helix plays a pivotal role in the structure and functions of the exchangeable apolipoproteins. Site-directed mutagenesis and other molecular biology-based techniques are now available for probing this structural motif. Although many reviews have been written on this subject, due to the availability of new techniques for studying exchangeable structure and function of apolipoproteins, literature on this is rapidly expanding and it has become necessary to review the published literature on the location and properties of these amphipathic helices in apolipoproteins and to compare these results with recently developed and ever-expanding computer methods for location and characterization. A. Methoh for Characterization of Amphipathic Helices

Several simple methods are available to identify the existence of the amphipathic a-helix motif. Two straightforward graphical techniques, the Schiffer-Edmundson helical wheel diagram (Shiffer and Edmundson, 1967) and the helical net (or grid) representation by Lim (1978), are the methods of choice for initial analysis. T h e “helical hydrophobic moment” was introduced by Eisenberger et al. (1982) as a more quantitative method to describe an amphipathic helical sequence, a numerical way of expressing the helical amphipathicity of a protein segment. This method consists of the vector sum of the hydrophobicity values of the amino acids, taking into account their specific periodic orientation in the a helix, i.e., one residue every 100”o r 3.6 residues in a turn. T h e hydrophobic moment analysis, usually performed in a windowing fashion, is most often combined with the calculation of average hydrophobicities. A plot of both values for every n-residue-long segment in a protein, often referred to as an Eisenberg plot, enables the detection of different kinds of helices as they cluster into specific regions of the plot. Transmembrane helices have a low helical hydrophobic moment and high hydrophobicity, surface-seeking helices have an average hydrophobicity and high helical hydrophobic moment, and the helices of most globular proteins have both average hydrophobicity and helical hydrophobic moment characteristics (Eisenberg et al., 1984a).

THE AMPHlPATHlC a-HELIX MOTIF

31 1

An extension of the above method consists of calculating hydrophobic moments considering all possible side-chain orientations, i.e., "hydrophobic moments" using periodicity angles between 0" and 180". This is equivalent to performing a Fourier analysis (Finer-Moore et al., 1989; Bazan et al., 1987). This is one method of choice for a good localization of of an amphipathic helix (Boguski et al., 1985). Because the moment calculation is quantitative, it allows the estimation of the probability that a set of residues would form an amphipathic structure by chance. A simple Monte Carlo-type approach provides the answer: scramble a certain sequence, while conserving the relative frequencies of its amino acid residues, and compare the results of the real sequence with the distribution values obtained by the randomizations (Cornette et al., 1987; von Heijne, 1986). There are now a wide range of methods available to detect domains with amphipathic helicial characteristics at the residue level. However, new methods for describing the amphipathic nature of protein segments at the atomic level are still under development. Attempts to incorporate atomic hydrophobicity values (Cornette et al., 1987; Tanford, 1978) in describing the amphipathic nature of peptides or other molecules have been described (Eisenberg and McLachlan, 1986). In the future, however, side-chain flexibility, effective solvent-accessiblesurfaces, electrostatics, and molecular dynamics will have to be included to obtain an accurate description of the amphipathic nature of these protein fragments at an atomic level. T o study the location and further classification of amphipathic helices, we recently developed five computer implementations of the helical wheel and the helical net algorithms (Jones et al., 1992).The helical wheel program (WHEEL) creates a Schiffer-Edmundson helical wheel diagram (Shiffer and Edmundson, 1967) of a given sequence of amino acids arranged as an ideal a helix (100" rotation per residue) seen down the long axis. This simple representation of the helical wheel provides many other properties of a given putative amphipathic a helix: (1) T h e residues are projected onto a circular figure that is rotated so as to orient the nonpolar face toward the top of the page; i.e., the hydrophobic moment (Eisenberg et al., 1982) points toward and perpendicular to the top of the page. (2) By specifying a program option, WHEEL/SNORKEL, the wheel orientation to the page is realigned so that the nearest positive residues to either side of the hydrophobic moment are placed on a line parallel to the top of the page. (3)Any hydrophobicity scale can be used to describe the helical wheel representation of an amphipathic helical sequence. In the results reported here we have used a normalized version of the GES scale (Engelman et al., 1985).

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The helical net program (HELNET) creates a diagram by the method of Lim (1978) of the a helix seen as a cylinder cut along the center of the polar face and flattened. The center of the nonpolar face, determined by the hydrophobic moment, lies in the center of the figure and is oriented to rise out of the page. Analysis of multiple helical wheels can be done using the COMBO program, which superimposes and averages the wheels for specified sets of amino acid sequences. Before the helices are superimposed, each helix is rotated so that the nonpolar face points toward the top of the page. The residues are projected onto two circular figures. A similar program for the summation of helical net representations of a set of sequences (COMNET) superimposes and averages the helical nets for specified sets of amino acid residues. The residues selected are represented by small filled circles. The helical nets are superimposed so that the midpoint of each helix coincides. The COMBO program summarizes only the polar face of a given set of amphipathic helical sequences. T o better describe the average properties of a set of sequences, a program called CONSENSUS was developed. This program superposes the helices in the same fashion as COMBO. However, unlike COMBO, this takes into account the properties of all of the naturally occurring amino acid residues. Thus, a single figure classifies the amino acid residues into five physical-chemical groups: positive (Arg, Lys), negative (Glu, Asp), polar (Am, Gln), neutral (Tyr, Pro, His, Ser, Gly, Thr, Ala), and hydrophobic (Cys, Trp, Val, Leu, Ile, Met, Phe). CONSENSUS uses a graduated shaded contour to plot, at 20"intervals, the scaled radial distribution of these five classes of amino acid residues. Also a consensus amino acid is shown for each 20" position if there is an amino acid residue that occurs at that position most often and at least one-third of the time. In order to develop a comparison database with which to analyze the amphipathic helices of the apolipoproteins, we used COMBO, COMNET, and CONSENSUS to analyze five of the seven originally described classes of amphipathic helices (Fig. 4). Classes A, L, and H are included because these three represent surface-active amphipathic helices with measurable lipid affinity. Furthermore, as will be described later, a direct comparison of the properties of these two groups with those of class A has enabled us to hypothesize and identify many hitherto unknown properties of apolipoproteins and other class A amphipathic helical peptide analogs. Class M is included because it has significant lipid affinity (although it is not surface active), and class G is included because it is similar to certain types of nonclass A amphipathic helices also found in apolipoproteins.

THE AMPHIPATHIC U-HELIX MOTIF

313

B . Amphipathic Helix Classes The amphipathic a-helix motif has been described in exchangeable apolipoprotein sequences, but is also present in certain polypeptide hormones (Kaiser and Kezdy, 1983, 1984; Taylor et al., 1984), polypeptide venoms (Bernheimer and Rudy, 1986; Argiolas and Pisano, 1985), polypeptide antibiotics (Zasloff et al., 1988; Soravuia et al., 1988), complex transmembrane proteins (Engelman et al., 1980; Engelman and Zaccai, 1980), and the human immunodeficiency virus glycoprotein (Eisenberg and Wesson, 1990; Segrest et al., 1990). In addition, amphipathic helices involved in both intra- and intermolecular protein-protein interactions have been described in a number of proteins, including globular proteins (Perutz et al., 1965), calmodulin-regulated protein kinases (Kretsinger, 1980), and coiled-coil-containing proteins (Crick, 1953; Cohen and Parry, 1986). Although many of the biologically active peptides and proteins possess this common structural motif, it is clear that they differ in their biological activity. The concept of amphipathic helix classes was derived from the fact that amphipathic helices differ in both structure and function. In a review article from this laboratory, naturally occurring amphipathic helices were grouped into seven distinct classes (A, apolipoproteins; H, polypeptide hormones; L, lytic polypeptide; G, globular proteins; K, calmodulin-regulated protein kinases; C, coiled-coil proteins; and M, transmembrane proteins). These groupings were based on a detailed analysis of physical-chemical and structural properties using helical wheel projections (Segrest et al., 1990). The primary determinant of class was found to be a characteristicof the polar face: charge, charge density, charge distribution, and angle subtended. I . Class A Amphipathic Helices There is a considerable degree of variation between the helical domains of the different apolipoproteins. The class A amphipathic helix, like the other classes of surface-active amphipathic helices (classes H and L), has a high mean hydrophobic moment. Class A differs, however, from classes H and L in three significantways in the structure of the polar face (Fig. 4). (1) The most distinctive feature of class A is the unique clustering of positively charged residues at the polar-nonpolar interface and of negatively-charged amino acid residues at the center of the polar face. (2) Arg and Lys residues cluster within 20" of either side of the 180" polar-nonpolar plane; negatively charged residues subtend a radial angle of 100"centered on the polar face (Fig. 4). (3)Although classes H and

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FIG.4. Amphipathic helix classes analyzed by COMBOIQUALITY, COMNEI', and CONSENSUS computer programs. Five of seven amphipathic helix classes originally described (Segrest et ol., 1991) are shown: class A (apolipoproteins). class 1. (lytic polypeptide venoms), class H (polypeptide hormones), class C (globular proteins). and class M (transmembrane proteins). Class A: A total of 28 amphipathic helical sequences were analyzed at the following residue positions: apolipoproteins A-1 (44-65, 66-87. 99- 120, 121-142, 143-164, 165-186, 187-208, 220-241), A-I1 (18-30, 39-47, 52-66), A-IV (40-61,62-83,95-1l6,1l7-l38,139-160,161-182,183-204,205-226.227-248,249269,270-288,289-310,311-332). C-1 (7-14, 18-29.33-53). and C-Ill (40-67). Class L: A total of 13 amphipathic helical sequences were analyzed full length as amphipathic helices from the following lytic peptides; magainin I and 2; bombolitin I, 111, I V , and V; crabrolin; mastoparan M, X, A, 11, C, and polistes mastopraran). Class H : A total of 12 amphipathic helical sequences, representing the 1 I-mer window with the highest hydrophobic moment, were analyzed at the following residue positions for the following polypeptide hormones: calcitonin (14-24). corticotropin-releasing factor ( 13-23), P-endorphin (20-30). glucagon (17-27), secretin (14-24), vasoactive intestinal peptide (18-28). neuropeptide Y (23-33). growth hormone-releasing factor (19-29). parathyroid hormone 1-34 (10-20). adrenocorticotropin hormone (5- 15). pancreatic polypeptide (24-34). calcitonin gene-related p e p tide (10-20). Class G: A total of 12 four-helix bundle amphipathic a-helical domains were analyzed from worm myohemerythrin [ 18-38, 40-62, 69-87, 93- 1 101, bacterial cytochrome b-562 [2-19,24-45.62-86, 88-1081, worm hemerythrin [21-37, 41-64, 69-86, 90-1031, and bacterial cytochrome c3 [5-23,42-54, 79-100, 106-1 171. Class M: A total of 59 transmembrane amphipathic helical sequences were analyzed. In order to exclude the possibility of charged residues at either end of each transmembrane sequence "snorkeling" out of the hydrophobic interior of the membrane, the N- and C-terminal four residues were omitted from each putative transmembrane sequence analyzed. Transmembrane helices analyzed were as follows: /3-adrenergic receptor (helices A-C), bacteriorhodopsin (helices A-G), Band I11 (helices 1- lo),y-adreneric receptor A (helices 1-4). y-adreneric receptor B (helices 1-4), glucose transporter (helices 1-12), sodium transporter (helices l-8), and rhodopsin (helices A-G). The program for addition of helical wheels (COMBO/ QUALITY, top figures), a variation on the the COMBO program described elsewhere (Jones et al., 1992), superimposes and averages the helical wheels (WHEEL) for specified sets of amino acid sequences. Before the helices are superimposed, each helix is rotated so that the nonpolar face points toward the top of the page. The residues are projected onto a single circular figure. The closed circles represent the count of positively charged residues and the open circles represent the count of negatively charged residues. The program for addition of helical nets (COMNET, middle figures) superimposes and averages the helical nets for specified sets of amino acid residues. The residues selected are represented by small filled circles. The nets are superimposed so that the midpoint of each helix coincides. The consensus wheel program (CONSENSUS, bottom figures) superimposes the helices in the same fashion as COMBO and a single figure classifies the amino acid residues into five physical-chemical groups: positive (Arg, Lys), negative (Glu, Asp), polar (Asn, Gln), neutral (Tyr, Pro, His, Ser, Gly, Thr, Ala), and hydrophobic (Cys, Trp, Val, Leu, Ile, Met, Phe). CONSENSUS uses a graduated shaded contour to plot, at 20" intervals, the scaled radial distribution of these five classes of amino acid residues. Also, a consensus amino acid is shown for each 20"position if there is an amino acid residue that occurs at that position most often and at least one-third of the time.

FIG4, Class A

PositivelNegativeResidues

Positive Residues

kesidues horn Center

Negative Residues

0 Positive Negative Polar Neuual Hydrophobic

FIG.4. (continues)

PmitivdNegativeResidues

Positive Residues

Residues tiom Center

Negative Residues

Q Positive W Negative w Polar w Neutral w Hydrophobic

FIG.4. (continues) 316

PositivdNegntive Residues

- 5 -

-

tesidues from Centei

Negative Reaidues

0 Positive Negative Polar Neutral Hydrophobic

FIG.4. (continues) 317

Positive Residues

Residues from Center

Negative Residues

0 Positive Negative Polar Neutral Hydrophobic

FIG. 4. (continues) 318

FIG4,Class M

. *. I

I Positive Residues

Residues from Center

0

319

Negative Residues

Positive Negative Polar Neutral Hydrophobic

320


L have positively charged residues at the center of the polar face and high ratios of positive-to-negative charges, class A helices are zwitterionic in nature. Class A shares two of the distinctive features of class G helices (globular proteins): (1) a zwitterionic polar face that subtends a mean radial angle, perpendiculak to the long axis of the helix, of 180"or slightly greater; and (2) the polar face of the class A amphipathic helix has the second highest mean charge density of the seven classes; in an average 22-mer amphipathic helix of class A there are four Lys/Arg and four Glu/Asp residues. Many exchangeable apolipoproteins have tandem repetitive class A amphipathic helical domains. Such an arrangement of apolipoproteins is thought to be responsible for the size and structure of a particular lipoprotein species. 2. C h s L and C h s H Amphipathic Helices Class L (lytic) amphipathic helices include venoms such as bombolitins and mastoparan from Hymenoptera that are hemolytic (Argiolas and Pisano, 1983);antibiotics such as magainins, isolated from Xenopus laevis skin (Zasloff et al., 1988); and seminal plasmin from semen (Sitaram and Nagaraj, 1989). As the name implies, these peptides disrupt artificial phospholipid bilayers, although magainin and seminal plasmin are not hemolytic. Unlike the apolipoproteins and peptide hormones (class H), each peptide of this class consists entirely of an amphipathic helix. Members of class H include P-endorphin (Taylor and Kaiser, 1986), calcitonin (Epand et al., 1983), secretin (Robinson et al., 1982), glucagon (Wu and Yang, 1980), and adrenocorticotropic hormone (Verhallen et al., 1984). Peptides of this structural class average 36 residues in length, with the longest member being the 84-residue parathyroid hormone (Epand et al., 1985).The helical domains make up only a portion of these peptides, often inset several residues from the N terminus, and cover approximately 18-20 residues. Classes L and H have several similar properties (Fig. 4). Both have mean hydrophobic moments >0.35 and are highly positively charged. In addition, both have intermediate charge densities and have polar faces that subtend an average angle of 100" or less perpendicular to the long axis of the helix. There are three significant differences between the two classes. (1) Most strikingly, the peptide hormones have a mean Lys/Arg ratio of 0.7, whereas class L peptides have a mean Lys/Arg ratio of 46. (2) The class H peptides have a higher hydrophobic moment but a lower nonpolar face hydrophobicity than do the class L helices. (3)Class L has a bimodal cluster of positively charged amino acid residues; class H has only a single cluster. An important similarity between the two surface-active amphipathic

T H E AMPHIPATHIC a-HELIX MOTIF

32 1

helical motifs, class L and class A, is that both contain bilaterally symmetric clusters of positively charged amino acid residues. In class A, the two clusters are at the polar-nonpolar interface approximately 180”apart; in class L, the two clusters are midway between the polar-nonpolar interface and the center of the polar face, approximately 90” apart. Later in this review (Section II,C), using a concept dubbed the “snorkel” hypothesis, we suggest a structural basis for this bilateral symmetry of basic residues in classes A and L that is related to the topography of phospholipid-water interfaces. 3. Class M Amphipathic Helices

Complex transmembrane proteins belonging to class M can be considered “inside-out’’proteins with polar as well as charged groups within the hydrophobic membrane-embedded sequences. Amphipathic alignment of residues gives rise to helical packing interfaces that are polar, rather than nonpolar as for membrane proteins. The term “amphipathic” carries a slightly different meaning for membrane proteins, wherein charged residues are scarce and polar uncharged groups are largely responsible for the amphipathic nature of transmembrane helices. Not surprisingly, class M amphipathic helices differ the most in their physical-chemical properties compared to the other classes of amphipathic helices (Fig. 4). They have a very low charge density and are the only class with a net negative charge. Class M has the narrowest polar face, subtending a mean radial angle, perpendicular to the long axis of the helix, of less than 60”. Class M helices, like class G helices, have a low mean helical hydrophobic moment (0.12 per residue). Finally, class M has a high mean nonpolar face hydrophobicity comparable to that found in both class A and class L. 4 . Class G Amphipathic Helices

Class G amphipathic helices from multiple a-helix-containing globular proteins, such as myoglobin and hemerythrin, have several properties similar to those of the class A helices from plasma apolipoproteins (Fig. 4).Both, on average, have zwitterionic polar faces with moderate to high mean charge densities and both have wide polar faces that subtend an angle, perpendicular to the long axis of the helix, of 180”or greater. The major differences in their physical properties, and presumably the reasons why most globular proteins generally do not interact with lipids, are (1) a random distribution of negative and positive charges around the perimeter of the polar face, without the marked clustering seen in class A, (2) a lower mean nonpolar face hydrophobicity (0.65 versus 0.74 per residue), and (3)a slightly lower mean hydrophobic moment.

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C . Amphipathic a Helices in Exchangeable Apolipoproteins 1 . Evolutionaly Origzn As described earlier, many of the exchangeable apolipoproteins possess tandem repetitive class A amphipathic helical domains in their sequences. This periodic pattern of an a helix with well-demarcated polar and nonpolar faces is encoded into the genomic structure of the exchangeable apolipoproteins. All the human exchangeable apolipoprotein genes have been cloned and sequenced. All except apoA-IV show a remarkable similarity in having four exons and three introns (Luo et al., 1986). In addition, several of these genes are located close to each other on the genome (Driscoll and Getz, 1986). T h e most striking feature of these exchangeable apolipoproteins is the presence of internal 11residue-long amino acid repeats (Fitch, 1977). In apoA-I, apoA-IV, and apoE, the ll-mer repeats have evolved into multiple 22-mer tandem repeats. When analyzed using the programs described previously, most of these 22-mer repeat units have the characteristics of amphipathic a helices (Segrest et al., 1990). Many of the 22-mer repeats have prolines at the first, and only the first, position. These 22-mer repeats appear predominantly in exon 4 and their number ranges from 13 in apoA-IV to 1 in apoC-111. Based on their degree of homology and pattern of internal repeats, an evolutionary tree has been proposed (Luo et al., 1986) for the exchangeable apolipoproteins. It is hypothesized that, through gene duplications, a single gene has evolved to produce the current multigene family of apolipoproteins. T h e 1l-mer/22-mer evolutionary pathway for apolipoproteins can be explained as a result of the 3.6 amino acid residues per turn periodicity of an a helix; 1 I residues form three complete turns of an a helix. Consequently, tandem duplication of an 1l-residue amphipathic a helix produces a 22-residue amphipathic a helix (Fig. 5 ) . There is little twist (20" or less) between the polar and nonpolar faces of the two identical 1l-mer halves (Segrest et al., 1990; Anantharamaiah et al., 1990a). This 1 1-mer/ 22-mer motif also means that continuous amphipathic helices significantly longer than 22 residues can exist; e.g., there will be a twist of 40"or less between the polar and nonpolar faces of two tandem identical 22-mer amphipathic helices (Fig. 5 ) .

2. Physical-Chemical Properties a. Charged Amino Acid Side Chains. Since the discovery of the amphipathic helix motif for explaining the lipid-associating properties of the exchangeable apolipoproteins, our laboratory has focused on the question of whether the positions of charged residues on the polar face


323

FIG. 5. Structural relevance of 1 I-mer/22-mer tandem repeats in the exchangeable apolipoproteins.WHEELENORKEL analysis of a 22-mer formed from a tandem repeat of the 1 I-mer class A amphipathichelix sequence, ELLEALKAKLA.The thin arrowsindicate the angle (20") between tandem repeated residues within the 22-mer (e.g., residue 1 to residue 12); the thick arrows indicate the angle (40") between tandem repeated residues between two successive 22-mers (e.g., residue 1 to residue 23).

play a role in lipid affinity for class A amphipathic helices. T h e unique charged residue distribution found in apolipoprotein amphipathic helical domains was observed even in the first space-filling models of the amphipathic helical domains of apoC-I and apoC-111 (Segrest et al., 1974). Peptides analogs were therefore designed to mimic the amphipathic helical domains of apolipoproteins with respect to the distribution of charged residues, with positively charged residues at the polarnonpolar interface and negatively charged residues at the center of the polar face. To address the importance of this charge distribution, these mimics were compared with peptide analogs with reversed charge distribution; i.e., in these analogs the negatively charged amino acids were at the polar-nonpolar interface and the positively charged residues were at the center of the polar face. It was shown that peptide analogs with the reversed charge distribution have decreased lipid affinity relative to the class A mimics (Kanellis et al., 1980; Anantharamaiah et al., 1985; Chung et al., 1985; Epand et al., 1987; Anantharamaiah et al., 1991). Our explanation for these results is based on the possession by the positively charged Lys and Arg residues of four and three methylene units, respectively, in their side chains, with the amino or guanidino group attached to

324


the terminus of the methylene units. Thus, the bulk of the van der Waals surface areas of the positively charged residues are hydrophobic and are thus amphipathic. Therefore, it is not unreasonable to suggest that these amphipathic basic residues, when associated with phospholipid, extend (“snorkel”) toward the polar face of the helix to insert their charged moieties into the aqueous milieu. Thus, essentially the entirety of the uncharged van der Waals surface of the amphipathic helices of the apolipoproteins can be buried within the hydrophobic interior of a phospholipid monolayer (Fig. 6A). T o confirm this initial hypothesis further, we have incorporated charged unnatural amino acids with varying alkyl chain lengths into de novo-designed peptide analogs. We have shown that, independent of the charge on the amino acid residue, increased alkyl chain lengths of residues located at the polar-nonpolar interface resulted in increased lipid affinity; increased alkyl chain lengths of residues located in the middle of the polar face had no effect on lipid affinity (Anantharamaiah et al., 1991; Venkatachalapathi et al., 1990; Segrest et al., 1991). Three additional mechanisms have been suggested to explain the charge clustering specific to class A amphipathic helices: (1) The charge distribution is complementary to the charge distribution in the polar head group region of phospholipid monolayers (Segrest et al., 1974). (2) T h e charge distribution is important in initiating association of the apolipoprotein with the surface of phospholipid monolayers via electrostatic interactions (Segrest, 1976). (3) T h e charge distribution gives tandem amphipathic helices the potential to associate laterally in an antiparallel conformation via electrostatic interactions (Segrest, 1976); Nolte and Atkinson ( 1992), using molecular modeling techniques, have placed particular emphasis on the possible importance of this mechanism for the association of tandem amphipathic helices with the edge of phospholipid discs. The Nolte and Atkinson (1992) paper and the structure of discoidal phospholipid : apolipoprotein complexes will be discussed later. T h e original model for the amphipathic helical domains of the apolipoproteins was class A in its structural motif (Segrest et al., 1974). Subsequently, however, detailed analyses of the structural motifs of each of the amphipathic helical domains of the exchangeable apolipoproteins showed considerable diversity from class A motif. That is, the positivenegative charge-clustering motif found in class A amphipathic helices does not exist to the same extent in all apolipoproteins. Even in a given apolipoprotein, helical domains differ in their adherence to the class A motif. It is clear that lipid affinity varies perceptibly between the different exchangeable apolipoproteins (Sparrow and Gotto, 1982; Pownall and Massey, 1982; Scanu et al., 1980) and between different regions within a

FIG.6. Schematicdiagram of the snorkel model of amphipathic helices of (A) the class A motif and (B) the class L motif showing postulated insertion into a phospholipid monolayer. Long axis of the amphipathic helix is perpendicular to the plane of the page. Dimensions are approximately to scale. Note snorkeling to the aqueous surface by the interfacial amphipathic Lys residues in the class A helix and by the central Lys residues in the class L helix. In the class A model the shorter negatively charged residues are localized to the center of the polar face because of the close proximity of this portion of the helix edge to the aqueous surface. Note the depth of burial of the center of each a-helix backbone relative to the phospholipid head groups.

APoA-I1

5

N)

-10

15

20

25

5

APoC-I

NI P

10

15

20

Residue number 35 40 45

.

0.39

b


0.54

5

W

N UY

APoC-I1

N) P

10 P P

50

55

60

40

45

-

70

:75 c

50

20

*

0.41

55

)C

0.50

--15

65

I -

0.44

0

30

25

30


50

55

60

65

70

75 C

P

0.35

0.38

0.31

b

Residue number

5

10

15

20

25

30

35

40

45

50

55

60

65

70

75

on0 0 1 038

0 40

034

P

P

055

P

P

042

P

P

P

nrn-mm=rn=m-mm 04V

= ~~~

~~

035

043

033

057

032

04b

034

034

P 041

P 037

023

~

Class A amphipathic helix

0.~1

053

P

0Class G* amphipathic helix

Hydrophobic momenthsidue

c)Lipid-binding fragment

Class Y amphipathic helix

-

Non-lipid-bindingfragment

FIG.7. Diagrammatic representation of the distribution of experimentally and analytically derived properties along the amino acid sequence of the exchangeable apolipoproteins (A-I, A-11, A-IV, C-I, C-11, C-111, and E). T h e following features are represented: ( 1) locations of lipid-associating and non-lipid-associating domains suggested by experiment, (2) computer-derived locations of amphipathic helices of class A, class G*. and class Y,(3) positions of all Pro residues, and (4) hydrophobic momenthesidue calculated using a normalized GES hydrophobicity scale (Engelman el al., 1985) for the tandem repeats and predicted amphipathic helical domains.

n

I

n

m


329

given apolipoprotein (Sparrow and Gotto, 1980). It is therefore possible that this lipid affinity is correlated with the extent to which a given amphipathic helical domain in an apolipoprotein sequence fits the class A snorkel motif. A computer-based strategy was used to analyze amphipathic helix diversity in apolipoproteins. Amino acid sequences for the apolipoproteins analyzed in this study were obtained from the National Biomedical Research Foundation (NBRF) database (Bethesda, MD). The single-sequence programs WHEEL and HELNET were used to analyze individual amphipathic helices (Jones et al., 1992), and the multiplesequence programs COMBO, COMNET, and CONSENSUS were used to analyze groupings of amphipathic helices (Jones et al., 1992). Putative amphipathic helical domains in apolipoproteins A-I, A-11, A-IV, C-I, C-11, C-111, and E were identified from the NBRF database using the WHEEL and HELNET programs via a defined search and select algorithm described elsewhere (Segrest et al., 1992). Finally, all five computer programs were used to analyze and classify each domain (Jones et al., 1992). Diagrammatic representations of the results of the computer-based analysis of the amphipathic helical domains in the exchangeable apolipoproteins are shown in Fig. 7. T o facilitate comparison, Fig. 7 also includes information on the location of lipid-associating and non-lipidassociating domains. Based on the properties of their class A amphipathic helices, the exchangeable apolipoproteins fall into three separate groups: apoA-11, apoC-I, apoC-11, and apoC-111, with well-defined class A amphipathic helical domains defined as class A2 domains; apoA-I and apoE, FIG. 8. WHEELISNORKEL AND HELNETISNORKEL analyses of typical examples of a class A2. a class A,, a class G*, and a class Y amphipathic helix. The helical wheel program (WHEEL) creates a helical wheel diagram of a given sequence of amino acids arranged as an ideal a helix (100' rotation per residue) seen down the long axis. The residues are projected onto a circular figure that is rotated so as to orient the nonpolar face toward the top of the page; i.e., the hydrophobic moment points toward and perpendicular to the top of the page. By specifying a program option, WHEELISNORKEL, the wheel orientation to the page is realigned so that the normal to the top of the page bisects the nearest positive residues to either side of the hydrophobic moment. The helical net program (HELNET) creates a diagram of the a helix seen as a cylinder cut along the center of the polar face and flattened. The center of the nonpolar face, determined by the hydrophobic moment, lies in the center of the figure (dotted line) and is oriented to rise out of the page. (A) WHEELISNORKEL analysis and (B) HELNETISNORKEL analysis of apoC1[7-321 (A2 ); (C) WHEELISNORKEL analysis and (D) HELNETISNORKEL analysis of apoA-I[ 165-1861 (A, ); (E) WHEELlSNORKEL analysis and (F) HELNETISNORKEL analysis of apoE[91-116] (G*); (G) WHEEL/SNORKEL analysis and (H) HELNET/ SNORKEL analysis of apoA-1[99-120] (Y).

330

JERE P. SEGKEST ET Al..

PositivdNegative Residues

THE AMPHIPATHIC u-HELIX MOTIF

33 1

with typical but less well-defined class A amphipathic helical domains, are defined as class A, domains, and apoA-IV, with atypical class A amphipathic helical domains, are defined as class A4 domains. Eight separate class A amphipathic helical domains were identified in the four apolipoproteins containing class A2 domains; three in apoA-11, one in apoC-111, and two each in apoC-I and apoC-11. Figure 8A and B depicts WHEEL and HELNET analyses of residues 7-32 from apoC-I as an example of an individual class A amphipathic helical domain from this group. COMBO and COMBO/SNORKEL analyses, respectively, of the eight sequences making up class A2 are shown in Fig. 9A and B. T h e positive/ negative charge-clustering motif in these eight amphipathic helical domains is exact; that is, the midpoints of the positive charge clusters are symmetrically distributed at & 1OO", and the separation between the charge clusters is virtually complete. Unlike apoA-I and apoE (see below), there is relatively little difference in the results whether helical wheel orientation is by the hydrophobic moment algorithm or the snorkel algorithm. COMNET analysis shows that there is no significant charge clustering along the length of the helical axis (Segrest et al., 1992). The degree of charge separation for amphipathic helices in apoA-11, apoC-I, apoC-11, and apoC-111 is well demonstrated by the program CONSENSUS/SNORKEL (Fig. 10A). This algorithm also defines several meaningful elements of a consensus sequence for class Az. Four Lys residues cluster at and below (on the polar side of) the polar-nonpolar interface and three Glu residues cluster in the center of the polar face. Table I11 is a compilation of physical and chemical properties derived from COMBO analyses of the different sets of potential amphipathic helical classes from all exchangeable apolipoproteins. T w o of these physical and chemical properties distinguish the class A2 amphipathic helices from the rest: (1) both the mean hydrophobic moment ( ( p H ) ) and the FIG. 9. Analysis of the potential amphipathic helices in the exchangeable apolipoproteins by the COMBO/QUALITY and COMBOIQUALITYISNORKEL programs. The following sequences were analyzed: class AP,apoA-11[7-30.39-50,51-711, apoC-1[732, 33-53], apoC-I1[14-39, 44-55], apoC-II1[40-67]; class Al, apoA-1[44-65, 66-87, 121-142, 143-164, 165-186, 187-2081, apoE[ 161-182, 203-2661; class G*, apoA-I[8331, apoA-IV[7-31], apoE[25-51, 52-83, 91-1 16. 135-160, 268-2851, apoC-II[60-76], apoC-III[8-29]; class Y, apoA-I[88-98, 99-120, 209-2 19, 220-2411. apoA-IV[40-61, 62-94, 139- 160, 183-204, 227-248, 249-288, 289-310, 31 1-3321. COMBO/ QUALITYlSNORKEL is the sum of multiple WHEELBNORKEL analyses. (A) COMBO/ QUALITYISNORKEL analysis for class A,; (B) COMBO/QUALITY analysis for class A2; (C) COMBO/QUALITY/SNORKEL analysis for class A l ; (D) COMBO/QUALITY analysis for class G*; (E) COMBO/QUALITY/SNORKEL analysis for class Y.

332

JERE P. SECREST E T AL.

FIG. 10. CONSENSUS/SNORKEL analyses of classes AP, A,, G*, and Y. (A) CONSENSUS/SNORKEL analysis of class AS;(B) CONSENSUS/SNORKEL analysis of class A , ; (C) CONSENSUS analysis of class G*; (D)CONSENSUS/SNORKEL analysis of class Y.

hydrophobicity of the nonpolar face are maximal, and (2) Lys residues are almost five times more prevalent than Arg residues. ApoA-I and apoE were identified as having six and two potential class A amphipathic helical domains, respectively. COMBO/SNORKEL analysis of these eight class A amphipathic helices is shown in Fig. 9C. Several


333

features distinguish these domains from the class A2 described earlier: (1) The class A motif is typical in these two apolipoproteins; the mean angles of the two positive clusters are at precisely +go"; (2) the positivenegative charge-clustering motif is noticeably less well defined than for the class A2 apolipoproteins; and (3)when the helical wheel is oriented by the hydrophobic moment algorithm, the positive charge clustering is still visually apparent (COMBO; not shown) but considerably less well defined than when the orientation is by COMBO/SNORKEL. The major features defined by CONSENSUS/SNORKEL analysis of class A1 (Fig. 10B) are two Arg residues at the polar-nonpolar interface and four Leu residues in the center of the nonpolar face. From Table 111, class A1 has a nonpolar face hydrophobicity comparable to that of class A2,but the mean hydrophobic moment is considerably lower, and, unlike class A l ,Arg residues are twice as prevalent as Lys residues; Fig. 11 shows a COMBO analysis for the distribution of Lys versus Arg for class A2 versus class AI. A typical example of the class A1 domain, apoA-I[1651861, is shown in Fig. 8C and D. The class A amphipathic helical domains present in apoA-IV are unique in their properties. Only four potential class A amphipathic helical domains have been identified in apoA-IV; these are rather atypical compared to class A domains in the other exchangeable apolipoproteins (Segrest et al., 1992). Not all amphipathic helical domains in exchangeable apolipoproteins fit the class A motif. As seen in Fig. 7, each of the exchangeable apolipoproteins, except apoA-I1 and apoC-I, was identified as having putative amphipathic helical domains that cannot be classified as class A in their radial arrangement of positive and negative residues. Detailed examination of the individual domains suggested that they fall into two basic types: (1) The first type of amphipathic helix is present in five of the seven apolipoproteins and is distinguished by a random radial arrangement of positive and negative residues. These amphipathic helices are similar but not identical to the class G amphipathic helices found in globular proteins and thus we call them class G*. (2) The second type of amphipathic helix is present in only two of the seven apolipoproteins and is distinguished by a radial clustering of positive and negative residues into a pattern unlike that of class A; because of the presence of positively charged residues at the polar-nonpolar interface as well as at the center of the polar face, we term this the class Y motif. A total of nine class G* amphipathic helical domains are located in five of the seven exchangeable apolipoproteins, five in apoE and one each in apoA-I, apoA-IV, apoC-11, and apoC-111 (Fig. 7). Four of these five domains in apoE are located in the amino-terminal half of the molecule

TABLE 111 Properties of Amphipalhit Helital Domains of Apolipoproteins Hydrophobicity per AA of nonpolar faceb.' Class A2

A1 G*

Y

Apolipoprotein or protein A-11, C-I, C-11, C-111 A-I, E A-I, A-IV, E, C-11, C-111 A-IV, A-I

Mean ( p H )per AAa*b 0.43 0.34 0.44 0.37

CON"

CON/SNKd

+AA"*'

-AAQX

Lys : Arg ratio"

0.74 0.7 1

0.75 0.72 0.61 0.67

1.9 2.0 1.6 1.9

1.8 2.1 1.6 2.2

4.8

0.69 0.65

Data derived from CONSENSUS analyses. AA, Amino acid. Calculated using a normalized (unitless) GES hydrophobicity scale (Engelman et al., 1985). Includes only the six residues centered on the nonpolar face. Data derived from CONSENSUS/SNORKEL analyses. ' Per 11 amino acids. a

0.5 1

.o

1.2


335

LyslArg Residues

FIG. 1 1. COMBO/QUALITY analysis of the distribution of Lys versus Arg for class A2 versus class A , . (A) Class A,; (B) class A , . Shaded circles represent the count of Lys residues and open circles represent the count of Arg residues.

and the fifth is at its carboxy-terminal end. All but apoC-I1 contain an amino-terminal class G* domain largely derived from the first two of the three tandem 1 1-mer repeats located in exon 3. In apoC-11, the single class G* domain is located at the carboxy terminus and is largely derived from the second of the two tandem 1 1-mer repeats located in exon 4. Figures 9D and 1OC show COMBO and CONSENSUS analyses, respectively, of the nine domains classified as class C* amphipathic helices. Consistent with the lack of charged residue clustering, the only consensus feature identified is a cluster of four Leu residues on the nonpolar face. It is apparent from Table 111 that the class G* amphipathic helices have a high mean hydrophobic moment and a moderately high nonpolar face hydrophobicity. A typical example of a class G* amphipathic helical domain from apoE[91-116] is shown in Fig. 8E and F. The class Y motif is seen in exchangeable apolipoproteins apoA-I and apoA-IV-8 of the 13 putative amphipathic helical domains in apoA-IV and 4 of the 1 1 domains in apoA-I are of the Y class. Figures 9E and 10D

336

JERE P. SECREST ET AL.

are COMBO/SNORKEL and CONSENSUS/SNORKEL analyses, respectively, of these 12 amphipathic helices. It is readily apparent from these figures that the basic features of the class Y motif are two negative residue clusters on the polar face separating the two arms and the base of the Y motif formed by three positive residue clusters. The class Y amphipathic helices have both an intermediate mean hydrophobic moment and an intermediate nonpolar face hydrophobicity compared to the other amphipathic helical classes in the exchangeable apolipoproteins (Table 111). A typical example of a class Y amphipathic helical domain from apoA-I[99-1201 is shown in Fig. 8G and H. In humans, apoA-IV is found primarily in the free protein (nonlipoprotein) portion of plasma. Although the reason is not clear, it is possible that the lack of class A motif in the amphipathic helical domains of human apoA-IV causes it to associate poorly with the lipoprotein surface. In rats, however, apoA-IV is seen on HDLs. Examination of individual amphipathic helical domains of rat apoA-IV does show the presence of the class A motif in its structure, thus supporting our hypothesis that the class A motif is essential for binding of apolipoproteins to lipoproteins.

6. Hydrophobic Amino Aczd Side Chains. The measurement of nonpolar face hydrophobicity is therefore relevant to the snorkel concept. For all of the class A amphipathic helical subclasses (except apoA-IV), including those in the insect apoLp-111, the hydrophobicity of the nonpolar face is higher when the helices are oriented by the snorkel algorithm than when oriented by the hydrophobic moment algorithm (Table 111); this is also true for the unusual class Y amphipathic helical domains found in apoAIV and apoA-I, which presumably also represent lipid-associating domains. On the other hand, for amphipathic helical domains with weak lipid affinity (the class G* found in the apolipoproteins and the class G amphipathic helices found in four-helix bundle proteins), the hydrophobicity of the nonpolar face is higher on average when the helix is oriented by the hydrophobic moment algorithm than when oriented by the snorkel algorithm. These results are remarkable in that they suggest that the snorkel orientation more accurately reflects the orientation of lipidassociating helices than does orientation by the hydrophobic moment. c. Helix Length. One property of exchangeable apolipoproteins that must be explained is their ability to target specific lipoprotein particles; e.g., apoA-I associates almost entirely with HDLs, which have a smaller radius of curvature than other lipoproteins. ApoE associates invariably


337

with TG-rich lipoproteins and large HDL subspecies, both of which possess larger radii of curvature. It seems probable that this targeting is controlled at least in part by apolipoprotein sensitivity to variations in the properties of lipoprotein surfaces, such as variations in surface pressure, lipid and apolipoprotein composition, and surface curvature. Because all of the apolipoproteins possess amphipathic helical domains, it is possible that structural variations in the amphipathic helical domains play a role in the lipid targeting of apolipoproteins. As previously noted, apoA-I contains Pro-punctuated tandem repetitive amphipathic helical domains. In apoE, however, the major lipidbinding domain maps to a class A amphipathic helix motif (residues 202-266; see Fig. 7) with no Pro punctuations (Fig. 12A).Thus, this is by far the longest unbroken amphipathic helix among the exchangeable apolipoproteins (65 residues) and one in which the polar-nonpolar interface is in register throughout its length due to a four-amino-acid deletion compared with apoA-I. In light of this single, long amphipathic helix in apoE, because apoA-I targets smaller lipoprotein particles than does apoE, it is significant that apoA-I, known to have multiple lipid-binding domains, is composed of a series of tandem repeating 22-residue-long proline-punctuated amphipathic helices. Further, the polar-nonpolar faces of adjacent amphipathic helices in apoA-I are 40" out of register (Fig. 12B), and almost all repeats are punctuated by Pro residues. These structural features would seem suited to allow the association of a long, continuous amphipathic helix to the highly curved surface of high-density lipoprotein particles; Pro residues in the trans configuration produce a bend (-30" ) toward the helix face opposite the Pro, and thus curvature, and a right-handed twist (-40" ) to the continuous helix [these values come from typical protein crystallographic data given by Richardson and Richardson (1989)l. This twist would thus allow the hydrophobic faces of adjacent helical domains to remain aligned. Based on these observations, we propose a model in which amphipathic helix length, at least in part, affects lipoprotein targeting. We suggest that longer class A amphipathic helices (i.e., apoE) preferentially target larger triglyceride-rich lipoprotein particles, and shorter helices (i.e., apoA-I) preferentially target smaller high-density lipoproteins. The model can be tested by peptide synthesis strategies and site-directed mutagenesis. d. Helix Cross-Sectional Shape. By forging a conceptual link to both phospholipid shapes and class A amphipathic helices, we have been able to marshal evidence for a unifying molecular principle that relates cross-

338


B A

FIG. 12. HELNETlSNORKEL analyses of (A) apoE[202-2661 and (B) apoA-I[ 1211861. Hydrophobic residues are highlighted (bold outline) and Pro residues are marked by arrowheads. Note the straight nonpolar face in the apoE segment and the twist to the nonpolar face in the apoA-I segment.


339

sectional shapes of amphipathic helices to effects on biological membrane bilayer stability (Tytler et al., 1993). Phospholipids, the predominant lipid components of biological membranes, can exist in several different phases, including micellar, bilayer ( L a ) , inverted hexagonal (HII), inverted cubic (QII)(Cullis and Hope, 1985), and other intermediate inverted phases (Ellens et al., 1989). Inverted phases have been implicated in both membrane leakage and membrane fusion (Ellens et al., 1989). The phase that a phospholipid prefers to adopt at a particular temperature is determined by its dynamic molecular shape (Cullis and Hope, 1985). Based on shapes of longitudinal sections, phospholipids have been placed into three classes: (1) Coneshaped phospholipids (wedge-shaped in cross section) include lysophospholipids and detergents; these contain relatively large polar head groups favoring a positive surface curvature and the micellar phase. (2) Cylindrical-shaped phospholipids include phosphatidylcholine (PC); these contain equal polar head group and fatty acyl chain cross-sectional areas that favor a flat surface and the La phase. (3)Inverted cone-shaped phospholipids (inverted wedge-shaped in cross section) include phosphatidylethanolamine (PE); these contain relatively large fatty acyl chain cross-sectional areas favoring a negative surface curvature and inverted lipid phases (Cullis and Hope, 1985). Although more quantitative theories are being developed (Gruner, 1985), the shape concept of lipid packing provides a good qualitative description of lipid-phase propensity. Importantly, combinations of wedge and inverted-wedge phospholipids are known to form stable L, bilayer structures (Hauser et al., 1981). A series of peptide analogs of class A and class L amphipathic helices were synthesized and studied (Tytler et al., 1993). Two of these peptides, designated 18A and 18L, were modeled as idealized Q helices, energy minimized, and displayed in cross section (Tytler et al., 1993). T h e overall cross-sectional shape of a class A amphipathic helix in the snorkel orientation was found to be that of a wedge with a polar base and a hydrophobic apex, schematically illustrated in Fig. 6A. In contrast, the shape of a class L amphipathic helix in cross section is reciprocal to that of a class A amphipathic helix; i.e., an inverted wedge with its apex at the polar face and its base buried in the lipid (Fig. 6B). T h e results of our physical-chemical studies of the peptide analogs of class A and class L amphipathic helices (Tytler et al., 1993) can be summarized as follows: Class A peptides stabilize model and biological membranes so as to inhibit membrane lysis by class L polypeptides. Class L peptides induce inverted nonbilayer structures in model membranes and increase significantly the binding of class A peptides to erythrocyte membranes. Increasing the bulkiness of the hydrophobic face andfor decreas-

340


ing the width of the polar face for class L amphipathic helices, thus broadening the inverted wedge shape, increases the ability of the analogs to lyse membranes. Increasing the bulkiness of the charge moieties of the basic residues for a class L amphipathic helix, and thus narrowing the inverted wedge shape, blocks the ability of the analog to stabilize inverted nonbilayer phases and thus to lyse membranes. The dramatic sensitivity of the properties of class L amphipathic helices to the bulkiness of the charged moieties of basic residues may explain why 45 of 47 basic residues in the 16 known examples of class L lytic peptides are Lys and only 2 are Arg (Segrest et al., 1990). Increasing the bulkiness of the charge moieties of the basic residues for a class A amphipathic helix, thus broadening the wedge shape, increases the ability of the analog to destabilize inverted nonbilayer phases, and thus increases its ability to stabilize membranes and to inhibit membrane lysis by class L peptides. Based on these results, we have concluded that the inverted wedge-shaped cross sections of class L amphipathic helices produce membrane lysis following membrane insertion by stabilization of inverted nonbilayer lipid phases. We also conclude that the wedge-shaped cross sections of class A amphipathic helixes stabilize bilayers following membrane insertion by inhibiting inverted nonbilayer lipid phases; we further suggest that the reciprocally shaped class A amphipathic helices inhibit class L-dependent membrane lysis by binding to and “filling in” membrane bilayer defects created by the class L amphipathic helices.

3. X-Ray Structure Determination of Apolipopfoteins One of the prerequisites for the determination of structure of a protein or a peptide by the X-ray method is that they be crystallized. Because of the amphipathic helical nature of apolipoprotein sequences, these proteins are not easily crystallized. Progress has therefore been impeded in determining the three-dimensional structure of exchangeable apolipoproteins. T h e first apolipoprotein from insects, apolipophorin, or apoLp-111, has been crystallized and X-ray structure has been described at 2.5 A resolution (Breiter el al., 1991). Insects have only one major kind of lipoprotein, called lipophorin, which contains two apoB-like apolipoproteins, apolipophorin I and I1 (Shapiro et al., 1988); several of the insects also have a third apolipoprotein of 18-20 kDa (Shapiro et al., 1988), apolipophorin 111 (apoLp111). T h e suggested function of apoLp-111 is to help move lipid from the fat body to flight muscles during prolonged flight. ApoLp-111 in the resting insect circulates in the hemolymph in the form of a lipid-free globular protein monomer. During flight, apoLp-111 is postulated to bind


34 1

to hydrophobic defects created by expansion of the lipophorin particles, allowing continued lipid loading (Shapiro et al., 1988). Holden and co-workers (Breiter et al., 1991) have described a molecular structure for apoLp-111 from hemolymph of the African migratory locust, Locusta mzgrutoria, determined at 2.5 8, resolution. T h e structure determined for apoLP-111 is that of five long amphipathic a helixes connected by short loops to form a five-helix bundle globular protein. A hinged movement of two helices, as shown in Fig. 13A, was proposed to account for the partitioning of apoLp-111 from its solution-phase fivehelix bundle globular structure to a lipid-bound unfolded conformation on the surface of lipophorin. This proposed conformational change is quite similar to the hinged-domain movement (Fig. 13B) postulated for apoA-I (Cheung et al., 1987; Brouillette et al., 1984; Anantharamaiah et al., 1990b), as discussed later. The five actual amphipathic helical domains of the insect apolipoprotein Lp-111 (Breiter et al., 1991) analyzed by COMBO/SNORKEL and CONSENSUS/SNORKEL are shown in Fig. 14A and B, respectively. From these analyses it is apparent that the amphipathic helical domains of apoLp-111 are similar in several important ways to the class A amphipathic helical domains of the apolipoproteins: the amphipathic helical domains of apo Lp-111 have a high mean hydrophobic moment (0.38 per residue), a high mean nonpolar face hydrophobicity (0.71 per residue), and a well-defined negative charge cluster. There also appears to be some interfacial positive charge clustering in the amphipathic helical domains of apoLp-111, but the clustering is weak. As is commonly found in the exchangeable human apolipoproteins, three of the a helices (helices 2,4, and 5) are punctuated by Pro. The major difference between the amphipathic helical domains of apoLp-111 and those of the exchangeable apolipoproteins is that the amphipathic helical domains of apoLp-111 have a very low charge density. The 191-amino acid residue amino-terminal segment of human apoE (E-22; see below) has been crystallized and its structure determined at 2.5 8, resolution. This fragment of apoE, containing the LDL receptorbinding domain, was determined to have a four-helix bundle structural motif (Wilson et ul., 1991). The up and down helices of this globular domain were located at residues 24-42, 54-81, 87-122, and 130-164, with the latter containing the LDL receptor-binding domain (Rall et al., 1986). A fifth a helix at residues 44-53 forms a short link between the first two a helices of the four-helix bundle. The E-22 fragment of apoE is known to associate only weakly with lipid (Gianturco et al., 1983). With the exception of apoA-I1 and apoC-I, all apolipoproteins analyzed contain one or more amphipathic helical domains of the class G*.

342


0 HINGED

B

TANDEM AMPHIPATHIC HELIXES OF APOLIPOPROTEIN 1-1

HINGED DOMAIN

FIG.13. Models for hinged domains: (A) in insect apoLp-I11 (adapted from Breiter el aL, 1991, Copyright 1987 American Chemical Society) and (B) in apolipoprotein A-I. (Adaptedwithpermission from Cheung etaL, 1987.) The arrowsindicatea transition from the hingeclosed to the hinge-open conformation.

ApoA-I, apoA-IV, apoC-111, and apoE contain one or more class G amphipathic helical domains at their amino terminus. As shown in the amphipathic helix map (Fig. 7), the amino-terminal class G* amphipathic helical domains of apoE, with a relatively weak lipid affinity (Gianturco et al., 1983), correspond closely to the position of a


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PositivdNegative Residues

FIG. 14. COMBOIQUALITYISNORKEL (A) and CONSENSUSISNORKEL(B) analyses of the five amphipathic helical domains of the insect apolipoprotein Lp-111. The am129-1561. phipathic helical domains in insect apoLp-I11 are [7-32.35-66,70-86,95-121,

four-helix bundle globular structure determined by X-ray structural analysis (Rall et al., 1986). The type of amphipathic helix found in the amino-terminal domain of apoE is similar, in the first approximation, to the class G amphipathic helix found in four-helix bundle globular proteins (Segrest el al., 1990) in that both have wide zwitterionic polar faces. There are differences, however. T h e amino-terminal amphipathic helices of apoE have a higher mean hydrophobic moment (0.44 versus 0.32), a higher mean nonpolar face hydrophobicity (0.68 versus 0.62), and a higher mean charge density (3.9 versus 3.0 charged residues per 11-mer) than do €our-helix bundle amphipathic helices (Table 111). It is

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therefore tempting to speculate that the amino-terminal domain of apoE, including the LDL receptor-binding region, may be lipid-associated under certain conditions and globular under other conditions; the receptorbinding region has been shown to be much more active when associated with lipid than when lipid free (Rall et al., 1986). We further suggest that the other single class G* amphipathic helical domains located in apoC-11, apoC-111, apoA-I, and apoA-IV interact with either lipids or proteins in a manner regulated by local environmental conditions. Consistent with this model, the class G* domain in the carboxy-terminal region of apoC-I1 is involved in the activation of lipoprotein lipase by this apolipoprotein (Catapano et al., 1979; Vinio et al., 1983). T h e amphipathic helix map (Fig. 7) suggests a second and perhaps related possibility for the lipid association of the amino-terminal domain of apoE. The class A amphipathic helix located between residues 181192 is disordered in the crystal structure (Wilson et al., 1991) but might associate with lipid when lipid is present; this region has been shown to be less protease sensitive when the amino-terminal domain is lipid bound (Rall et al., 1986). 4 . Conformational Analysis

Given the widespread occurrence of sequences in apolipoproteins that evidently code for amphipathic helices, it is not surprising that many workers have attempted to identify possible secondary structural elements in apolipoproteins and to predict possible tertiary interactions and overall arrangements of secondary structure elements when these proteins are bound to lipids (Edelstein et al., 1979). Here we discuss models for apoA-I and apoE-3 developed by Nolte and Atkinson (1992). These models resulted from an examination of the primary sequence of human plasma and apoA-I and apoE-3 using a variety of approaches, and an integration of the resulting data into unified predictions for the secondary structures of those molecules. To begin with, Nolte and Atkinson analyzed sequence homologies among previously identified 11-amino acid repeats within apoA-I and apoE-3, to place each helix in either the A or B class, as proposed by Li el al. (1988). Using mutation-based criteria in these homology calculations, Nolte and Atkinson changed several of the assignments previously given by Li el al. Next, they used four different methods (Chou and Fasman, 1978; Levitt, 1978; Gilbrat et al., 1987; Gascuel and Golmard, 1988) to establish independently preferences for the helix, extended, and coil structures along the primary sequences. These preferences were supplemented with physicochemical analyses on the primary sequences, using windowed averages of physical properties (Kyte and Doolittle, 1982),


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average hydrophobic moment calculations (Eisenberg et al., 1984b),onedimensional Fourier transformations of physical properties to find sequential periodicities (Finer-Moore and Stroud, 1984), and the calculation of the depth of penetration of putative amphipathic helices into the water-lipid interface, using an algorithm developed for this work (Nolte and Atkinson, 1992). One important experimental result was available, the quantitative measurement of the fraction of each secondary structural element by circular dichroism (CD) on purified lipid-protein complexes. This provided a constraint that allowed a careful evaluation of the secondary structure predictions derived from the various approaches, some of which were developed for water-soluble proteins and therefore of uncertain reliability for proteins in a lipid environment. The data from these analyses were combined using an integrated prediction method to arrive at a consensus secondary structure model for each protein. The integrated method involved 36 steps, with independent predictions at each step. The final model was based on an evaluation of the various predictions, with judicious intervention by the authors. As an aid to developing the appropriate weighting of all the data, they carried out the analysis for apoE-3 without reference to the available crystal structure (Wilson et al., 1991), then used the known structure of the HDL-binding amino-terminal domain of apoE-3 as feedback to reevaluate the weighting. The final model of apoE-3 contained 193 residues (65%) in the ahelical conformation, 22 residues (7%) of /3 sheet, and 84 residues (28%) as random coil or turn. These fractions compared very favorably with the CD-derived constraints, which were 61% helix, 11% sheet, and 28% randomlturn. For the 139 residues in the amino-terminal domain whose crystal structure is known, 118 (85%) were predicted to be in the same conformation as in the crystal. T h e model assigned a random coil to 37 residues that were present but not observed in the crystal structure, presumably because of disorder. In addition, the model predicted two short helical fragments (1-6 and 178-183) at the ends of the fragment, in regions that are disordered in the crystal structure. These two regions may be sensitive to environment, requiring either the full apoE-3 protein and/or bound lipid to form stable structures. The composite model for apoA-I contained 168 residues of helix (69%), 23 residues of sheet (9%), and 52 residues that are either turn or random coil (2 1%). Again, these values are very close to the experimental constraints for helix, sheet, and turn/coil (70, 10, and 20%, respectively). The model contains six major helical segments, in agreement with the minimum number of helices suggested by Jonas et al. (1989). T h e central domain is composed of multiple A/B tandem repeats, which Nolte and

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Atkinson suggest are folded back on each other in antiparallel fashion. There is longitudinal separation of charge along the helices, suggesting that oppositely charged regions could interact when folded in this motif. The pattern of breaks between the six helices would also allow the exchange of a pair of helices from the solvent to the surface of the lipid particle, as we have previously suggested (Brouillette et al., 1984; Cheung et al., 1987). D . Structure-Function Studies of Amphipathic Helices in Apolipoproteins Amphipathic helix variation is not only important in determination of lipid affinity and lipoprotein targeting, but is also important in the function of exchangeable apolipoproteins. However, amphipathic helix variation has not been correlated with the biological activity of a particular apolipoprotein. In this section we will review the literature for evidence linking amphipathic helices (and/or specific apolipoprotein sequences) with certain functional properties of the exchangeable apolipoproteins. The results of this review of amphipathic helix/function linkage will be compared with the map of amphipathic helix location and classification. We believe that this exercise should allow us to generalize the structural motifs required for a particular biological activity of a given exchangeable apolipoprotein. 1 . Studies with de Novo-Designed Synthetic Peptide Analogs

Studies of peptide analogs have suggested an important role in the structure-function relationship of biologically active peptides and proteins. A number of laboratories have utilized either protein fragments or peptide analogs to study the function of different regions of apolipoproteins. Whereas the studies of fragments addressed the role of specific sequences in their biological activity, the model peptide analogs provided details of the role of specific amino acids in determining the structure-function relationship of the amphipathic helical motif. The key structural features predicted for the amphipathic helix by the original model (Segrest et al., 1974) enabled three laboratories to study independently how amino acid variability determined the properties of the amphipathic helix (Kanellis et al., 1980; Fukushima et al., 1980; Sparrow et al., 1981). The strategy adapted by these investigators was based, not on the primary sequence of naturally occurring apolipoproteins, but on incorporating the periodicity of the secondary structural features of the amphipathic helix motif into the sequences of the peptide analogs.


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Kaiser and colleagues (Fukushima et al., 1980) showed that a 22residue peptide can be synthesized entirely from Glu, Lys, and Leu arranged periodically in the typical class A distribution to form an amphipathic cr helix with equal polar and nonpolar faces. Indeed, as studied by quantitative ultrafiltration, gel-permeation chromatography, and circular dichroism, the peptide associated effectively with phospholipid and mimicked some of the physical and chemical properties of apoA-I (Fukushima et al., 1980; Kaiser and Kezdy, 1983, 1984; Nakagawa et al., 1985). Because of the presence of tandem repetitive amphipathic helical domains in apoA-I, a dimer of this 22-residue peptide was later synthesized and found to more closely mimic apoA-I than the monomer; thus the study of peptide analogs supported the concept that 44-mers represent the minimal functional domain in apoA-I (Nakagawa et al., 1985). In a study by the Baylor group (Sparrow et al., 1981), lipid-associating peptides (LAPS), LAP-16, LAP-20, and LAP-24 (16, 20, and 24 amino acid residues long, respectively), were shown to associate with phospholipid. This investigation showed that peptide analogs of the amphipathic helix as short as 10 to 12 residues in length have the ability to interact with phospholipid (McLean et al., 1991). An interesting series of peptides with a variation of LAP-16 were also synthesized with the amino terminus of LAP-I6 blocked with fatty acyl chains of various lengths (Ponsin et al., 1986a). These peptides allowed the investigators to study the role of hydrophobicity in lipid-associating ability. The peptides interacted with lipid to form stable lipoprotein complexes and associated with high-density lipoproteins both in vitro and in uiuo (Ponsin et al., 198613). In uivo injection of reassembled HDLs containing a series of radiolabeled acylated peptides showed that the plasma half-life increased with the acyl chain length. These results support the notion that the rates of clearance of the exchangeable apolipoproteins are a predictable function of their lipid affinity (Ponsin et al., 1986b). This was confirmed by studies in our laboratory using peptide analogs of varying lipid affinity as determined by surface pressure measurements (Garber et al., 1992). Lipid affinity of these peptides was varied by modifying residue number and charge distribution on the polar face. Although peptides with lower lipid affinity partitioned between HDL and the free protein phase in plasma of rats injected with the peptides, those with higher lipid affinity associated with HDL alone and had slower rates of clearance. Organ distributions were determined for D- and L-18A peptides (analogs of the class A amphipathic helix made from D- and L-amino acids). Greater than 50% of nonthyroid radioactivity was recovered in the liver from rats injected with either peptide; the kidneys had

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the second greatest uptake (40 and 22%, respectively). Rats injected with ~ - 1 8 Ahad five times as much radioactivity in the urine as did animals injected with D-18A; all radioactivity in L- 18A-injected animals were in the form of free Iz5I.Unpublished studies (Anantharamaiah and Garber, 1991) with ~ ~ - 3 7 ppeptide A (a dimer of D- and ~ - 1 8 Awith higher lipid affinity than 18A peptide) demonstrated a much lower kidney uptake. These observations suggest that the kidney may be a secondary organ of clearance of non-lipoprotein-bound peptides, whereas the liver is the primary clearance site of HDL-bound peptides (and perhaps apolipoproteins). Studies of de novo-designed amphipathic peptides, excluding for the moment the question of the role of charged residues, have demonstrated the following features: (1) The degree of amphipathicity correlates with the ability of peptides to interact with phospholipid; e.g., an increase in the hydrophobicity of the nonpolar face increases the lipid affinity (Kanellis et al., 1980; Fukushima et al., 1980; Sparrow et al., 1981; Anantharamaiah et al., 1987). (2) Lipid association increases the a helicity of the peptides (Kanellis et al., 1980; Fukushima et al., 1980; Sparrow et al., 1981; Ponsin et al., 1986b). (3) Inclusion of a Pro within the sequence of the putative helix decreases the lipid affinity of the peptide (Pownallet al., 1987). (4) Stereospecificity for the lipid-associating ability of an amphipathic helix does not exist. This is proved by the fact that amphipathic helices synthesized entire from D-amino acids are as efficient in associating with lipid as are those synthesized from L-amino acids (S. K. Srinivas et al., 1990). Our laboratory focused on the importance of the charged residue arrangement on the lipid-associating ability of de novo-designed amphipathic helical peptide analogs. Some of the salient features of our findings using a set of these peptides have been mentioned in the discussion of the snorkel hypothesis. The de novo-designed peptides have been very useful in addressing particular questions, such as the requirement of a particular amino acid in a specific position for a specific functional activity. Thus, synthetic peptide analogs addressed the issue as to whether apoA-I is the major LCAT activator by localizing the major LCAT-activating domain (Anantharamaiah et al., 1990a).The study of a Pro-linked dimer of an 18-residue peptide (18A) has provided information about cooperatively between Pro-linked tanden repetitive amphipathic helical domains in apoA-I in determining lipid affinity (Anantharamaiah et al., 1985). Peptide analogs have also been used to study the snorkel hypothesis, which would predict that the Lys residues in 18A are in different microenvironments, whereas in 18R, Lys residues are in the same microenvironment. N M R studies of peptide-dimyristoly-

THE AMPHlPATHlC u-HELIX MOTIF

349

phosphatidylcholine (DMPC) complexes of ['3C-d2methyZ118A and its reversed-charge distribution analog were in agreement with this (LundKatz et al., 1993). Model synthetic peptide analogs have also been useful in understanding the putative HDL receptor binding, as we have recently described (Mendez et al., 1992). We believe that de novo-designed peptides should give information on the possible in vivo role of different classes and subclassesof amphipathic helical domains present in different apolipoproteins. 2. Studies of Native Apolipopotein Sequences

A number of laboratories have examined the lipid affinity of hydrolytic fragments and synthetic peptide analogs of apolipoproteins. Figure 7 contains a diagrammatic summary of the locations in the exchangeable apolipoproteins of lipid-associating domains (lines with double arrowheads) and non-lipid-associating domains (lines without arrowheads) suggested by these experiments. Human apoA-I1 is a homodimer of two 77-residue-long monomers. Both the dimer and the carboxymethylated monomer associate with lipid to form lipoprotein complexes (Jackson et al., 1973). Synthetic peptides apoA-II[47-771 and apoA-II[40-771 associate with phospholipid, whereas peptides apoA-II[65-771 and apoA-II[56-771 have essentially no lipid affinity (Ma0 et al., 1977). In other studies, peptide apoA-11[173 11 failed to associate with lipid, but the addition of five more residues, apoA-II[12-311, resulted in lipid association (Chen et al., 1979; Kroon et al., 1978). This suggests that there are at least two lipid-associating domains in apoA-I1 located at opposite ends of the molecule (Fig. 7). T h e amphipathic helical structures belong to the class A2 motif. About 5% of HDL and about 40 to 60% of VLDL protein are apolipoprotein C. ApoC-I is a 57-amino-acid residue polypeptide and is the smallest of the exchangeable apolipoproteins. CNBr treatment of apoC-I produced two fragments, apoC-I[ 1-38] and apoC-I[39-571; the aminoterminal fragment, apoC-I[ 1-38] had the stronger lipid affinity (Jackson et al., 1974). In another study, the synthetic peptide apoC-I[32-571 was found to associate with phospholipid (Sparrow et al., 1977). Therefore, there appear to be at least two lipid-associating domains in apoC-I located between residues 1-31 and 32-57 (Fig. 7). The amphipathic helical domains of apoC-I are good examples of class A2 amphipathic helices. This supports experimental results showing that this protein interacts with lipid avidly. ApoC-I1 is a small component of HDLs and is present in VLDLs in large amounts. T h e N-terminal region is probably not involved in lipid association because of the presence of three Pro residues, which might

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JERE P. SECREST E T AL.

disrupt any amphipathic helical nature in this region. Several synthetic peptide fragments of apoC-I1 have been examined for lipid association. ApoC-II[50-781 and shorter peptides did not associate with phospholipid, but apoC-II[43-781 was able to form phospholipid-peptide complexes (Catapano et al., 1979), suggesting that a lipid-associating domain was at least partially located between residues 43 and 50 (Fig. 7). No studies of the lipid-associating properties of the amino-terminal half of apoC-I1 have been reported. ApoC-111 is the most abundant of the C peptides. ApoC-I11 has a carbohydrate attached to Thr-74. The variations in the three isolated forms of apoC-I11 correspond to either the absence (apoC-111-0) or the presence of one (apoC-111-1) or two (apoC-111-2) sialic acid residues. Based on the proposal that the amphipathic helical region of apoC-I11 is located between residues 40 and 67 (Anantharamaiah et al., 1991), Sparrow et al. (1973) synthesized apoC-III[ 1-79], apoC-III[55-791, apoCIII[48-791, and apoC-III[41-791. Using fluorescence and circular dichroism changes in the presence of lipid, these authors concluded that the lipid-associating domain of apoC-111 is located between residues 4 1 and 79. In other supporting experiments, apoC-I11 was cleaved at the Arg40-Gly41peptide bond into two fragments using thrombin (Sparrow et al., 1977);apoC-III[41-791, but not apoC-III[ 1-40], was found to interact with phospholipid (Anne et al., 1977) (Fig. 7). T h e amphipathic helix at residues 40-67 corresponds to the class A2 motif and is likely to be the region representing the major lipid-associating domain of this protein. Residues 8-39 correspond to the G* motif. This is in agreement with experimental results in that the peptide corresponding to residues 40-79 interacts with the lipid and not the N-terminal half of the molecule (residues 1-39). ApoA-I is the major protein component of HDLs. CNBr fragmentation and studies using synthetic peptide corresponding to different regions of apoA-I have identified the lipid-associating domains to be in the C-terminal region of this protein. Our laboratory synthesized two amino-terminal peptide fragments, apoA-I[ 1-33] and apoA-I[8-331. Although a previous computer analysis suggested an amphipathic helix in this region (Segrest, 1977), the synthetic fragments associated weakly, if at all, with lipid (R. V. Srinivas et al., 1991). In the same study, apoA-I[661201 was found to associate well with phospholipid. Kroon and Kaiser (1978) synthesized apoA-I[ 147-1681 and apoA-I[ 158-1681 and found that only the longer peptide associated with phospholipid. Finally, it has been reported that synthetic peptides apoA-I[ 165- 1851, apoA-1[2 182431, apoA-I[202-2431, and apoA-I[ 195-2431 all associate with phospholipid, but apoA-I[225-2431 does not (Sparrow and Gotto, 1980) (Fig. 7).


35 1

Two out of eight 22-mer tandem repeats of apoA-I (helices 3 and 8) correspond to class Y and the other helices correspond to class Al. Among 22-mers, the 66-87 region of apoA-I does not possess a welldefined nonpolar face; two Glu residues appear at the nonpolar face and other nonpolar amino acids (Asn and Thr, two of each) are present in this face. This would therefore be a weak lipid-associating domain, in agreement with the hypothesis that the proposed hinged domain of apoA-I resides in this region. This region is thought to be lipid bound or unbound, depending on the circumstances. The lipid-unbound form may be responsible for protein-protein interaction on apoA-I-cell interaction. An examination of apoA-I sequences from other species and comparison to the 66- 120 region of human apoA-I indicated high sequence homology. The postulated major LCAT-activating domain is located in the 22-mer helices 2 and 3. The displaceability of apoA-I by apoA-I1 can be explained by the fact that apoA-I1 possesses a class A2 amphipathic helical domain whereas apoA-I has class A 1, a weaker lipid-associating motif compared with class A2. The 22-mers are punctuated by Pro residues, thus allowing apoA-I to fold on the surface of a particle with the radial curvature of HDL (Fig. 12B). ApoE is a polypeptide of 299 amino acid residues. Because of the ability of this protein to bind to lipoprotein receptors, it has been studied extensively. Proteolysis of apoE by thrombin treatment of hypertriglyceridemic VLDLs produces two apoE fragments designated E- 12 and E-22, with molecular weights of 12 and 22 kDa, respectively (Gianturco et al., 1983). Fragment E-22 corresponds to apoE[l-1911 and E-12 corresponds to apoE[ 192-2991. Although E- 12 remains associated with VLDLs, E-22 dissociates from thrombin-treated VLDLs. As noted previously, X-ray crystallograpy studies of the E-22 fragment indicate the presence of a four-helix bundle globular structure (Wilson et al., 1991). These results indicate that the lipid-associating domain(s) of apoE appear to be located on the carboxy-terminal half of the molecule (Fig. 7). The amphipathic helical domain corresponding to residues 202-266 is a 65-residue-long class A amphipathic helix, The helical net diagram shows that the hydrophobic face is straight, and thus the entire region can act as a long, single helix (Fig. 12A). The length of this helix may be the determining factor for the residence of this protein on the surface of the VLDL, a lipoprotein particle with greater surface radius of curvature than HDL. ApoA-IV is a protein of unknown function. It is a major protein of lymph, chylomicrons, and VLDLs. In human plasma the majority of apoA-IV is present unassociated with lipoproteins. N o experimental data have been published suggesting localization of the lipid-associating regions of apoA-IV.

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3. Studies of Intact Apolipopoteins a. Structure of Apolipoprotein-Lipid Complexes. Because of their surface activity, exchangeable apolipoproteins spontaneously interact with most phospholipid liposomes, coating the surface. With certain phospholipids, notably dimyristoylphosphatidylcholine, this reaction proceeds further, disrupting the liposomes until small ( class A, > class Y > class G*. This ranking does not, of course, take into account the number and cooperativity of amphipathic helices.

5. Lipoprotein Lipme Activation An amphipathic helix enhances the lipoprotein lipase activation capacity of apoC-I1 (Catapano et al., 1979; Vinio et al., 1983).Synthetic peptide studies localized the lipoprotein lipase activation site of this protein to residues 55-78. CNBr cleavage of this protein produced peptides apoC11[1-91, apoC-11[10-591, and apoC-I1[60-781. Lipoprotein lipase activation studies on these fragments indicate that the enzyme-activating domain is localized to apoC-II[60-781. Comparing these experimental results with the amphipathic helix map (Fig. 7), the lipoprotein lipase-activating domain of apoC-I1 corresponds precisely to the predicted carboxy-terminal class G* amphipathic helical domain (residues 60-76). 6 . Receptor Recognition

The receptor-binding domain of apoE has been localized to the region encompassing residues 130- 160, and the major lipid-binding domain resides in the carboxyl-terminal one-third of the polypeptide chain (Rall


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et al., 1986; Innerarity et al., 1987). From the amphipathic helix map (Fig. 7) the receptor domains and the lipid-binding domains of apoE correspond to class G* and class A amphipathic helical domains, respectively. Inhibiting monoclonal antibodies have been used to localize the heparin-binding of apoE to two sites at residues 142-147 and 243-272 (Weisgraber et al., 1986). The first site corresponds to the LDL receptorbinding domain of apoE and is recognized both in solution and when apoE is lipid bound. The second site is recognized only when apoE is lipid-free, suggesting that residues 243-272 are part of the major lipidbinding doman of apoE; from the amphipathic helix map (Fig. 7) the major lipid-binding domain is predicted to be the class A amphipathic helix located between residues 203 and 266.

7 . Lecithin-Cholesterol Acyltransferase Activation Lecithin-cholesterol acyltransferase is a water-soluble plasma enzyme that plays an important role in the metabolism of HDLs by catalyzing the formation of cholesteryl esters on HDLs through the transfer of fatty acids from the sn-2 position of phosphatidylcholine to cholesterol (Jonas, 1986). ApoA-I is the major cofactor of LCAT in HDLs and reconstituted lipoproteins (Fielding et al., 1972). Many laboratories have used techniques such as synthetic peptide analogs (Anantharamaiah et al., 1990a; Anantharamaiah, 1986), monoclonal antibodies (Banka et al., 1990), and recombinant HDL particles (Jonas and Kranovich, 1978) to attempt to identify the major LCAT-activating region of apoA-I. It is known that LCAT binds to interfaces, such as the surface of HDL. Because amphipathic helices are surface active, they have been suggested to play a role in the activation of LCAT (Fielding et al., 1972; Anantharamaiah, 1986). However, the enzyme does not require a cofactor for the hydrolysis of water-soluble substrates such as the p-nitrophenyl esters of fatty acids (Bonelli and Jonas, 1989).Because of this it has been suggested that the major role of amphipathic helices is to disrupt the waterphospholipid interface to expose the buried substrate to LCAT (Bonelli and Jonas, 1989). Although all of the exchangeable apolipoproteins contain amphipathic helices, apoA-I is the superior LCAT activator (Anantharamaiah et al., 1990a). Further (with the exception to be discussed below), all amphipathic helical peptides studied have an intrinsic upper limit to their ability to activate LCAT, approximately 30% of apoA-I (Anantharamaiah et al., 1990a; Ananthararnaiah, 1986). Therefore, simple water-phospholipid interface disruption by amphipathic helices may be necessary for LCAT activation, but is clearly not sufficient. Additional structural features must be involved.

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J E W P. SEGREST ET AL.

As shown in Fig. 7, structural analysis suggests the presence, starting at residue 44,of 10 tandem and structurally separate class A or class Y amphipathic helical domains in apoA-I. Several lines of evidence point to the amino-terminal region of these tandem amphipathic helices as the predominant LCAT-activating domain in apoA-I: 1. Our laboratory has suggested that the amphipathic helical domains between residues 66 and 120 are important for LCAT activation; this region represents two 22-mer helices and an intervening ll-mer. We proposed that a unique positioning of a Glu residue on the nonpolar face of helix 2 (residue 78 in apoA-I[66-871 and helix 3 (residue 111 in apoA-I[99-1201; see Fig. 7G) is responsible for the higher LCATactivating ability of apoA-I (Anantharamaiah et al., 1990a). Synthetic consensus peptides with the same positioning of Glu residues were found to be equipotent with apoA-I, on a weight basis, in activating LCAT. N o other synthetic peptides are known to be as active. 2. ApoA-I-specific monoclonal antibodies have been used in conjunction with synthetic peptides to suggest that part of the LCAT activation domain resides between residues 96 and 1 11 (Banka et al., 1990). 3. Earlier studies failed to identify a major LCAT-activating domain among the four fragments produced by CNBr hydrolysis of apoA-I (Fielding et al., 1972). It is likely that the functional importance of the 66-120 domain was missed in these earlier studies because two of the three methionines present in apoA-I are found at the end of helix 2 (residue 86) and in the middle of helix 3 (residue 112). It is likely that cleavage at these positions destroyed the conformation of the region required for LCAT activation. 4.Reconstituted apoA-I/cholesterol/palmitoyloleoylphosphatidylcholine discoidal complexes of homogeneous size activate LCAT to varying degrees, depending on the complex size. Fluorescence spectroscopy of the complexes suggests that the amino-terminal region of apoA-I (through helix 3) is not directly interacting with lipid in the complexes that are the poorest LCAT activators (Jonas et al., 1990). The mechanisms whereby the amino-terminal region of apoA-I might activate LCAT are not known but could involve a combination of increased substrate accessibility and stabilization of active intermediate compounds (Anantharamaiah et al., 1990a).

8. Putative High-Density Lipoprotein Receptor Activity It has been proposed that HDL binds to certain cells via high-affinity saturable binding sites (Gwynneand Strauss, 1982; Oram et al., 1981) and that this binding may be mediated by a membrane protein (Oram and


36 1

Graham, 1987). Alternatively, it has been suggested that HDL binding to cells may occur via amphipathic helix-mediated binding to membrane lipid. Several synthetic amphipathic peptide analogs were tested for their ability to promote cellular cholesterol efflux and compete with labeled HDL for binding to the high-affinity HDL-binding protein (HBP) (Handwerger et al., 1987). The 18-amino-acid residue peptide, 18A, capable of interacting with lipid did not interact with HBP. However, a dimer of 18A linked by Pro to form a 37-residue peptide interacted with HBP and independently caused cholesterol efflux (Mendez et al., 1992). These results suggest that a de novo-designed class A amphipathic helical motif common to the exchangeable apolipoproteins is sufficient to promote HDL receptor-mediated cholesterol transport from cells and raises the question as to the specificity of the HBP protein and even whether there is, in fact, a receptor involved in direct interactions with HDL. Human placental lactogen is a protein hormone, the release of which has been shown to be stimulated by incubation of HDL with an enriched fraction of cultured trophoblast cells: this biological activity is due to apolipoproteins A-I, A-11, and C-I (Handwerger et al., 1987).Jorgensen et al. (1989) have shown that the synthetic peptide analogs of the class A amphipathic helix mimic apolipoproteins in this biological activity. The degree of human placental lactogen release from trophoblasts by these peptide analogs is correlated with lipid affinity, thus suggesting that the role of apolipoproteins in human placental lactogen release may be mediated through an interaction of amphipathic helices with plasma membrane phospholipids (Jorgensen et al., 1989). A study by LeBlond and Marcel (1991) using monoclonal antibodies supports the concept of a biological role for the direct interaction of amphipathic helices with plasma membranes; these authors suggest that “the optimum uptake o f . . . HDL . . . requires the . . . cooperative binding of the amphipathic a helical repeats [of apoA-I] to HepG2 cell membranes.” Consistent with direct interaction of amphipathic helixes with plasma membranes is the fact that apoA-I, apoA-11, and apoA-IV bind equally well, even though apoA-I1 is made up almost entirely of class A amphipathic helical domains (Fig. 7). 9. Antiviral and Antiinflammatory Activities of Amphipathic Helices

Several novel functions of apolipoproteins have been suggested to be mediated by the amphipathic helical domains. Owens et al. (1990) tested the effect of apoA-I on HIV-mediated cell fusion, the major cytopathic effect in HIV infections. Both amphipathic peptides and free apoA-I, but not HDL, effectively inhibited the HIV-induced cell fusion (Owens et al., 1990). HSV-induced cell fusion was also inhibited by apoA-I and am-

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FIG. 16. Two amphipathic helical segements of serum amyloid A. (A) Residues 1-24; (B) residues 53-73.

phipathic peptide analogs and not HDL (R. V. Srinivas et al., 1990, 1991), thus indicating that rhe amphipathic helical regions of apoA-I are involved in the fusion inhibitory effect. Blackburn et al. (1991) have demonstrated nontoxic inhibition of neutrophil activation using physiologic concentrations of free apoA-I and not HDL. These results suggest that the lipid-associating sites of apoA-I are responsible for this biological activity. Consistent with this is the finding that the amphipathic helical model peptide analogs also inhibit neutrophil activation (Blackburn et al., 1991). Serum amyloid A (SAA),an acute-phase protein, is not a major protein component of normal HDL. However, during an acute-phase response, the concentration of this protein increases and it specifically associates with HDL (Olphin and Price, 1988). Therefore, it is necessary to understand the properties of this protein from the point of view of the nature of the amphipathic helix present in this protein. SAA associates with


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lipids in a manner similar to that of exchangeable apolipoproteins (Segrest et al., 1976). Several investigators have shown that in vitro SAA associates with HDL and displaces apoA-I (Coetzee et al., 1986). This process could have implications in the immunoregulatory role of HDL. There are two amphipathic helical domains in SAA (Fig. 16). Although no fragments have been described, several investigators have studied the lipid-associating properties of SAA and showed that those properties are similar to that of apoA-I; thus SAA also acts as a protein detergent. 111. CONCLUSIONS

Evidence for the presence of amphipathic a helices in all of the exchangeable apolipoproteins, except apoE, is indirect but convincing. First, circular dichroism data show that association of exchangeable apolipoproteins and their peptide analogs with phospholipid produces a substantial percentage increase in a helicity of the proteins or peptides (Jackson et al., 1975). Second, the amino acid sequences of the putative amphipathic helical domains have the periodic patterns of a helices containing sharply demarcated polar and nonpolar faces (Segrest and Feldmann, 1977). Taken together, these two observations provide strong evidence that the putative amphipathic helical domains are a helical and amphipathic when the apolipoproteins are bound to lipid. In summary, w e have shown that the predicted locations and properties of class A and class G* amphipathic helices shown in Fig. 7 are in good agreement with the existing experimental data. We suggest that the limits of lipid-associating amphipathic helical domains can be more accurately defined by helical wheel-based algorithms than can the limits of amphipathic helices involved in protein associations. In any case, the preliminary amphipathic helix map should prove useful as a guide for future experimentation. It provides, for example, a working model for the design of site-specific mutations to map formally the structure-function relationships of the different amphipathic helical domains in the exchangeable apolipoproteins.

REFERENCES Anantharamaiah, G. M. (1986). In “Methods in Enzymology” (J. Segrest and J. Albers, eds.), Vol. 128, pp. 626-668. Academic Press, Orlando, FL. Anantharamaiah, G . M., Jones, J. L., Brouillette, C. G., Schmidt, C. F., Chung, B. H., Hughes, T. A., Bhown, A. S., and Segrest,J. P. (1985).J.Eiol. Chem. 460,10248-10255. Anantharamaiah, G. M., Hughes, T. A., Iqbal, H., Gawish, A., Neame, P., Meadley, M. F., and Segrest, J, P. (1987).J. LipldRes. 49,309-318.

364


Anantharamaiah, G. M., Venkatachalapathi, Y. V., Brouillette, C. G., and Segrest, J. P. (1990a).Artffioscbrosi.~( D a h ) 10,95-105. Anantharamaiah, G. M., Brouillette, C. G., Engler, J. A., D e h f , H., Venkatachalapathi, Y.V., Boogaerts, J., and Segrest, J. P. (1990b).In “Molecular Biology of Atherosclerosis” (A. D. Attie, ed.), pp. 149-158. Elsevier, New York. Anantharamaiah, G. M., Brouillette, C. G., Engler, J. A., D e h f , H., Venkatachalapathi, Y. V., Boogaerts, J. L. (1991).Adv. Exp. Med. 485, 131-140. Anne, K. C.,Gallagher, J. G., Gotto, A. M., Jr., and Morrisett, J. D. (1977).Biochistry 16,

2151-2156. Argiolas, A., and Pisano, J. J. (1983).J.Biol. Chem. 258, 13697-13702. Argiolas, A., and Pisano, J. J. (1985).J.Biol. C h . 460, 1437-1441. Atkinson. D., Smith, H. M., Dickson, J., and Austin, J. P. (1976).Eur. J. Biochem. 64,

541-547. Atkinson, D., Small, D. M., and Shipley, G. G. (1980).Ann. N.Y. Acad. Sci. 348,284-298. Austin, M. A. (1989).Am.J. Ep2dnniol. 149,249-259. Banka, C. L., Smith, R. S., Bonnet, D. J., and Curtiss, L. K. (1990).“Localization of an Apolipoprotein A-I Epitope Critical for LCAT Activation,” 63rd Sci. Sess. American Heart Association. Dallas, TX. Baumstark, M., Wolfman, W.,and Werner, K. (1983).Biochim. Biophys. A c h 751, 108-1 14. Bazan, J. F., Fletterick, R. J., McKinley, M. P., and Prusiner, S . B. (1987).Protein Eng. 1,

125- 135. Bernheimer, A. W., and Rudy, B. (1986).Biochim. Biophrs A c h 864, 123-141. Blackburn, W. D., Dohlman, J. G., Venkatachalapathi, Y.V., Pillion, D. J., Koopman, W. J., Segrest, J. P., and Anantharamaiah, G . M. (1991).J. Lipid Res. 39, 191 1-1918. Boguski. M. S., Elshourbagy, N. A., Taylor, J. M., and Gordon, J. I. (1985).Proc. Natl. Acad. Sci. U.S.A. 84,992-996. Bonelli, F. S., and Jonas, A. (1989).J.Biol. C h . 464, 14723-14728. Brausser, R., DeMutter, J.. V a n h , B., Goormaghtigh, E., Ruysschaert, J. M., and Rosseneu, M. (1990).Biochim. Biophys. Actn 445, 1043-1050. Breiter, D. R., Kanost, M. R., Benning, M. M., Wesenberg, G., Kaw, J. H., Wells, M. A., Rayment, I., and Holden, H. M.(1991).Biochishy 30,603-608. Brouillette, C. G., Jones, J. L., Ng, T. C., Kercret, H., Chung, B. H., and Segrest, J. P. (1984).BiOChnnishy 43,359-367. Catapano, A. L.,Kinnunen, P. K. J., Breckenridge, W. C., Gotto, A. M., Jr.. and Sparrow, J. T. (1979).BiOChem. B e h y s . Res. Commun. 89,951-957. Chen, T. C., Sparrow, J. T., Gotto, A. M., Jr., and Morrisett, J. D. (1979).Biochemistry 18,

1617-1627. Cheung, M. C., Segrest, J. P., Albers, J. J., Cone, J. T., Brouillette, C. G., Chung, B. H., Kashyap, P. M., Glasscock, M. A., and Anantharamaiah. G. M. (1987).J.Lipid Res. 48,

913-929. Chou, P. Y.,and Fasman, G. D. (1978).Adu. Enzymol. 47,45-148. Chung, B. H., Anantharamaiah, G. M.,Brouillette, C. G., Nishida, T., and Segrest, J. P. (1985).J. Biol. Chnn. 460, 10256-10262. Coetzee, G. A., Strachan, A. F., van der Westhuyzen, D. R., Hoppe, H. C., Jeenah, M. S., and de Beer, F. C. (1986).J.Bwl.Chnn. 461,9644-9651. Cohen, C., and Parry, D. A. D. (1986).TrendCBioch. Sci. 11,245-248. Cornette, J. L., Cease, K. B., Margalit, H., Spouge, J. L., Berzofsky, J. A., and DeLisi, C. J. (1987).Mol. Biol. 195,659-685. Crick, F. H. C. (1953).Actn Clyshllogr. 6,689-697. Cullis, P. R., and Hope, M. J. (1985).In “Biochemistry of Lipids and Membranes” (D. E. Vance and J. E. Vance, eds.), p. 25.Benjamin/Cummings, Menlo Park, CA.


365

Curtiss, L. K., and Edgington, T. S. (1985).J.Biol. Chem.460,2982-2993. Curtiss, L. K., and Smit, R. S . (1988).J. Biol. Chem.463, 13779-13785. Driscoll, D. M., and Getz, G. S. (1986). In “Methods in Enzymology” (J. Segrest and J. Albers, eds.), Vol. 128, pp. 41-70. Academic Press, Orlando, FL. Edelstein, C., Kezdy, F. J., Scanu, A. M., and Shen, B. W. (1979).J.Lipid Res. 40, 143-153. Edelstein, C.,-Hatari, M., and Scanu, A. M. (1982).J. Biol. Chem.457,7189-7195. Eisenberg, D., and McLachlan, A. D. (1986). Nature (London) 319, 199-203. Eisenberg, D., and Wesson, M. (1990).Bqbolymers 49,171-177. Eisenberg, D., Weiss, R. M., and Tenvilliger, T. C. (1982). Nature (London) 499,371-374. Eisenberg, D., Schwartz, E., Komaromy, M., and Wall, R. (1984a).J. Mol. Biol. 179, 125142. Eisenberg, D., Weiss, R. M., and Tenvilliger, T. C. (1984b).Proc. Natl. Acad. Sci. U.S.A. 81, 140- 144. Ellens, H., Single, D. P., Alford, D., Yeagle, P. L., Boni, L., Lis, L. S., Quin, P. J., and Bentz, J. (1989).Biopolymrs 48,3692-3703. Engelman, D. M., and Zaccai, G. (1980). Proc. Natl. Acad. Sci. U.S.A. 77,5894-5898. Engelman, D. M., Henderson, R., McLachlan, A. D., and Wallace, B. A. (1980). Proc. Natl. Acad. Sci. U.S.A. 77,2023-2027. Engelman, D. M., Steitz, T. A., and Goldman, A. (1985).Annu. Reu. B w h y s . Bi0phys. C h . 15,321-353. Epand, R. M., Epand, R. F., Orlowski, R. C., Schuetter, R. J., Boni, L. T., and Hui, S. W. (1983).Biopolymers 44,5074-5084. Epand, R. M., Epand, R. F., Hui, S . W., He, N. B., and Rosenblatt, M. (1985). 1nt.J. P e p . Protein Res. 25, 594-600. Epand, R. M., Gawish, A., Iqbal, M., Gupta, K. B., Chen, C. H., Segrest, J. P., and Anantharamaiah, G. M. (1987).J. Biol. Chem.464,9389-9396. Fielding, C. J., Shore, V. G., and Fielding, P. E. (1972).Biochem. Bi0phys Res. Commun. 46, 1493- 1498. Finer-Moore, J., and Stroud, R. M. (1984).Proc. N a L Acad. Sci. U.S.A. 81, 155-159. Finer-Moore, J., Bazan, J. F., Rubin, J., and Stroud, R. M. (1989).In “Prediction of Protein Structure and the Principles of Protein Conformation” (G. Fasman, ed.), pp. 7 19-760. Plenum, New York. Fitch, W. M. (1977). Genetics 86,623-644. Forte, T. M., Nichols, A. V., Gong, E. L., Lux, S., and Levy, R. L. (1971). Biochim. Blophys. Acts 248,38 1-386. Forte, T. M., Gong, E. L., and Nichols, A. V. (1974).Biochim. Blophys. Acta 337, 169-174. Frey, S . , and Tamm, L. K. (1991).Bi0phys.J. 60,922-930. Fringeli, U. P., and Gunthard, H. H. (1981). In “Membrane Spectroscopy” (E.Grell, ed.), pp. 270-332. Springer-Verlag, Berlin. Fukushima, D., Yokoyama, S., Kroon, D. J., Ferenc, J. K., and Kaiser, E. T. (1980).J.Bwl. Chem. 255,10651-10657. Garber, D. W., Venkatachalapathi, Y. V., Gupta, K. B., Ibdah, J., Phillips, M. C., Hazelrig, J. B., Segrest, J. P., and Anantharamaiah, G. M. (1992). Arteriosck7osis Thromb. 14, 886-894. Gascuel, 0.. and Golmard, J. L. (1988). Comput. A#. Biosci. 4,357-365. Gianturco, S . H., Gotto, A. M., Jr., Hwang, S . C., Karlin, J. B., Lin, A. H. Y.,Prasad, S. C., and Bradley, W. A. (1983).J.Biol. Chem. 458,4526-4539. Gilbrat, J. F., Gamier, J., and Robson, B. (1987).J. Mol. Biol. 198,425-443. Glomset, J. A. (1968).J.Lzpul Res. 9, 155-167. Goldstein, J. L., Kita, T., and Brown, M. S. (1983).N . Eng1.J. Med. 309,288-296. Groener, T. E., van Gent, T., and van Tol, A. (1989). Biochim.Biophys. Acta 1004,93-100.

366


Gruner, S. M. (1985). Proc. Natl. Acad. Sci. U.S.A.82,3665-3669. Gwynne. J. T., and Straws, J. F. (1982). Endow. Rev. 3,299-329. Hamilton, R. L. (1984). In “Plasma Protein Secretion by the Liver” (H. Glaumann, T. Peters, Jr., and C. Redman, eds.), pp. 357-374. Academic Press, London. Handwerger, S., Quardfordt, S., Barrett, J., and Harman, L. (1987). J. Clin. Invest. 79, 625-628. Harrick, J. H. (1967). “Internal Reflection Spectroscopy.” Wiley, New York. Hauser, H., Pascher, J., Person, R. H., and Sundell, S. (1981). Biochim. Biophys. Acto 650, 21-51. Havel, R. J., Shore, V. G., Shore, B., and Dier, D. M. (1970). Circ. Res. 27,595-600. Havel, R. J., Kane, J. P., and Kashyap, M. L. (1973).j. Clin. Invest. 52,32-38. Holmgre, A., Johansson, L. B.-A., and Lindblom, G. (1987).J. Phys. Chem. 91,5298-5301. Hubner, W., and Mantsch, H. H. (1991). Bi0phys.J. 59,1261-1272. lnnerarity, T. L.,Weisgraber, K. H., Rall, S. C., and Mahley, R. M. (1987).Acta Med. S c a d . , SUpPl. 715,51-59. Jackson, R. L., Morrisett, J. D., Pownall, H. J.. and Gotto, A. M., Jr. (1973).J. Biol. Chem. 248, 52 18-5224. Jackson, R. L., Morrisett, J. D., Sparrow, J. T., Segrest, J. P., Pownall, H. J.. Smith, L. C., Hoff, H. F., and Gotto, A. M., Jr. (1974).J. Biol. Chem. 249,5314-5320. Jackson, R. L., Morrisett, J. D., Gotto, A. M., Jr.. and Segrest, J. P. (1975). Mol. Cell. Biochem. 6,43-50. Janus, E. D., Nicoll, A., Wootton, R., Turner, P. R., Magill, P. J., and Lewis, B. (1980).Eur.j. Clin. Invest. 10, 149-159. Jiao, S., Cole, T. G., Kitchems, R. T., Pfleger, B., and Schonfeld, G. (1991).Metab., Clin. Ex$. 39, 155-160. Johnson, W. J., Mahlberg, F. H., Chacko, G. K., Phillips, M. C., and Rothblat, G . H. (1988). J . Biol. C h . 263,14099-14106. Jonas, A. (1986).J. Lipul Res. 27,689-698. Jonas, A., and Kranovich, D. J. (1978).J. Biol.Chem. 453,5758-5763. Jonas, A., Kezdy, K., and Wald, J. (1989).j. Biol. Chem. 264,4818-4824. Jonas, A., Wald, J. H., Toohill, K. L. H., Krul, E. S., and Kezdy, K. E. (199O).J. Biol. Chem. 465,22123-22129. Jones, A. L., Hradek, G. T.. Hornick, C., Renaud, G., Windler, E. E. T., and Havel, R. J. (1984).j. LipuiRes. 25, 1151-1158. Jones, M. K., Anantharamaiah, G. M.,and Segrest, J. P. (1992).J. Lipid Res. 33, 287-296. Jorgensen, E. V.,Anantharamaiah, G. M., Segrest, J. P., Gwynne,J. T., and Handwerger, S. (1989).j. Bwl. C h . 264,9215-9219. Kaiser, E. T., and Kezdy, F. J. (1983). Proc. Natl. Acad. Sci. U.S.A. 80, 1 137-1 140. Kaiser, E. T., and Kezdy, F. J. (1984). Science 243,249-255. Kanellis, P., Romans, A. Y., Johnson, B. J.. Kercret, H., Chiovetti, R., Jr., Allen, T. M., and Segrest, J. P. (1980).J. Biol. C h . 255, 11464-1 1472. Kannel, W. B., Castelli, W. P., Gordon, T., and McNamara, P. M. (1971). Ann. Intern. Med. 74, 1-12. Koizumi, J., Inazu, A., Yagi, K., Koizumi, I., Uno, Y., Kajinami, K., Miyamoto, S., Moulin, P., Tall, A. R., Mabuchi, H., and Takeda. R. (1991). Atherosclerosis (Shannon,Irel.) 90, 189- 196. Kottke, B. A., Zinsmeister, A. R., Holmes, M. D., Jr,. Kneller, R. W., Hallaway, B. J., and Mao, S. J. (1986). MayoClin. Proc. 61, 313-320. Krebs, K. E., and Phillips, M.C. (1984). FEES Lett. 175,263-266. Kretsinger, R. H. (1980). CRC Cril. Rev. Biochem. 8, 119-174.


367

Kroon, D. J., and Kaiser, E. T. (1978).J. Org. C h . 43,2107-2113. Kroon, D. J., Kupferberg, J. P., Kaiser, E. T., and Kezdy, F. J. (1978).J. Am. C h . SOC.100, 5975-5977. Krul, E. S., and Schonfeld, G. (1986).In “Methods in Enzymology”(J.Segrest and J. Albers, eds.), Vol. 128, pp. 527-553. Academic Press, Orlando, FL. Kyte, J., and Doolittle, R. F. (1982).J. Mol. Biol. 157, 105-132. LeBlond, L., and Marcel, Y. L. (1991).J.Biol. C h . 266,6058-6067. Levitt, M. (1978). Biochemistry 17,4277-4285. Li, W.-H., Tanimura, M., Luo, C.-C., Datta, S., and Chan, L. (1988).J. L i e Res. 29, 243-27 1. Lim, V. I. (1978). FEES Lett. 89, 10-14. Lund-Katz, S., Anantharamaiah, G. M., Venkatachalapathi, Y. V., Segrest, J. P., and Phillips, M. C. (199O).J.B i d . C h . 265, 12217-12223. Lund-Katz, S . , Phillips, M. C., Segrest, J. P., and Anantharamaiah, G. M. (1993). In “Peptides: Chemistry and Biology”(R. Haughten, ed.). ESCOM Press, Leiden, The Netherlands. Luo, C. C., Li, W. H.,Moore, M. N., and Chan, L. (1986).J.Mol. Biol. 61,313-320. Mao, S . J. T., Sparrow, J. T., Gilliam, E. B., Gotto, A. M., Jr.. and Jackson, R. L. (1977). B i ~ ~ h i S t 16,4150-4156. ty Marcel, Y., Provost, P., Koa, H., Rafai, E., V u Dac, N., Fruchart, J. C., and Rassart, E. (1991).J.Biol. C h . 266,3644-3653. Marreo, H., and Rothschild, K. J. (1987). Biophys.J. 52,629-635. McLean, L. R., Hagaman, K. A., Owen, T.J., and Krstenansky,J. L. (1991). Biochemisty30, 31-37. Mendez, A. J., Anantharamaiah, G. M., and Oram, J. F. (1992). Circulation 86, 1-1, Abstract 7. Morton, R. E., and Zilversmit, D. B. (1982).J.L i e Res. 23, 1058-1067. Nakagawa, S. H., Lau, H. S. H., Kezdy, F. J., and Kaiser, E. T. (1985).J.Am.C h . SOC.107, 7087-7092. Nilsson-Ehle, P., Garfinkel, A. S., and Schotz, M. C. (1980). Annu. Rev. Biochem. 49,667693. Nolte, R. T., and Atkinson, D. (1992).Biophys.J.63, 1221-1239. Okamura, E., Umemura, J., and Takenaka, T. (1990). Biochim. Biophys. Actu 1025, 94-98. Olphin, S. E., and Price, C. P. (1988).Atherosclerosis(Shannon,Irel.) 69,93-94. Oram, J. F., and Graham, D. L. (1987).J. Biol. C h . 264,7349-7442. Oram, J. F., Albers, J. J.. Cheung, M. C., and Bierman, E. L. (1981).J. Biol. C h . 256, 8348-8356. Oram, J. F., Brinton, E. A., and Bierman, E. L. (1983).J. Clin. Invest. 72, 161 1-1621. Owens, R. J.. Anantharamaiah, G. M., Kalhone, J. B., Srinivas, R. V., Compans, R. W., and Segrest, J. P. (199O).J.Clin. Invest. 86, 1142-1 150. Parhofer, K. G., Hugh, P., Barrett, R., Bier, D. M.,andSchonfeld, G. (1991).J.LiPtdRes. 32, 1311-1323. Perutz, M. F., Kendrew. J. C., and Watson, H. C. (1965).J. Mol. Biol. 13,669-678. Piot, F., DeLoof, H., Vu Dac, N., Clavey, V., Fruchart, J. C., and Rosseneu, M. (1988). Bi~chim.Biophys. Acts 959, 160-168. Ponsin, G., Hester, L., Gotto, A. M., Jr., Pownall, H. J., and Sparrow, J. T. (1986a).J. Bzol. C h m . 261,9202-9205. Ponsin, G., Sparrow, J. T., Gotto, A. M., Jr., and Pownall, H. J. (1986b).J. Clzn. Invest. 77, 559-567.

368


Pownall, H. J., and Massey, J. B. (1982).Biophys.J. 37, 177-179. Pownall, H. J., Massey, J. B., Sparrow, J. T., and Gotto, A. M., Jr. (1987). In “Plasma Lipoproteins” (A. M. Gotto, Jr., ed.), pp. 95-127. Elsevier (Biomed. Div.), New York. Rall. S. C., Weisgraber, K. H., and Mahley, R. W. (1986).In “Methods in Enzymology” (J. Segrest and J. Albers, eds.), Vol. 128, pp. 273-287. Academic Press, Orlando, FL. Richardson, J. S., and Richardson, D. C. (1989).In “Prediction of Protein Structure and the Principles of Protein Conformation” (G. Fasman, ed.), pp. 1-98. Plenum, New York. Robinson, R. M., Blackeney, E. W., Jr., and Mattice, W. L. (1982). Biopolymers 21, 12171228. Rye, K. A., Garrety, K. H., and Barter, P. J. (1992).J.LzpidRes. 33,215-224. Scanu, A. M., Lagocki, P., Edelstein, C., and Byrne, R. (1980). Bi0phys.J. 37, 179-181. Segrest, J. P. (1976). FEES Lett. 69, 1 1 1-1 15. Segrest, J. P. (1977). C h . Phys. L i e 18,7-22. Segrest, J. P., and Feldmann, R. J. (1977). Biopolymers 16,2053-2065. Segrest, J. P., Jackson, R. L., Morrisett, J. D., and Gotto, A. M., Jr. (1974). FEES Lett. 38, 247-253. Segrest, J. P., Pownall, H. J.. Jackson, R. L., Glennar, G. G., and Pollack, P. S. (1976). Biochemisty 15,3187-3191. Segrest, J. P., DeLoof, H., Dohlman, J. G., Brouillette, C. G., and Anantharamaiah, G. M. (1990). PrOtn’W 8, 103-1 17. Segrest, J. P., Venkatachalapathi, Y. V., Srinivas, S. K., Gupta, K. B., DeLoof, H., and Anantharamaiah, G. M. (1991). In “Molecular Conformation and Biological Interactions” (P. Balaram and S. Ramachandran, eds.), pp. 597-609. Indian Academy of Sciences, Bangalore. Segrest. J. P., Jones, M. K., DeLoof, H., Brouillette, C. G., Venkatachalapathi, Y. V., and Anantharamaiah, G. M. (1992).J. LipldRes. 33, 141-166. Shapiro, J. P., Law, J. H., and Wells, M. A. (1988).Annu. Rev. Entomol. 33,297-318. Sherrill, B. C., Innerarity, T. L., and Mahley, R. W. (198O).J.Biol. Chem. 255, 1801-1807. Shiffer, J., and Edmundson, A. B. (1967). Bi0phys.J. 7, 121-135. Silberman, S. R., Bernini, F., Sparrow, J. T., Gotto, A. M., Jr., and Smith, L. C. (1987). B i o C h i ~ l ~26, y 5833-5843. Sitaram, N., and Nagaraj, R. (1989).J. Biol. C h . ,265, 10438-10442. Slotte, J. P., Oram, J. F., and Bierman, E. L. (1987).J.Bwl. C h . 465, 12904-12907. Soravuia, E., Martini, G., and Zasloff, M. (1988). FEES Lett. 228,337-340. Sparrow, J. T., and Gotto, A. M., Jr. (1980). Ann. N.Y.Acad. Sci. 348, 187-21 1. Sparrow, J. T., and Gotto, A. M., Jr. (1982). CRC Crit. Rev. B i o c h . 13,87-107. Sparrow, J. T., Gotto, A. M., Jr., and Morrisett, J. D. (1973).Proc. Natl. Acad. Sci. U.S.A. 70, 2 124-2 128. Sparrow, J. T., Pownall, H. J., Sioler, G. F., Smith, L. C., Soutar, A. K., and Gotto, A. M.,Jr. (1977). In “Peptides, Proceedings of the Fifth American Peptide Symposium” (M. Goodman and J. Meienhofer, eds.), pp. 149-152. Wiley, New York. Sparrow, J. T., Ferenz, C. R., Gotto, A. M., Jr.. and Pownall, H. J. (1981). In “Peptides: Synthesis-Structure-Function” (D. Roch and E. Gross, eds.), pp. 253-256. Pierce Chem. Co., Rockford, IL. Srinivas, R. V., Birkedal, B., Owens, R.J., Anantharamaiah, G. M., and Segrest, J. P. (1990). Virology 176,48-57. Srinivas, R. V., Venkatachalapathi, Y.V., Zheng, R., Owens, R. J.. Gupta, K. B., Srinivas, S . K., Anantharamaiah, G. M., Segrest, J. P., and Compans, R. W. (1991).J.Cell. Biochem. 45,224-237. Srinivas, S . K.,Venkatachalapathi, Y. V., Gupta, K. B., Segrest, J. P., and Anantharamaiah, G. M. (1990). FASEBJ. 4, A2148 (Abstr. 2633).


369

Steinmetz, A., and Uterman, G . (1985).J.Biol. Chem. 260,2258-2264. Tall, A. R., Small, D. M., Shipley, G. G., and Lees, R. S.(1975).Proc. Nutl. Acad. Sci. U.S.A.

72,4940-4942. Tall, A. R., Small, D. M.. Dekelbaum, R. J., and Shipley, G. G. (1977).J.Biol. Chem. 252,

4701-4711. Tall, A. R., Sammett, D., Vita, G . M., Deckerbaum, R., and Olivecrona, T. (1984).J.Biol. Chem. 259,9587-9594. Tanford, C. (1978).Science 299, 1012-1018. Tatami, R.,Mabuchi, H., Ueda, K., Ueda, R., Haba, T., Kametani, T., Ito, S., Koizurni,J.. Ohta, M., Miyamoto, S., Nakayama, A., Kanaya, H., Oiwake, H., Genda, A., and Takeda, R. (1981).Circulation 64,1174-1 183. Taylor, J. W., and Kaiser, E. T. (1986).Phann. Rev. 38, 291-319. Taylor, J. W., Osterman, D. G., Miller, R. J.. and Kaiser, E. T. (1984).J. Am. Chem. SOC.103,

6965-6966. Tytler. E. M., Segrest, J. P., Epand, R. M., Nie, S., Epand, R. F., Mishra, V. K., Venkatachalapathi, Y. V., and Anantharamaiah, G. M. (1993).J.Biol. Chem. 268,in press. Venkatachalapathi, Y. V., Gupta, K. B., DeLoof, H., Segrest, J. P., and Anantharamaiah, G . M. (1990).In “Peptides: Chemistry and Biology” (J. Rivier, ed.), pp. 672-673. ESCOM Press, Leiden. Verhallen, P. F. J., Demel, R. A,, Zwiers, H., and Gispen, W. H. (1984).Biochim. Biophys. Acta

775,245-254. Vinio, P., Virtanan, J. A., Kinnunen, P. K. J., Gotto, A. M., Jr., Sparrow, J. T., Pattus, F., Bugis, P.,and Verger, R. (1983).J. Biol. Chem. 258,5477-5482. von Eckardstein, A,, Funke, H., Walter, M., Altland, K., Benninghoben, A., and Assman, G. ( 1990).J . Biol. Chem. 265,8610-86 17. von Heijne, G. (1986).E M B O J . 5, 1335-1342. Wald, J. H.,Goormaghtigh, E., DeMutter, J., Ruysschauer, J. M., and Jonas, A. (1990). J. Biol. Chem. 265,20044-20050. Weinberg, R. B., and Jordan, K. J. (199O).J.Biol. Chem. 565,8081-8086. Weinberg, R. B., and Spector, M. S.(1985).J.Biol. Chem. 260,4914-4921. Weisgraber, K., Rall, S. C., Mahley, R. W., Milane, R. W., Marcel, Y. L., and Sparrow, J. T. (1986).J. Biol. Chem. 261,2068-2076. Wilson, C., Wardell, M. R., Weisgraber, K. H., Mahley, R. W., and Agard, D. A. (1991). Science 252,1817-1822.

Wilson, P. W. F., Abbott, R. D., and Castelli, W. P. (1988).Arteriosclerosis ( D a l h ) 8,737-741. Wlodawer, A., Segrest, J. P., Chung, B. H., Chiovetti, R., Jr., and Weinstein, J. N. (1979). FEES Lelf. 104,231-235. Wu, C . 4 . C.,and Yang, J. T. (1980).Biochemistry 19,2117-2122. Zasloff, M., Martin, B., and Chen, H. C. (1988).Proc. Natl. Acad. Sci. U.S.A. 85,910-913.


LIPOPHORIN: THE STRUCTURE OF AN INSECT LIPOPROTEIN AND ITS ROLE IN LIPID TRANSPORT IN INSECTS By JOSC L. SOULAGES and MICHAEL A. WELLS Department of Blochemlstry and Center for insect Sclence, Bloiogkal Sciences West, University of Arizona, Tucson, Arlzona 86721

1. 11.

111.

1V.

V.

VI. VII.

Introduction ....................................................... Lipid and Apolipoprotein Composition of Lipophorins ................. A. Lipid Composition of Lipophorins ............................... B. Apolipoproteins ................................................ Size, Molecular Weight, Heterogeneity, and Shape of Lipophorins ....... Organization of Lipids and Proteins in Lipophorins .................... A. Apolipoproteins ................................................ B. Location of Phospholipids ....................................... C. Location of Hydrocarbons ....................................... D. Location of Diacylglycerols ...................................... E. Lipophorin Models ............................................. Metabolism ........................................................ A. Biosynthesis ................................................... B. Physiological Roles of Lipophorin ................................ Metabolic Implications of Lipophorin Structure ........................ Concluding Remarks and Future Directions ........................... References .........................................................

371 372 373 375 384 388 389 390 390 390 391 393 394 397 405 408 409

I. INTRODUCTION Vertebrate, especially mammalian, lipoproteins have been extensively studied. In the invertebrate world, only insect lipoproteins have received serious attention. Whereas vertebrates rely on a battery of lipoproteins (chylomicrons, very low-density lipoproteins, low-density lipoproteins, and high-density lipoproteins) to effect lipid transport, insects use primarily a single type of lipoprotein, lipophorin, for lipid transport. Lipophorin is both more versatile than vertebrate lipoproteins in terms of the diverse lipids it transports and more efficient than vertebrate lipoproteins in that, for the most part, it delivers lipids to tissues without being internalized and destroyed. We believe that new insights can be obtained from an understanding of insect lipoproteins, and in this article we review the current state of knowledge about the structure and metabolism of lipophorins. Insects are the dominant terrestrial group of animals. Several hundred thousand species of insects have been described, and some investigators ADVANCES IN PROTEIN CHEMISTRY. Val. 45

37 1

Copyright Q1994 by Academic Press. Inc. All righu of reproduction in any form reserved.

372

JOSL L. SOULAGES AND MICHAEL A. WELLS

believe the total number of insect species may number in the millionsseveral times the total number of all other animal species. Insects first appeared about 350 million years ago and have evolved to utilize for food almost every available organic resource on the planet. Humans usually regard insects as adversaries, carriers of disease, and pillagers of food sources. However, life as we know it could not exist without insects: pollinating activities, scavenging and recycling refuse, and serving as a food source for other animals are only a few of the essential roles played by insects. Because of their extraordinary diversity, insects also have considerable potential as subjects for research; it is likely that new and novel solutions to biological problems can be found among insects. Generally, insect biochemistry and physiology resemble that of vertebrates: basic metabolic pathways and their control are similar and endocrine control is exerted by equivalent mechanisms, although the structures of the hormones differ. Insects have an open circulatory system in which blood (hemolymph) is enclosed by basement membranes that surround all tissues. Generally, unlike hemoglobin, hemolymph does not contain oxygen-carrying molecules; oxygen diffuses to the tissues through a network of trachea and tracheoles that are open to the atmosphere. A specialized tissue characteristic of insects is the fat body. Dispersed throughout the insect body, the fat body combines many of the functions of vertebrate liver and adipose tissue. T h e fat body is perhaps the most versatile metazoan tissue: it is the site of synthesis of most of the hemolymph proteins; it stores fat, glycogen, waste material, and specialized proteins; it produces most of the components that make u p the insect egg yolk and in some insects contains specialized endosymbionts. Detailed studies on insect lipoproteins have been carried out only in the last decade or so (for previous reviews, see Chino, 1985; Beenakkers et al., 1985; Shapiro et al., 1988; Law and Wells, 1989; Ryan, 1990; Van der Horst, 1990; Law et al., 1992). For the most part our understanding of insect lipoproteins and their metabolism have been derived from studies on only two species: Manduca sexta and Locusta migratoria. When other species have been studied, the picture developed using M. sexta and L. migratoria has been generally confirmed. However, more than 99.9% of all insect species have not been investigated and, considering the diversity in food sources and life histories, it would not be surprising if the analysis of more species revealed new and exciting variations on the theme developed in this review. 11. LIPIDAND APOLIPOPROTEIN COMPOSITION OF LIPOPHORINS

Insect lipoproteins are generally isolated by single-step ultracentrifugation in a density gradient. In all insects studied to date, the majority

LlPOPHORlN LIPID TRANSPORT IN INSECTS

373

of hemolymph lipids have been found associated with a single lipoprotein particle. Thus, even though there is a considerable variation in lipid content and composition among the insect lipoproteins, the common name lipophorin has been given to all insect lipoproteins (Chino et al., 1981a). Lipophorins are further named according to their buoyant density range: low-density lipophorin (LDLp), high-density lipophorin (HDLp), and very high-density lipophorin (VHDLp) (Beenakkers et al., 1988). Lipophorins can be further identified by indicating the life stage from which they were isolated: HDLp-L, high-density lipophorin from larvae; HDLp-A, high-density lipophorin from adults, etc. Two apolipoproteins have been observed in all lipophorins: apolipophorin I (apoLp-I) with a molecular mass of 230-250 kDa and apolipophorin I1 (apoLp-11)with a molecular mass of 70-85 kDa. In LDLp and some adult HDLp a third apolipoprotein is found, apolipophorin 111 (apoLp-111) with a molecular mass of 18-20 kDa. A. Lipid Composition of Lipophorins

The lipid composition of lipophorin and its concentration in hemolymph are functions of the age and developmental stage of the insects (Ziegler, 1984; Wheeler and Goldsworthy, 1983; Prasad et al., 1986a; de Bianchi et al., 1987; Telfer et al., 1991; Gonzalez et al., 1991). Significant changes in the lipid content and composition are also observed during starvation (Mwangi and Goldsworthy, 1977a; Tsuchida et al., 1987; Ziegler, 1991) and flight (Beenakkers, 1973; Justum and Goldsworthy, 1976; Van der Horst et al., 1978; Ziegler and Schulz, 1986). Table I contains a compilation of the lipid composition found in lipophorins. An obvious hallmark of lipophorins is their high content of diacylglycerol (DG), which in most cases is the main neutral lipid. sn-1,2Diacylglycerol accounts for most of the DG found in lipophorin. The small amounts of 1,3-DG occasionally reported probably represent an isomerization product of sn-1,2-DG (Tietz and Weintraub, 1980). Another unique characteristic of lipophorins is the presence of long chain ( C ~ O - ~normal, O), and methyl-branched aliphatic hydrocarbons, which in some insects or metabolic stages represents a significant proportion of the nonpolar lipid components (Katase and Chino, 1982; Blomquist et al., 1987; Katagiri et al., 1985; Katagiri and de Kort, 1991). Compared with vertebrate lipoproteins, lipophorins show a virtual absence of triacylglycerols and cholesterol esters, as well as a very low content of free sterols. The only common major lipid component of vertebrate and insect lipoproteins is phospholipid (PL). Lipophorin PLs have been characterized in only a few insect species (Table 11). The major PLs are phosphati-

374

JOSk L. SOULAGES AND MICHAEL A. WELLS

TABLE I L i e Compositwn and Density of Lipophminp Lipid

Insect Having lipophorins without apoLp-111 Acheta domesticus (A) A@ meUi@a (L) Diotrca grandiaulla (L) Drmophila melanogask-r(L) Lglinotarsa &cemlincata (A) Locurla migrolmia (A) Manduca smta Larvae Prepupal- 1 Prepupal-2 Pupae Larvae Larvae Musca domesfica (A) Periplaneta americana (A) P h i h m i a cynfia (L) Podisus maculivmtris (A) Tipula trivittafa (A) T d o m a infestam (A) Having lipophorins with apoLp-Ill Acheta domeslicus LDLp (A) Locurta migroforia LDLp (A) Manduca scxta HDLp (A) Manduca sexta LDLp (A)

DC

PL

HC

18.0 13.3 15.4 7.4 2.2 13.4

14.0 12.8 13.0 23.1 18.7 14.8

4.5 2.0

15.7 20.2 12.5 17.5 30.3 21.4 6.0 8.0 24.8 16.7 21.6 19.5

16.7 23.3 18.9 21.6 20.8 17.7 20.0 22.8 11.4 14.9 12.0 14.9

36.6 26.1 25.0 46.9

8.9 10.9 14.0 7.1

ST

(9%)

Density

Ref!

-

-

3.9 3.2 5.4

6.0 6.4 0.7 1.2 3.2

42.8 41.0 38.0 37.5 44.9 41.0

1.106 1.130 1.110 1.16 1.09 1.12

1 2 3 4 5 6

1.151 1.128 1.177 1.139 1.155 1.144 1.145 1.12

7

1.5 5.7

2.2 1.0 2.7 0.5 5.8 - ND 3.6 7.3 1.9 2.1

37.3 46.9 34.8 46.4 47.5 45.3 34.2 49.6 44 32.0 51.4 47.0

1.10

11 12 13 14

3.5 6.4 3.5 2.3

ND 0.5 2.5 1.7

ND 2.4 1.3 0.7

52.2 46.3 51.5 62.2

1.061 1.065 1.08 1.03

1 15 16 16

-

TG

0.9 20.4 8.7

0.7

2.8

1.1

0.6

1.1 1.0

0.5 0.4 2.3 3.1 6.0 15.0 0.6

-

-

1.0 0.7 2.1

-

1.2 1.8 1.8 2.8 0.6 1.0

-

1.16 1.117

7 7 7 8 9 10 11

a DG, Diacylglycerol; PL, phospholipid; HC, hydrocarbon, TG, triacylglycerol; ST, sterol; L, larval stage; A. adult stage; ND, not determined. Other lipophorins partially characterized: Bombyr mon' (Miura and Shimizu, 1989a,b),Rhodniusprolh (Gondim ct al., 1989a), C h i n a nwrsilans (Ochanda et al., 1991). Key to references: (1) Strobel et al., 1990; (2) Robbs et al., 1985; (3) Dillwith el al., 1986; (4) Fernando-Warnakulasuriya and Wells, 1988; (5) de Kort and Koopmanschap, 1987; (6)Chino and Kitazawa, 1981;(7) Prasad etal., 1986a;(8) Pattnaik etal., 1979; (9)Tsuchida etal., 1987; (10) Capurro. 1988; (11) Chinoetal., 1981b; (12) Haunerlandetal., 1992; (13) Neven et al., 1989; (14) Rimoldi et al.,1991; (15) Chino et al., 1986; (16) Ryan el al., 1986a.

dylethanolamine (PE) and phosphatidylcholine (PC). With the exception of L. mipatoria lipophorin, all insect lipophorins have a high content of PE. Lipophorin from the freeze-tolerant cranefly Tiplla trivzttutu contains significant amounts of phosphatidylinositol (PI) (Neven et al., 1989), which has not been detected in any other lipophorin. The presence of PI is essential for the ice nucleation activity of this lipophorin, an activity not shown by any other lipophorin. Low amounts of sphingomyelin, lyso-PC, lyso-PE, and acidic phospholipids have also been detected in lipophorin.

375

LIPOPHORIN LIPID TRANSPORT IN INSECTS

TABLE I1 Phospholipid Composition of Lipoplrorinp Insect

PC

PE

Diatraea grandiosello Leptinotarsa decemlineatu Locwta migratoria Mandwa sex& Periplaneta americana Philosomia cyntia Tipulo triuittatu Triatom infestans

40.0 54.4 95.0 34.3 68.0 48.0 16.0 35.9

51.0 45.6 5.0 54.4 32.0 32.0 62.0 64.1

SPH

PI

9.0

-

11.3 20.0 10.5

-

Ref. Dillworth ef al. (1986) Katagiri and de Kort (1991) Chino and Downer (1982) Pattnaik et al. (1979) Chino and Downer (1982) Chino and Downer (1982) Neven el al. (1989) Rimoldi et al. (1991)

PC, Phosphatidylcholine; PE, phosphatidylethanolamine; SPH, sphingomyelin; PI, phosphatidylinositol.

Although PLs are common components of vertebrate and insect lipoproteins, the high PE : PC ratio found in liphophorins distinguishes them from vertebrate lipoproteins, where PC is the predominant polar lipid (Chapman, 1986). Free fatty acids are also found in the hemolymph of most insects, presumably bound to lipophorin (Dillwith et al., 1986; Beenakkers, 1973; Van Marrewijk et al., 1984; Rimoldi et al., 1991; Pattnaik et al., 1979; Ryan et al., 1986a). Analysis of the fatty acid composition of lipophorin lipids has been performed in only a few insect species and myristate, palmitate, stearate, oleate, and linoleate were detected as the main fatty acids of PLs and DG (Beenakkers et al., 1985; Katagiri and de Kort, 1991; Miura and Shimizu, 1989a,b; Fichera and Brenner, 1982; Fernando- Warnakulasuriya and Wells, 1988; Kuthiala and Chippendale, 1989; Neven et al., 1989). B . Apolapopoteans 1 . Structural Proteins

There is one molecule each of apoLp-I and apoLp-I1 in each lipophorin molecule (Shapiro et al., 1984; Surholt et al., 1992). A similar amino acid composition, as well as the presence of oligosaccharide chains of the high-mannose type, has been observed for apoLp-I and apoLp-I1 from different insect species (Pattnaik et al., 1979; Ryan et al., 1984; Shapiro et al., 1984; Kashiwazaki and Ikai, 1985; Dillwith et al., 1986; Nagao and Chino, 1987; Rimoldi et al., 1991). Both apolipoproteins are water insoluble when separated from lipophorin, and in this regard resemble vertebrate apoB. No amino acid sequence data, either from direct or cDNA

376

JOSE L. SOULAGES AND MICHAEL A. WELLS

sequencing, has been published for either apoLp-I or apoLp-I1 from any insect. It has been shown that apoLp-I and apoLp-I1 are not immunologically related (Ryan et al., 1984;Schulz et al., 1987).Polyclonal antiserum against M. sexta apoLp-I1 showed cross-reactivity to the apoLps-I1 of seven insect orders, whereas antiserum against M. sexta apoLp-I did not cross-react with apoLp-I from any of the insect species tested (Ryan et al., 1984). On the other hand, monoclonal antibodies prepared against L. mzgratoria apolipophorin I, 11, or I11 did not cross-react with the corresponding apolipoproteins from three other species of insects (Schulz et al., 1987). In lipophorin, apoLp-I is much more susceptible to proteolytic cleavage than is apoLp-11, and iodination of native lipophorin results in iodination of apoLp-I but not apoLp-I1 (Pattnaik et al., 1979;Shapiro el al., 1984; Kashiwazaki and Ikai, 1985). Polyclonal antibodies raised against lipophorin, when analyzed by Western blotting, generally show stronger reactivity toward apoLp-I than toward apoLp-I I. Antibodies raised against purified apoLp-I will react with native lipophorin, whereas antibodies raised against purified apoLp-I1 generally do not react with native lipophorin (Shapiro et al., 1984).On the other hand, cross-linking experiments show that the two apolipoproteins are in close proximity in lipophorin (Kashiwazakiand Ikai, 1985),and monoclonal antibodies can be raised against apoLp-I1 when immunization is carried out with lipophorin (Schulz et al., 1987). These data suggest that apoLp-11, or a portion of apoLp-11, is somehow “sequestered” from the aqueous environment in native lipophorin. Of course, the environment and/or the folding of apoLp-I1 in lipophorin might render it protease resistant or unreactive with iodine without requiring that the protein be buried in the outer lipid layer of lipophorin. ApoLp-I and apoLp-I1 can be dissociated from lipophorin in 8 M guanidinium chloride (GdmCI) and isolated by gel-permeation chromatography (Kawooya et al., 1989).Under these conditions, apoLp-I1 is isolated as a lipid-free protein, which is, however, insoluble when GdmCl is removed, even in the presence of detergent. ApoLp-I isolated by this procedure has associated with it all the lipid present in lipophorin, but the lipid can be removed by extracting the GdmCl solution with ethanol : ether to yield a lipid-free protein solution. A soluble preparation of lipid-free apoLp-I can be obtained after removal of GdmCl if a detergent that forms large micelles is present, e.g., Triton X-100or lysophosphatidylcholine. In the presence of detergents that form small micelles, e.g., cholate or deoxycholate, apoLp-I does not remain in solution, in the absence of GdmCI. In this regard it differs from apoB. The detergent, preferably lysophosphatidylcholine, can be replaced by phosphati-


377

dylcholine to regenerate a particle about the same size as lipophorin (Kawooya et al., 1989).These data have been interpreted to indicate that apoLp-I plays the major structural role in stabilizing lipophorin. At present, there is no indication as to the role of apoLp-11. 2 . Exchangeable Apolipoproteins Only one water-soluble, exchangeable, apolipoprotein has been found associated with lipophorin, apolipophorin 111 (apoLp-HI), which has a molecular mass of 18-20 kDa. Generally, apoLp-111 is only found in insects that use lipid to fuel flight (Table HI), although some exceptions have been noted, e.g., the flightless grasshopper Balytetth psolus and the house cricket Acheta domesticus. ApoLp-111 has not been found in cockroaches (order Dictyoptera), bees (order Hymenoptera), or flies (order Diptera). In the family Acrididae, order Orthoptera (grasshoppers and locusts), apoLp-111 is glycosylated, but in all other cases reported to date, TABLE 111 Distribution of Apolipophorin III among Insects Species L M W migraftma ~ Bartlrtlu psolllr Mchnopw dflerc-nhal Casfnmargur a/ncanur Arhrla d m s f t r u r Crtllw inttgcr

T h a w ncubngulu Acanlhoccphala grnnulosa LIIhoccm m c d w A W u r herbmi Ranalra quadndentaro Podtsur macuhvmfru Rhodntllr prolcru Tnnloma tn/tsfaru Dtrobrorhu gminafur Coltnu lrxana Manduca scxb H y b hwab Achrronlia a h o p o s Hyalophora rccrofna Bombyx mon

Family

Order

Comments

Acrididae Acrididae Acrididae Acrididae Gryllidae Gryllidae Coreidae Coreidae Belastomatidae Belastomatidae Nepidae Peniatomidae Reduviidae Reduviidae Cerambycidae Scarabaeidae Sphingidae Sphingidae Sphingidae Saturnidae Bombycidae

Orthoptera Orthoptera Orthoptera Orthoptera Orthoptera Orthoptera Hemiptera Hemipiera Hemiptera Hemiptera Hemiptera Hemiptera Hemiptera Hemiptera Coleoptera Coleopiera Lepidopiera Lepidopiera Lepidoptera Lepidoptera Lepidoptera

Glycorylatedcomplete sequence, crystal structure Glycorylaied: N-terminal sequence Glycorylaied; N-terminal sequence Glycosylaied Complete sequence

1 2 2 3 4

N-terminal sequence

6 7

-

N-terminal sequence

-

-

Ref.”

5

8 7 7 9 10 II

Complete squence Complete sequence

N-terminal sequence

-

-

7 7 12 7 13 14 15

a Key to references:( I ) Chino and Yazawa, 1986; Van der Horst et al., 199 1; Kanost et al., 1988; and Breiter et al., 1991; (2) Ryan etal., 199Od; (3) Haunerland etal., 1986; (4)Strobel et al., 1990; A. F. Smith and M. A. Wells, unpublished; (5) A. Hendrick, unpublished; (6) Wells et al., 1985; G . J. P. Fernando and M. A. Wells, unpublished; (7) A. F. Smith and M. A. Wells, unpublished;(8) M. K. Kanost and M. A. Wells, unpublished;(9) Haunerland et al., 1992; (10) Condim et al., 1989a; (11) J. L. Soulages, unpublished; (12) Kawooya et al., 1984; and Cole et al., 1987. (13) Surholt et al., 1992; (14) W. H. Telfer, unpublished; (15) Miura and Shimizu, 1989a,b.

378

JOSk L. SOULACES A N D MICHAEL A. WELLS

apoLp-I11 is not glycosylated. ApoLp-111 from L. mipatoria has been shown to contain N-linked oligosaccharide chains of the high-mannose type (Nagao and Chino, 1987). The function of apoLp-111is to facilitate transport of lipid from sites of storage in the fat body to sites of utilization in certain metabolic situations, e.g., flight. The triacylglycerolstores of the fat body are converted to DG, which leaves the fat body and becomes associated with preexisting HDLp in the hemolymph. In the process, HDLp is converted to LDLp and several molecules of apoLp-I11 become associated with LDLp. LDLp moves to the flight muscle, where the DG is hydrolyzed by a lipoprotein lipase. As the DG is removed, LDLp is converted back to HDLp and apoLp-111 dissociates. The HDLp and apoLp-I11 then cycle back to the fat body to carry more DG (see Section V for details). In contrast to the situation in vertebrates, in which most of the exchangeable apolipoproteins are bound to lipoproteins, most apoLp-I11 in insect hemolymph is free and represents a major component of lipophorin-free hemolymph (Wheeler and Goldsworthy, 1983; Kawooya et al., 1984; Van der Horst etal., 1984; Chino and Yazawa, 1986).Again in contrast to vertebrate apolipoproteins, apoLp-111 shows no tendency to self-associate in solution and this property has allowed extensive hydrodynamic characterization of the protein (Kawooya et al., 1986). ApoLp111binds to lipid surfaces with high affinity (Kawooya et al., 1986; Demel et al., 1992)and forms a stable complex with LDLp, which can be isolated by density gradient ultracentrifugation (Shapiro and Law, 1983) or by gel-permeation chromatography (Wheeler and Goldsworthy, 1983). However, apoLp-I11has a lower affinity for lipoprotein surfaces than do vertebrate apolipoproteins, because human apoA-I can displace apoLp111 from LDLp, whereas apoLp-I11 does not displace apolipoproteins from human HDL (Liu et al., 1991). The molecular structure of apoLp-111 from L. mipatoria has been determined at 2.5 8, (Breiter et al., 1991) (Fig. 1). The protein is composed of five long a helices connected by short loops and would be predicted to behave as a prolate ellipsoid in solution-a prediction in accord with hydrodynamic data suggesting that apoLp-I11 is a prolate ellipsoid with an axial ratio of about 3 (Kawooya et al., 1986).The helices are distinctly amphipathic with the hydrophilic side chains pointing toward the aqueous phase and the hydrophobic residues pointing into the interior of the protein. A similar structural motif, containing four a helices, was found in the N-terminal22-kDa receptor-binding domain of apoE (Wilson et al., 1991). Although the amphipathic a helix has been extensively discussed as a structural motif in exchangeable apolipoproteins (Segrest et al., 1992), these structural results are the first


379

FIG. 1. Molecular structure of apoLp-111. Ribbon drawing of the Locus& migratoria apoLp-I11 structure as determined by X-ray crystallography (Breiter el al., 1991). The helices are numbered beginning at the amino-terminal end of the protein. Reprinted with permission from Breiter el al. (1991).Copyright 1991 American Chemical Society.

demonstration that the amphipathic a helix is the predominant secondary structural element in such apolipoproteins. The complete amino acid sequences of apoLp-111 from M. sextu (Cole et ul., 1987), L. mzgrutoriu (Kanost et ul., 1988), Achetu domesticus (A. F. Smith and M. A. Wells, unpublished), and Derobruchus geminatus (A. F. Smith and M. A. Wells, unpublished) have been. determined from cDNA sequences and these sequences are aligned in Fig. 2;also shown in Fig. 2 are the location of the five a helices in the L. migrutoriu structure. The percent identities and similarities for pairwise comparisons between these sequences is shown in Table IV. A striking feature of these data is the low degree of sequence identity among these four apoLp-111 sequences, averaging only 23.3%. Note that the percentage identity between apoLps-111 from the fairly closely related species A. domesticus and L. mzgrutoriu is slightly higher than the percentage identity between apoLps111 from the very distantly related species L. migrutoriu and M. sextu. The overall low degree of sequence identity among the apoLps-111 is surprising considering the fact that all these apoLps-111can be found associated with LDLp, and it has been shown that M. sex& and L. migrutoriu apoLps111 are functionally equivalent in an in vitro system (Van der Horst et ul., 1988). Perhaps sequence identity is not as important for the function of apoLp-111 as is conservation of amino acids with similar physical properties, as might be suggested by the fact that percentage similarity between the four sequences is moderately high, averaging 43%. When the amino acid sequences in Fig. 2 are simplified by grouping the amino acids into

380

JOSt L. SOULACES AND MICHAEL A. WELLS

50 0

Map03 Dgapo3 Wapo3 hap03

0

0

DAGTTGADPN KVAEKS.QLQ DAPAGGNAFE DAA.GBVNIA

0

0

0

0

0

SLPEAAQRHF QNLTATIQNA ELAANAQQIV NNVTQTLQGN EMEKHAKEFQ KTFSEQF.NS EAVQQLNHTI VNAAHELEET helix 1

LPSQ...EEV LPDS...KKV LVNSKNTQDF LGLPTPDEAC

---------- I

I--------

0

RTQLQTHAQT VEVLNTNAQN NKALKDGSSS N.LLTEQANA

I--------------

100 0

Map03 Dgapo3 Mmapo3 hp03

0

0

0

PANNLQAAAT LANSVQSVVD VLQQLSAFSS FKTKIAEVTT helix 2

0

0

QFNEKAAELS KIKTEIKNNQ SLQGAISDAN SLKQEAEKEQ

----------- I

0

0

0

0

0

0

GDAQTAVRQA AQQLEQQVSN GEIDNvLlrQV SSKLSETAAE GKAKEALEQA RQNVEKTAEE GSVAEQLNAF ARNLNNSIHD I---- helix 3 ----I

0

LRQQF.PDGA LQKQLGPEGQ LRK.AHPDVE A......ATS 150

0

M a p 03 Dgapo3 MSap03 hp03

0

QAADKLKASIE KQAKEIKANLD KEANAFKDKLQ LNLQDQLNSLQ

I---------

0

SALAEV... KGLKDAVAQ AAVQTTVQE SALTNVGHQ helix 4

QEAEARRVQP VEKLTKAIEP SQKLAKEVAS WQDIATKTQA

--------- I

HADAVAESLK ETAKLKADLT NMEETNKKLA SAQEAWAPVQ

0

0 0

TAARTAVEQA NAAKTFLDQI PKIKQAYDDF SALQEAAEKT

I------------

170 0.

Adapo3 Dgapo3 Hsapo3 map03

0

TVITNQVQQS VEVSNNVQQQ VKBAEEVQKK KEAAANLQNS helix 5

0

VQQAANAH.. VRATLDEKH. LBEAATKQ.. IQSAVQKPAN

------- I

FIG. 2. Alignment of apoLp-111 sequences. The amino acid sequences of apo1.p-Ill from Ache& domesticus (Ad, house cricket), Derobrachus geminatus (Dg, palo verde beetle), Locusta migratoria (Lm), and Mandwa sexla (Ms) were aligned by the method of Feng and Doolittle (1987). The positions of the five helices in the L. migratoria apoLp-I11 are indicated. ( 0 )Conserved residues; (0)hydrophobic residues.

five groups, i.e., neutral, acids and their amides, basic, hydrophobic, and proline, the results (Fig. 3) obtained show a much higher degree of “identity”between the sequences. These results suggest that it is indeed the properties of the side chain of the amino acid that are conserved TABLE 1V Painuise Identities and Similarities for Insect Apolipophorin 111

Percentage identity Percentage similarity Achetu domesticus Derobrahus geminatus Locustu migratoria Mandwa sexta

A. domesticus

45.0 48.6 41.0

D. geminatzcs

L. migratoria

M . sexla

25.0

33.8 15.7

21.1 19.1 25.5

36.5 42.0

45.4

38 1


** Adapo3 Dgapo3 Msapo3 hap03

**

**

DIGGGGIDID GIIDIIDHHI DDIGIGIDDI HIIDHG.DID DIIIDIDDII DDIGDGIDGD DIPIGGDIID DIDHHIHDID HGIGDD1D.G DII-GHIDII DIIDDIDHGI IDIIHDIHDG I-------- helix 1 IIDDIDIIIG DIDDHIIDIG IIDHIDGIID HIHGDIHDDD IIDDIGIIGG GIDGIIGDID IHGHIIDIGG GIHDDIDHHD helix 2 I

-----------

** Adapo3 Dgapo3 Msapo3 Lmapo3

DIIDHIHIGI HDIHDIHIDI HDIDIIHDHI IDIDDDIDGI (---------

**

I...PGDDDI I...PDGHHI IIDGHDGDDI IGIPGPDDII

DGIIID...I DDIDIHHIDP DHGIHDIIID IDHIGHIIDP DIIIDGGIDD GDHIIHDIIG DGIIGDIGHD IDDIIGHGDI helix 4 I

---------

BGDIDGHIDG IDIIDGDIDD DHIIBDGGDG D-IIGDDIDI

I_---------___-

100

*******

GDIDGIIHDI IDDIDDDIGD GDIDDIIHDI GGHIGDGIID GHIHDIIDDI HDDIDBGIDD GGIIDDIDII IHDIDDGIHD I---helix 3 ----I

** *

* * * *.

* * * * * * * *** Adapo3 Dgapo3 Msapo3 hap03

* * ***

**

50

***.**

**

---------- I

* * * * * * * **** Adapo3 Dgapo3 Msapo3 hap03

** ***

*.*

*

HIDIIIDGIB DGIHIHIDIG DIDDGDHHII GIDDIIIPID

IBDDI-PDGI IDHDIGPDGD 1HH.IHPDID I......IGG 150

***.***.

GIIHGIIDDI DIIHGIIDDI PHIHDIIDDI GIIDDIIDHG

I------------

170

GIIGDDIDDG IDDIIDIH.. IDIGDDIDDD IHIGIDDHH. IHHIDDIDHH IHDIIGHD.. HDIIIDIDDG IDGIIDHPID helix 5 ------- I

FIG. 3. Alignment of simplified apoLp-111 sequences. The sequences in Fig. 2 were simplified by grouping amino acids with similar side-chain properties: G = G , S, T; D = D, E, N, Q; H = H, K, R; I = A, L, I, M, V, F, Y,W; P = P. The asterisks indicate residues with similar side-chain properties found in all four sequences. Species acronyms as in legend to Fig. 2.

among these proteins, not the exact amino acid. Helical wheel analysis suggests that four or five amphipathic a helices are present in each protein, again emphasizing the importance of this secondary structural element in apolipoproteins. When spread as a monolayer at the air-water interface at low surface pressure, the molecular area of M. sexta apoLp-I11 (3800 A*) is nearly twice as large as would be predicted for a prolate ellipsoid of axial ratio 3 lying on its side (Kawooya et al., 1986). As the monolayer of apoLp-I11 is compressed, it undergoes a phase transition in which the area occupied by the protein is reduced to 480 A2.Under these conditions, the area occupied by the protein is about that predicted if apoLp-I11 were binding to the surface via one end of the ellipsoid. Demel et al. (1992) have criticized the monolayer results of Kawooya et al. (1986), claiming that their monolayer results with L. rnzgrutoria and M.sex& apoLps-111, which showed a molecular area for apoLp-I11 of 2300 A, do not support the

382

JOSI? L. SOULAGES AND MICHAEL A. WELLS

suggestion that apoLp-111 occupies an area larger than its cross-sectional area at the air-water interface. However, Demel et al. (1992) chose to measure the area occupied by apoLp-111 at the collapse pressure of the film, whereas Kawooya et al. (1986) measured the area at low surface pressure. Although there is obvious disagreement about the behavior of apoLp-111 at the air-water interface, it is the behavior of apoLp-111 on lipid surfaces that is relevant to its binding to lipoproteins. The area occupied by apoLp-111on PL- or DG-coated polystyrene beads was determined to be 4300 A2(Kawooya et al., 1986). Analysis of the area occupied by apoLp-111 on lipophorins showed two states: in one, the area, 630 A', was about that expected if the protein bound via one end; in the other state, the area, 3500 A2,was about twice as large as predicted from the largest cross-sectional area of the protein (Wells et al., 1987). Demel et al. (1992) reported the binding of apoLp-111 to a DG monolayer and suggested that each apoLp-111binds to 92 molecules of DG. Unfortunately, interpretation of these results is ambiguous. On one hand, the data can be interpreted to suggest that apoLp-111 covers the area occupied by 92 DG molecules, which, with a surface area of 58 A2 for a DG molecule, means that apoLp-111 occupies a surface area of 5300 A2,an area even larger than that found by Kawooya et al. (1986) and Wells et al. (1987). On the other hand, the data can be interpreted to mean that apoLp-111 binds to the free air-water interface generated as the DG film is compressed by adsorption of apoLp-111, i.e., the protein forces its way between DG molecules. The data of Demel et al. (1992) show that the increase in surface pressure caused by adsorption of apoLp-111 would reduce the area per DG molecule by 7.3 A*, and for 92 DG molecules this would correspond to an area of air-water interface occupied b apoLp-111 (Malcolm, of 670 A'. If the minimal area per amino acid residue is 15 1973)) this would mean that only 45 residues, 28% of the protein, or about two helices, have actually penetrated to the air-water interface. This is a much lower value than that found for vertebrate apolipoproteins (Weinberg et al., 1992). These data suggest that, if apoLp-111 unfolds on the lipid surface, only a fraction of the protein actually penetrates between the polar head groups. Clearly, more data are needed to confirm this suggestion. Studies employing lipase treatment of apoLpIII-containing lipoproteins (Kawooya et al., 1991) and the characteristics of mixed apoLp-III-DG monolayers (Demel et al., 1992) suggest that apoLp-111 shows some specificity in binding to DG in the surface. These data are the basis for a model for the binding of apoLp-111 to lipophorin; the model suggests that apoLp-111binds to DG in the surface of lipophorin via one end and then spreads on the surface as depicted in Fig. 4 (Kawooya et al., 1986; Breiter et al., 1991). This model predicts an

i2


383

FIG.4. A model showing the proposed unfolding of apoLp-111on a lipoprotein surface, showing only the a-carbon backbone of the protein. It is proposed that the protein binds to the surface via one end and that helices 3 and 4, and helices 1,2, and 5 , then move relative to each other (filled arrows) around hinges in the loops connecting helices 4 and 5 , and 2 and 3 (unfilled arrows). The hydrophobic side chains, which are in the interior of the protein in the folded state, face the lipoprotein surface in the unfolded state.

initial binding via hydrophobic residues located in the loops between helices 1 and 2 and helices 3 and 4. These loops are relatively nonpolar and contain the only hydrophobic residues in the protein that are not buried-two leucines, which are conserved in all apoLp-111 sequences. According to the model, the protein spreads on the surface, without loss of its helical structure, via hinges located in the loops between helices 2 and 3 and helices 4 and 5. In essence, the hydrophobic interactions that hold the five helices together are replaced by energetically equivalent interactions that hold the apolipoprotein on the surface. T h e gene encoding M. sexta apoLp-I11 has been sequenced (Cole et al., 1990). It is composed of four exons; the first exon contains most of the signal sequence and the second exon contains the rest of the signal sequence, a Pro segment and some of the coding region for the mature protein; exons 3 and 4 contain the remainder of the coding region for the mature protein. This gene organization has some similarity to that of vertebrate apolipoprotein genes (Li el al., 1988), except that the coding region is divided among three exons in apoLp-I11 and only two exons in vertebrate apolipoproteins. What, if anything, can be said about the evolutionary relationship between apoLp-I11 and exchangeable vertebrate apolipoproteins? Both bind to lipoproteins and seem to have similar structural motifs. Whether apoLp-I11 is the ancestor of the vertebrate proteins can not be stated with

384

JOSE L. SOULAGES AND MICHAEL A. WELLS

any certainty. The evolutionary relation between insect apoLps-111, which are strictly orthologous proteins, is not easily demonstrated at the amino acid or nucleotide sequence level, therefore it is not surprising that insect apoLps-111 have little sequence homology to vertebrate apolipoproteins. If, as seems likely, amino acid sequence conservation is not strongly selected for in apolipoproteins, it may prove difficult to establish an evolutionary relationship between apoLp-111and the vertebrate apolipoproteins. It is also possible that apoLp-111 and the vertebrate apolipoproteins represent an example of convergent evolution and that there is not an ancestral relationship between apoLp-111 and the vertebrate apolipoproteins. In this regard, one must also consider the possibility that all insect apoLps-111 did not arise from a common ancestor, because some think that the ability to fly developed more than once among the insects. 111. SIZE,MOLECULAR WEIGHT,HETEROGENEITY, AND SHAPE OF LIPOPHORINS

Although lipoproteins are apparently easily purified by ultracentrifugation in a density gradient, it is actually impossible to obtain a chemically and physically homogeneous lipoprotein preparation. Even in a narrow density range a large polydispersity is expected and has been found when studied. The accurate determination of the molecular weight of a purified soluble protein can be achieved in several straightforward ways, e.g., gel electrophoresis, analytical ultracentrifugation, cDNA sequencing, and several other less commonly used methods. T h e shape of a pure protein in solution can be determined by spectroscopic techniques such as small-angle X-ray scattering (SAXRS) or can be inferred from its hydrodynamic properties. Some of the same techniques, e.g., sedimentation equilibrium, flotation rates, gel-filtration chromatography, electron microscopy, SAXRS, and gradient gel electrophoresis, can be used in determining the average molecular weight and shape of lipoproteins. Except for sedimentation equilibrium, most of the other experimental approaches are based on determination of an average radius from which, provided the density is known, the molecular weight can be estimated. However, as in any polydispersed system, the value of the average molecular weight determined depends on the method used. For example, electron microscopy gives a number-average radius from which one calculates a number-average molecular weight, whereas light scattering gives a Z-average radius from which one calculates a Z-average molecular weight. T h e ratio of the Z-average molecular weight to the number-average molecular weight can be significantly greater than one depending on the polydispersity of the system.

385


In order to compare the results of size and shape determinations for lipophorins and to discuss the differences observed using different methodologies, we have compiled most of the size-related data obtained for lipophorins in Table V. The data for some vertebrate lipoproteins are also included to permit a general discussion. Table V shows that there are large differences among the radii reported for lipoproteins and, at the same time, a systematic variation of the radius for any particular lipoTABLE V Comparison of Lipoprotein Sires Deduced Using Different Techniques" Volume ratios Lipoprotein

R,,

R,,,

Re,,

L. rnigratoria HDLpb P. amerzcana HDLp' P. cyntia H D L ~ ~ L. decemlineata HDLp' M . sexfa HDLP-LJ M. sexfa HDLp-A" M. sexta LDLph L. migratoria LDLp' HDL2 (human)' HDLs (human)' LDL, (human)h LDLy (porcine)h

59 59.5 59 67.3 85.2

90.3 90.3 77.4 98.1 -

78 80 65 82.5 80 57 85 67 128 147 50 44-49 38 39-41 110 107 -

-

50.5 40.8 86 81

60.7 54.1 137 129

R,,,

RdL sas:dc

em:dc em:gge

58.3 61.7 59.2 59.3 60 65 87.1 69.9

3.7 3.1 2.3 4.5 -

2.4 2.2 1.3 2.7 2.4 2.2 3.1 9.3

-

1.7 2.3 4.0

-

-

4.0

-

-

2.8 2.0

1 .o

0.8 2. I 2.3

" R,,, , Radius determined by analytical ultracentrifugation; Rsa,,radius determined from SAXRS; R,,, , radius determined by electron microscopy on negatively stained samples; R,, , radius determined by nondenaturing gradient electrophoresis; R&, radius obtained from the density and lipid content of the lipophorin, assuming one molecule of apoLp-1 and apoLp-I1 per particle. For those lipophorins containing apoLp-111, two, sixteen, and nine molecules of apoLp-I11 per particle were used, respectively, for M . sexta HDLp-A and LDLp, and L. migratoria LDLp. bRS,, (Katagiri el af., 1987); R,,,, (Nagao and Chino, 1991); R,, (Chino et al., 1981b). Re", = 85 was reported by Van Antwerpen et al. (1988). ' R,,, (Katagiri el a/., 1987);R,,, and Ra, (Chino ef al., 1981b). 'RSaJ(Katagiri etnl., 1987);R,,,, (Chino and Kitazawa, 1981). ' R,,, and R,,,, (Katagiri et af., 1991). /RaU (Pattnaik ef af., 1979);Re,,, (Ryan et al., 1990a; Kawooya et al., 1991); R,, (J. L. Soulages and M. A. Wells, unpublished). " R , , (Wells el af., 1987); Re,, (Ryan et a / . , 1990a; Kawooya ef al., 1991); R,,, 0. L. Soulages, and M. A. Wells, unpublished). R,, (Wells el af., 1987). ' R,,,, (Nagao and Chino, 1991);a similar value (140)was reported by Van Antwerpen et al. ( 1 988). Laggner and Muller (1978);Laggner (1982). R,,, (Jurgens el al., 1981);R,, (Jackson ef al., 1976).

'

386

JOSk L. SOULACES A N D MICHAEL A. WELLS

protein, depending on the technique used. With the exception of the human HDLs, it is clear that radii obtained by electron microscopy and SAXRS are the largest values, with SAXRS giving the largest dimensions. The large differences in the values for radii obtained with these two techniques, compared to those obtained by gradient gel electrophoresis and analytical ultracentrifugation, become even more pronounced if they are translated into volume or, what is equivalent, employed to estimate the molecular weight. It is noteworthy that in some cases up to a fourfold difference in the estimated molecular weight is observed when, for example, data from analytical ultracentrifugation are compared to those obtained with SAXRS or electron microscopy. Because such differences are sometimes overlooked and data obtained by different techniques are indiscriminately utilized to draw conclusions about lipoprotein structure, it seems appropriate to discuss the methodologies briefly. Although SAXRS has proved to be a powerful and accurate technique when applied to the structure of homogeneous, monodispersed particles in solution-or nonhomogeneous, but monodispersed, complexes of two components (Feigin and Sverdgun, 1987; Clatter and Kratky, 1982)-its application to the study of lipoprotein structure requires assumption of monodispersity. However, heterogeneity in terms of size has been found in most lipoprotein preparations analyzed by electron microscopy (Chapman, 1986). In addition, all lipoprotein fractions that have been analyzed by gradient gel electrophoresis show marked polydispersity characterized by populations that differ in size as well as in chemical composition (Nichols et al., 1983; Krauss and Burke, 1982; Cheung and Albers, 1984). Similar polydispersity has also been observed in lipohorin preparations from L. mzgratoria, using electron microscopy (Nagao and Chino, 1991), and M. sexta, using gradient gel electrophoresis (J. L. Soulages and M. A. Wells, unpublished). Thus, polydispersity is likely to be a common characteristic of insect lipoproteins. The extreme sensitivity of SAXRS to sample heterogeneity makes obtaining accurate data about the geometry, size, and internal organization of a lipoprotein a difficult, if not impossible, task. The natural heterogeneity of lipophorins and the presence of discreet aggregates, which are not uncommon, might account for the large radii estimated by SAXRS. In addition, the lack of a correction to infinite density contrast for the data on lipophorins might affect the calculated value of the radius of gyration (Laggner and Muller, 1978).Extensive treatments of the theoretical and experimental aspects of the study of lipoproteins and other particles by SAXRS have been published (Laggner, 1982; Feigin and Sverdgun, 1987). Electron microscopy is the other technique that is widely employed to


387

assess the size and geometry of lipoprotein particles. Among the methods employed, negative staining of the lipoprotein preparation has been most often utilized, but some studies involving cryofixation (freeze-fracture and etching) have been reported (Forte and Nordhausen, 1986). In principle, electron microscopy can give a number-average radii, and also provide a direct measure of sample heterogeneity. Unfortunately, the analysis of negatively stained samples by electron microscopy requires observation of samples in an environment in which the hydration shell is removed. This seems to be one of the sources of artifacts in samples containing lipids. The most important artifact, because it cannot be readily corrected, is a possible flattening of the lipoprotein particles, which increases as the lipid content of the lipoprotein increases (Forte and Nordhausen, 1986). It is probably for this reason that the sizes of lipoprotein particles are often overestimated by electron microscopy. The potentially more powerful freeze-fracture-etching technique has many technical problems and associated artifacts, e. g., sample heterogeneity makes conclusions about particle morphology essentially impossible (Forte and Nordhausen, 1986; Aggerbeck and Gulik-Krzywicki, 1986). Low-temperature electron microscopy is the most recently developed technique to observe frozen hydrated biological macromolecules (Adrian et al., 1984). This technique has the advantage that the sample solution is not fractured and the molecules can be observed in their hydrated states. It has been suggested that this technique might be useful in determining lipoprotein size (Aggerbeck and Gulik-Krzywicki, 1986), but no reports have yet appeared. In preliminary studies of adult M. sextu HDLp in a hydrated frozen sample, the average diameter obtained was very close to that obtained by analytical ultracentrifugation or gradient gel electrophoresis, and smaller than that obtained in negatively stained samples (R. Van Antwerpen, unpublished). The close agreement for the values of radii observed by analytical ultracentrifugation, gradient gel electrophoresis, and, in some cases, gel-filtration chromatography is consistent with a consensus basic structure for lipophorin consisting of one molecule each of apoLp-I and apoLp-11, and a variable lipid content. In some insect stages apoLp-111 binds to this basic structure in different amounts depending on the content of diacylglycerol. Assuming that this is the correct description of lipophorin structure, then from the molecular weights of the apolipoproteins and their relative amounts, and the lipid content and density of the lipoprotein, a compositional molecular weight can be calculated. As can be observed in Table V, the radii of the lipophorin particles calculated in this way show very close agreement to those determined by analytical ultracentrifugation and gradient gel electrophoresis. This con-

388

JOSB

L.

SOULACES AND MICHAEL A. WELLS

sistency gives further support to the previous comments about the necessity for analyzing carefully data obtained by electron microscopy and SAXRS. For example, in a recent report using electron microscopic observations of LDLp and HDLp of adult L. mzgratoria, it was concluded that the LDLp particles are the result of intermolecular fusion of HDLp particles (Nagao and Chino, 1991). This conclusion might be correct. However, because this conclusion was based on the comparison of sizes obtained by electron microscopy, which for HDLp gives a molecular weight twice the value obtained by analytical ultracentrifugation, it is reasonable to suggest that the large differences between the reported sizes of HDLp and LDLp are the effect of some artifact, such as flattening, which would markedly increase with the increase in lipid content. An independent measure of the size of LDLp will be required before 'the existence of fusion can be evaluated. It is generally agreed that lipoproteins have a spherical or sphericallike structure. However, there are not unambiguous methods to confirm this assumption. Two suitable techniques might be SAXRS and electron microscopy; however, for the reasons previously discussed, the accuracy and power of these techniques are obscured by the presence of artifacts. Electron microscopy of phosphotungstate negatively stained samples has been used to suggest that changes in the lipid content of lipophorin were accompanied by modifications in the morphology of the lipoprotein particles (Ryan et al., 1992). If this is the case, the assumption of spherical particles would not be appropriate for those lipophorins with a low lipid content. However, the features reported by Ryan et al. (1992) were not observed in uranyl acetate negatively stained samples of lipophorins (Ryan et al., 1990a; Kawooya et al., 1991). Technical artifacts, such as positive staining, might be present in either of these preparations, which would obscure interpretation of the data. Further work on this important point is needed. OF LIPIDS AND PROTEINS I N LIPOPHORINS IV. ORGANIZATION

T h e elucidation of the structure of any lipoprotein, i.e., the organization of the protein and lipid components, is a challenging problem, because it relies on several techniques that give only partial and approximate information. Structural models should be consistent with experimental data that characterize the physiological role and the physicochemical properties of the lipoprotein and its components. Considerable effort has been expended to fit experimental data to structural models of mammalian lipoproteins (Zilversmit, 1965; Sata el al., 1972; Schneider et al., 1973; Havel, 1975; Verdery and Nichols, 1975; Shen et al., 1977;


389

Edelstein et al., 1979; Oeswein and Chun, 1981). The concept of a lipoprotein core of nonpolar lipids surrounded by an outer shell of phospholipids and proteins is a common theme in all these studies. T h e existence of the lipid core is supported by the properties of lipids, SAXRS studies, and is apparent for large lipoproteins with high lipid contents. On the basis of SAXRS studies this concept has also been applied to lipoproteins with low lipid contents, such as human HDL (Scanu, 1972; Laggner, 1982). In addition to the lipid core, experimental support for a surface localization of phospholipids and cholesterol has been obtained from NMR studies (Henderson et al., 1975; Yeagle et al., 1978; Lund-Katz and Phillips, 1986). Among the problems involved in constructing a general model for vertebrate lipoprotein structure, some of the more important are the heterogeneity in size and apolipoprotein composition and the small number of examples of lipoproteins containing similar apolipoprotein composition, which can be used to validate the model. Thus, in most cases, data as elementary as the stoichiometry of the apolipoproteins cannot be included in the models (Shen el al., 1977) or, if stoichiometry data are included, only a partial fit of the data to the model is observed (Edelstein et al., 1979). One of the most intriguing features of lipophorin composition is the large variation in lipid content and composition that can be accommodated without modifications in the apolipoprotein composition of the particles (Table I). This feature makes lipophorin a good system in which to analyze the structure of lipoproteins and the physicochemical factors that govern their structure and properties. In addition to the previously discussed data on the size and shape of lipophorins, several studies on other aspects of lipophorin structure have been performed and need to be discussed before describing models for lipophorin structure.

A . Apolipoproteim From the average circular dichroism (CD) spectra of locust and cockroach lipohorins a high content of P-sheet secondary structure was estimated for apoLp-I and apoLp-I1 (Kashiwasaki and Ikai, 1985). A similar result was obtained by Kawooya et al. (1989) with M.sexta lipophorin, where a secondary structure composed of 58% /3 structure, 30% a helix, and 12% random coil was estimated. As described earlier, most data are consistent with a surface localization of apoLp-I, whereas the localization of apoLp-I1 has been described as sequestered. However, apoLp-I1 can not be completely “buried” because water-soluble crosslinking reagents react with both apolipoproteins (Kashiwasaki and Ikai,

390

JOSk L. SOULACES AND MICHAEL A. WELLS

1985) and it is possible to generate monoclonal antibodies against apoLpI1 when immunization is carried out with intact lipophorin (Schulz et al., 1987). Although a portion of both apoLp-I and apoLp-I1 must be exposed to the aqueous media, the low solubility of both apolipoproteins in water (Pattnaik et al., 1979; Kawooya et al., 1989) and the stability of the lipid-apoLp-I interaction in GdmCl indicate a strong lipid-protein interaction in the native lipophorin particle. In this regard, the ordering effect of apoLp-I and apoLp-I1 on the lipid components of lipophorin, as measured by anisotropy of fluorescence (Soulages et ad., 1988a). which is unaffected after extensive proteolytic cleavage of apoLp-I (Rimoldi et al., 1991), supports the idea of a lipid-embedded localization, for at least a portion of the apolipoproteins.

B . Location of Phospholipids "P NMR studies on locust lipophorin indicate that a large proportion of the PLs reside on the surface of lipophorin (Katagiri et al., 1985). In addition to this study, the susceptibility of lipophorin phospholipids to hydrolysis by phospholipase A2 is consistent with a surface localization of PLs (Katagiri et al., 1985; Kawooya et al., 1991). Although the amphipathic nature of PLS makes this a logical conclusion, it has to be pointed out that the use of hydrolytic enzymes to demonstrate the location of any component would be valid only if a rearrangement of the lipid components in lipophorin does not occur on the time scale of the experiment. Because this condition is never met, a cautious interpretation of such data is necessary. C . Location of Hydrocarbons Experimental evidence for a core localization of hydrocarbon was obtained from 'C NMR, calorimetry, and SAXRS studies (Katagiri et al., 1985, 1987). Thus, from the similarity in thermotropic behavior of pure hydrocarbons isolated from locust lipophorin, and the same hydrocarbons in the native lipoprotein, it was concluded that hydrocarbons are partially segregated from the other lipophorin components forming a hydrocarbon-rich cluster. The presence of an internal region of low electronic density was observed by SAXRS in lipophorins containing hydrocarbons, suggesting that hydrocarbons form part of an inner lipid core, which is not exposed to the aqueous environment. D . Location of Diacylglycerols Another major component of lipophorin is DG. Because the hydroxyl group of DG gives some polarity to an otherwise highly nonpolar mole-


39 1

cule, and because of the active metabolism of DG, a partial exposure of lipophorin DG to the aqueous media has been proposed (Pattnaik et al., 1979; Soulages et al., 1988a; Shapiro et al., 1988; Van der Horst, 1990). However, it was also proposed that, due to the possible destabilizing effect of DG on the highly curved surface of lipophorin, a surface localization of DG is unlikely in those lipophorins without apoLp-111 (Soulages and Brenner, 1991). Kawooya et al., (1991) concluded that DG stabilizes the lipophorin particle: after removal of PLs from M . sexta LDLp, the integrity of the particle was maintained. However, because LDLp contains apoLP-111,it is also possible that apoLp-I11 stabilized the particle. When DG-labeled LDLp, produced in vivo, was treated with lipase, radioactive DG was hydrolyzed more rapidly than unlabeled DG (Kawooya et al., 1991). These data were interpreted as showing the existence of at least two, nonequilibrating, DG pools in lipophorin. However, the same result would be obtained if there were two or more species of lipophorin in the sample that had been labeled to different DG specific activities during in vivo labeling-once again lipophorin heterogeneity clouds interpretation of data. In order to study the effect of DG content on the structure and properties of lipophorin, we have recently developed a method that allows the specific modification of the DG content of lipoproteins employing sn-l,2dioctanoyl glycerol (diC8)(Soulagesand Wells, 1994a).We observed that the loading of lipophorin with diC8 promotes the binding of apoLp-111, indicating that the main requirement for apoLp-111binding to lipophorin is an increase in the content of surface DG. DiCs loading occurred in the presence or absence of apoLp-111, but, in the absence of apoLp-111, the DG-loaded particles are not stable and spontaneously aggregate. A sharp decrease in the degree of order of the lipid phase of the lipoproteins was observed by anisotropy of fluorescence of diphenyhexatriene as the content of diC8 was increased. A comparison of the lipid order of artifically loaded lipophorins, in the presence and absence of apoLp-111, natural lipophorins which differ in their DG content, and mammalian lipoproteins, clearly showed that DG has a extraordinary perturbing effect on the lipid order of lipoproteins. The result of this recent work provides further evidence for the destabilizing role of DG and we have suggested that the lipid disordering effect of DG plays a prominent role on the lipophorin surface in such a manner as to permit the binding of the weak lipid-binding apoLp-111 to the lipoprotein surface. E. Lipophorin Models There have been only a few attempts to fit composition and structure data for lipophorin to a model. Based on SAXRS studies of lipophorin

392

JOSr L. SOULACES AND MICHAEL A. WELLS

from L . mipatoria and Periplaneta americana (Katagiri et al., 1987), and Leptanotursa decemlineata (Katagiri et al., 1991), the distribution of PLs, apolipoproteins, hydrocarbons, and DG in each of the lipophorins was deduced. Employing electron density and composition data, the authors fit the distance distribution functions to a three-layer centrosymmetrical lipophorin model. This model proposes that lipophorins are composed of three radially symmetrical layers-an outer layer that contains apoLpI and PL, a middle layer that contains apoLp-I1 and DG, and a core that contains hydrocarbon. The exact composition of each layer depends on the lipophorin under consideration. However, in both studies, the authors used spherical radii that were large enough to accommodate from two to four times the number of electrons or molecules that constitute the lipophorin particles. Thus, the dimensions of the particles are clearly inconsistent with the molecular weight and radii of lipophorins (Table V), and, therefore, an accurate picture of the lipophorin particle cannot be inferred from these studies. Another approach has been to use lipophorin composition and size to develop a model. Two models were developed based on the paradigm of Shen et al. (1977) for mammalian lipoproteins. Pattnaik et al. (1979), on the basis of the compositional data for larval M. sexta lipophorin, concluded that the surface layer of lipophorin contains substantial quantities of DG in addition to PL. Shapiro et al. (1988), employing the same paradigm to study the organization of L. mipatoria and M . sexta HDLps and LDLps, also concluded that substantial amounts of DG would be in the surface layer, particularly in LDLp, where it might interact with apoLp-I1I. The last model for lipophorin was constructed on the basis of the density-composition data for 12 lipophorins that do not contain apoLp111 (Soulages and Brenner, 1991). The composition data were fit quantitatively to a lipophorin model wherein the particles were assumed to be spherical and to contain a hydrophobic lipid core composed of hydrocarbon, DG, and TG and a surface layer composed of apolipoproteins, PL, sterol, and small amounts of DG. This study showed good correlations between the proposed structure, the composition of lipophorins, and the space-filling requirements of the lipoprotein components. Thus, in spite of the apparently random variations in lipid content and composition of lipophorins, a model emerged that is consistent with the presumed properties of the lipids and apolipoproteins, and the physiological role of lipophorin. Among the findings of that study are the following conclusions: (1) as the size of the lipophorin increases, the content of PL increases, suggesting a fundamental role for PL in defining the size of the lipophorin surface layer (Fig. 5) and hence the volume of the particle; (2) in agreement with the known perturbing effect of DG on natural and

LIPOPHORIN LlPlD TRANSPORT IN INSECTS

75

393

125 150 175 200 moles PL/mole Lipophorin

100

FIG.5. Dependence of lipophorin surface area on the mole of phospholipid per particle. The data are from 15 lipophorins without apoLp-111, and calculations were done according to Soulages and Brenner (1991). The surface area of the lipophorin was related to the surface area occupied by apoprotein and lipid by the equation 4 P RZ - WCHOL ACHOL) = NPL(APL +

~

D

G

+) APROI..

where R2 is the radius of the particle, N is the number of molecules of lipid per particle, A is the molecular area of the component, and a is the number of DG molecules per molecule of PL on the surface. CHOL, Cholesterol; PL, phospholipid; DG, diacylglycerol; PROT, apolipoprotein. Assuming a molecular area of 20 A' for cholesterol, a Y intercept value, which is surface area occupied by the apolipoprotein, of 32,000 f 2000 A' is obtained. Thus, depending on the size of the lipophorin, apoLp-I and apoLp-I1 occupy from 62 to 82% of the surface area. From the slope of the plot (73.5 f 15.8 A'), and assuming that the molecular area of a PL may have a value between 75 and 96 A', a value for a near 0 is calculated, which suggests a virtual absence of DG on the lipophorin surface.

artificial membranes (Dawson et al., 1983, 1984; Das and Rand, 1984, 1986; Hamilton el al., 1991), only small amounts of DG seem to reside in the surface layer; (3) the apolipoproteins occupy a large proportion of the lipophorin surface, from 60 to 80%; (4) in conjunction with PLs, the apolipoproteins form an outer shell about 20 A thick; ( 5 ) the area occupied per apolipoprotein amino acid at the lipoprotein surface would be about 10 A2,which would be consistent with a high content o f p structure; and (6) a small proportion (5-10%) of the apolipoprotein could be embedded in the inner lipid core.

V. METABOLISM The key to understanding lipoprotein metabolism in insects was the discovery that lipophorin functions as a reusable shuttle (reviewed by Chino, 1985). Thus, lipophorin can be described as a basic apolipoprotein-phospholipid complex, which can carry a variety of lipids in

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JOSk L. SOULACES AND MICHAEL A. W E U S

its core and deliver these lipids to various tissues without internalization of the lipophorin particle and without destruction of the basic apolipoprotein-phospholipid complex. Electron microscopicevidence in support of this hypothesis has been presented for locust flight muscle (Van Antwerpen et al., 1988) and the midgut of the dragonfly, Aeshna cyanea, (Bauerfeind and Komnick, 1992a): lipophorin does not enter tissues to which it delivers lipid or from which it derives lipid. Although it was shown that lipophorin entered the fat body of A. cyanea, it was proposed that this intracellular lipophorin may be aged material in the process of degradation (Bauerfeind and Komnick, 1992b). Other hallmarks of lipophorin metabolism are its ability to deliver specific lipids to specific tissues (Chino and Kitazawa, 1981) and the fact that the same basic apolipoprotein-phospholipid complex changes its metabolic roles during the life of the insect. A. Biosynthesis

Lipophorin is biosynthesized only in the fat body, with no synthesis occurring in the midgut (Prasad et al., 1986b; Bauerfeind and Komnick, 1992b).In A. cyanea, immunocytochemical data suggest that the secretion of lipophorin follows a normal pathway for a secreted protein: lipophorin was found in the endoplasmic reticulum, Golgi bodies, and secretory vesicles (Bauerfeind and Komnick, 1992b). 1 . Larvae

In M. sexta larvae, a nascent lipophorin consisting of apoLp-I, apolp11, and phospholipid is assembled in the fat body and secreted into hemolymph as a very high-density lipophorin (Prasad et al., 1986b). A

similar VHDLp can be isolated from hemolymph of larvae fed a fat-free diet. When larvae raised on a fat-free diet are fed a bolus of triolein, and lipophorin is isolated 6 hr later, a lipophorin of normal density can be isolated in which diolein is the predominant DG (Prasad et al., 1986b). These data suggest that the nascent lipophorin secreted from the fat body is converted into mature lipophorin by picking up DG from the midgut. Bauerfeind and Komnick (1992a) arrived at a similar conclusion based on immunocytochemical studies on A. cyanea. FernandoWarnakulasuriya et al. (1988) showed in M. sexta larvae that lipophorin biosynthesis was independent of the amount of lipid in the diet: on a fat-free diet larvae produce a lipophorin with a very low lipid content; on a high-fat diet larvae produce a lipophorin with a higher than normal lipid content. However, in neither case was the lipophorin concentration in hemolymph different from that in normal larvae. It should be noted


395

that insects raised on a fat-free diet deposit significant amounts of TG in their fat bodies, presumably derived from dietary carbohydrate (Fernando-Warnakulasuriya et al., 1988). Therefore, the lack of a DGrich lipophorin in the hemolymph of such insects is not due to lack of a DG precursor in the fat body, but must be due to lack of a DG precursor in the diet. These results show two distinct features of lipophorin biosynthesis during the larval stage. First, the nascent lipophorin produced in the fat body by de nouo synthesis is an apolipoprotein-phospholipid complex that derives its transported lipids from the midgut. Second, lipophorin biosynthesis is not coupled to fat intake, as is the case with vertebrates. These processes are illustrated in Fig. 6 and fit observations made on lipid storage in larvae. Thus, it has been shown that more than 70% of the fatty acids in the diet are stored as TG in the larval fat body (Tsuchida and Wells, 1988). Although the fat body can convert carbohydrates to fatty

FIG.6. Model for delivery of dietary lipid from the midgut to the fat body. Dietary lipid (triacylglycerol,TG) is hydrolyzed to fatty acid (FA) in the lumen of the midgut, absorbed into midgut epithelial cells, and used to synthesize diacylglycerol (DG). The DG is picked up by lipophorin via a mechanism that does not involve internalization of the lipophorin. Two cases are shown. In one case, newly synthesized (nascent) lipophorin (nLp), which is secreted from the fat body, picks up DG from the midgut and is converted to a DG-loaded HDLp. The DG-loaded HDLp then moves to the fat body, where it delivers the DG without internalization and is converted to a DG-unloaded HDLp. In the second case, the DGunloaded HDLp travels to the midgut, where it picks up DG and is converted to a DGloaded HDLp.

396


acids (Horie and Nakasone, 1971), it would be inefficient for the fat body to produce and secrete a mature lipophorin, if most of the DG is going to be returned to the fat body. By synthesizing a nascent lipophorin particle, the larva produces the most efficient vehicle for transport of DG from midgut to fat body. The only other larval system that has been studied is the southwestern corn borer, Diatraea grandiosella (Venkatesh et al., 1987; Bergman and Chippendale, 1989). In both diapausing and feeding-stage larvae, it was shown that the fat body incubated in vitro released a mature, i.e., lipid-loaded, lipophorin. However, the data suggest a very low rate of lipophorin biosynthesis, and this may indicate that the fat body was not producing lipophorin at its in vivo rate, which could complicate interpretation of the results. High rates of protein synthesis and secretion are difficult to maintain in in vitro fat body incubations unless the system is kept well oxygenated (Noriega and Wells, 1992).

2. Pupae At the end of the larval stage, M . sexta enters the pupal stage, during which adult metamorphosis occurs. During the pupal stage there is no lipophorin biosynthesis (Prasad et al., 1987); however, lipophorin continues to play a central role in lipid transport, but now lipid is transported from fat body to developing adult tissues (Tsuchida and Wells, 1988). Therefore, lipophorin biosynthesized in the larval stage is important for lipid transport in both the larval and pupal stages: this may be the reason that lipophorin biosynthesis is uncoupled from fat intake. If the amount of lipophorin made during the larval period depended on the amount of fat in the diet, it is likely that pupae might not have sufficient lipophorin to support adult development. 3. Adults Lipophorin biosynthesis in adult insects has been studied in the house fly, Musca domestica (Capurro and d e Bianchi, 1990b), in L. migratoria (Weers et al., 1992), and in M . sexta (S. V. Prasad and M. A. Wells, unpublished). In each of these cases in vitro incubations of fat body resulted in the release of a lipophorin whose density and lipid composition closely resembled that of mature lipophorin. An important difference between larvae and adults is the rate of lipophorin biosynthesis. For example, in M . sexta larvae, which are rapidly growing, the amount of lipophorin per animal can increase u p to 10-fold in 3 days (Prasad et al., 1987), requiring a prodigious rate of lipophorin synthesis. In adults, lipophorin synthesis need only replace that which is lost from the hemolymph due to turnover: the half-life of lipophorin in adult L. migratm-ia is


397

several days (Downer and Chino, 1985). In the adult, this low rate of lipophorin biosynthesis would not significantly deplete the fat body of lipid if a mature lipophorin were produced, whereas, in larvae, during the peak period of lipophorin synthesis, the fat body lipid stores would be signficantly depleted, if a mature lipophorin were produced. It is possible that lipophorin can be made either as a nascent particle o r as a mature particle, depending on the developmental stage and/or insect under investigation. A thorough understanding of the mechanism of lipophorin biosynthesis during insect development will require additional work. T o finish this duscussion on lipophorin biosynthesis we will mention studies on the origins of PLs, hydrocarbons, sterols, and carotenoids. It has been reported that in adult M.sexta and Rhodnius prolixzts PL can be transferred from fat body to lipophorin (Van Heusden et al., 1991; CorrCa et al., 1992). This transfer of PL is independent of de novo synthesis of lipophorin; however, the mechanism by which it occurs is unknown. Hydrocarbon transport by lipophorin has been studied only in P . americana. Katase and Chino (1982) have shown, in in vitro incubations, that a fat body rich in oenocytes, one type of cell in the hemolymph, which is the major site of hydrocarbon biosynthesis (Diehl, 1975), can release labeled hydrocarbon to lipophorin. It was also shown, using in vitro incubations, that the labeled hydrocarbon in lipophorin was delivered to the epidermis, the normal site of hydrocarbon deposition in insects. T h e sterols and carotenoids that are present in lipophorin must arise from the diet, because insects cannot biosynthesize either sterols or carotenoids de nova Chino and Gilbert (197 1) have shown that sterol can be transferred from the midgut to lipophorin, and the same is most likely true for carotenoids. The mechanism by which hydrocarbons, sterols, and carotenoids are transferred from either oenocytes or midgut epithelial cells to lipophorin is unknown.

B . Physiologacal Roles of Lipophorin In this section we describe what is known about lipophorin metabolism. First we discuss the lipid transfer particle, which may play an important role in transferring lipids to and from lipophorin, then we describe the different roles of lipophorin in lipid delivery. 1 . Lipid Transfer Particle

The lipid transfer particle (LTP) is a very high-density lipoprotein having a molecular mass greater than 650 kDa. LTP contains about 15% lipid and three apolipoproteins with M, = 320,000,85,000, and 55,000. First discovered in M. sexta (Ryan et al., 1986a), LTP has been purified

398

JOSd L. SOULAGES AND MICHAEL

A. WELLS

from M. sexta (Ryan et al., 1986b, 1988a), L. mipatoria (Hirayama and Chino, 1990), and M . domestica (Capurro and de Bianchi, 1990a) hemolymph. LTP catalyzes lipid exchange and net transfer between different lipophorins (Ryan et al., 1986a,b, 1988a,b; Hirayama and Chino, 1990) and between lipophorin and human HDL (Ryan et al., 1990b). A model of LTP has been developed based on electron microscopy (Ryan et al., 1990~).LTP differs from vertebrate lipid transfer proteins (Tall, 1986) in both its size and ability to catalyze net lipid transfer between lipoproteins. It has been shown that LTP can catalyze carrier-mediated transfer of DG between lipoproteins (Blacklock et al., 1992). It was speculated that LTP may function in vivo to catalyze lipid transfer between cells and lipophorin (Ryan et al., 1988a),but only one study has attempted to elucidate the physiological role of LTP (Van Heusden and Law, 1989). In this study, using the adult M. sextu fat body in an in vitro lipid transfer system, it was shown that antibodies against LTP inhibited transfer of DG from fat body to lipophorin, but had no effect on the transfer of DG from lipophorin to fat body. Thus, at present the only known physiological function of LTP is to facilitate transfer of DG from fat body to lipophorin and little is known about the mechanism by which LTP catalyzes either lipid exchange or transfer, except that the transferred lipid moves through the lipid pool of LTP (Ryan et al., 1988a). 2. Lipid Transport from Midgut to Fat Body

Fatty acids released from dietary lipids by midgut lipases (Bollade et al., 1970; Weintraub and Tietz, 1973, 1978; Hoffman and Downer, 1979a; Rimoldi et al.,1985; Tsuchida and Wells, 1988) are absorbed into midgut epithelial cells and transformed, by as yet uncharacterized reactions, to DG. Midgut of M . sexta contains fatty acid-binding proteins, which may play a role in fatty acid absorption by the epithelial cells (Smith et al., 1992). When labeled fatty acids, either as free fatty acids or TG, are fed to insects or placed in midgut sacs, which are then incubated in vitro in a lipophorin-containing medium, virtually all of the labeled fatty acid that leaves the midgut is found in lipophorin as DG (Chino et al., 1981b; Chino and Kitazawa, 1981; Rimoldi et al., 1985; Tsuchida and Wells, 1988; Bauerfeind and Komnick, 1992a). For example, using fatty acidlabeled triolein, Tsuchida and Wells (1988) showed in M . sextu larvae that during a 4-hr period nearly 90% of the fed fatty acid was absorbed, and, of that absorbed, more than 70% was found in the fat body as TG. In the hemolymph, more than 95% of the labeled fatty acids was present as DG, and all the hemolymph DG was present in lipophorin. When midgut sacs, containing labeled triolein, were incubated in vitro, it was shown that essentially no labeled lipid left the midgut unless lipophorin was present


399

in the incubation medium, and then only labeled DG was found in the media. When lipophorin, containing labeled DG, was injected into M. sexta larvae, the DG disappeared from the hemolymph with a half-life of about 50 min, and after 4 hr about 60% of the injected label was found in fat body as T G (Tsuchida and Wells, 1988).When DG-labeled lipophorin was incubated in vitro with fat body, more than 95% of the label was taken up by fat body within 4 hr; the half-life of labeled DG in the incubation medium was about 60 min, a value comparable to the in vzvo measurement. During the incubation, after 95% of the DG had entered the fat body, there was no detectable loss of lipophorin apolipoproteins from the incubation medium. However, the density of the lipophorin in the incubation medium increased substantially, consistent with the loss of DG. These results are compatible with the hypothesis that lipophorin functions as a reusable, noninternalized shuttle. A few reports have appeared describing lipophorin receptors. Evidence for high-affinity lipophorin binding was demonstrated in adult L. migratoria flight muscle (Hayakawa, 1987;Van Antwerpen et al., 1990) and fat body (Van Antwerpen et al., 1989).A lipophorin receptor from larval M. sexta fat body has been purified and characterized (Tsuchida and Wells, 1990).The receptor has a molecular mass of 120 kDa, requires Ca2+ for activity, and is inhibited by suramin. In these properties the lipophorin receptor is similar to the human LDL receptor, although the insect receptor does not bind human LDL. T h e purified receptor has a single, high-affinity binding site for lipophorin. The fat body lipophorin receptor shows an 8-fold higher affinity for DG-rich lipophorin than for DG-poor lipophorin. In preliminary experiments, a lipophorin receptor from the midgut has been partially characterized (K. Tsuchida and M. A. Wells, unpublished). The midgut receptor differs from the fat body receptor in molecular mass (140kDa) and the fact that it does not require metal ions for activity. The midgut receptor has 12-fold higher affinity for DG-poor lipophorin than for DG-rich lipophorin. The mechanism by which these two receptors distinguish between a DG-rich and a DG-poor lipophorin is unknown. It should be remembered that both types of lipophorin have identical structural apolipoproteins (apoLp-I and apoLp-11), and differ only in lipid content. It may be that the two types of lipophorin have different surface areas, caused by different lipid core volumes, and that these different surface areas may result in different conformations of apoLp-I and apoLp-11, which are recognized by the two types of receptors. In this regard, it has been reported that apoLp-I and apoLp-I1 are more susceptible to proteolysis in DG-poor lipophorin than in DG-rich lipophorin (Ryan et al., 1992).It has also been reported

400


that monoclonal antibodies specific for apoLp-I1 inhibit DG uptake by the fat body in vitro (Hiraoka and Hayakawa, 1990) and that lipophorin binds fat body proteins (Schulz et al., 1991), which might suggest that some epitope on apoLp-I1 is recognized by the receptor. The different affinities of the midgut and fat body receptors for DGrich and DG-poor lipophorins suggest a model that explains why DG is transported from the midgut to the fat body (Fig. 7). This model proposes that a DG-poor lipophorin, e.g., the nascent lipophorin secreted from the fat body or a recently DG-depleted lipophorin, binds to the midgut receptor, which facilitates transfer of DG from the midgut to the DG-poor lipophorin, producing a DG-rich lipophorin. The DG-rich lipophorin dissociates from the midgut receptor and binds to the fat body receptor, which facilitates transfer of DG from the DG-rich lipophorin to fat body, producing a DG-poor lipophorin. The DG-poor lipophorin then dissociates from the fat body receptor and cycles back to the midgut receptor.

Receptor-I Fat Body

Receptor-It

1

\

J

Midgut DG '.I

FIG.7. Proposed role of midgut and fat b o d y receptors in lipid transport. The midgut receptor specifically recognizes diacylglycerol (DG)-poor lipophorins. When lipophorin is bound to the midgut receptor, DG is transferred from the midgut to lipophorin, producing a DG-rich lipophorin. The DG-rich lipophorin dissociates from the midgut receptor and binds to the fat body receptor, which specifically recognizes a DG-rich lipophorin. When lipophorin is bound to the fat body receptor, DG is transferred from lipophorin to the fat body, producing a DG-poor lipophorin. The DG-poor lipophorin dissociates from the fat body receptor and cycles back to the midgut. It is proposed that the differences in DG content between the two types of lipophorin translate into some structural differences between the apolipoproteins on the surfaces of the two types of lipophorins, which are recognized by the two receptors.


40 1

3. Lipid Transportfrom Fat Body to Flight Muscle This is the most intensively studied aspect of lipophorin metabolism. Lipid, mobilized from the fat body, is the primary substrate used by insects to fuel long-term flight (Beenakkers et al., 1984).The mobilization of lipid from the fat body is regulated by adipokinetic hormone (AKH), a peptide hormone released from the corpus cardiaca (Orchard, 1987). AKH can also activate glycogen phosphorylase in the fat body (Beenakkers et al., 1984): whether AKH activates lipolysis of glycogenolysis depends on the developmental stage of the insect. The details of the signal transduction pathways that are involved in AKH-dependent activation of lipolysis in the fat body are unknown, but the available information suggests that the effect of AKH on the fat body has many parallels with the effect of glucagon on vertebrate adipose tissue and liver: (1) AKH has been shown to elevate CAMPlevels in the fat body (Spencer and Candy, 1976; Gade and Holwerda, 1976; Gade and Beenakkers, 1977; Asher et al., 1984; Wang et al., 1990), (2) cyclic nucleotides have been shown to stimulate protein kinase (Pines and Applebaum, 1977) and lipase (Pines et al., 1981) activities in fat body homogenates, and (3) evidence has been presented to show that AKH treatment leads to phosphorylation and activation of a lipase in the fat body (E. L. Arrese and M. A. Wells, unpublished observations). It is also known that the activating effect of AKH is dependent on extracellular Ca2+ (Lum and Chino, 1990; Wang et al., 1990; Van Marrewijk et al., 1991) and that a Ca2+ionophore can mimic the effects of AKH on lipid mobilization both in vivo and in vitro (Lum and Chino, 1990; Wang et al., 1990). Developing these preliminary observations into a complete description of the regulatory pathways is clearly a fertile area for future research. The pathway for formation of DG from TG in the fat body is unknown. The lipolytic activity in fat body homogenates from L. migratoria (Tietz and Weintraub, 1978) and M . sextu (Arrese and Wells, 1992) converts T G primarily to free fatty acids (FFAs), whereas in the desert locust Schistocerca gregaria (Spencer and Candy, 1976) and the cockroach P. americana (Hoffman and Downer, 1979b)the end products were DG and FFA. Microsomes from the L. migratoria fat body can acylate 2-MG to produce DG (Tietz et al., 1975). It has been shown that the DG released from the fat body has the sn-1,2 configuration (Lok and Van der Horst, 1980; Tietz and Weintraub, 1980), therefore either a pathway involving de novo synthesis of DG via phosphatidic acid or the stereospecific hydrolysis of TG could be involved. Several studies have characterized the AKH-induced formation of LDLp, both in vivo and in vztro (Mwangi and Goldsworthy, 1977b, 1981;

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JOSr L. SOULAGES AND MICHAEL A. WELLS

Van der Horst et al., 1979, 1981, 1984, 1987; Wheeler and Goldsworthy, 1983; Shapiro and Law, 1983; Shapiro et al., 1984; Kawooya et al., 1984; Van Heusden et al., 1984, 1987a; Goldsworthy et al., 1985; Chino et al., 1986, 1989; Wells et al., 1987; Surholt et al., 1988; Strobel et al., 1990; Nagao and Chino, 1991). Although there are some minor differences in details, all of these studies support the suggestion that the mechanism of AKH-induced formation of LDLp is the same in all insects. Indeed, it has been shown that mixtures of components, i.e., fat body, HDLp, and apoLp-111, from different insects will form normal LDLp (Van der Horst et al., 1988; Ziegler et al., 1988; Van Heusden and Law, 1989; Chino et al., 1992). Our current understanding of the formation and metabolism of LDLp during flight is summarized in Fig. 8. AKH induces DG formation from T G stores and the DG leaves the fat body, by an unknown mechanism, but with the assistance of LTP (Van Heusden and Law, 1989). and is delivered to HDLp. ApoLp-111 assists in the uptake of DG by HDLp by

FIG. 8. Role of lipophorin in DG delivery to flight muscle. Adipokinetic hormone (AKH) is released from the corpus cardiacum and binds to the fat body, where it cause production of CAMP and entry of Ca2+. These second messengers activate lipolysis of triacylglycerol(TG) and production of diacylglycerol (DG). The DG leaves the fat body with the assistance of a lipid transfer particle (LTP) and is taken up by HDLp. The capacity of HDLp to carry DG is increased by binding of apoLp-Ill to the surface. Ultimately, LDLp is formed and moves to the flight muscle, where a lipoprotein lipase hydrolyzes the DG to produce fatty acid (FA) and regenerate HDLp and apoLp-111. The FA enters the flight muscle, where it is oxidized to produce the ATP required to power flight. HDLp and apoLp-111 circulate back to the fat body to complete the cycle.


403

binding to incipient hydrophobic patches on the lipoprotein surface caused by expansion of the lipoprotein as a result of the increased DG content of the core. Additional insight into LDLp formation has resulted from studies on insects that do not form LDLp in response to AKH, even though all of the hemolymph components necessary for formation of LDLp are present. Larval L. mipatoria produce only small amounts of LDLp when injected with AKH (Van der Horst et al., 1987). Furthermore, in vitro incubation of larval fat body in adult hemolymph did not lead to production of LDLp, although larval fat body has ample lipid to support formation of LDLp. On the other hand larval fat body does respond to AKH by elevating CAMP(Gade and Beenakkers, 1977) and glycogen phosphorylase is activated by AKH (Van Marrewijk et al., 1984). The flightless grasshopper Barytettix:psolus also does not form LDLp following injection of AKH (Ziegler et al., 1988). Yet, the hemolymph contains HDLp and apoLp-111, both of which showed normal function when tested in another grasshopper, Melanopus differentials, which does form LDLp. AKH stimulates glycogen phosphorylase in B. psolus fat body. In M . sexta larvae, AKH stimulates glycogen phosphorylase, but does not cause LDLp formation, although larval hemolymph contains HDLp and apoLp-111 (Ziegler et al., 1990). Because AKH causes activation of glycogen phosphorylase in all of these cases, it is unlikely that the AKH receptor is missing or that AKH does not cause elevation of cellular CAMP.The lack of a lipolytic response to AKH may involve the AKH-sensitive lipase: either the enzyme is missing or it cannot be activated. The cockroach P. americana does not form LDLp in response to AKH (Chino et al., 1992). In this case the hemolymph lacks apoLp-111, which would limit the amount of DG that could be carried by lipophorin. A particularly interesting result came from comparison of the solitary and gregarious phases of L. mipatoria (Chino et al., 1992). Solitary-phase locusts do not fly; gregarious-phase locusts are strong fliers and this is the phase that has been extensively studied. The transformation from solitary to gregarious phase occurs when the locusts are crowded, as occurs when food supplies become diminished. Solitary-phase locusts do not form LDLp when injected with AKH, although they have HDLp and apoLp-111 in their hemolymph. Solitary-phase locusts have less fat body (15%) and a lower TG content per milligram of tissue (5%) compared to gregarious-phase locusts. One possibility is that solitary-phase locusts cannot form LDLp because of insufficient TG stores. However, it is also known that juvenile hormone titers are higher in solitary-phase locusts than in gregarious-phase locusts and this may be an important factor in determining the metabolic state of the insect. It is also possible that the solitary locust fat body lacks AKH receptors.

404


How does LDLp deliver fatty acids to the flight muscle? Flight muscle from L. migratoria (Wheeler et al., 1984,1986; Wheeler and Goldsworthy, 1985; Van Heusden et al., 1986, 1987a,b) and M. sexta (Van Heusden, 1993) has been shown to contain a membrane-bound lipoprotein lipase that hydrolyzes DG to FFA. T h e lipase has higher activity against LDLp than against HDLp, which would account for its selective hydrolysis of DG in LDLp. Once produced, the fatty acids are presumed to diffuse into the flight muscle cell, where they are oxidized. In this regard it should be noted that fatty acid-binding proteins have been identified in the flight muscle of L. mipatoria (Haunerland and Chisholm, 1990) and M. sexta (M. C. Pape and M. C. Van Heusden, unpublished). These fatty acidbinding proteins could play a role in uptake of fatty acids by flight muscle. Considering the high degree of analogy between lipid mobilization in insects and vertebrates, why d o insects use DG, instead of free fatty acids, to transport fatty acids from fat body to flight muscle? Insects are quite capable of metabolizing hemolymph FFA (Stanley-Samuelson et al., 1988): the half-life of hemolymph FFA in adult Triatoma infestam (Soulages et al., 1988b) and M. sexta (Soulages and Wells, 1994b) is only 2-3 min. In both insects FFAs are carried by lipophorin and no evidence could be found for an albumin-like molecule, which transports FFA in vertebrates. In adult M. sexta 75% of the FFA in hemolymph is reesterified into DG, TG, and PL in the fat body and the rest is oxidized. The fact that hemolymph FFAs are taken into the fat body so rapidly, coupled with the fact that insects have an open circulatory system, suggests that FFA would be a poor form in which to transport fatty acids to flight muscle, because most of the fatty acid released from fat body would be rapidly taken back into the fat body and reesterified, whereas with DG, the delivery of fatty acids to flight muscle is very efficient because flight muscle contains lipoprotein lipase. 4 . Lipid DeliveT to Developing Oocyte Eggs from Hyalophora cecropia (Telfer, 1960), Samina Cynthia (Chino et al., 1977), M . sexta (Kawooya et al., 1988), and R. p-olixus (Gondim et al., 1989b) have been shown to contain a very high-density lipophorin, VHDLp-E, which is derived from HDLp-A in the hemolymph by a receptor-mediated process (Kawooya et al., 1988; Telfer and Pan, 1988; Kulakosky and Telfer, 1990); this represents the only known exception to the generalization that lipophorin delivers its lipids to tissues without internalization. The conversion of HDLp-A to VHDLp-E involves removal of DG, which is catalyzed by a lipoprotein lipase found in the yolk body of the egg (Van Antwerpen and Law, 1992). However, in spite of the presence of lipophorin in the egg, 90% of the lipid in the egg is


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delivered by a noninternalizing mechanism, involving LDLp as the lipid carrier (Kawooya and Law, 1988; Telfer et al., 1991). At present it is not known whether a lipase, either the one described above or another, is involved in removal of DG from LDLp, a mechanism analogous to the situation in flight muscle, or whether some other mechanism exists to deliver DG to the ovary. 5. Changes in Lipophorin Metabolism during Development When M . sexta larvae reach the end of the larval period, they enter the prepupal stage, during which they stop eating, void their midgut, and, if in the wild, burrow in the ground or leaf litter to pupate. During the 3- to 4-day prepupal period, striking changes in lipophorin metabolism occur. Within 48 hr, the larval lipophorin (HDLp-L) is first converted to a higher density form, which is relatively depleted of DG (HDLp-Wz), and then to a lower density form (HDLp-W1), which is relatively enriched in DG (Prasad et al., 1986a). During this period apolipoprotein synthesis ceases (Prasad et al., 1987). The conversion of HDLp-L to HDLp-W2 seems to reflect continued delivery of DG to fat body, or other tissues, in the absence of feeding. The conversion of HDLp-W2 to HDLp-W1 may reflect a switch in metabolism in the fat body: it changes from a lipidstoring tissue to a lipid-mobilizingtissue. This hypothesis is supported by in vitro and in viuo experimental data demonstrating that fat bodies from prepupae did not take up labeled DG from lipophorin (Tsuchida and Wells, 1988). Little is known about lipophorin metabolism during the pupal period, except that there is a lipophorin whose density and lipid composition (HDLp-P)differ from those of lipophorins isolated from other life stages (Prasad et al., 1986a).However, it is known that HDLp-P is derived from HDLp-L by alterations in lipid composition,because no new lipophorin is produced by the fat body in the prepupal or pupal stages. It is likely that lipophorin metabolism is controlled by the changing hormonal milieu during the prepupal and pupal periods, but nothing is known of the details. VI. METABOLICIMPLICATIONS OF LIPOPHORIN STRUCTURE

The lipid composition of a lipophorin depends on the steady-state relationship between thermodynamic and kinetic factors that dictate the movement of lipids between tissues and lipophorin. The thermodynamic factors are the metabolic state of the tissue and the stability of the lipophorin, and the lipoprotein will incorporate or release lipid depending on the magnitude and sign of the difference between these two thermo-

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dynamic contributions. However, the steady-state composition will also depend on the rate at which the lipid composition of lipophorin can be changed. For example, if a thermodynamically stable lipophorin takes up DG from the fat body, the DG content of the particle will be displaced from equilibrium. The particle can reestablish equilibrium by delivering the excess DG to some tissue, but the rate at which that occurs will depend on the rate of lipid transfer. It would not be possible to fit the widely varying lipid compositions of lipophorins without apoLp-111 to a single structural model, unless, in vivo, the lipid composition and content of lipophorins were determined primarily by the physiochemical requirements for particle stability. This suggests that lipophorins transport lipids in such a manner as to maintain a near-equilibrium composition, which could be accomplished by a rapid rate of lipid transfer or by transporting only a small amount of lipid in any one cycle, e.g., between the fat body and the tissue. In this regard, the data of Tsuchida and Wells (1988) on the half-life of DG in feeding M. sexta larvae, coupled with the content of lipophorin per animal (Prasad et al., 1987)and the composition of HDLpL (Prasad et al., 1986a), can be used to calculate that the rate of DG delivery from lipophorin is 20 nmol/min. This amounts to only 1% of the DG in HDLp-L delivered per minute, a rate that should not cause the lipid composition of HDLp-L to deviate very far from equilibrium. Assuming that the in vivo composition of lipophorin is not far from equilibrium, we can use the results of the composition-structure model to make some predictions about the role of PL and DG in regulating lipophorin metabolism. The PL content of lipophorin controls its core volume and hence its lipid-carrying capacity. In other words, the PL content determines an optimal value for the core volume that corresponds to the maximum stability of the particle. An increase in core volume above the optimal value would destabilize the particle, but stability can be reestablished by getting rid of DG, or some other core lipid; a decrease in core volume below the optimal value would also destabilize the particle, but stability can be reestablished by taking up DG, or some other core lipid. Although a lipophorin particle with a reduced core volume is less stable, there seem to be no physiological mechanisms to rapidly adjust the PL content in order to establish a new optimal core volume, because, in vivo, lipophorin PL turnover is slow. In fact, a slow adjustment of the PL content of lipophorin would be necessary if the optimal core volume is an important determinant in directing the movement of DG, or other core lipids, into or out of lipophorin. Even though the adjustment of the PL content of lipophorins is slow, there must be some mechanism to accomplish such an adjustment, as is most clearly demonstrated by the changes in PL content in lipophorin that accompany


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the changes in core lipid composition and content during the transition from the larval to pupal stage in M. sexta, a time when de nouo synthesis of lipophorin has stopped. Of particular importance to lipophorin metabolism are the properties of DG. The strongly perturbing effect of DG on phospholipid bilayers has been attributed to its relatively small polar head group in comparison to the volume occupied by the fatty acyl chains. This geometric disproportion, accentuated by the high radius of curvature of lipophorin particles, would promote the exposure of the hydrocarbon chains of DG and neighboring lipids to water, which would destabilize the particle due to the hydrophobic effect. In this context the high concentration of PE found in most of the lipophorins may also be important. It is well known that, although PC adopts bilayer structures, PE is a promoter of nonlamellar lipid structures, in a manner similar to that displayed by DG. Thus, it can be speculated that the presence of substantial amounts of PE on the surface of lipophorins would enhance the hypothesized destabilizing effects of DG. The combined destabilizing effects of DG and PE on the lipophorin surface would create a situation in which transfer of lipid from the particle would be energetically favorable, while at the same time, the disorder on the surface could lower the activation energy, and hence increase the rate, of processes involved in lipid transfer. For example, a small increase in the concentration of DG on the lipophorin surface, produced after DG loading in the midgut, would cause a strong hydration of the surface. This increased hydration could promote activation of lipolytic enzymes, or activation of lipid transfer proteins and/or spontaneous release of the DG for the surface. Any or all of these activities could play a role in reestablishing the stability of the lipophorin surface. Such considerations might account for the efficient lipid transport system of insects, which does not use a lipid transfer mechanism involving uptake and/or degradation of the whole lipoprotein particle. In LDLp, the relative excess of DG, compared to PL, must result in accumulation of DG on the surface of the particle. The surface disorder caused by the presence of DG and the increase in surface free energy caused by the hydrophobic effect could promote insertion of apoLp-111 into the lipoprotein surface. The increased hydration and disorder in the surface of LDLp might also be responsible for the fact that LDLp is a better substrate for muscle lipoprotein lipase than is HDLp, in which the surface content of DG is small. The accumulation of DG in LDLp suggests that the fat body is producing DG more rapidly than it can be consumed by the flight muscle. This probably occurs because HDLp is a poor substrate for lipoprotein lipase and does not deliver fatty acids to flight muscle very effectively. Therefore, when AKH stimulates produc-

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tion and release of DG from the fat body, the DG can not be delivered to flight muscles, but instead accumulates in HDLp, causing the production of LDLp. The net result is to produce, in the hemolymph and without requiring de nova synthesis, a lipophorin particle that carries much more DG and at the same time is an excellent substrate for the flight muscle lipoprotein lipase. A totally different rationale has to be applied to the transport of hydrocarbons. These extremely hydrophobic compounds seem to reside in the interior of the lipophorin particles. A mechanism involving uptake and degradation of the lipoprotein might be possible for the transport of hydrocarbons to the epidermal cells. This type of mechanism might be important in certain stages of insect development, when the lipoprotein could deliver amino acids and other lipid components necessary for the construction of the cuticle. A similar process may also exist for the delivery of carotenoids and sterols. A poorly understood aspect of lipophorin metabolism is tissue-specific delivery of lipids. At present mechanistic details are lacking, but some properties of the system are apparent. Lipid seems to be transferred from lipophorin to tissue only in those cases in which the tissue can carry out some additional reaction with the lipid, e.g., in the fat body DG is converted to TG, and in the epidermis hydrocarbon is secreted onto the outer surface of cuticle. In other cases there may be intracellular lipidbinding proteins that, by binding the lipid, drive lipid uptake into the tissue. These or other processes would drive the equilibrium of lipid transfer in favor of the tissue, irrespective of the mechanism by which lipid delivery occurs. VII. CONCLUDING REMARKS AND FUTURE DIRECTIONS Major progress has been achieved in understanding the structure and function of apoLp-111, including the amino acid sequence of the protein from four species and the molecular structure of one of them. These results are of significance not only for insect biochemistry but for an understanding of the function of apolipoproteins in general. T h e natural variability in the amino acid sequence of apoLps-111, coupled with the possibility for selective site-directed mutagenesis and the ability of the protein to exist in water-soluble or lipid-bound states, make this apolipoprotein an excellent model with which to analyze the details of lipidprotein interactions. Lipid metabolism in insects represents a system with many unique characteristics, which although complex, still appears simpler than that of vertebrates. Insects offer an especially attractive system in which to study


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the metabolism of DG, a lipid effector of importance in vertebrates. Because DG plays such a central role in lipid metabolism in insects, it is expected that the enzymes regulating DG metabolism might be more readily studied in insects. Although such studies are of importance for understanding lipid metabolism in insects, they may also provide information of relevance to the regulation of DG metabolism in vertebrates. With regard to lipoprotein metabolism in insects there remain several fundamental questions which need to be addressed. By what mechanism does DG move between lipophorin and tissues? What is the in uivo role of LTP? What are the structure and function of lipophorin receptors? How are apoLp-I and apoLp-I1 organized on the surface of lipophorin? What is the pathway for synthesis and secretion of lipophorin? What is the mechanism of LDLp formation? What are the details of how LDLp delivers DG to tissues? How does lipophorin achieve tissue-specific delivery of lipid? In addition, future studies should be directed toward refinement of the structural model of lipophorin. Such studies should take further advantage of the diversity of insects by obtaining more compositional data, and, perhaps more importantly, should involve collecting more, and better, physical data. As stated before, the fact that so many compositionally divergent lipophorins exist, which use the same basic apolipoprotein-phospholipid matrix, offers a unique opportunity to define the structure of a lipoprotein in considerable detail. ACKNO w LEDGMENTS We thank Drs. Estela Arrese, Carolina Barillas-Mury, Don Frohlich, John Law, Ann Peterson, Alan Smith, Rik Van Antwerpen, and Randi Van Heusden for helpful comments and criticisms during preparation of this review. Unpublished work from the authors’ laboratory was supported by NIH Grant HL 391 16.

REFERENCES Adrian, M., Dubochet, J.. Lepault, J., and McDowall, A. W. (1984). Nature (London)308, 32-34. Aggerbeck, L. P., and Culik-Kryzwicki, T. (1986). In “Methods in Enzymology”(J. Segrest and J. Albers, eds.), Vol. 128, pp. 457-472. Academic Press, Orlando, FL. Asher, C., Moshitzky, P., Ramachandran, J.. and Applebaum, S. W. (1984). Gen. Comp. Endocrinol. 55, 167-173. Bauerfeind, R., and Komnick, H. (1992a).J. Insect Physiol. 38, 147-160. Bauerfeind, R., and Komnick, H. (1992b).j. Insect Physiol. 38, 185-198. Beenakkers, A M. T. (1973). Insect Biochem. 3,303-308. Beenakkers, A. M. T., Van der Horst, D. J., and Van Marrewijk, W. J. A. (1984). Insect Biochm. 14,243-260. Beenakkers, A. M. T., Van der Horst, D. J., and Van Marrewijk, W. J. A. (1985). Prog. L i e Res. 44, 19-67.

410

JOSk L. SOULACES AND MICHAEL A. WELLS

Beenakkers, A. M. T., Chino, H., and Law, J. H. (1988).Insect Biochem. 18, 1-2. Bergman, D. K., and Chippendale, G. M. (1989).Insect Biochem. 19, 361-365. Blacklock, B. J., Smillie, M.. and Ryan, R. 0. (1992).J. Biol. Chem. 267, 14033-14037. Blomquist, G. J., Nelson, D. R., and de Renobales, M. (1987).Arch. Insect Biochem. Physiol. 6, 227-265. Bollade, D., Paris, R., and Moulins, M. (1970).J. Insect Physiol. 16,45-53. Breiter, D. R., Kanost, M. K., Benning, M. M., Wesenberg, G., Law, J. H., Wells, M. A., Rayment, I., and Holden, H. M. (1991).Biochemistty 30,603-608. Capurro, M. de L. (1988). Doctoral Thesis, University of Sao Paulo, Brazil. Capurro, M. de L., and de Bianchi, A. G. (1990a).Comp. Biochem. Physiol. B 97B, 649-653. Capurro, M. de L., and de Bianchi, A. G. (1990b).Comp. Biochem. Physiol. B 97B, 655-659. Chapman, J. M. (1986).In “Methods in Enzymology” (J. Segrest and J. Albers, eds.), Vol. 128, pp. 70-144. Academic Press, Orlando, FL. Cheung, M. C., and Albers, J. J. (1984).J. Biol. Chem. 259, 12201- 12209. Chino, H. (1985).I n “Comprehensive Insect Physiology, Biochemistry, and Pharmacology” (G. A. Kerkut and L. I. Gilbert, eds.), Vol. 10, pp 115-136. Pergamon, Oxford. Chino, H., and Downer, R. G. H. (1982).Adu. Biophys. 15,67-92. Chino, H., and Gilbert, L. 1. (1971).Insect B i o c h . 1,337-347. Chino, H., and Kitazawa, K. (1981).J. LiPidRes. 22, 1042-1052. Chino, H., and Yazawa, M. (1986).J. Lipd Res. 27,377-385. Chino, H., Downer, R. G. H., and Takahashi, K. (1977).Biochim. Biophys. Actu487,508-516. Chino, H., Downer, R. G. H., Wyatt, G. R., and Gilbert, L. I. (1981a).InsectBiochem. 11,491. Chino, H., Katase, H.,Downer, R. G. H.,andTakahashi, K. (1981b).J. LipdRes. 22,7-15. Chino, H., Downer, R. G. H., and Takahashi, K. (1986).J. LipidRes. 27,21-29. Chino, H., Kiyomoto, Y., and Takahashi, K. (1989).J. Lipid Res. 30,57 1-578. Chino, H., Lum, P. Y., Nagao, E., and Hiraoka, T. (1992).J. Comp. Physiol. B 162, 101-106. Cole, K. D., Fernando-Warnakulasuriya, G. J. P., Boguski, M. S., Freeman, M.. Gordon, J. I., Clark, W. A., Law, J. H., and Wells, M. A. (1987).J. Biol. Chem. 262,11794-1 1800. Cole, K. D., Smith, A. F., and Wells, M. A. (1990).Insect Biochem. 20, 381-388. C o d a , G., Gonsim, K. C., and Masuda, H. (1992).Arch. InsectBiochem.Physiol. 19,133-144. Das, S., and Rand, R. P. (1984).Biochem. Biophys. Res. Commmun. 124,491-496. Das, S., and Rand, R. P. (1986).Biochemistty 26,2882-2889. Dawson, R. M. C., Hemington, N. L., and Irvine, R. F. (1983).Biochem. Biophys. Res. Commun. 117, 196-201. Dawson, R. M. C., Irvine, R. F., Bray, J., and Quinn, P. J. (1984). Biochem. Biophys. Res. Commun. 125,836-842. de Bianchi A. G., Capurro, M. de L., and Marinotti, 0.(1987).Arch. Insectdiochem.Physiol. 6, 39-48. de Kort, C. A. D., and Koopmanschap, A. B. (1987).Arch. InsectBiochem.Physiol. 5,255-269. Demel, R. A., Van Doorn, J. M., and Van der Horst, D. J. (1992).Biochim. Biophys. Acta 1124, 151- 158. Diehl, P. A. (1975).J. Insecf Physiol. 21, 1237-1246. Dillwith,J. W., Lenz, C.J., and Chippendale, G. M. (1986).J. Comp. Physiol. B 156,783-789. Downer, R. G. H., and Chino, H. (1985).Insect Biochem. 15,627-630. Edelstein, C., KCzdy, F. J., Scanu, A. M., and Shen, B. (1979).J. LipldRes. 20, 143-153. Feigin, L. A., and Sverdgun, D. I. (1987).In “Structure Analysis by Small Angle X-ray and Neutron Scattering” (G. W. Taylor, ed.). Plenum, New York and London. Feng, D. F., and Doolittle, R. F. (1987).J. Mol. Evol. 24,351-360. Fernando-Warnakulasuriya, G. J. P., and Wells, M. A. (1988).Arch. Insect Biochem. Physiol. 8, 243-248.


41 1

Fernando-Warnakulasuriya, G. J. P., Tsuchida, K., and Wells, M. A. 1988).Insect Btochem.

18,211-214. Fichera, L., and Brenner, R. R. (1982).Acta Physiol. Latinoam. 32,21-29. Forte, M. T., and Nordhausen, R. W. (1986).In “Methods in Enzymology” (J. Segrest and J. Albers, eds.), Vol. 128,pp. 442-457. Academic Press, Orlando, FL. Gade, G., and Beenakkers, A. M. T. (1977).Gen. Comp. Endocrinol. 32,481-487. Gade, G., and Holwerda, D. A. (1976).Insect Biochem. 6,535-540. Clatter, O., and Kratky, 0. (1982).“Small Angle X-ray Scattering.” Academic Press, London. Goldsworthy, G. J., Miles, C. M., and Wheeler, C. H. (1985).Physiol. Entomol. 10, 151-164. Gondim, K. C., Oliveira, P. L., Coelho, H. S. L., and Matsuda, H. (1989a).InsectBiochem. 19,

153- 161. Gondim, K. C., Oliveira, P. L., and Matsuda, H. (1989b).J.Insect Physiol. 35, 19-27. Gonzalez, M. S., Soulages,J. L., and Brenner, R. R. (1991).Insect B i o c h . 21,679-687. Hamilton, J. A., Bhamidipati, S. P., Kodali, D. R., and Small, D.M. (1991).J. Biol. C h . 266,

1177-1 186. Haunerland, N. H., and Chisholm, J. M. (1990).Biochem. Biophys. Actn 1047,233-238. Haunerland, N. H., Ryan, R. O., Law, J. H., and Bowers, W.S. (1986).Insect Biochem. 5,

797-802. Haunerland, N. H., Ortego, F., Strausfeld, C. M., and Bowers, W. S . (1992).Arch. Insect Biochem. Physiol. 20,49-59. Havel, R. J. (1975).Adu. Exp. Med. Biol. 63,37-59. Hayakawa, Y. (1987).Biochim. Biophys. A c h 919,58-63. Henderson, T. O.,Kruski, A. W., Davis, L. G., Glonek, T., and Scanu, A. M. (1975). Biochemistry 14, 1915-1920. Hiraoka, T., and Hayakawa, Y. (1990).Insect Biochem. 20,793-799. Hirayama, Y.,and Chino, H. (199O).J.Lipid Res. 31,793-799. Hoffman, A. G. D., and Downer, R. G. H. (1979a).C a n . J . Zool. 54, 1165-1171. Hoffman, A. G. D., and Downer, R. G. H. (1979b).Lipid 14,893-899. Horie, Y.,and Nakasone, S. (1971).J.Insect Physiol. 17, 1441-1450. Jackson, R. L., Taunton, 0. D., Segura, R., Gallagher, J. G., Hoff, J. H., and Gotto, A. M. (1976).Comp. Biochem. Physiol. B 53B,245-259. Jurgens, G., Knipping, G. M. J.. Zipper, P., Kayushina, R., Degovivics, G., and Laggner, P. (1981).Biochemistry 20,3231-3237. Justum, A. R., and Goldsworthy, G. J. (1976).J.Insect Physiof. 22,243-249. Kanost, M. K., Boguski, M. S., Freeman, M., Gordon, J. I., Wyatt, G. R., and Wells, M. A. (1988).J.Biol. Chem. 263, 10568-10573. Kashiwazaki, Y., and Ikai, A. (1985).Arch. Biochem. Biophys. 237, 160-169. Katagiri, C.,and de Kort, S. (1991).Comp. Biochem. Physiol. B 100B, 149-152. Katagiri, C., Kimura, J., and Murase, N. (1985).J.Biol. Chem. 260, 13490-13495. Katagiri, C., Sato, M., and Tanaka, N. (1987).J.Biol.C h . 262, 15857-15861. Katagiri, C., Sato, M., de Kort, S., and Katsube, Y.(1991).Biochemistty 30,9675-9681. Katase, H., and Chino, H. (1982).Biochim. Biophys. Acta 710,341-348. Kawooya,J . K.,and Law, J. H. (1988).J.Biol. C h . 263,8748-8753. Kawooya, J. K., Keim, P. S., Ryan, R. 0..Shapiro, J. P., Samaraweera, P., and Law, J. H. (1984).J.Biol. Chem. 259, 10733-10737. Kawooya, J. K.. Meredith, S. C., Wells, M. A., Kkzdy, F. J., and Law, J. H.(1986).J.Biol. Chem. 261,13588-13591. Kawooya,J. K., Osir, E. 0..and Law, J. H. (1988).J. Biol. Chem. 263,8740-8747. Kawooya, J. K., Wells, M. A., and Law, ,I. H. (1989).Biochemistry 28,6658-6667.

412

JOSf L. SOULAGES AND MICHAEL A. WELLS

Kawooya,J. K., Van der Horst, D. J., Van Heusden, M. C., Brigot, B. L. J., Van Antwerpen, R., and Law, J. H. (1991)./.LiPidRes. 32, 1781-1788. Krauss, R. M., and Burke, D. J. (1982)./.LipUi Res. 23,97-104. Kulakosky, P. C., and Telfer, W. H. (1990).Arch. Insect Biochem. Physiol. 14,269-285. Kuthiala, A., and Chippendale, M. G. (1989).Arch. Insect Biochem. Physiol. 12, 123-131. Laggner, P. (1982).In “Small Angle X-ray Scattering” (0. Clatter and 0. Kratky, eds.), pp. 329-359. Academic Press, London. Laggner, P., and Muller, K. W. (1978).Q. Rm. Biophys. 11,371-425. Law, J. H., and Wells, M. A. (1989)./.Biol. Chem. 264,16335-16338. Law, J. H., Ribeiro, J. M. C., and Wells, M. A. (1992).Annu. Rev. Biochem. 61,87-111. Li, W.-H., Tanimura, M., Luo, C.-C., Datta, S., and Chan, L. (1988)./. Lipid Res. 29,

245-27 I. Liu, H., Malhotra, V., and Ryan, R. 0. (1991).Biochem. Biophys. Res. Commun. 179, 734-

740. Lok, C. M., and Van der Horst, D. J. (1980).Biochim. Biophys. act^ 618,80-87. Lum, P. Y., and Chino, H. (199O).J.Lipid Res. 31,2039-2044. Lund-Katz, S.. and Phillips, H. C. (1986).Biochemistry 25, 1562-1568. Malcolm, B. R. (1973).Prog. Surf.Membr. Sci. 7, 183-229. Miura, K., and Shimizu, I. (1989a).Comp. Biochem. Physiol. B 89B,95-103. Miura, K., and Shimizu, I. (1989b). C m p . Biochem. Physiol. B 94B, 197-204. Mwangi, R. W., and Goldsworthy, G. J. (1977a).J.Insect Physiol. 23, 1275-1280. Mwangi, R. W., and Goldsworthy, G. J. (1977b).J.Comp. Physiol. 114, 177-190. Mwangi, R. W., and Goldsworthy, G. J. (1981)./.Insect Physiol. 27,47-50. Nagao, E.,and Chino, H. (1987)./.Lipid Res. 28,450-454. Nagao, E.,and Chino, H. (1991)./.LipidRes. 32,417-422. Neven, L. G., Duman, J. C., Low, M. G., Schl, L. C., and Castellino, F. J. (1989)./.Comp. Physiol. B 159,71-82. Nichols, A. V., Blanche, P. J., and Gong, E. L. (1983).Handb. Electrophoresis 3,29-46. Noriega, F. G., and Wells, M. A. (1992).Insect Biochem. 22,585-590. Ochanda, J. O.,Osir, E. O., Nguu, E. K., and Olembo, N. K. (1991).Comp. Biochem. Physiol. B

99B,811-814. Oeswein, J. Q., and Chun, P. W. (1981).Biophys. Chem. 14,233-245. Orchard, I. (1987)./.Insect Physwl. 33,451-463. Pattnaik, N. M., Mundall, E. C., Trambusti, B. G., Law, J. H., and Kkzdy, F. J. (1979).Comp. Biochem. Physiol. B 63B,469-476. Pines, M., and Applebaum, S. W. (1977).Insect B i o c h . 8, 183-187. Pines, M., Tietz, A., and Applebaum, S. W. (1981).Gen. Comp. Endocrinol. 43,427-43 I. Prasad, S.V., Ryan, P. O., Law, J. H., and Wells, M. A. (1986a).J.Biol. Chem. 261,558-562. Prasad, S . V., Fernando-Warnakulasuriya,G.J. P.,Sumida, M., Law, J. H., and Wells, M. A. (1986b).J. B b l . Chem. 261,17174-17176. Prasad, S. V., Tsuchida, K., Cole, K. D., and Wells, M. A. (1987).I n “Molecular Entomology” (J.H. Law, ed.), pp. 267-273. Alan R. Liss, New York. Rimoldi, 0.J., Peluffo, 0. R., Gonzalez, M. S., and Brenner, R. R. (1985).Comp. Biochem. Physiol. B 82B, 187-190. Rimoldi, 0.J., Soulages, J. L., Gonzalez, M. S., Peluffo, 0. R., and Brenner, R. R. (1991). Actu Physiol. Phannacol. Latinoam. 40,239-255. Robbs, S. L., Ryan, R. O., Schmidt,J. 0.. Keim, P. S., and Law, J. H. (1985)./.LipldRes. 26,

241-247. Ryan, R. 0.(1990)./.LipUiRes. 31, 1725-1739. Ryan, R. O., Schmidt, J. O., and Law, J. H. (1984).Arch. Insect Biochem. Physiol. 1,575-383.


413

Ryan, R. O., Prasad, S. V., Henriksen, E. J.. Wells, M. A., and Law, J. H. (1986a).J. Biol. Chem. 261,563-568. Ryan, R. O., Wells, M. A., and Law, J. H. (1986b). Biochem. Biophys. Res. Commun. 136, 260-265. Ryan, R. 0..Senthilathipan, K. R., Wells, M. A., and Law, J. H. (1988a).J. Biol. Chem. 263, 14 140- 14145. Ryan, R. O., Haunerland, N. H., Bowers, W. S., and Law, J. H. (1988b). Biochim. Biophys. Acta 962, 143-148. Ryan, R. O.,Van Antwerpen, R., Van der Horst, D. J., Beenakkers, A. M. T., and Law,j.H. (1990a).J. Biol. Chem. 265,546-552. Ryan, R. O., Wessler, A. N.. Price, H. M., Ando, S., and Yokoyama, S . (1990b).J. Biol. Chem. 265,10551-10555. Ryan, R. O., Howe, A., and Scraba, D. G. (199Oc)..j.Lipld Res. 31,871-879. Ryan, R. O., Ziegler, R., Van der Horst, D. J., and Law, J. H.(1990d). Insect Biochem. 20, 127-133. Ryan, R. O., Kay, C. M., Oikawa, K., Liu, H., Bradley, R., and Scraba, D. G. (1992).J. Lipid Res. 33,55-63. Sata, T., Havel, R. J., and Jones, A. L. (1972).J. LipldRes. 13,757-768. Scanu, A. M. (1972). Biochim. Biophys. Acta 265,471-508. Schneider, H., Morrod, R. S., Colvin, J. R., and Tattrie, N. H. (1973). Chem. Phys. Lip& 10, 328-353. Schulz, T. K. F., Van der Horst, D. J., Amesz, H., Voorma, H. 0.. and Beenakkers, A. M. T. (1987). Arch. Insect Biochem. Physiol. 6,97-107. Schulz, T. K. F., Van der Horst, D. J., and Beenakkers, A. M. T. (1991). Biol. Chem. Hoppe-Seyler 372,5- 12. Segrest, J. P., Jones, M. K., DeLoof, H., Brouillete, C. G., Venkatachalapathi, Y. V., and Ananthatarnaiah, G. M. (1992).J. LipidRes. 33, 141-166. Shapiro, J. P.. and Law, J. H. (1983). Biochem. Biophys. Res. Commun. 115,924-931. Shapiro, J. P., Keim, P. S., and Law, J. H. (1984).J. Biol. Chem. 259,3680-3685. Shapiro, J. P., Law, J. H., and Wells, M. A. (1988). Annu. Rev. Entomol. 33,297-318. Shen, B. W., Scanu, A. M., and Kkzdy, F. J. (1977).Proc. Natl. Acad. Sci. U.S.A. 74,837-841. Smith, A. F., Tsuchida, K., Hanneman, E.,Suzuki, T. C., and Wells, M. A. (1992).J. Biol. Chem. 267,380-384. Soulages, J. L.. and Brenner, R. R. (1991).J. LipidRes. 32,407-415. Soulages,J. L., and Wells, M. A. Biochemistry, (1994a). Soulages, J. L., and Wells, M. A. (1994b). Insect Biochem. 24, in press. Soulages, J. L., Riomoldi, 0. R., and Brenner, R. R. (1988a).J. LipulRes. 29, 172-182. Soulages,J. L., Rimoldi, D. J., Peluffo, 0. R., and Brenner, R. R. (1988b). Biochem. Biophys. Res. Commun. 157,465-471. Spencer, 1. M., and Candy, D. J. (1976). Insect Biochem. 6,289-296. Stanley-Samuelson, D. W., Jurenka, R. A., Cripps, C., Blomquist, G. J., and de Renobales, M. (1988). Arch. Insect Biochem. 9, 1-33. Strobel, L. M., Kanost, M. K., Zeigler, R., and Wells, M. A. (1990). Insect Biochem. 20, 859-863. Surholt, B., Schulz, T. K. F., Goldberg, J., Van der Horst, D. J.. and Beenakkers, A. M. T. (1988). Insect Biochem. 18, 117-126. Surholt, B., Van Doorn, J. H., Goldberg, J., and Van der Horst, D. J. (1992). Biol. Chem. Hoppe-Sqler 373, 13-20. Tall, A. R. (1986).J. Lipid Res. 27,361-367. Telfer, W. H. (1960). Biol. Bull. (Woods-Hole, Mass.) 118,338-351.

414

JOSd L. SOULAGES AND MICHAEL A. WELLS

Telfer, W. H., and Pan, M.-L. (1988).Arch. Insect Biochem. Physiol. 9,339-355. Telfer, W. H., Pan, M.-L., and Law, J. H. (1991).InsectBiochem. 21,653-663. Tietz, A., and Weintraub, H. (1978).Insect Biochem. 8, 11-16. Tietz, A., and Weintraub, H. (1980).Insect Biochem. 10,61-63. Teitz, A., Weintraub, H., and Peled, Y.(1975).Biochim. Biophys. Actu 388, 165-170. Tsuchida, K., and Wells, M. A. (1988).Insect Biochem. 18,263-268. Tsuchida, K., and Wells, M. A. (199O).J.Biol. Chem. 265,5761-5767. Tsuchida K., Prasad, S. V., and Wells, M. A. (1987).Insect Biochem. 17, 1139-1 141. Van Antwerpen, R., and Law, J. H. (1992).Arch. Insect Biochem. Physiol. 20, 1-12. Van Antwerpen, R., Linnemans, W. A. M., Van d e r Horst, D. J., a n d Beenakkers A. M. T . (1988).Cell Tissue Res. 252,661-668. Van Antwerpen, R., Wynne, H. J. A., Van d e r Horst, D. J., a n d Beenakkers, A. M. T. (1989).Insect Biochem. 19,809-814. Van Antwerpen, R., Beekwilder, J., Van Heusden, M. C., Van der Horst, D. J., and Beenakkers, A. M. T. (1990).Biol. Chem. Hoppe-Seyler 371, 159-165. Van d e r Horst, D. J. (1990).Biochim. Biophys. Acta 1047, 195-21 1. Van d e r Horst, D. J.. Baljet, A. M., Beenakkers, A. M. T., and Van Handel, E. (1978).Insect Biochem. 8,369-373. Van d e r Horst, D. J., Van Doorn, J. M., and Beenakkers, A. M. T. (1979).Insect Biochem. 9,

627-635. Van der Horst, D. J., Van Doorn, J. M., DeKeijzer, A. N., and Beenakkers. A. M. T . (1981). Insect Biochem. 11,717-723. Van der Horst, D. J., Van Doorn, J. M., and Beenakkers, A. M. T. (1984).Insect Biochem. 14,

495-504. Van der Horst, D. J., Beenakkers, A. M. T., Van Doorn, J. M., Gerritse, K., and Schulz, T. K. F.(1987).Insect Biochem. 17, 799-808. Van d e r Horst, D. J., Ryan, R. O., Van Heusden, M. C., Schulz, T. K. F., Van Doorn, J. M., Law, J. H., and Beenakkers, A. M. T . (1988).J.Biol. Chem. 263,2027-2033. Van der Horst, D. J., Van Doorn, J. M., Voshol, H., Kanost, M. K., Ziegler, R., and Beenakkers, A. M. T. (1991).Eur.J. Biochem. 196,509-517. Van Heusden, M. C. (1993).Insect Biochem. 23,785-792. Van Heusden, M. C., and Law, J. H. (1989).J.Biol. Chem. 264, 17287-17292. Van Heusden, M. C., Van d e r Horst, D. J., and Beenakkers, A. M. T. (1984).J.Insecf Physiol.

30,685-693. Van Heusden, M. C., Van der Horst, D. J., Van Doorn, J. M., Wes, J., a n d Beenakkers, A. M. T. (1986).Insect Biochem. 16,517-523. Van Heusden, M. C., Van der Horst. D. J., Voshol, J., and Beenakkers, A. M. T. (1987a). Insect Biochem. 17,771-776. Van Heusden, M. C., Van d e r Horst, D. J., Van Doorn, J. M., and Bennakkers, A. M. T. (1987b).Comp. Biochem. Physiol. B 88B, 523-527. Van Heusden, M. C., Van d e r Horst, D. J., Kawooya, J. K., and Law, J. H. (1991).J.Lipid Res. 32,1789-1794. Van Marrewijk, W. J. A., van de Broek, A. T. M., Van der Horst, D. J., a n d Beenakkers, A. M. T. (1984).InsectBiochem. 14, 151-157. Van Marrewijk, W. J. A., Van den Broek, A. T. M., and Beenakkers, A. M. T. (1991).Insect Biochem. 21,375-380. Venkatesh, K., Lenz, C. J., Bergman, D. K., and Chippendale, G . M. (1987).Insecf Biochem. 17, 1173-1 180. Verdery, R. B., a n d Nichols, A. V. (1975).Chem. Phys. Lipids 14, 123-134. Wang, Z., Hayakawa, Y.,and Downer, R. G . H. (1990).Insecf Biochem. 20,325-330.


415

Weers, P. M. M., V an der Horst, D. J., Van Marrewijk, W. J. A., Van den Eijnden, M. Van Doorn, J. M. V., and Beenakkers, A. M. T. (1992).J.Lipid Rex 33,485-49 1. Weinberg, R. B., Ibdah, J. A., and Phillips, M. C. (1992).J. B i d . C h . 467,8977-8983. Weintraub, H., and Teitz, A. (1973).Biochim. Biophys. A c h 306,31-41. Weintraub, H., and Teitz, A. (1978). Insect Biochem. 8,267-274. Wells, M. A., Ryan, R. 0..Prasad, S. V., and Law, J. H. (1985).Insect B i o c h . 15,565-571. Wells, M. A., Ryan, R. O., Kawooya, J. K., and Law, J. H. (1987).J . Biol. C h . 264, 4 172-4 176. Wheeler, C. H., and Goldsworthy, G. J. (1983).J.Insect Physiol. 29,339-347. Wheeler, C . H., and Goldsworthy, G. J. (1985).B i d . Chem. Hoppe-Seyler 366, 1071-1077. Wheeler, C. H., Van der Horst, D. J., and Beenakkers, A. M. T. (1984).Insect B i o c h . 14, 261-266. Wheeler, C. H., Boothby, K. M., and Goldsworthy, G. J. (1986).B i d . C h . HopPe-Sqrler367, 1127-1 133. Wilson, C., Wardell, M. R., Weisgraber. K. H., Mahley, R. W., and Agard, D. A. (1991). Science 252, 18 17- 1822. Yeagle, P. L., Martin, B., Pottenger, L., and Langdon, R. G. (1978). Biochemistty 17,27072710. Ziegler, R. (1984).Gen. Comp. Endocrinol. 54, 51-58. Ziegler, R. (1991).J.Comp. Physiol. 161, 125-131. Ziegler, R., and Schulz, M. (1986).J. Insect Physiol. 32,903-908. Ziegler, R., Ryan, R. O., Arbas, E. A., and Law, J. H. (1988).Arch. Insect Biochem. Physiol. 9, 255-268. Ziegler, R., Eckart, K., and Law, J. H. (1990). Peptides(N.Y.) 11, 1037-1040. Zilversmit, D. B. (1965).J.Clin. Invest. 44, 1610-1622.


AUTHOR INDEX Numbers in italics refer to the pages on which the complete references are listed.

A Aalto-Set?ila, K., 267,299 Abbott, R.D., 307,369 Abdo, Y., 162,196 Abergel, C., 44,50 Abita,J.-P., 60,80 Abousalham, A., 26,27,46 Abrahamsson, S., 69.80 Achari, A., 58,63,64,66,69,70,80,86 Adams, G. H., 216,244 Adams, P. A., 177,196 Adamson, G. L., 222, 223,246 Adelman, J., 156, 199 Aden, R. A., 17,50 Adinolfi, A., 154,196 Adinolfi, M., 154,196 Adrian, M., 387,409 Agard, D. A., 213,248,285,286,287, 288,289,290,295,301,302, 341, 344,345,351,369,378,415 Agellon, L. B., 262,296 Aggerbeck, L. P., 215, 226,244,246,247, 257,258,259, 260,264,285,295, 302,387,409,41I, 412 Ahlstedt, S., 183,196 Ailhard, G., 2,20,48 Akanuma, Y., 267,302 Akoi, K., 171,196 Alagon, A. C., 57,86 Albers, J. J., 250,296, 308, 341, 342,346, 355,357,360,364,367,386,410 Alden, R. A., 11,50 Aldini, R., 177,201 Alexander, C., 240,242,243,244 Algelico, F., 209,248 Allen, F. H., 15.47 Allen,J., 39.48 Allen, T. M., 323,346,348,366

Alord, D., 339,365 Alpers, D. H., 133,151 Altland, K., 357,369 Altschul, S. F., 101, 150 Amaki, I., 162, 163,200 Ameis, D., 41, 49 Amesz, H., 376,390,413 Amin, S., 93, 104,149 Amit, B., 268,269,271,298 Anantharamaiah, G. M.,287,293,300, 308,309,311,313,314,322,323, 324,329,33 1,333,339,340,341, 342,343,346,347,348,350,353, 354,355,357,359,360,361,362, 363,364,365,366,367,368,369, 378,413 Ancell, H., 152,196 Andalibi, A., 41,49 Andersen, S., 186,197 Anderson, I., 18.50 Anderson, L. J., 250,295 Anderson, R. A., 44,47 Ando, S., 398,413 Andrade, J. D., 161, 176,198 Andreasen, F.,47 Anfinsen, C. B., 40,47, 154,201 Angal, S., 43,47, 48 Annand, R.,57,58,82 Anne, K. C., 350,364 Antonian, E., 2,47 Antonio, R.,209,248 Anundi. H., 138,150 Aoyama, S., 39,47 Appelt, K., 98,150 Applebaum, S.W.,401,409,412 Aqvist, J., 92, 115, 138, 139,148,150 Arai, K., 162, 163,196, 199,200 Arakawa, F.,269,299 Arakawa, K., 269,299 417

418

AUTHOR INDEX

Araki, K., 269,299 Araujo, P.S . , 58,80 Arbas, E.A., 402,403,415 Archibald, F. M.,225,248 Areas, E. P.G., 58,80 Argiolas, A., 313,320,364 Argraves, W.S.,264,300 Argyres, M.,209,246 Arias, I. M.,90,150 Arita, H.,57,84 Arnold, K. S.,207,247,263,271,272, 284,297,299 Ashbrook, J. D.,178,202 Ashcom, J. D.,264,300 Asher, C.,401,409 Ashizawa, E.,39.50 Ashley, G.,11,33,50 Assefbarkhi, N.,269,300 Assmann, G.,44,50,260,261,269,277, 298,299,300,357,369 Atkinson, D.,209,210,235,244,245, 324,344,345,352,364,367 Atsma, W.,76,87 Attie, A. D.,210,247 Austin, J. P.,352,364 Austin, M.A., 307,364 Avignon, A., 121,122,141,149 Avila, E. M.,250,295

Baartels, K. S., 99,I51 Baba, T., 45,47 Bachovchin, W.W.,11.50 Backhart, B. D.,235,236,237,248 Baillargeon, M.W.,19,47 Bajszar, G., 195,199 Baker, E. N.,11,47 Baker, M. E.,21 1,244 Baldwin, G . S., 156,202 Balestra, M.E.,210,245 Baljet, A. M.,373,414 Ban der Horst, D. J., 404,415 Banaszak, L.J.. 42,50,93,98,104,110, 113. 115, 116,120,121, 123, 126, 128,129,130,133, 134,135, 142, 144,145,147,149,150, I51 Banka, C. L., 360,364 Bar-Tana. J., 147,148 Baratti, J., 2,49

Barbara, L.. 177,201 Barclay, M.,225,248 Barclay, R. K.,225,248 Barlow, P. N.,69,80 Barlow, P.,66,69,80 Barrett, A. J., 38,50 Barrett, J., 309,361,366 Barrett, R.,307,367 Bartels, C. F.,45,49 Barter, P.J., 308,368 Barton, G. J . , 169,203 Barynin, V. V.,99,151 Bass, N. M.,90,91,93,104,133,148, 150,151 Basu, P. K., 180,200 Basu, S. K.,195,200 Basu, S. P., 167,201 Batenburg, M.,6,39,47 Bates, M.L.,133,151 Bauerfiend, R.,394,398,409 Baulard, A., 39,47 Bauman, A., 141, 142,143,149 Baxa, C.A., 120,148 Baynes, J. W.,155, 183,197,198 Bazan, J. F.,31 1,364,365 Beacham, 1.R.,39,49 Beaven,G. H.,172,177, 192,196,198 Bech, I. M.,44,47 Beckmann, E.,69,82 Becquart, J., 195,203 Beekwilder, J., 399,414 Beenakkers, A. M.T., 372,373,375,376, 377,378,379,385, 388,390,394, 396,399,400,401,402,403,404, 409,410,411,413,414,415 Behrens, P. Q.. 156,196 Beisiegel, U.,255,275,295,301 Bekkers, A. C.A. P. A., 64,80 Belfield, G. P.,195,202 Belfrage, P.,45,46,47,49, 50 Bell, J. D.,58,81 Ben-Avram, C.M.,41,47 Ben-Zeev, O.,41,42.47,48 Benassayag, C.,177,201 Bendedouch, D.,161,196 Bengtsson, G., 45.50 Bengtsson-Olivecrona, G., 18,41,42.48, 50,275,295 Benjkamin, G.S.,171,199 Bennetech, S . L.,130,150

AUTHOR INDEX

Benning, H. M., 93, 104, 113, 122, 131, 132, 133, 148, 287,296, 341, 342, 357,364, 377,378,379,383,410 Benninghoben, A,, 357,369 Bensadoun, A., 42,49 Bentz, J., 339.365 Berenson, G. S., 275,300 Berg, 0. G., 76,78,81,83 Berg, 0..71.83 Bergeron, J. M., 156, 203 Bergman, D. K., 396,410,414 Bergors, T., 108, 122, 126,149 Berka, T. R.,39,48 Berman, M. C., 177,196 Bernheimer, A. W., 313,364 Bernier, M., 128, 148, 149 Bernini, F., 356,368 Bernlohr,D.A.,93, 104, 110, 115, 116, 120, 122, 126, 127, 128, 129, 130, 134, 135, 142, 144, 145, 147,148, 149,150,151 Bersot, T. P., 262, 269,278, 284,295, 297,299,300 Bersot, T., 269,297 Bertics, S. J., 208,248 Berzofsky, J. A., 31 1,364 Betsholtz, C., 207,246 Bhamidipati, S. P., 186,198, 393,411 Bhown, A. S., 323,348,353,354,363 Bier, D. M., 307,367 Bierman, E. L., 307,308,360,367,368 Biltonen, R. L., 58.81 Binsbergen, J. V., 76,87 Birdsall, N. J. M., 79,81 Birkedal, B., 309, 362,368 Birkenmeier, E. H., 133, 151 Birkett, D. J., 179, 181, 182,202 Birktoft, J. J.. 11,50 Birktoft, J., 11, 17,47, 50 Birnbaum, E. R., 165, 169,198 Bjorgell, P., 45.50 Bjorkling, F., 3, 5, 17, 19, 22,47, 78,81 Bjursell, G., 208, 209, 210, 212,245,247 Blackburn, W. D., 309,362,364 Blackhart, B. D., 208,209,244,246 Blackhart, B., 210,211,213,232,246 Blacklock, B. J., 398,410 Blackney, E. W., Jr., 320,368 Blagoev, B., 57,85 Blanche, P. J., 386,412

419

Blankenship, D. T., 274,276,296 Bleibaum, J., 18,50 Blomquist, G. J., 373,404,410, 413 Blomqvist, P., 80,86 Bloomfield, V., 161, 173,196 Blow, D. M., 11,47, 167,201 Blow, D., 74,81 Blowman, H. L.,2,47 Blum, C. B., 265,269,295,299 Blum, C., 256,277,302 Blum, Kaelin, D., 42,48 Blumberg, B. S., 163,198,202 Blumenstock, F. A., 181, 189,201 Blundell, T. L., 32,50 Bobbitt, J. L., 64,87 Bock, S. C., 156,199 Bocskei, Z., 92,93, 140,148 Bodmer, M. W., 43,47,48 Boel, E., 2, 6, 11, 35,47,49 Boelens, R.,58,66,82 Boerwinkle, E.,209,244, 257,280,297 Boguski, M. S., 311,364, 377,379,410, 411 Bohme, J., 138,150 Bohmer, F.-D., 93, 104,148 Bolin, J., 58,63,81 Bollade, D., 398,410 Bolognesi, M.,92,93,98, 139,150 Bomalaski, J. S., 58,81 Bondjers, G., 209,210,212,247 Bonelli, F. S., 359,364 Boni, L. T., 320,365 Boni, L., 339,365 Bonicel, J., 2,48 Bonner, J., 156,199,201 Bonnert, D. J., 360,364 Bonsen, P. P. M., 60,80 Boogaerts, J., 323,324,341,350,355,364 Boothby, K.M., 404,41S Boren, J., 240,245 Borenstein, S., 252,297 Borg, K. O., 179,199 Borgstrom, B., 9,47 Borisov, V.V., 99, 151 Borst, G. C., 161,196 J. M.,166,196 Bos, 0. Boston, M., 39,50 Bostrom, K., 209,210,212,245,247 Boswell, D. R.,162,196 Bothner-By, A. A., 79,85

420

AUTHOR INDEX

Bott, R., 39.50 Boucher, F., 195,203 Boudouard, M.,2,48 Boundy, K.L., 120,148 Bourdeaux, M.,178,197 Bowden, G. T., 90,93, 104,149 Bowers, W. S., 377, 398,411, 413 Bowman, B. H., 156, 160,202,203 Boyle, E., 40.47 Boyles, J. K., 250,268,269, 271,295,297, 298 Bradley, R., 388,399,413 Bradley, W. A., 210, 211,214, 215,245, 248, 258,266,272,296, 341, 342, 351,365 Brady, I., 3,4, 17, 47 Brand, J. G., 179,196 Brand, S., 162,196 Branden, C., 98,148 Branden, C.-I., 33, 49 Brandenberg, N. P., 60,82 Branner, S., 44,47 Brasseur, R., 209,245 Bratt, J. M.,93, 104, 110, 134, 135, 151 Braunitzer, G., 139, 148 Bray, J., 393,410 Brecht, W. J., 275,280,297 Breckenridge, A,, 178,196 Breckenridge, W. C., 2 1 1,246, 344, 350, 358,364 Breddam, K., 5,30, 31, 33, 38,44,47,49 Breiter, D. R., 287,296, 341, 342, 357, 364,377,378,379,383,410 Brennan, S. O., 162, 163,196,200, 269, 301 Brenner, R. R.,373,375, 390,391,392, 393,398,404,411,412,413 Brenner, S., 2,7,47 Breslow, J. L.,207, 208,209,245,246, 251,255,256, 257,261,267, 277, 282,296,299,301,302 Brewer, H. B., 210,246 Brewer, H. B., 111,269,298 Brewer, H. B., Jr., 207, 208,210,245, 246, 260,261,267,269,282,297, 298,299,300 Briand, C., 178,197 Brinton, E. A., 307,367 Broadhurst, M.J., 58,87 Brock, A., 178,197

Brockerhoff, H., 2,47 Brockman, H. L., 45,49,410 Brodersen, R., 177, 186,197 Brogstrom, B., 27,48 Brooks, A. R.,208,209,245,246 Broom, M. B., 160,161,167, 168,197 Brouillette, C. G., 212,248, 287, 293,300, 3 W 3 13,322,323,324,329,33 1, 333,340,341, 342, 343,346, 348, 350, 352,353,354,355,357, 359, 360,363,364,368,378,413 Brown, A., 164,200 Brown, J. B., 183,197 Brown, J. H., 253,262,267,268,298 Brown, J. R., 155, 156, 160, 161, 166, 171,181,186,190,191,196,197,201 Brown, K. E., 179,197 Brown, K. F., 178, 179,197,202 Brown, M.A., 2,41,47 Brown, M.L., 262,296 Brown, M.S., 44,48, 207,246, 250,256, 262,263,264,265,266,271,277, 295,296,297,300,301,302,307,365 Brown, R. K., 40,47 Brown, W. M., 156,197 Browning,J. L., 58,59,60,62,63,67,71, 72, 73, 78,81,86 Bruckett, E., 214, 218, 223,225, 226,245 Brune, J. L., 156, 160,202 Brunie, S., 55,58,59,63,66,67,69,80, 81,82, 85 Bruno, G. A. P., 207.246 Bruno, G., 207,245 Bryan, P., 17,47 Brzozowski, A. M.,3, 5, 17, 19,22, 39.47, 48, 49, 78,81 Brzozowski, A., 3,4, 17,47 Bubois-Dalcq, M.,126, 151 Buchwald, P., 42,48 Buelt, M.K.,120, 128, 129, 130, 144, 148, 149 Bugg, C. E., 167,197 Bugis, P., 344,358,369 Bullock, T., 5, 30.31, 33, 38,49 Bunn, H. F., 186,201 Burgess, W. H., 264,300 Burgoyne, R. D., 53,81 Buringame, A. L., 90, 93, 104, 150 Burke, C. W., 177,197 Burke, D. J., 216,246,386,412

AUTHOR INDEX

Burman, K. D., 161,196 Burns, G. A. P., 209,246,251,296 Burton, S.J., 195,201 Butler, E.,209,248 Butler, R., 208,209,245,247,248 Butler-Brunner, E., 208,247 Byers, M.,207,246 Byrne, R., 324,368 Byrnes, L., 156,197

C Cai,S.-J., 210,211,245 Caiati, L., 208,209,244,246 Callan, W. M.,180,197 Callard, I. P., 211,247 Cambillau, C., 3,4,5, 10, 18,22,27,28, 33,41,44,48,50,197 Camejo, G., 290,296 Camerman, A., 189,197 Camerman, N., 189,197 Camp, S., 45,50 Campbell, M. M.,58,81 Candy, D. J.. 401,413 Canioni, P., 26,47 Cann, H., 207,245 Cannistraro, S., 169,197 Canzler, H., 277,279,301 Cappelleri, G., 177,201 Capurro, M.de L., 373, 374,396,398, 410 Cardin, A. D., 274,276,296 Carew, T. E., 262,296,299 Carlsson, P., 208,209,2 10,212,245,247 Carrell, R. W., 162,196, 269,301 Carriere, F., 44,50 Carson, M.,175, 193,197 Carter, D. C., 155, 160, 161, 164, 167, 168, 182, 183, 185,197,198,200 Carter, P., 74,75,81 Casey, M.L., 271,302 Cassel, D. L., 259,296 Casteli, W. P., 307,369 Castelino, F. J., 374, 375,412 Castellani, A. A., 162, 163,198,200 Castelli, W. P., 307,366 Castle, C. K., 252,299 Catapano, A. L., 344,350,358,364 Cavaggioni, A., 92,93, 140,148 Cease, K. B., 3 11,364

42 1

Ceve, G., 77,81 Chacko, G. K., 308,366 Chahinian, H.,43,48 Chaillan, C., 26, 27,46 Chajek-Shaul, T., 273,300 Chambon, P.,134,149 Chan, I., 3,50 Chan, L., 2,41,42,48,47, 207,209,210, 211,212,213,214,226,244,245, 247,248,251,252,253,296,298, 322,344,367,383,412 Chang, C.-C., 76,87 Chang, D. J., 251,299 Chang, L.-S.,76,87 Chang, T., 66,87 Chao, Y.-S.,266,302 Chaouleas,J. G., 54,84 Chapman, D., 58,84 Chapman, J. M.,375,386,410 Chapman, M.J., 205,214,215,218,223, 225,226,245,246 Chappell, D. A., 250,296 Chapus, C., 26,27,46 Chapus, S., 10,47 Charlton, E., 19,47 Charlwood, P. A., 164,197 Chatterton, J. E.,227,231, 232,235,236, 237,238,240,243,245,248 Chaut, J. C., 41,49 Cheah, E., 33,37,50 Chen, C. H., 250,296,323,365 Chen, G. C., 207,226,245,296 Chen, H. C., 313,320,369 Chen, L., 208,247 Chen, M.C., 169,197 Chen, P.-F., 212,245 Chen, R., 139,148 Chen, S. H., 161,196 Chen, S.-H., 2,41,47, 177, 192,196, 207, 210,211,212,214,226,245,248 Chen, T. C., 349,364 Chen, W.-Q., 57,58,82 Chen-Liu, L. W., 235,236,237,238,240, 243,248 Cheng, L., 121, 122, 141, 142, 143,149 Cheung, M.C., 308,341,342,346,355, 357,360,364,367,386,410 Chignell, C. F., 178, 179,197 Chinander, L. L., 120,148 Chincarini, C. C., 183,197

422

AUTHOR INDEX

Chino, H., 372,373, 374,375, 377, 378, 385, 386, 388,394,397,398,401, 402,403,404,410,411,412 Chiovetti, R., Jr., 323,346,348,352, 354, 366,369 Chippendale, G. M., 374, 375,396,410, 414 Chippendale, M. G., 375, 412 Chisholm, J. M., 404,411 Chiyoko, S., 162, 163,202 Cho, M. J., 178,197 Cho, W., 57,58,81,85 Chock, S. P., 53,81 Choi, S. Y., 266,296 Chothia, C., 33,49 Chou, P. Y., 253,254,296,344,364 Chow, E. P., 57,58,84 Christef, N.,177,201 Christiansen, L., 3,4, 17, 47 Christiansen, M., 2, 6, 11, 35, 47 Christophe, J., 57,83 Chu, M.-L., 208,209,244 Chuang, S. S . , 208,248 Chuck, S. L., 240,245 Chun, P. W., 389,412 Chung, B. H., 308,323,341,342,346, 348,352,353, 354,355, 357,363, 364,369 Chytil, F., 91, 121, 141, 149, 151 Cistola, D. P., 113, 121, 123, 133, 141, 142,143, 147,149,150, 186,200 Cladaras, C., 209, 210,245 Clancy, L. L., 167,197 Clark, A. B., 250,275,296,298 Clark, J. D., 54,81 Clark, W. A., 377,379,410 Clarke, A. R.,275,296 Clarke, H.R. G., 208,248, 278,300 Clavey, V.,356,367 Clawson, D. K., 64.87 Cleansby, A., 6,39,47 Coassolo, P., 178, 197 Coelho, H. S. L., 377,404,411 Coetzee, G. A., 363,364 Cohen, B. L., 154,200 Cohen, C., 313,364 Cohn, E. J., 166, 197 Colburn, K. A., 212,247 Cole, G., 16,50 Cole, K. D., 377, 379, 383, 396,405, 406, 410.412

Cole, T. G., 308,366 Coleman, R. D., 212,245 Colson, C., 39,47 Colvin, J. R., 388,413 Compans, R.W., 309,350,361,362,367, 368

Cone, J. T., 308, 341, 342,346,355,357, 364

Constants, J., 156, 160,201 Cooke, N. E., 156,197 Cooper, A. D., 266,296 Cooper, D. L., 156,198 Cornette, J. L., 31 1,364 Cornish-Bowden, A., 184,198 Correh, G., 397,410 Cowan, S. W., 92,93,97, 104, 108, 110, 126, 127, 130, 134, 135, 138, 139, 140,149 Cox, L. A., 252,297 Craik, C. S., 16,50, 75,81,86 Cramer, K. D., 75,81,88 Crameri, R., 57,58,82 Craven, B. M., 167,200 Crenne, J.-Y., 195,203 Crepy, 0.. 178,200 Crick, F. H. C., 313,364 Cripps, C., 404,413 Crooks, M. J.. 178, 179,197,202 Crowl, R. M., 58.84 CshzAr, A., 257,280,297 Cserpaan, I., 195,199 Cuccia, I. A., 19,49 Cullis, A. F., 166, 201 Cullis, P. R., 339, 364 Culwell, A. R., 294,300 Cunningham, B. A., 410 Cupples, R. L., 156, 160,202 Curtiss, L. K., 207, 208, 227, 231, 232, 245,247,248, 250,274,296,299, 356,360,364,365 Cusanovich, M., 99,150 Cygler, M., 3,4,6,7,8, 16, 18, 22,29, 33, 37,48,50

D’Albis, A., 177, 192, 196 DArcy, A., 3,4,9. 10, 18, 20,42,50, 98, 151

D’Athis, P., 178, 179,202 d’Auriol, L., 41,49

AUTHOR INDEX

dAvignon, A., 121, 122, 134, 136,149 Dahlen, B., 69,80 Darby, G., 167,197 Darnal, D. W., 165, 169,198 Darnfors, C., 208,245 Dartois, V., 39, 47 Das, D., 93, 104,151 Das, H. K., 208,245,248, 251,296 Das, S., 393,410 Datta, S., 2,41, 47, 251,298, 344,367, 383,412 Dauaberman, J., 39,50 Dauber-Osguthorpe, P.,58,86 Daughaday, W. H., 177,197 Dautrevaux, M., 162,196 David, E. V., 156,197 Davidson, N. W., 210,245 Davies, D. R., 166, 199 Davies, G. E., 251,299 Davies, H. M., 18,50 Davies, R. C., 41,42,50 Davignon, J., 255,257, 269,280,282, 296,299 Davis, C. G., 271,302 Davis, E., 162, 163,200,201 Davis, L. G., 389,411 Davis, P.D., 58,87 Davis, R. A., 236,248 Davis, R. C., 41,42,47, 275,296 Davis, R. W., 252,253,296 Dawson, R. M. C., 393,410 Day, J. F., 155, 183, 197, 198 Day, J., 39, 49 Dayhoff, M.O., 190,197 de Bianchi, A. G., 373,398,410 De Caro. J., 2,48 De Geus, P.,4, 10,33,50, 58,66,82,84 De Gier, J. J., 176, 198 de Haas, G. H., 53,54, 55,58,60,63,64, 66,69, 74, 75, 76, 77, 78,80, 82,83. 84,85,86,87 de Knijff, P.,269,300 de Kort, C. A. D., 374,410 de Kort, S., 373,375, 385,392,411 De Loof, H., 21 1,213,245,287,293,300 De Martino, A. G., 178,200 de Renobales, M.,373,404,410, 413 de Silva, H. V., 261, 267,296 Dean, R. T., 215,232,247 Deber, C. M., 188,199 DeBony, J., 79,81

423

Deckelbaum, R.J., 18,48, 216,245 Deckerbaum, R., 308,369 Degovivics, G., 385,411 Deguchi, K., 205,246 deHaas, G. H., 43,48 DeHaas, G . H., 76,83 Deisenhofer, J., 11,50 DeKeijzer, A. N., 402,414 Dekelbaum, R. J., 352,369 Dekker, B. M. M.,195,202 Dekker, N., 58.66.82 Del Ben, M.,209,248 Del Vale, J., 93, 104, 149 DeLisi, C. J., 311,364 DeLoof, H., 210,211,212,214,248,313, 314, 322,324,329,331,333,340, 343,356,367,368,369,378,413 DeLoof, J. A., 323,324,341, 350, 355, 364 DeLucas, L. J., 167,197 Demaret, J.-P., 58.82 Demel, R. A., 320,369, 378,381,382,410 Demmer, L. A., 210,245 Dempcy, R., 57, 58,82 Denfors, I., 184,199 Deng, T., 58,74,75,82 Dennis, E. A., 53,58,69,79,80,81,82, 84,85 Dennison, 0. E., 156,198 Derewenda, U., 3 , 5 , 6 , 17, 19,22,30,32, 33,47,48, 78,81 Derewenda, Z. S., 3 , 5 , 6 , 17, 19,22, 28, 30,32,33,37,39,41,47,48,49,50, 78,81 Derewenda, Z., 3,4, 17,47 Desbiens, N., 179,200 Deslypere, J. P.,210, 211,245 Desnuelle, P.,1, 2,9, 20, 26,48, 50 Deutzman, R.,140,149 Dezdy, F. J., 215,244 Dickerson, R. E., 166, 190, 191,197, 199 Dickson, J., 352,364 Diderichsen, B., 39,49 Diehl, P. A., 397,410 Dier, D. M., 305,366 Dijkman, R.,69,76, 78,86,87 Dijkstra, B. W., 5, 33, 48, 55, 58, 59,60, 60, 66,67, 69, 74, 76, 78,82,84, 85, 86,87 Dijkstra, B., 33,37,50 374,375,410 Dillwith,J. W.,

424

AUTHOR INDEX

Dinitz, H. M., 166,199 DiPersio, L. P., 45,48 Dkon, J. W.,188, 191,197 Dobson, D. E., 127,149 Dobson, E., 3,4, 17,47 Dobson, G. G., 3,5,6, 17, 19,22,39,47, 48,49, 78,81

Dobson, G., 3,4, 17,47 Docherty, A. J. P., 43,48 Dodsworth, N.,195,201 Doers, T. M., 210,247 Dohlman, J. G., 309,313,322,340, 343, 362,364,368

Dolan, P.,275,300 Dolle, P., 134,149 Donaldson, D., 162,196 Dong, L.-M., 269,296 Donner, J., 27,48 Doolittle, M. H., 210,247, 275,296 Doolittle, R. F., 156, 160, 190, 192, 198, 344,367,380,410

Doom, J. H., 375,377,413 Dorset, D. L., 69,82 Dow, E. R., 64,87 Downer, R. G. H., 373,374,375,385, 397,398,401,404,410,411,414

Downs, D., 45,47,50 Drabble, K., 72,86 Drenth, J., 11,47,49, 55,58,59,60,66, 67,69,74,76,78,82,84,85,86,87

Driscoll, D. M., 322,365 Drisko, J., 236,248 Dubochet, J., 387,409 Dubois, B. W.,208,248 Dudler, T., 57,58,82 Dueland, S., 236,248 Dufton, M. J., 54,82 Dugaiczyk, A., 156, 160,198, 199,200 Duman, J. G., 374,375,412 Dunning, A. M.,209,245 Dupureur, C. M., 74,86 Dupureur, C., 74,75,82 Dyer, C. A., 274,296 Dziegielewska, K. M., 156, 197

E Eaker, D., 64,87 Ealick, S. E., 167, 197 Ebert, D. L., 210,247

Eckart, K., 403,415 Eddy, R., 207,246 Edelstein, C., 324,344,357,365,368, 389,410

Edgington, T. S., 356,365 Edmundson, A. B., 310,311,368 Edwards, Y.H., 210,247 Efimov, A. V.,22,32,48 Eglof, M.-P., 3.5, 10, 18,22, 27,28,50 Egmond, M. R., 6, 39,47,60,82 Egner, U., 130,150 Ehlenz, K., 269,300 Ehnholm, C., 250,269,296,301 Eiferman, F., 156, 198 Eil, C., 161,196 Einspahr. H. M., 167,197 Eisenberg, D., 310, 311,313,345,365 Eisenberg, S., 18,48, 273,282,300,301 Eisenthal, R., 184, 198 Eklund, H., 64,87 Eliopoulos, E. E., 92,93,98, 139,150 Elkarim, A. A., 183,198 Ellens, H., 339,365 Elofsson, R., 179, 199 Elovson, J., 209, 210,212,235,236,237, 238,240,243,245,247,248

Elshourbagy, N.A., 3 11,364 Elso, A. B., 69,78,86 Eltze, C., 269,300 Emerson, T. E., Jr., 154, 194,198 Emi, M., 269,296 Erntage, J. S., 43,48 Endo, N., 162,202 Endo, S., 205,246 Enerback, S., 41,50 Engberg-Pedersen, H., 178,198 Engelrnan, D. M.,31 1,313,327,334,365 Enghild, J., 250,300 Engler, J. A., 323, 324,341, 350, 355, 364 Epand, R. M., 320,323,339,365,369 Epp, O., 16,50, 92,93, 140, 149 Era, S., 186,198 Erdos, E. G., 38,50 Ericksson, U., 93, 104,151 Eriksson, U., 92, 138, 139,150 Erlanson-Albertsson. C., 9.47 Eshourbagy, N.A., 251,299 Esser, V.. 264,265,266,297 Etcheverry, T., 195,198 Etienne, J., 41,49

AUTHOR INDEX

Etzold, G., 93, 104, 148 Evans, H. J., 66.84

425

Fisher, M.,267,299 Fisher, W. R., 211,214,215,222,226, 245,248

F Fakaki, F., 210,247 Falk, H., 140,149 Fan, C., 18,50 Farber, G. K., 33,48 Farquhar, M.G., 275,300 Farr-Jones, S., 11.50 Fasman, G. D., 253,254,296,344,364 Faustinella, F., 3,41,42,48,50 Fazio, S., 279,280,296,297 Feeney, J., 79,81 Fehske, K. J., 155,198 Feigin, L. A., 386,410 Feil, S., 90,93, 104,149 Feld, R. D., 162, 163,200 Feldhoff, R. C., 172, 173,198,201 Feldman, R. J., 354,363,368 Feller, G., 39,48 Fencl, V., 152,198 Feng, D. F., 380,410 Feng, L., 161, 176,198 Fenton, M.J., 21 1,247 Ferenc, J. K., 346,347,348,365 Ferenz, C. R., 346,347,348,368 Fernanado, G., 294,300 Fernando-Warnakulasuriya, G. J. P., 374, 375,377,379,394,395,410,411,412

Ferrato, F., 44,50 Ferri, G., 162, 163,198,200 Ferri, S. R., 37,50 Ferri, S., 18,48 Fetzer, V. A., 225,248 Fichera, L., 375,411 Fielding, C. J., 273,297, 359,360,365 Fielding, P. E., 273,297. 359,360,365 Fiess, H.A., 189, 198 Figge, J., 152,198 Fiil, N. P., 2,6, 11, 35,47 Fijii, S., 78,86 Findlay, J. B. C., 92,93, 139, 140,148, 150

Finer-Moore, J., 75,86, 31 1 , 345,365 Finlayson, J. S., 155, 171, 187,201 Fiorella, P. D., 120, 134, 149 Fischer, M.J. E., 166, 196 Fisher, J., 58,66,82,86

Fitch, W. M., 322,365 Fleer, E. A. M.,64,82 Fleer, R., 195,203 Fletcher, J. E., 177, 178, 186,202 Fletterick, R.J., 16,50, 75,86 Fletterick, R.,31 1,364 Flores-Riveros,J. R., 128,149 Flower, D. R., 92,93, 140,148 Fliickiger, R., 186,199 Fogh-Andersen, N., 180,198 Foglizzo, E., 26, 27,46 Fojo, S. S., 207,245 Fong, L. G., 266,296 Fontaine, R. N., 45,49 Fontencilla-Camps, J. C., 44,50 Foreman, R. C., 156,197 Forgez, P., 214,218,223,225,226,245, 250,300

Forrester, W., 195,198 Forte, M.T., 387,411 Forte, T. M.,226,245,267,300, 352,354, 365

Fortier, C., 208, 209,246 Foster, J. F., 155, 172, 173, 176,198,203 Fourest, E., 209,244 Francis, B.. 58,82 Frangione, B., 250,302 Frank, S., 57,83 Franke, A. E., 156,199 Franken, P. A., 60,64,66,74,75,80, 84,86

Franken, S. M.,33, 37,50 Franken, S., 5,33,48 Franklin, S. G., 163,198 Frants, R. R., 269,300 Fraser, R., 269,301 Fredrikson, G., 45,46,47,49 Fredriksson, A., 138,150 Free, M.L., 39,49 Freeman, J. A., 250,300 Freeman, M.,377,379,410,411 Freer, S. T., 11,50 Frey, S., 354,365 Friedander, E. J., 258,270, 271,274,297 Friedberg, F., 180,200 Frieden, C., 144, 145,150 Friedl, W., 209,246

426

AUTHOR INDEX

Friedman, T., 45,50 Friedrich, E.A., 194,202 Fringeli, U. P., 353, 354,365 Frolow, F., 5, 10,33, 37,50 Fruchan, J. C., 356,367 Fruchart, J.-C., 215,232,247 Fry, D. L., 262,298 Fujita, M., 162, 163,202 Fukazawa, C., 252, 267,299,300 Fukushima, D., 346,347,348,365 Fukuzaki, H., 21 1,248 Fuller, M., 2 10,2 11,2 13,232,246 Funahashi, T., 263,296 Funke, H., 269,300,357,369 Fiirestenberger, G., 90,93, 104,149

G Gabay, S., 179, 180,198, 199 Gablian. N., 58.80 Gade, G.,401,403,411 Gago, F., 71.85 Galibert, F., 41,49 Gallagher, J. G., 350,364, 385.41 1 Gallego, J., 71,85 Galliano, M., 162, 163, 196, 198, 199,200, 201

Gambao, G., 64,87 Gandour, R. D., 15,48 Cannon, F., 156,197 Gantz, I., 93, 104, 149 Garber, D. W., 347,365 Garcia, Z. C., 210,245 Garfinkel, A. S., 40,42,48,50, 307,367 Gargouri, Y.,43,48,50 Garlick, R. L., 186,201 Garman, E., 6,39,47 Gamier, J., 344,365 Garrety, K. H., 308,368 Garten, S., 178, 179, 198 Gascuel, 0.. 344,365 Gatmaitan, Z., 90,150 Gausepohl, H., 264,297 Gawish, A., 323,363,365 Gebicke-Haerter, P. J., 250,295 Gebicke-Hgrter, P. J.. 250,297 Gedde-Dahl, T.,Jr., 251,299 Geis, I. G., 190, 191, 197 Geisow, M. J., 172, 195, 198,201

Gelb, M. H., 54, 57,58, 59,60, 61.62, 63, 64,65,67,68, 69, 70, 71,72, 73, 76, 77,78, 79,82,83, 85,86,87, 88 Gemborys, M. W., 178,200 Genda, A., 307,369 Genest, J., 177,201 George, P. M., 269,301 George, P., 162,196 Gerday, C., 39,48 Gerig, J. T., 183,198 Gernert, K. M., 160, 161, 167, 168,197 Gerritse, K., 402,403,414 Gerwirth, D., 58,63,81 Getz, G. S., 252,299, 322,365 Getzoff, E., 99,150 Gevers Leuven, J. A., 269,300 Ghiselli, G., 267, 300 Ghomashchi, F., 7 1,83 Gianturco, S. H., 210, 211,214,248, 258, 266,272,296,341,342,351,365 Gibson, B., 90.93, 104,150 Giguere, V., 93, 104,149 Giladi, H., 268,269,271,298 Gilbert, L. I., 373, 397, 410 Gilbrat, J. F., 344,365 Giller, T., 42, 48 Gilliam, E.B., 258,296, 349,367 Gispen, W. H., 320,369 Glackin, C., 156, 199 Glassock, M. A., 308, 341,342, 346,355, 357,364 Clatter, O., 386,411 Glatz, J. F. C., 91, 149 Glennar, G . G., 363,368 Glomset, J. A., 266,296, 307, 308,365 Glonek, T., 389,411 Gmachi, M., 55,57, 58.82, 84 Goetinck, P. F., 95, 151 Goldberg, J., 375, 377, 402, 413 Goldberger, G., 25 1,302 Goldman, A., 5, 10,33,37,50, 311, 327, 334,365 Goldstein, J. L., 44,48, 207.246, 250, 256, 262,263.264, 265, 266,271, 277,295,296,297,300,301, 302, 307,365 Goldstein, S., 205, 215,245,246 Goldsworthy, G . J.. 373, 378, 401, 402, 404,411,412,415 Golmard, J. L., 344,365

427

AUTHOR INDEX

Gomez, F., 57,83 Gondim, K.C.,374,377,404,411 Gong, E.L.,352,354,365,386,412 Gonsim, K. C.,397,410 Gonzalez, M.S., 373,375,390,398,411, 412 Good, J.-J., 225,248 Goodey, A. R., 195,202 Goodman, D.S., 137,138,149,177,198 Goodson, T., Jr., 64,87 Gordon, D.A.,210,246 Gordon, J. I., 42,50,91,93,104,110, 113. 115, 119. 120,121,122,123, 133,134,136,142,144,145,147, 149,150,151,210,245,311,364, 377,379,410 Gordon, J., 121, 122, 141,149 Gordon, T., 307,366 Gordon, V.,259,296 Gorecki, M.,267,268,269,271,298,299 Gorin, M.B.,156,198 Gorrnsen, E.,47 Gotoda, T., 267,300 Gotohda, T., 267,302 Gotto, A. M.,385,411 Gotto, A. M.,Jr., 207,210,211,212,213, 214,215,226,245,248, 258,259, 266,272,294,296,300, 309,310, 323,324,329,341,342,344,346, 347,348,349,350,35 1, 356,358, 363,364,365,366,367,368,369 Gotz, F., 39,48 Goulinet, S.,214,218,223,225,226,245 Graham, D. L.,360,367 Graham, L.,240,245 Granade, M. E.,214,222,245 Grandour, R. D.,83 Grant, S. M.,210,246 Gratzer, W.D., 177,192,196 Gray, G., 39,50 Gray, J . E.,156,160,190,192,198 Grebenko, A. I., 99,151 Green, H. O.,178,198 Green, R.,37,50 Greenfield, M.R.,250,299 Greenman, B.,267,299 Greenwood, A.,39,49 Gregg, R. E.,255,257,267,269,280, 282,296,297,299,300 Griffin, R. G., 11,50

Griffith, O., 77,87 Griffiths, G.,90,93,104,150 Grigot, B. L.J., 382,388,390,391,412 Grochulski, P.,29,48 Groenendijk, C.,269,300 Groener, T.E.,308,366 Groom, C. R.,92,93,140,148 Gros, P.,58,66,83 Gross, E.,253,262,267,268,298 Gross, V., 171,199 Grosse, R.,93,104,148 Grundy, S. M.,208,248,339,366 Gu, Z.-W., 207,210,211,214,226,245, 248 Guidoni, A., 2.48 Guilhot, S.,41,49 Gulik-Kryzwicki, T.,226,246,387,409, 411 Gunsteren, W.F., 58,66,83 Gunthard, H. H., 353,354,365 Guo, L. S. S., 259,275,296,297 Gupta, K. B.,314,323,324,347,348, 350,362,365,368,369 Giirber, A.,166,198 Guthrow, C.E.,155,198 Gutierrez, J. M.,58,82 Guttler, F., 178,198 Guyer, M.,207,245 Gwynne, J. T.,250,300,309,360,361, 366

H Haas, M. J., 39,48 Haba, T., 307,369 Haberland, M.E., 217,246 Habib,G.,210,211,245 Hadvary, P.,48 Hadzopoulou-Cladaras, M.,209,210,245 Haefiger, D.N.,156,198 Hagag, N., 165,169,198 Hagarnan, K. A., 347,367 Hahn, P. F.,40,48 Haling, P.J., 2,50 Hallaway, B.J., 308,366 Halliwell, B., 194,198 Hallman, D.M.,257,280,297 Hamann, U.,264,297 Hamilton, J. A., 186,198,200,393,411

428

AUTHOR INDEX

Hamilton, R. L., 240,242,243,244, 256, 260,275,296,297,307,366 Hamlin, R., 75.86 Hammond, M. G., 245 Hamosh, M.,43,48 Hamsten, A., 209,245 Han, J. H., 45,49 Han, Q,,41,42,47 Hancock, A. J., 69,84 Handelmann, G. E., 250,297 Handwerger, S., 309,361,366 Hanks, J., 266,300 Hanneman, E., 104, 121, 131,151, 398, 413 Harad, W., 11,50 Harada, K., 267,300 Harder, K. J., 270,271,301 Harding, G. B., 177,202 Hardman, D. A., 207,210,226,235,245, 246 Harel, M., 5, 10,33,37,50 Harman, L.,309,361,366 Harmony, J. A. K., 250,262,274,276, 295,296,297,302 Harrick, J. H., 353,366 Harrington, W. F.. 176,198 Harris, T. J. R., 43,47, 48 Harrison, G. B., 240,247 Hart, B. J. T., 176,198 Hart, R. G., 166,199,260,300 Hartley, B., 11.47 Hartsuck, J. A., 167,201 Hanvood, J., 2,49 Hashimoto, Y.,39,49 Hasler-Rapacz, J. O., 208,247 Hasler-Rapacz,J., 210,247 Hassanien, F. R., 2,49 Hastrup, S., 44,47 Hata, Y.,2,7,49,50 Hatari, M., 357,365 Haunerland, N. H., 377,398,404,411, 413 Hauser, H., 69,83,217,246,339,366 Haut, S. M.,121,150 Havekes, L. M., 269,279,280,284,296, 300,301 Havel, R. J., 216,225,240,242,243,244, 247, 256,260, 266,269, 275,277, 297, 300,302, 305, 307,366, 388, 411, 413

Hawkins, D. J., 18,50 Hayakawa, Y.,399,400,401,411,414 Hayashi, K., 78,86 Hayasuke. N., 195,200 Hayek, T., 267,299 Hayes, G., 57,58,84 Hayes, S . B., 262,296 Hazelrig, J. B., 347,365 Hazzard, W. R., 256,277,302 He, N. B., 320,365 He, X.-M., 155, 160, 161, 164, 167, 168, 182,197,198,200 Heartsuck, J., 40,45,50 Hees, M., 255,277, 279,301 Hegele, R. A., 269,296 Heinrikson, R. I., 30,50 Heinrikson, R. L.,2,50, 54,57, 58.81, 83,85,86 Heinzmann, C., 41, 49, 207, 210,246,247 Heizmann, C., 45,46,49 Heldin, C.-H., 93, 104,148 Hellman, U., 93, 104, 148 Hernington, N. L., 393,410 Hemmings, B. A., 38,50 Henderson, R., 16,49,313,365 Henderson, T. O., 389,411 Hendrickson, W.A., 171,202 Hendrickson, W.,99,149 Hendriks, R., 66, 74,75,84 Henriksen, E.J., 374,375, 397,398,412 Hensel, C., 38,50 Herrnann, R. B., 64,87 Herr, F. M., 134, 146,149 Hershberg, R. D., 54,83 Herz, J., 264,265,266,296,297,301 Herzog, R.,57,83 Hesler, C. B., 262,296 Hession, C., 57, 58, 84 Hester, L., 347,348,367 Heuckeroth, R. O., 91,151 Hider, R. C., 54,82 Higliorini, M., 264,300 Higuchi, K., 208,210,246 Hilderman, H., 250,299 Hille, J. D. R., 76,87 Hinchcliffe, E., 195, 198 Hiraga, S., 269,298 Hiramatsu, K., 171, 196 Hiramori, K., 269,302 Hiraoka, T., 400,402,403,410,411

429

AUTHOR INDEX

Hirayama, Y., 398,411 Hirel, P.-H., 195,203 Hirose, N., 274,276,296 Hiroyuki, H., 210,247 Hitchcock, P. B., 69,83 Hitzemann, R. A., 195,198 Hixson,J. E., 252,297 Ho, J. X., 160, 183, 185,198 Ho, W. K. L., 90,150 Hoekema, A., 195,202 Hoekstra, W. P. M., 58,82 Hoff, H. F., 349,366 Hoff, J. H., 385,411 Hoffman, A. G. D., 398,401,411 Hoffman, R. D., 128,149 Hofmann, B., 6,39,49 Hofteenge, J.. 38,50 Hol, W. G. J., 58,60,66,76, 78,82,83 Holbrook,J. J., 275,296 Holcombe, K. S., 214,247, 260,261,298 Holden, H. M., 93, 104, 113, 122, 131, 132, 133,148, 341,342,357,364, 377,378,379,383,410 Holden, H., 92,93, 140,149 Holdsworth, G., 250,295 Holley, B., 38,50 Holm, C., 45,46,49 Holm, L., 192, 199 Holmes, M. D., Jr., 308,366 Holmgre, A., 353,366 Holowachuk, E. W., 156, 160, 183, 185, 198,199 Holwerda, D. A., 401,411 Honig, B., 16,50, 78,86 Honort, B., 187, I99 HWk, M., 275,300 Hope, M. J., 339,364 Hoppe, P. C., 133, I51 Hoppern, A. T., 178,200 Horie, Y., 279,280,296,297, 396,411 Horjales, E.. 197 Hornick, C., 307,366 Hors-Cayla, M. C., 207,246 Hospattankar, A. V.,210,246 Hospattankar, A., 210,246 Hosteenge,J., 38,50 Houck, C. M., 156,199 Howe, A., 398,413 Hradek, G. T., 307,366 Hresko, R. C., 128,149

Hsiao, H.-Y., 17,47 Hu, C. Z., 161, 176,198 Huang, B., 74,86 Huang, L.-S., 209,246 Huang, P. C., 179, 180,198,199 Hubbard, R. E., 39,49 Huber, R., 11,50, 92,93, 140, I49 Hubl, S . T., 209,232,248, 252,253,296 Hubner, W., 353,366 Hudson, B. S., 63, 77,84 Huge-Jensen, B., 2,6, 11, 35,47 Hugh, P., 307,367 Hugh-Jensen, B., 3,4,5, 17, 19,22,39, 47,49, 78,81 Hughes, T. A., 323,348,353,354,363 Hughes, W. L., 155, 161, 166,199 Hughes, W. L., Jr.. 166,197 Hui, D. Y.,45,48,49, 250,262,271,278, 295,298,297,302 Hui, J., 58,86 Hui, S . W., 320,365 Hultmark, D., 179,199 Humphries, S. E., 207,246, 269,299 Humphries, S., 209,248 Hunkapillar, M. W., 74,83 Hunt, C. R.,127,149 Hunziker, W., 3, 4, 9, 10, 18, 20, 42,48, 50,98,151 Huss, K., 162, 163,196,199 Hussain, M. M.,264,266,267,275,280, 297,298,299 Hutchinson, D. W., 162,196, 199 Huttunen, J. K., 209,248 Hwang, J.-K., 11,50 Hwang, K. Y., 6,39,49 Hwang, S. C., 341,342,351,365 Hwang, S.-L. C., 266,272,296

I Iadarola, P., 162, 163,.198,199,200 Iandolo, J. J., 39,49 Ibdah, J. A., 207,247,291,297 Ibdah, J., 347,365 Iberg, N., 186,199 Ichio, I., 205,246 Ignatius, M. J., 250,295,297 Ijzerman, A. P., 174,202 Ikai, A., 375,376,389,411 Ikeda, K., 78,86, 182,199,200,250,299

430

AUTHOR INDEX

lkehara, Y., 38,50 Imada, T., 187,202 Inada, Y., 187,202 Inazu, A., 262,296, 308,366 Ingraham, R.,54.84 Innerarity, T. L., 207,210,211,213,232, 245,246,247, 250,253, 257,258, 260, 261,262, 263,264, 266, 267, 268, 269,270, 27 1, 272, 274, 275, 277, 278,280, 282, 284, 290,297, 298,299,300,301,307, 359,366,368 lnoue, K., 57,84 lnouye, S., 39,47 lqbal, H., 363 Iqbal, M., 323,365 Irons, L.. 217,246 lrvine, R. F., 393,410 lsbir, T., 163, 198 Ishibashi, S., 267,302 lshikawa, Y.,21 1,248, 273,297 Ishioka, N., 163, 199 Isobe, T., 162, 163,202 Itakura, H., 210,247 lto, S., 307,369 ltoh, H., 262,297 Itoh, S., 225,248 Iwai, M., 2, 7,49 lwanaga. S., 58,66,84,88

J Jackson, C. L., 207,246 Jackson, K. W., 45,47 Jackson, R. L., 250,259,274, 276,295, 296,300,309,310,323, 324,346, 349,363,366,367,368,385.41 1 Jacobsen, C., 169, 199 Jacobsen, J., 171, 177, 188,199,201 Jacobsen, S., 180,200 Jacobson, S. F., 207,210,211,213,232, 246 Jaeschke, M., 261,277, 279,301 Jagodzinski, L. L., 156,199 Jain, M. H., 76,85 Jain, M. K., 58,63, 71, 76, 78,81,83 Jain, M.-K., 71,83 Jain, M., 43,48 Jakobs, C., 282,300 Jakoby, M. G., 141, 142, 143,149 James, M. N. G., 11,50, 61,63,85,86

James, S. J., 42,48 Jansen, E. H. J. M., 66,87 Janssen, L. H. M., 166, 179,196, 199 Janus, E. D., 269,301, 307,366 Jarnagin, A., 39,50 Jarvis, B. W., 128, 149 Jeanpierre, M., 207,246 Jensen, G. L., 42,49 Jeon, H. S., 6,39,49 Jessup, W. K., 215,232,247 Ji, 2.-S., 264,266,275,280,297 Jiang, R.-T., 74,86 Jiao, S., 308,366 Johansen, B.,57,58,84 Johansson, L. B.-A., 353,366 Johnson, B. J.. 323,346,348,366 Johnson, D. F., 210,245,247 Johnson, D., 210,211,213,232,246 Johnson, 1. D., 63,77,84 Johnson, L. A., 39.49 Johnson, L. K., 55, 57,58,83, 86 Johnson, L. L., 45,49 Johnson, P. F., 287.298 Johnson, P., 176,198 Johnson, R.A., 195,201 Johnson, W. J., 308,366 Jollts, J.. 156, 160,201 Jollts, P., 156, 160,201 Jonas, A., 250,302, 345, 352,355,356, 359,360,364,366 Jones, A. L., 216,225,247, 307,366, 388, 413 Jones, A., 130,150 Jones, J. L., 308, 323, 341,346,348, 352, 353,354,355,363,364 Jones, M. K., 212,248, 287, 293,300, 31 1, 314,329, 331,333,366,368, 378,413 Jones, N. D., 64,87 Jones, T. A., 92,93,97, 104. 108, 110, 115, 122, 126, 127, 130, 134, 135, 138, 139, 140,148,149,150 Jordn, K. J., 357,358,369 Jorgensen, E. V., 309,361,366 Jorgensen, S., 39,49 Jornval, H., 41.50 Jost, P. C., 63, 77,84,87 Julien, R., 26, 47 Junien, C., 207,246 Jurenka, R. A., 404,413

AUTHOR INDEX

Jurgens, G., 385,41I Justum, A. R.,373,411

Kabanov, A. V., 75,85 Kaffarnik, H., 250,269,282,300 Kahlon, T . S., 222, 223,246 Kaiser, E. T., 312, 313, 320, 346, 347, 348,349,350,355,365,366,367,369 Kaiser, I. I., 58,82 Kajinami, K., 308,366 Kakizoe, T., 154, 194, 199 Kakudo, M., 2,7, 11,49,50 Kalhone, J. B., 309, 361,367 Kalk, K. H., 5,33, 11,48, 49, 60,69, 74, 76,78,82,86 Kalman, M., 195,199 Kamada, M.,171,196 Kametani, T., 307,369 Kamphuis, L. G., 11,49 Kanai, M.,138, 149 Kanaya, H., 307,369 Kanda,T., 93,104,120,149,150 Kane, 246 Kane, C. D., 120, 142, I51 Kane, J . P.,207,210,226,235,245,246, 259, 260,269,277,296,297,300, 301, 305,366 Kaneko, S., 269,299 Kanellis, P.,323,346, 348,366 Kannel, W. B., 307,366 Kanost, M. K., 374,377,378,379,383, 402,410,411,413,414 Kanost, M.R.,287,296, 341, 342, 357, 364 Kao, H., 356,367 Kaplan, D., 262,302 Kaptein, J. D., 252,299 Kaptein, R.,58,66,82 Karathanasis, D. K., 207,245 Karathanasis, S. K., 251,296 Karathanasis, S., 251,302 Karlin, J. B., 266,272,296, 341, 342, 351, 365 Karplus, M., 69,85 Kashiwazaki, Y., 375,376,389,411 Kashyap, M. L., 305,366 Kashyap, P.M.,308, 341,342, 346, 355, 357.364

431

Katagiri, C., 373,375, 385,390, 392,411 Katase, H.,373, 374, 385, 397,398, 410, 411 Katchalski, E., 171,199 Katsube, Y., 7,50, 385,392,411 Katsuki, M., 267,300 Kaur, S . , 90,93, 104,150 Kaw, J. H.,341,342,357,364 Kawai, Y., 257,259,264,291,302 Kawakami, M.,267,302 Kawano, Y., 58.80 Kawasaki, H., 250,299 Kawashirna, S., 205,246 Kawauchi, S., 78,86 Kawooya, J. K., 376,377,378,381, 382, 383, 385,388,389,390,391,397, 402,404,405,411,412,414,415 Kay, C. M., 388,399,413 Kayser, H., 140,149 Kayushina, R., 385,411 Kazlauskas, R.J., 19.49 Kececioglu, J. D., 101, 150 Keim, P.S., 374,375,376,377,378,402, 411,412,413 Kelley, M. J., 58,84 Kempner, E. S., 42,48 Kendall, F. E., 167,199 Kendrew, J. C., 166,199,212,247, 310, 3 13,367 Kennedy, D. F.,58,84 Kenney, E., 195, I98 Kercret, H., 308, 341,346,352,355,364 Keresztes-Nagy, S., 179,199 Kerfelec, B., 26,27, 46 Kerret, H., 323,346,348,366 Kkzdy, F. J., 2,30,47,50, 57,58,81,85, 86, 216, 218,248, 312,313, 344, 347, 349, 355,365,366,367, 374, 375, 376, 378,381,382,383, 385, 388, 389,390,391,392,410,411,412,413 Kezdy, K., 345,352,355,356,360,366 Khan, M.Y., 172, 173,199 Khmelnitsky, Y. L., 75,85 Kihara, H., 58,76,87,88 Kim, H. K., 6,39,49, 122,149 Kim, S., 6,39,49 Kim,T. W., 207,211,212,215,226,245, 248 Kimura, J., 373,390,411 King, D. J., 43,47

432

AUTHOR INDEX

King, T. P., 173, 184,199 Kini, R. M., 66,84 Kinnunen, P. K. J., 77,80,86, 344, 358, 369 Kinnunen, P. K., 344,350,358,364 Kirby, A. J., 15,47 Kirchgessner, T. G., 41,45,46,49, 210, 246 Kirven, M. J., 266,296 Kisak, 1. V.,207,247 Kissel, J. A., 45,49 Kita, T., 307,365 Kitagawa, Y., 78.86 Kitamura, K., 90, 120, 126, 127,151 Kitamura, N., 57,84, 374,385, 394, 398, 410 Kitchems, R. T., 308,366 Kiyomoto, Y., 402, 410 Kjkllen, L., 275,300 Klatzmann, D., 195,203 Klibanov, A. M.,2.50 Klickstein, L. B., 38,49 Klinkert, M.,90,93, 104,150 Klopfenstein, W. E., 177, 199 Kloss, J., 55,58,86 Klotz, I. M.,189, 198 Klyacho, N. L., 75.85 Kneller, R.W., 308,366 Knipping, G. M.,385,411 Knopf, J. L., 54,81 Knott, T. J., 207,208,209,210, 211,213, 232,244,246,247 Knott, T., 208,209,246 Knudsen, J., 147,149 Kobayashi, K., 38,49 Kobori, S.,269,298 Kodali, D. R., 393, 411 Kodama, T., 210,247 Koh, S.-N., 183, 186,199 Koizumi, I., 308,366 Koizumi, J., 262,297, 307, 308,366 Kolattukudy, E.,39,50 Kolzumi, J., 262,296 Komada, M.,57,84 Komaromy, M.C.,41.49 Komaromy, M.,310,365 Komnick, H., 394, 398, 409 Kondo, T., 182,199 Koopman, W. J., 309,362,364 Koopmanschap, A. B., 374,410

Kordel, M.,6,39,49 Korn, E., 39,48 Korthals, J. S., 15,50 Koschinsky, T., 262,296,299 Kosik, I. M., 250,295 Kostka, V., 156,200 Kostner, G. M.,216,246 Koszelak, S., 167,197 Kotite, L., 269, 277,297,300 Kottke, B. A., 308,366 Kovanen, P. T., 277,300 Kovar, M.,294,300 Kowal, R. C., 264,265,266,296,297,301 Kraft, R., 93, 104,148 Kragh-Hansen, U., 155, 166, 180, 187, 199 Kramer, K. D., 15,50 Kramer, R. M.,54,57,58,64,84,87 Kranovich, D. J., 359,366 386,411 Kratky, 0.. Kraulis, P. J., 139, 150 Krauss, R. M., 216,246, 386, 412 Kraut, J., 11, 16, 17,49, 50 Krebs, K. E.,290, 291,297,299, 354,366 Kreil, G., 55,84 Kremer, J. M.H., 174,202 Kretsinger, R. H., 313,367 Kreusch, A., 99,151 Krieg, P., 90, 93, 104, 149 Krisans, S., 275,297 Kristofferson, A., 183, 196 54, 81 Kriz, R. W., Kroon, D. J., 346,347,348,349, 350,365, 367 Krstenansky, J. L., 347,367 Kruck,T. P. A., 180, 181,199 Krul, E. S., 272,298, 345, 356,360,366, 367 Kruski, A. W., 389,411 Kuchler, K., 55.84 Kudo, I., 57.84 Kugimiya, W., 39.49 Kuhne, C., 57,85 Kuipers, 0. P., 69,78,86 Kuipfers, 0. P., 66, 74, 75,84 Kuipfers, O., 58,82 Kulakosky, P. C., 404,412 Kunusoki, M.,11,50 Kuperberg, J. P., 349,367 Kurihara, H.,57.84

AUTHOR INDEX

Kurono, Y., 182,199,200 Kurtz, A., 93,104,148 Kuthiala, A., 375,412 Kuusi, T., 269,301 Kwak, J.-G,74, 75,82 Kyger, E. M., 45,49 Kyle, R. A., 162,163,200 Kyte, J., 344,367

433

Laure, C. J., 58,80 Laurel, C.-B., 162,196 Lauwereys, M., 4,10,33,50 Law, A., 212,215,232,246,247 Law, J. H., 2,47,340,341,368, 372,373,

374,375,376,377,37a,379,381, 382,383,385,388,389, 390,391, 392,394,397,398,402,403,404, 405,406,410,411,412,413,414,415 L Law, J.,92,93,140,149 Law, S. E.,210,246 Labro, J.F. A., 166,196 Law, S. W., 156,160,198,199,200,207, Lack, L., 178,198 208,210,245,246 Lackner, D., 210,246 Lawn, R. M., 156,199 Lagercrantz, C.,184,I99 Lawson, D. M., 3,5,17,19,22,39,47, 48, Laggner, P.,216,246,385,386,389,411, 49,78,81 412 Lawton, P.,58,81 Lagocki, P., 324,368 Laxer, G.,209,245 Lagrand, G., 2,49 Lazdunski, M.,60,80 Lagrange,D.,214,215,218,223,225, Lebherz, H.G., 210,246 226,245 LeBlond, L.,36 1,367 Laird, D.M.,128,148,149 Ledbetter, D. H.: 2,41,47 Lakusta, H., 188,199 Ledden, D. J., 173,198 Lalazar, A., 268,269,271,273,298 Lee, A. G., 79,81 Lally, P. A., 156,203 Lee, B. R.,210,211,245 Lalouel, J.-M., 269,296 Lee, B., 212,215,248,252,298 Land, M. D., 128,149 Lee, C.Y.,39,49 Landais, D., 195,203 Lee, D. M., 217,226,246 Landis, B. A., 250,275,298 Lee, F.-S., 210,211, 214,248 Landschulz, W. H., 287,298 Lee, N. S.,42,50 Lane, M. D., 128,148 Lee, N., 208,210,246 Lang, J.H.,179,199 Lee, P., 21 1,246 Langdon, R.G., 215,226,247,248, 389, Lee, S.-M., 235,237,238,248 415 Lees, R.S., 369 Lange, L. G., 45,49 Leff, T., 208,245 Langen, A., 93,104,148 Lefkowith, J., 121, 122,141.149 Langenbeck, U., 255,301 Lehmann, W., 93,104,148 Langlais, J.,54,84 LeMaitre, M.,195,203 Laplaud, P. M., 214,218,223,225,226, Lenz, C.J.. 374,375,396,410,414 245 Leob, G . I., 173,199 Laposata,M., 119,121, 133, 144,150 Lepault, J., 387,409 Largman, C.,75.81 Leroy, P., 134,149 LaRosa, J.C.,269,298 Lesczcynski,J. F., 20,49 Larson, S.,39,49 Lesk, A., 33,49 Larsson, T., 184,199 Lessof, M. H., 154,196 Laser, E. C.,179,199 Letiza, J. Y.,207,247 Lau,H.S. H., 347,355,367 Levanon, A. Z., 268,269,271,298 Lau, K., 207,226,245 Levashov, A. V.,75,85 Lau, S.-J., 180,181,199 Levin, M. S., 120, 121, 122, 134,149,210, Lauer, S. J., 252,253,261,267,296 245

434

AUTHOR INDEX

Levine, Y. K., 79,81 Levitt, D. G . , 110,113, 149 Levitt, M., 344,367 Levy, R. L., 352,365 Levy-Wilson, B., 207,208,209,210,21 1,

Lohse, P.. 269,298

Lok, C.M.,372,401,412

Lornonte. B.,58,82 Long-Fox, J.,58,81 Longsworth, L. F., 164,199 213,232,244,245,246,247,271,298 Lord, R.C., 169,197 Lewin, J., 166,199 Lorscheider, F. L., 164,165,169,200 Lewin-Velvert, U.,273,300 Loscalzo, J.,154,188,202 Lewis, B.,177,197,307,366 Low, B.W., 166,200 Li, E.,120,121, 122, 134,136, 141,142, Low, M. G., 374,375,412 143,149 Lowe, J. B.,119,121,133, 144,150 Li, W. H.,322,367 Lowe, M. E.,3, 16.25,41,50 Li, W.-H., 2,41,47,207,210,211,212, Lowe, P.A., 43,47,48 213,214,226,245,247,248, 251, Lown, J. S.,212,247 252,253,296,298,344,367,383,412 Luc, G.,214,218,223,225,226,245 Li, Y., 3,4,6,7, 8, 18,22,50 Lucey, M.,93,104,149 Liao, D.-I., 5,30,31,33,38,49 Ludescher, R. D., 63,77,84 Liao, K., 128,149 Ludwig, E.H., 208,209,244,246,248 Liberatori, J.,139,150 Ludwig, E.,250,296 Lie, E.,2,49 Luetscher, J., 172,200 Lirn, V. I., 310,312,367 Luft, A. J., 164,165,169,200 Lin, A. H.Y., 266,272,296,341,342, Lukas, D. S., 178,200 35 1,365 Lurn, P.Y.,401,402,403,410,412 Lin, A. Y.,54,81 Lund-Katz, S.,207,247,291,292,302, Lin, G.,57,69,84 349,353,367,389,412 Lin, I.-W., 38.49 Luo, C. C., 322,367 Lin, L.-L., 54,81 LUO,C.-C., 2,41,47,251,252,298,344, Lindblorn, G., 353,366 367,383,412 Lindgren, F. T.,222,223,246, 257,259, Luscornbe, M., 275,296 260,264,295 Lusis,A. J.,41,49, 207,210,211,213, Lindup, W. E.,181,199 232,245,246,247,252,299 Linehan, L. A., 156,203 Lusis, A. L., 57,83 Lingappa, V. R., 240,245 Lusis, L. J., 45,46,49 Linnernans, W.A. M., 385,394,414 Lussier-Cacan, S.,269,299 Linton, M. F.,235,236,237,248 Luthi-Peng, Q,, 26,50 Lioyd, R.S.,121,141,151 Lux, S., 352,365 Liprnan, D. J..101,150 Luzzati, V., 226,247,412 Lipowsky, R.,69,84 Lyn, S.,93,104,I49 Lis, L. S.,339,365 Lyons, A., 43,47,48 Lister, M. D., 69.80.84 Liu, H., 378,388,399,412,413 Liu, H.-C., 66,87 Liu, S.-W., 207,21 1, 226,248 Ma, Y.,207,209,247,248 Liu, S.-Y., 58,88 Maatman, R.G.H.J., 91,151 Ljungstedt, I., 169,202 Mabuchi, H., 262,297,296,307,308, Lo, T.B.,55,87 366,369 Locke, B., 120,121,134,149 MacDonald, P. N.,120,121, 122,144,150 Lockridge, D.,45,49 MacPhee-Quigley, K., 45,50 Loffredo, W., 57,69,84 Macrae, A. R.,19,47 Lofgren, H., 69.80 MacRae, I. C., 39.49

AUTHOR INDEX

Madison,J., 162, 163,196, 199,200,201 Maeda, H., 269,298 Maeda, N., 210,247,267,302 Magill, P. J., 307,366 Mahlberg, F. H., 308,366 Mahley, R. M., 359,366 Mahley, R. W., 207, 208,209,210,211, 213,214,232,244,246,247,248, 250,25 1,252,253,256,257,258, 259,260,261,262,263,264,265, 266,267, 268,269,270,271,272, 273,274, 275,276,277,278,280, 282,284, 285,286,287,288,289, 290,295,296,297,298,299,300, 301,302, 307, 341,343,344,345, 351,358,359,368,369,378,415 Maier-Borst, W., 194,202 Maig, R. F., 179, 199 Mailly, F., 269,299 Majumdar, S., 195,200 Maki, A. H., 76,84 Malcolm, B. R.,291,299 Malhotra, V., 378,412 Maliwal, B. P., 76,83 Mammi. M., 93, 104,150 Mandel, A., 78,86 Manely, R. V., 167,201 Manetta,J., 54,84 Mann, C., 262,296 Mann, W. A., 269,298,299 Manning,J. A., 90,150 Mantsch, H. H., 353,366 Mao, G. D., 195,200 Mao, S. J. T., 349,367 Mao, S. J., 308,366 76,84 Mao, S.-Y., Maraganore,J. M., 2,30,50, 57,85 Marcae, A. R.,2,50 Marcel, Y.L., 215,227,231,232,245, 247, 262,270,271,274,276,277, 296,301,359,361,367,369 Marcel, Y.,210, 211,213,232,246, 356, 367 Margalit, H., 3 11,364 Margolin, A. I., 2.50 Margolis, S., 215, 226,247 373, 410 Marinotti, 0.. Marotti, K. R.,252,299 Marquart, M., 11.50 Marreo, H., 354,367

435

Marsh, J. B., 205,247 Marshall, A., 38,50 Martin, B. M., 57,86 Martin, B., 313,320,369,389,415 Martin, R.B., 215,248 Martinek, K., 75,85 Martinez, C., 3,4,5, 10, 18, 22.27.28, 33,50 Martinez, P. F., 38,50 Martini, G., 313,368 Marz, L., 57,85 Mas-Oliva,J., 264,266,297 Mase, T., 3, 18, 39,40,50 Mason, R. W., 179,200 Mason, R.,69,83 Massaer, M., 45,50 Massey,J. B., 324,348,367,368 Masuda, H., 397,410 Matarese, V.,120, 127, 147, 148, I50 Matejschuk, P., 162,199 Mathyssens, G., 4, 10,33,50 Matsubara, Y.,93, 104,149 Matsuda, H., 374,377,404,411 Matsuda, Y.-I.,162, 163,200 Matsumoto, A., 210,247 Matsunaga, A., 269,299 Matsura, Y.,2, 7,49 Matsushima, T., 252,299 Matsuura, Y.,11,50 Matsuzawa, Y.,262,302 Matthews, B.W., 32,50 Matthews, D. A,, 11,50 Mattice, W. L., 320,368 Mauldin,J. L., 214,222,245 Maulet, Y.,45,50 Maurice, R., 232,247 Maury, I.,195,203 Maxfield, F. R.,264,266,297 Mayaux, J.-F., 195,203 Mayr, I., 92,93, 140,149 Mazzarella, B.,209,248 McCarthy, B. J., 208,209,210,211,213, 232,235,236,237,244,246,248, 278,300 McCarthy, S.G., 19,47 McClure, D. B., 64,87 McClure, R.J., 167,200 McConathy, W.J., 40,50 McCray, P., 57,58,84 McDowall, A. W., 387,409

436

AUTHOR INDEX

McGrath, M. E., 16,50 McGuire, C.B., 250,300 Mckeel, D. W.,133,151 McKeever, B. M., 167,197 McKenzie, H. A., 139,150 McKinley, M. P., 31 1,364 McKnight, S.L.,287,298 McLachlan, A. D.,156,160,169,192,

200,311,313,365 McLean, J. W.,251,252,253,296,299 McLean, L.R., 347,367 McMeekin, T. L., 166,200 McMenamy, R. H., 178,200 McNamara, P. M., 307,366 McPherson, A., 39,49,167,197 McPherson, J., 251,296,302 McQueen, E. G., 179,200 McQuillan, J. J., 119, 121,133, 144,150 McRee, D.,99,150 Meadley, M. F.,363 Means,G. E., 178, 182,183, 186,198,

199,202 Medzihradszky, D., 90,93,104,150 Medzihradszky, K., 90,93,104,150 Meglin, N.,210,246 Mehrabian, M., 207,247 Meighen, E. A., 18,37,48,50 Meister, W.,48 Melchoir, G.W.,252,299 Melik-adamyan, W.R., 99,151 Meloun, B., 156,200 Menarad, R., 17,50 Mendez, A. J., 342,361,367 Meng, M. S.,269,298 Menge, U., 3,4,17,47 Mengel, M. C.,245 Menju, M., 269,300 Menzel, H. J.. 257,280,297 Menzel, J., 261,277,301 Mercken, L.,45.50 Meredith, S. C.,30,50, 252,299,378,

381,382,383,411 Merritt, E., 99,149 Merutka, G., 57.85 Messerschmidt, A.,92,93,140,149 Metcalfe, J. C.,79,81 Metz-Boutigu, M.-H., 156,160,201 Meyer, E., 16,50 Meyer, T., 99,150 Meyers, W.C.,250,275,298

Meyers, W., 266,300 Michishita, I., 262,297 Michl, H., 57.85 Mihelich, E. D.,64,87 Milane, R. W.,359,369 Miles, C.M., 402,411 Mill, G. L.,205,247 Miller, C.G., 45,46,49 Miller, J. A.,55,58.86 Miller, J. M., 252,298 Miller, K. R., 121, 133,141, 142,143,

149, 150 Miller, K. W., 219,247 Miller, N. E.,262,299 Miller, N. G., 209,248 Miller, P. A.,209,246 Miller, R. J., 313,369 Miller, T.Y.,160, 161,167,168,197,200 Mills, G. I., 215,245 Mills, G. L.,205,2 15,227,23 1, 232,245,

246,247,262,270,271,274,276, 277,296,301 Milne, R., 210,211,213,232,246,247 Milona, N.,54.81 Mims, M. P., 252,273,298,299 Min, H. Y.,127,149 Minchiotti, L.,162,163,198,199,200 Minghetti, P. P., 156,200 Miranda, R. D.,275,280,297 Mishkin, S.,90,150 Mishra, V. K., 339,369 Misumi, Y., 38,50 Mitchel, A. G., 178,197 Miura, K.,373,375,377,412 Miyamoto, S.,307,308,366,369 Miyata, T., 58.88 Mizushima, H., 57,84 Mohandas, T., 45,46,49,207,247 Mok, T., 217,226,246 Mokuno, H.,267,302 Molin, G.,2,49 Monaco, H. L.,92,93,98,139,150 Monaco, H., 93,104,I50 Monge, J. C., 208,246 Montay, G., 195,203 Moore, M.N.,322,367 Moravek, L.,156,200 Moreau, H., 43,44,48,50 Morgan, A., 53.81 Mori, H.,269,298

AUTHOR INDEX Morinaga, T., 156, 160,200 Moritz, J., 178,198 Moriyama, K., 269,299 Morris, M. A., 155, 198 Morrisett, J. D., 252,273,298,299, 309, 310, 324,346, 349,350,363,364, 366,368 Morriss-Kay, G., 134, 149 Morrod, R. S., 388,413 Morton, R. E., 308,367 Moser, D., 90,93, 104,150 Moser, P., 161, 169,200,202 Moshitzky, P., 401,409 Moskaitis, J. E., 156, 198 Motzny, S.,282,300 Moulin, A., 43,50 Moulin, P., 308,366 Moulins, M., 398,410 Mudge, G. H., 178, 179,200 Muirhead, H., 166,201 Mukherjee, K. D., 2,49 Muller, K. W., 385,386,412 Muller, R.,140,149 Muller, T., 93, 104,148 Muller, W. E., 155, 178, 184,198,200 Muller-Fahrnow, A., 130,150 Mundall, E. C., 374, 375,376, 385, 390, 391,392,412 Muiioz, V., 290,296 Murase, N., 373,390,411 Murase, T., 250,267,302 Murgita, R.A., 154,200 Murry-Brelier, A., 195,203 Musacchio, A., 99,150 Mwangi, R. W., 373,401,412 Myant, N. B., 250,299 Myers, R. L., 269,296 Myklebost, 0.. 264,297 Myles, T., 162,196

N Naberhaus, K. H., 156, 160,202 Naeme, P., 363 Nagabhushan, T. L., 167,197 Nagao, E., 375,378,385,386, 388,402, 403,410,412

Nagaraj, R.,320,368 Nagase, S., 154,200 Nagashki. T., 225,248

437

Nagoaka, S., 171,196 Nagy, B. P., 208,247 Naik, D. V., 179,200 Najarian, R. C., 156,199 Nakagawa, S. H., 347,355,367 Nakagawa, Y., 195,200 Nakai, H., 207,246 Nakarnura, H.,269,298 Nakano, T., 57,84 Nakasone, S., 396,411 Nakayarna, A., 307,369 Namba, Y., 250,299 Narnbu, S., 269,302 Nandedkar, A. K. N., 180,200 Nanjee, N., 209,248 Naoka, 76,87 Napoli, J. L., 120, 134,149 Naray-Szabo, G., 11,50 Naud, M., 93,104,149 Navia, M. A., 167,197 Naylor, S. I., 38,50 Naylor, S. L., 156, 160,202, 207,245 Nazareth, R. I., 178,200 Neel, J., 162, 163,202 Neeley, J. R., 130,150 Nelen, T. H. A., 179,199 Nelson, D. A., 178,201 Nelson, D. R.,373,410 Nelson, G., 167,197 Nestel, U., 99,151 Nestruck, A. C., 269,299 Neven, L. G., 374,375,412 Newcomer, M. E.,92,93,97, 104, 108, 110, 115, 126, 127, 130, 134, 135, 138,139,140,148,149,150 Newhouse, Y. M.,278,300 Newrnan, M., 210,247 Newton, M.,45,50 Ng, T. C., 308,341,346,352,355,364 Nguu, E. K., 374,412 Nichols, A. V., 226,245,352,354,365, 386,388,412,414 Nicoll, A., 307,366 Nicosia, M.,210,246 Nie, S., 339,369 Nielsen, S. U., 186,197 Nikazy, J., 41,42,47, 48,50 Niki, H., 269,298 Nilsen, 0. G., 180,200 Nilsson, S. F., 138,150

438

AUTHOR INDEX

Nilsson, S., 45,46,49 Nilsson-Ehle, P., 307,367 Nishida, T., 323,363,364 Nishioeda, Y., 269,302 Niviera, P., 180,200 Nixon, J. S., 58,87 Noble, M., 99, 150 Noel, J. P., 58,74, 75.82 Noel, J., 57,69,84 Noel, J.-P., 43.50 Nolte, R. T., 209,210,245, 324,344,345, 367 Norden, J. J., 250,300 Nordhausen, R. W.,387,411 Nordlund, P., 64,87 Noriega, F. G., 396,412 Norskov, L.,3,4, 17.47 North, A. C. T., 92,93, 139, 140,148, 150, 166,201 Northwehr, S. F., 93, 104,149 Norton, E. J., 160, 183, 185, 198 Norton, S. E., 45,49 Norton, S., 39,50 Nowaczynski, W.,177,201 Numson, S. H.,160, 161, 167, 168,197 Nuney, E., 177,201 Nurse, C. E., 180,200 Nye, E. R.,269,301

0 OhUigin, C., 210,211,247 OKonski, C. T., 161,202 OKonski, T. 0.. 169,200 Obrink, B., 137,150 Ochanda, J. O., 374,412 Ocklind, C., 137, 150 Ockner, R.K., 90, 133,150, 151 Oda, 76,87 Odani, S., 93, 104, 149, 162,202 Odera, M., 39,50 Odya, C. E., 38,50 Oefner, C., 5, 10, 33,50 Oers, N. S. C., 154,200 Oester, Y.T., 179, 199 Oeswein, J. Q., 389,412 Ogata, S., 38,50 Ogura, T., 269,298 Ohlendorf, D. H.,167,197

Ohno, M., 58, 76,87,88 Ohsawa, N.,267,300 Ohta, M., 307,369 Oikawa, K., 388,399,413 Oiwake, H.,307,369 Okabayashi, K., 195,200 Okada, M., 269,298 Okamura, E., 353,367 Olaisen, B.,251,299 Olembo, N. R., 374,412 Olivecrona, T., 18,41,42, 45.48, 50, 308, 369 Oliveira, P. L., 374, 377,404,411 Oliver, J., 39,49 Ollis, D. I., 33, 37, 50 Ollis, D., 5, 11, 33, 50 Olofsson, S . V., 209,210,212,247 Olofsson, L O . , 208,240,245 Olphin, S. E., 362,367 Omori-Satoh, T., 78,86 Onasch, M.A., 208,209,244,247 Oncley, J. L., 164, 178,200 0ng.D. E.,91, 120, 121, 122, 134, 144, 146,149,150 Ono, T., 93,104,120,149,150 Oram, J. F., 130,150, 307,308,342,360, 361,367,368 Oratz, M., 155,201 Orchard, I., 401,412 Ordovas, J. M., 267,300 Oriaku, E., 58.86 Orlowski, R. C., 320,365 Ortego, S., 377,411 Ortiz, A. R., 71.85 Osborne, J. C., 42,50 Osguthorpe, D. J., 58,81,86 Osir, E. O., 374,404,411, 412 Osterman, D. G., 313,369 Otani, Y.,39,49 Otomo, E., 250,299 Ottewill, R. H., 176, 198 Otto, A., 93, 104,148 Otwinowski, Z., 54, 59,60,61,62,64,65, 66,67,68,69,70, 71, 72,73, 76.77, 78, 79,80,85,87 OU,S.-H., 273,298 Owen, T. J., 347,367 Owens, R. J., 309,350,361,362,367,368 Ozdemir, Y., 163,198 Ozeki, J. L., 182,200

AUTHOR INDEX

P Pahler, A,, 99,149 Paik, Y.-K., 251,299 Paksay, K. S . , 210,245 Palm, W., 11,50 Palmer, L., 179,199 Pan, M.-L., 373,404,413 Pantoliano, M. W., 17,47 Panveliwalla, D., 177,197 Papiz, M. Z., 139,150 Parhofer, K. G., 307,367 Paris, R., 398, 410 Parks, J. S., 186,200 Parry, D. A. D., 313,364 Pasad, S. V., 374, 375, 397,398,412 Pascher, I., 69,80,83,69,85 Pascher, J., 339,366 Pastor, R. W., 69,85 Pathak, D., 5, 11,33,50 Patkar, S. A., 3,5, 17, 19,22,47, 78,81 Pattnaik, N. M., 374,375,576,385, 390. 391,392,412 Pattus, F., 344,358,369 Paubel, J.-P., 180,200 Paul, W. L., 179,200 Paulus, H. E., 210, 235,246 Paulweber, B., 208,247 Pauptit, R., 99,150 Peach, R. J., 162, 163,196,200 Peach, R., 162,196 Peacock, R., 209,245 Pearlman, W. H., 178,200 Pearson, R. H., 69,83,85 Pearson, W. R., 169,200 Pease, R. J., 207,210, 211,213,215,232, 240,246,247 Pease, R., 208,209,244 Pedersen,A. 0.. 186, 187,197,199 Pedersen, K. O., 171, 180, 188,201 Pedersen, P. V., 179,202 Peeters, R. A., 91, 120,150, 151 Peled, Y.,401,414 Peluffo, 0. R., 375,390, 391,398,404, 412,413 Pepe, M. G., 250,299 Pepinsky, R. B., 57,58,84 Perez, L. E., 21 1,247 Pericak-Vance, M., 250,300 Pernarowski, M., 178.197

439

Perrin, J. H., 178,201 Person, R. H.,339,366 Persson, A., 2,49 Persson, B.,41,50 Perutz, M. F., 166, 167, 168,202, 310, 313,367 Perutz, M.,212,247 Peters, T., Jr., 154, 155, 156, 160, 164, 166, 167, 171, 172, 180, 181, 186, 187, 188,189,191,201 Peterson, P. A., 92, 137, 138, 139,150 Peterson, P., 93, 104,151 Peuko, G. A., 33,48 Pfleger, B., 308,366 Pflugrath, J., 92,93, 140,149 Philips, H. C., 389,412 Phillips, D. C., 166,199 Phillips, M. C., 207,247, 259,290,291, 292,296,297,299,302, 308, 347, 353,354,365,366,367 Phillips, M. L., 227,231,232,235,237, 238,245,,247,248 Phillips, S. E. V., 92,93, 140,148 Phizakerley, R., 99,149 Piedrahita, J. A,, 267,302 Pieroni, G., 43,47,48,50 Pierotti, V. R., 208,209,244,246, 252, 253,296 Pies, A., 269,300 Pieterson, W. A., 54,60,80,85 Pillion, D. J., 309,362,364 Pilon, C., 41,49 Pines, M., 401,412 Piot, F., 356,367 Pirjo, P., 209,248 Pisabarro, M. T., 71,85 Pisano, J. J.. 313,320,364 Pitas, R. E., 250,253,262,263,267,268, 290,295,297,298,299 Pitzner, R., 208,248 Plant, S., 55,58,86 Plump, A. S.,267,299 Pohling, C., 38,50 Polgar, I., 38,50 Polites, H. G., 252,299 Pollack, P. S., 363,368 Pollard, H., 226,247 Ponsin, G., 347,348,367 Popp, F., 39,48 Poppenhausen, R. B.,90,150

440

AUTHOR INDEX

Porta, F. A., 162,201 Porta, F., 162,198,201 Possani, L.D.,57,86 Pottenger, L.,215,248,389.415 Poulos, T.,17,47 Poulose, A.J., 39,50 Powell, K.,167,197 Powell, L.M.,207,210,211,213, 232,

245,246,247 Powell, L., 208,209,244 Power, S., 39,50 Pownall, H.J.. 40,50,207,211,226,248,

250,259,300.302,324,346,347, 348,349,350,363,366,367,368 Poznansky, M.J., 195,200 Prack, M.M.,250,300 Prasad, S . C.,266,272,296,341,342, 351,365 Prasad, S.V.,373,374,377,394,396, 405,406,412,414,415 Price, C. P., 362,367 Price, H. M.,398,413 Priestley, L.M.,207,246 Primrose, W.U.,58,66,72,82,86 Protter, A. A.,209,210,212,247 Provost, P., 215,232,247,356,367 Pruin, N.,277,279,301 Prusiner, S.B., 31 1,364 Pruzanski, W., 55,57,58,83,86 Puijk, W.C.,55,77,85,87 Puppione, D.L.,214,235,237,238,247, 248 Putnam, F. W., 162,163,196,199,200, 202 Putnam, J. E.,54,64,84,87 Puyk, W.C.,66.87

Qian, S., 121,122,141,149 Qian, S.-J., 121,122,134,136,149 Quardfordt, S.,309,361,366 Quarfordt, S. H., 250,251,260,266,274,

275,296,298,299,300 Quarles, R. H.,126,151 Quill, S. G.,17,47 Quin, P. J., 339,365 Quinn, D.M.,67,88 Quinn, P. J.. 393,410 Quirk, A. V., 195,201

R Raag, R., 98, I50 Radhakrishnamurthy, B., 275,300 Radhakrishnan, R., 16,50 Radke, S. E., 18,50 Radvanyi, F.,76,87 Rafai, E.,356,367 Rajavashisth, T.B., 252,299 Rakusch, U.,216,246 Rall, L. B., 207,246 Rail, S. C., 341,343,344,358,359,366,

368,369 Rall, S. C.,Jr., 207,210,211,213,232, 246,251,252,253,257, 258,259, 267,268,269,270,271, 272,274, 276,277,278,279,280, 284,296, 297,298,299,300,301,302 Ramachandran, J., 401,409 Ramakrishnan, B., 74.86 Ramakrishnan, R., 18,48 Ramdas, L., 252,298 Ramesha, C.S., 54,81 Ramirez, F.,76,85 Ranadive, G.B.-Z., 58,83 Rand, R. P., 393,410 Randolph, A., 54,85 Ransac, S., 43,48,50 Rao, S. N., 167,201 Rapacz, J., 208,210,247 Rapp, G.W.,179,201 Rappaport, A. T.,19,49 Rask, L.,92,93,104,137,138,139,150, 151 Rassart, E., 232,247,356,367 Raston, G.B., 139,150 Rathe, J., 57.83 Rathelot, J., 26,47 Rawlings, N. C.,38,50 Raykova, D.,57,76,85,87 Rayment, I., 92,93,140,149,341,342, 357,364,377,378,379,383,410 Raz, A., 138,149 Read, R. J., 11.50 Read, R., 61,85 Reardon, C. A.,25 1,299 Reardon, I. M.,58,86 Reddick, R. L.,267,302 Reddy, M.N.,30,50 Reed, G . H.,54,83

AUTHOR INDEX

Reed, R. G., 186, 186,198,201 Reinheimer, J. D., 183, 198 Remington, S.J., 5,30,3 1,33,38,37, 49,50 Renaud, G., 307,366 Renetseder, R., 55,58, 59,60,67,78, 82.85 Renges, H. H., 209,245 Rennex, D., 38,50 Requandt, C., 39,50 Reuben, M. A., 209,210,212,245, 247 Reue, K. L., 252,299 Reyland, M. E.,250,300 Reynolds, J. A., 217,246 Rhee, S. G., 53,81 Rhonde, K., 93,104,148 Ribeiro, J. M. C., 372,412 Richards, E. G., 259,296 Richards, F. M., 166, 169,200,201 Richards, J. H., 74,83 Richardson, D. C., 108,150,337,368 Richardson, J. S.,35.50, 108,150, 337, 368 Richardson, K. S. C., 177,201 Rietsch, J., 26,50 Rimoldi, 0.J., 375,390,398,412 Riomoldi, 0. R., 390,391,404,413 Riomoldio, D. J., 390,413 Ripps, M. E., 209,246 Riviere, C., 43,47, 48 Ro, J. H.-S., 127,149 Robbins, J., 178,202 Robbs, S. L., 374,412 Roberts, C. J., 38,50 Roberts, E. F., 54,84 Roberts, G. C. K., 58,66, 72,82,86 Roberts, K. D., 54,84 Roberts, M. F., 58,79,85 Robertson, E., 207,246 Robertson, M. A., 269,296 Robertus, J. D., 17,50 Robinson, D. H., 72.86 Robinson, M. S., 178,200 Robinson, R. M., 320,368 Robson, B., 344,365 Roczniak, S.,75,81 Roda, A., 177,201 Roda, E., 177,201 Roelosen, B., 76,87

44 I

Rogers, J., 71,76,78,81,83 Rogne, S.. 264,297 Romans, A. Y.,323,346,348,366 Ronne, H., 137,138,150 Ropson, I. J., 144, 145,150 Rosa, J. J., 59,60,62,63,67,71,72,73, 78,86 Rose, G. D., 20.49 Rose, G., 147,148 Rosen, A.. 178, 179,196,201 Rosenbaum, L. M., 44,50 Rosenblatt, M., 320,365 Roses, A. D.,250,300 Ross, P. D., 155, 171, 187,201 Rosseneu, M.,.210,211,213,214,245, 248,356,367 Rossi, A., 162,201 Rossing, T. H., 152,198 Rossman, M. G., 167,201 Rossmann, M. G., 99,151, 166,201 Rostron, P., 259,296 Roth, J., 178,202 Roth, R. I., 259,300 Rothblat, G. H., 259,296,308,366 Rothman, J. H., 38,50 Rothschild, C., 121, 122, 141,149 Rothschild, K.J., 354,367 Rothschild, M.A., 155,201 Rotolo, F. S.,250,275,298 Rousseaux, J., 162,196 Rovery, M., 2, 10,47.48 Rozeboom, H. J., 5,33,48 Ruberte, E., 134,149 Rubin, E. M., 267,299 Rubin, J., 31 1,365 Rubingh, D., 39,49 Rucknagel, D. L., 163,202 Rudel, H., 130,150 Rudy, B., 313,364 Rugni, N., 3,5, 10, 18, 22,27,28, 50 Russell, D.W., 271,302 Russi, E., 154,201 Rutter, W. J., 45,49, 75,81,86 Ryan, R. O., 372,374,375,376,377,378, 379,382,385,388,397,398,399, 402,403,405,406,410,411,412, 413,414,415 Rye, K. A., 308,368 Rypniewski, W., 92,93,140,149

442

AUTHOR INDEX

Sacchetti, F.,169,197 Sacchettini, J. C., 42,50, 91, 93, 104, 110, 113, 115, 119, 121, 123, 133, 134, 142, 144,145, 147,149,150,151 Saenger, W., 130,150 Saha, N., 209,245,257,280,297 Saiansbury, M., 58,81 Saita, S., 58,88 Saito, Y.,187,202 Sakaguch, A. Y.,156,203 Sakaguchi, A. Y.,38,50,207,245 Sakai, K.. 19.50 Sakai, M., 156, 160,200 Sakai, N., 262,302 Sakai, T., 262,297 Sakai, Y.,262,297 Sakamoto, Y.,162, 163,200,201 Sakmar, T. P., 54,85 Salahuddin, A., 173,199 Salemme, F. R., 167, 197 Salvesen, G. S . , 250,300 Samaraweera, P., 377,378,402,411 Samejima, Y.,78,86 Sammett, D., 308,369 Sandblom, P., 115, 138,148 Sander, S., 192,199 Sandholzer, C., 257,280,297 Sando, G. N., 44,47,50 Sanny, C. G., 167,201 Santos, E. C., 178,202 Saraste, M.,99,150 Sarda, L., 2,3,5, 10,20,26,27,41,47, 48,50 Sargent, T. D., 156,199,201 Sari, J. C., 178,197 Sarkar, B.J., 188, 191,197 Sarkar,B., 180, 181, 188, 189,197,199 Sarma, R.,39,50 Sasak, W. V., 212,247 Sasaki,J., 269,299 Sasaki, N., 250,295 Sam, T., 216,225,247,260,297,388,413 Sato, K., 171,196 Sato, M., 385,390,392,41 I Satow, Y.,99,149 Saunders, A. M.,250,300 Saunders, N. R., 156,197 Savu, L., 177,201

Sawinski, V. J., 179,201 Sawyer, L., 92,93,98, 139,150 Scanu, A. M.,215,216,218,226,244, 247,248, 290,291,300, 324, 344, 357,365,368, 388,389,392,410, 411,413 Scapin, G., 92,93, 104, 1 10, 130, 133, 134, 138,139,145,150,151 Scatchard, G., 164, 176, 180, 184,200, 201 Schaefer, E. J., 267, 269,282,297,300, 301 Schanck, K., 39,47 Scheraga, H. A., 173,199 Schilling, J. W., 207,226,245, 250,297 Schilling, U.,194,202 Schiltz, E., 99, 151 Schl, L.C., 374,375,412 Schlaper, P., 209,248 Schleier, K. H.,39, 48 Schmauder-Chock, A., 53,81 Schmechel, D., 250,300 Schmid, R.,6, 39,49 Schmidt, C. F., 323, 348,353,354,363 Schmidt, J. O., 374, 375, 376,412 Schmitz, G., 44,50 Schneider, H.,388,413 Schneider, M., 92,93, 140,149 Schneider, W. J., 271,277,300,302 Schoenberg, D.R., 156,198 Schoenborn, W., 277,279,301 Schoentgen, F., 156, 160,201 Schomburg, D., 6,39,49 Schonfeld, G., 256, 277,302, 308,356, 366,367 Schotz, M.C., 40,41,42,45,46,47, 48, 49.50,272,275,298,296,307,367 Schrag, J. D., 3,4,6,7,8, 16, 18, 22, 37,50 Schrag, J., 29, 33,37,48,50 Schramrneijer, B., 195,202 Schrank, B., 139,148 Schrenk, H.H.,194,202 Schuetter, R. J., 320,365 Schulman, S. G., 174, 179,200,202 Schultz, T. K. F., 379, 402, 403, 413, 414 Schulz, G. E.,99,151 Schulz, M.,373,415 Schulz, T. K. F., 376,390,400,413

AUTHOR INDEX

Schumacher, M.,45,50 Schumaker, V. N., 207,208,209,214, 216,217,226,227,231,232,235, 236,237,238,240,243,244,245, 247,248 Schwartz, E., 310,365 Scott, D. L., 54,59,60,61,62,63,64,65, 67,68,69,70, 71, 72,73,76, 77,78, 79,85,86,87 Scott, D., 58,64,66,69,80 Scott, J., 207, 208, 209,210,211,212, 213,215,232,240,244,245,246,247 Scott, T. M.,208,248 Scraba, D. G., 388,398,399,413 Sebille, B., 178,201 Sedzik, J., 108, 122, 126, 149 Seebart, K. E., 4 1,42,50 Seeburg, P. H.,156,199 Segrest, J. P.,212,248, 287,293,300, 308,309, 310,311,313,314,322, 323, 324,329,331,333,339, 340, 341, 342, 343,346,347, 348,349, 350,352,353,354,355,357,359, 360,361,362,363,364,365,366, 367,368,369,378,413 Segura, R., 385,411 Sehayek, E., 273,300 Seilhamer, J. J., 55,57,58,83,86 Sekharuda, C., 74,86 Semenkovich, C. F., 41,48 Semeriva, M.,9,50 Senadhi, S. E., 167,197 Senadhi, V.-K., 167,197 Senthilathipan, K. R., 398,413 Sessions, R. B., 58,81,86 Setzer, D., 212,245 Sha, R. S., 120, 142,148,151 Shaklai, N., 186,201 Shapiro, B., 147,148 Shapiro, J. P., 340,341,368, 372,375, 376,377,378,391,392,402,411,413 Sharp, A. M.,3,37,48,50 Sharp, J. D., 64,87 Sharp, P.M.,207,211,226,248 Sharples, D., 179,201 Shaw, C. F., 188,201 Shaw, D. C., 139,150 Shaw, E., 38.50 Shechter, I., 210,245 Shekels, L.L.,128,149

443

Shelburne, F. A., 250,251,260,266,274, 299,300 Shen, B. W., 216,218,248,290,291,300, 344,365,388,389,392,413 Shen, B., 389,410 Shen, M. M.S., 222,223,246 Sherrill, B. C., 266,300,307,368 Shevitz, R. W.,64,87 Shew, B., 39,50 Shibasaki, Y.,210,247 Shieh, T.-C., 76,87 Shields, H. M.,133,151 Shiffer, J., 310, 311,368 Shimada, M.,267,300 Shimada, Y.,7,50 Shimamune, K., 154,200 Shimano, H., 267,300,302 Shimizu, A., 162, 163,196 Shimizu, I., 373,375,377,412 Shinto, L. H., 284,285,301 Shipley, G. G., 69,83,216,245, 352,364, 369 Shireman,'R. B., 21 1,248 Shively,J. E., 41,45,46,47,49 Shockley, P.,155, 160, 161, 166, 181, 186, 191,197,201 Shooter, E.M.,250,295,297 Shore, B., 255,259,260,300, 305,366 Shore, C., 166,199 Shore, V. G., 255,260,300, 305, 359, 360,365,366 Shore, V., 259,260,300 Shotz, M. C., 41,47 Shows, T. B., 207,246 Shrake, A., 155,171,187,201 Shreiber, S. S., 155,201 Shumiya, S., 154,200 Sibanda, B. I., 32,50 Sidler, W.,48 Sigler, P.B., 54,55,58,59,60,61,62,63, 64,65,66,67,68,69, 70.71, 72, 73, 76, 77, 78, 79,80,81, 85,86,87,88 Sijmons, P. C., 195,202 Silberman, S. R., 210.21 1,245, 356,368 Silman, I., 33,37,50 Silman, L., 5, 10,33,50 Simons, M.J., 45,50 Sing, C. F., 255,257,280,282,296 Singel, D.J., 154, 188,202 Single, D. P., 339,365

444

AUTHOR INDEX

Sinn, H., 194,202 Sioler, G. F., 349, 350,368 Sippl, M. J., 55,84 Sisson, P., 250,266,280,301 Sitaram, N., 320,368 Siu, C., 93, 104, 149 Sivaprasadarao, R., 139, I50 Sjoholm, I., 27.48, 169,202 Skarlatos, S. 1.. 269,298 Skene, J. H. P., 250,297 Skipski, V. P., 225,248 Skov, K. W., 39,49 Slaich, P. K., 72,86 Slayter, E. M., 164,202 Sleep, D., 195,202 Slotboom, A. J.. 53,54,58,60,64,66,76, 77,82,83,84, 87 Slotte, J. P., 308,368 Small, D. M., 186,200, 216, 217,219, 245,247,248,352,369,393,411 Smallcombe, S. H., 74,83 Smillie, M., 398,410 Smit, M., 269,300 Smit, R. S., 356,365 Smith,A. F., 93, 104, 113, 121, 122, 131, 132, 133,148,151,383,398,410,413 Smith, A. J., 120,148 Smith, C. D., 167, 197 Smith, H. M., 352,364 Smith, J. A., 38,49 Smith, J. D., 267,299 Smith, J., 99,149 Smith, L. C., 3,40,41,42,48,50, 349, 350,356,366,368 Smith, R. S., 274,296, 360,364 Smith, S. O., 11.50 Smith, W., 167,197 Smithies, O., 210,247 Snipes, G. J., 250,300 Snyder, R. S., 167,197 Sokoloski, T. D., 178,200 Sollene, N. P., 178, 182, 186,202 Soma, M.R., 273,299 Somerharju, P., 80,86 Soravuia, E.,313,368 Soria, L. F., 208,248 Soria, L., 209,246 Sosa, B. P., 57,86 Soulages,J. L., 373,390,391, 392,393, 404,411,413

Soutar, A. K., 349,350,368 Spadon, P., 92,93,98, 104, 130, 138, 139, 150,151 Sparkes, R. S., 45,46,49, 207, 209,247 Sparks, C. E., 290,299 Sparrow, D. A., 210,211,214,248, 294, 300 Sparrow, J. T., 210,211,214,248, 270, 271,274,276,277, 294,300,301, 324,329,344,346,347, 348,349, 350,356,358,359,364,366,367, 368,369 Specher, D. L., 269,299 Spector, A. A., 177, 178, 186,202 Spector, M.S., 357,369 Spencer, I. M.,401,413 Spencer, M., 184,199 Spener, F., 130,150 Spiegelman, B. M., 127,149 Spiekerman, A. M., 156,196 Spink, C., 27,48 Spouge, J. L., 311,364 Sprang, S., 75,86 Sprecher, D. L., 262,302 Spring, D. J., 235,236,237,238,240, 243,248 Squire, P. G., 161, 169,200,202 Srinivas, R. V., 309,350,361,362,367, 368 Snnivas, S. K., 314, 324, 348.350, 362, 368 Srinivasan, S. R., 275,300 Stable, H. Z., 57,69,84 Stalenhoef, A. F. H., 269,300 Stalfors, P.,45,47 Stamler, J. S., 154, 188,202 Stammers, D., 167,197 Standing, T., 75.86 Stangl, A., 139,148 Stanley, K. K., 264,297 Stanley-Samuelson, D. W., 404,413 Stark, D. H., 64,87 Stark, D., 282,297 Stein, E. A., 269,298 Stein, L., 90, 150 Steinberg, D., 262,296,299 Steiner, R. F., 178,202 Steinmeu, A., 250, 255,269, 277,279, 282,300,301,354,369 Steitz, T. A., 3 11,327,334,365

445

AUTHOR INDEX

Stemberg, M.J. E., 169,203 Stevens, T. H., 38,50 Stibitz, G. R., 178, 179,200 Stiers, D. L., 217,226,246 Stievenart, M.,57,83 Stoffel, W., 208,248 Stone, S. R.,38,50 Stoppini, M.,162, 163, 198,200 Storch, J., 93, 104, 110, 122, 134, 135, 147,149,151

Storer, A. C., 17,50 Stow, J. L., 275,300 Strachan, A. F., 363,364 Stralors, P., 45.50 Strandberg, B. E., 166,199 Stratowa, C., 45,49 Straws, J. F., 360,366 Strickland, D. K., 264,300 Strittmatter, W. J.. 250,300 Strobel, L. M.,374,377,402,413 Stroud, R. M.,75,86,3 11,345,365 Strynadka, N. C., 63,86 Stump, D. G., 121, 141,151 Stuyt, P. M.J., 269,300 Sudlow, G., 179, 181, 182,202 Suemune, H., 19,50 Sugihara, A,, 2,7,49,50 Sugimura, T., 154, 194,199 Sugita, O., 162,202 Suh, S. W., 6,39,49 Sultzman, L. A., 54,81 Sumida, M.,394,412 Sumiyoshi, T., 269,302 Sundarlingam, M.,74,86 Sundelin, J., 93, 104,151 Sundell, S., 69,80,83, 339,366 Sunderman, F. W., Jr., 180,197 Surholt, B., 375,377, 402,413 Sussmaan, J. L., 5, 10,33,50 Sussman, F., 11,50 Sussman, J., 33,37,50 Suter, F., 140, 149 Suter, M., 57,58,82 Sutton, A. D., 195,201 Suzuki, M.,90, 120, 126, 127,151 Suzuki, T. C., 104, 121, 131,151, 398, 413

Svenson, K. L., 45,46,49, 210,246,247 Svenson, K., 41,49 Sverdgun, D. I., 386,410

Swarte, M. B. A., 11,49 Sweester, D. A., 133,151 Sweet, R. M.,5,6,30,31,33,38,39,49 Sweetster, D. A., 91,151 Swenson, L., 37,50 Swilens, S., 4 5 5 0

T Tabaqchali, S., 177,197 Tabas, I., 264,266,297 Tainer, J., 99,150 Tajima, S., 257, 259,264, 269, 291,300, 302

Takabayashi, K., 187,202 Takada, Y.,39,49,269,299 Takahashi, K., 374,385,398,402,404, 410

Takahashi, N., 162, 163,199,202 Takahashi, Y.,162, 162, 163,202 Takaku, F., 267,300,302 Takasu, N., 57,84 Takayama, M.,225,248 Takeda, M.,262,297 Takeda, R., 262,296,297,307,308,366, 369

Takenaka, T., 353,367 Takeuchi, K., 39,50 Tall, A. R.,262,296,301, 308,352,366, 369,398,413

Talmud, P. J., 209,248,269,299 Tamaoki, T., 156, 160,200 Tamm, L. K., 354,365 Tanaka, N., 2,7,49, 385,390,392,411 Tanford, C., 31 1,369 Tang, J., 45,47 Tang, L. C., 53,81 Taniguchi, T., 21 1,248 Tanimizu, I., 225,248 Tanimura, M.,210,211,212,214,245, 248,251,298,344,367,383,412

Tapia, O., 115, 138,148 Tardieu, A., 226,247,412 Tarnoky, A. L., 161, 162, 172,198,202 Tarui, S., 262,302 Tatarnim, R., 307,369 Tattrie, N. H.,388,413 Taunton, 0.D., 385,411 Taylaur, C. E., 205,215,245,247 Taylor, E. W.,226,247

446

AUTHOR INDEX

Taylor, G., 167,197 Taylor, J. M., 251,252,261,267, 271, 296,298,299,3 1 1,364

Taylor, J. W., 313,320,369 Taylor, P., 45,50 Taylor, S . M., 235,236, 237,248 Taylor, S. S., 45,50 Taylor, W. H., 210,211,213, 232,246 Taylor, W. R., 169,203 Teater, C., 64,87 Teeter, M. M., 171,202 Teisberg, P., 251,299 Teitz, A., 398, 414, 415 Tekeuchi, K., 3, 18,39,50 Telfer, W. H.,373,404,412, 413 Tendler, M., 90,93, 104,150 Tennekoon, G., 128,149 Terwilliger, T. C., 3 10,3 1 1, 345,365 Teshima, K., 78,86 Theolis, R.,Jr., 232,247 Thim, L., 2,s. 4, 5 , 6 , 11, 17, 19.22, 35, 47, 78,81

Thiry, M., 39.48 Thomas, K. M., 69,83 Thomas, T., 207,247 Thompson, M. R., 161,202 Thornton, J. M., 32,50 Thorpe,S. R., 155,183,197,198 Thrift, R. N., 236,248 Thuaud, N., 178.201 Thunnissen, G. M., 66,84 Thunnissen, M. M. G. M., 60,69, 74,

Tornqvist, H., 45.47 Toxopeus, E.,64,80 Trambusti, B. G., 374,375,376, 385,390, 391,392,412

Trapp, B. D., 126,151 Trawick, J. D., 236,248 Triantaphylides, C., 2,49 Triplett, R.B., 215,248 Troxler, R.F.,251, 252,301 Tsai, LH., 55, 66,87 Tsai, M.-D., 57,58,69, 74, 75.82, 84, 86 Tsonis, P. A., 95, 151 Tsuchida, K., 104, 121, 131, 151, 373. 374, 394,395, 396,398, 399,405, 406,411,413, 414 Tsujisaka, Y., 2, 7, 49 Tsujita, T., 410 Tsunemitsu, M., 21 1,248 Tuchida, K., 396,405,406,412 Tuck, C. W., 45.49 Tucker, E. M., 161,202 Tulkki, A.-P., 77,86 Tun, P., 269,297 Turkenburg, J. P., 3, 5, 17, 19,22,47, 78,81 Turkenburg, J., 3, 4, 17,47 Turner, P. R.,307,366 Twigg, P. D., 160, 161, 167, 168, 183, 185,197,198 TWU,J.-S., 42.50 Tybring. L., 178,198 Tytler, E. M., 339,369

78,86

Thuren, T., 77,80,86, 250,266,280,301 Tietz, A., 373,401,412, 413, 414 Tikkanen, M. J., 209,245,248,272,298 Tilghman, S. M., 156,198 Tillement, J.-P., 178, 179,201,202 Tizard, R.,57,58,84 Toh-e, A., 39,50 Toker, L.,5, 10, 33,50 Tolley, S., 3,4, 17,47 Tomasselli, A. G., 58,81,86 Tominaga, Y., 7,50 Tomoi, M., 93, 104, 149 Tomonga, M., 250,299 Toner, J. J., 141, 142, 143,149 Twhill, K. L. H., 345, 356, 360,366 Twze, J., 98,148 Toribara, T. Y., 179,196

Ueda, K., 262,297,307,369 Ueta, N., 205,246 Uhlin, U., 64,87 Umemura, J., 353,367 Unge, T., 108, 122, 126,149 Unger, E.,275,300 Unger, W. G., 177,202 Uno, Y., 308,366 Urdea, M. S., 207,246 Urquhart, J., 177,203 Uterman, G., 354,369 Utermann, G., 250, 251,255,256,257, 259, 260,261, 277, 279,280,282, 296,297,300,301,302 Uyemura, K., 90, 120, 126, 127,151

AUTHOR INDEX

V Vadas, P., 55, 58,86 Vades. P., 57,83 Vagin, A. A,, 99, 151 Vahlquist, A., 138,150 Vainio, P., 80,86 Vainshtein, B. K., 99, 151 Valette, G., 177,201 Valivety, R. H., 2.50 Vallner, J. J., 178,201 Van Antwerpen, R.,382,385,388,390, 391,394,399,404,412,413,414 Van Beeumen, J., 99,150 van Bielen, J., 39,50 Van Camp, S. L., 121,150 van de Broek, A. T. M., 375,414 van de Rijn, P., 156,198 van Deenen, L. L. M., 53,87 van denBergh, C. J., 58,64,82,87 Van den Broek, A. T. M., 401,414 Van den Eijnden, M., 396,414 van den Maagdenberg, A. M. J. M., 269, 300 vander Elzen, P. J. M., 195,202 Van der Horst, D. J., 372,373,375,376, 377, 378,379,381,382,385,388, 390,391,394, 396, 397, 399,400, 401,402,403, 404,409, 410,412, 413, 414 van der Kooij-Meis, E., 269,300 van der Westhuyzen, D. R., 363,364 Van der Wiele, F. C., 76.87 Van Doorn, J. M. V., 396,414 Van Doorn, J. M., 378,379,381,382, 402,403,404,410,414 van Driel, I. R.,263,301 van Cent, T., 308,366 vancunsteren, W. F., 115, 138,148 Van Handel, E., 373.414 Van Heusden, M. C., 379,382,388,390, 391,397,398,399,402,404,412,414 Van Kessel, A. G., 120,150 van Kimmenade, A,, 39.50 van Linde,*M.,76.87 Van Marrewijk, W. J. A., 372,375,396, 401,409,414 van Nes, G. J. H., 60,82 van Oort, M. G., 76,87 van Scharrenburg, G. J. M., 77,82,87

447

van Tilbeurgh, H., 3,5, 10, 18,22,27, 28, 41,50 van Tol, A,, 308,366 Vandermeers, A., 57,83 Vandermeers-Piret, M.-C., 57,83 VanTuinen, P.,2,41,47 Vasquez, J. R.,16,50 Vassart, G., 45,50, 178, 179,202 Vaughen, T. A., 45,49 Vauhkonen, M., 21 1,248 Vaz, W. L. C., 63,76,83 Veerkamp, J. H., 91,92,93, 104, 120, 130,149,150,151 Vega, G. L., 208,248 Venable, R. M., 69,85 Venkatachalapathi, Y. V., 287,293,300, 309,314, 322,323,324,329,331, 333,339,341,347,348,350,353, 355,357, 359,360,362,364,365, 367,368,369,378,413 Venkatesh, K., 396,414 Verdery, R. B., 388,414 Verdonck, L., 178,202 Verger, R.,3, 5, 10, 18.22, 26,27, 28,41, 43,44, 47, 48,50, 344, 358,369 Verhalen, P. F. J., 320,369 Verheij, H. M., 53,55,58,64,66,69,74, 75,78,80,82,83,84,85,86,87

Vermeulen, A., 178,202 Verschueren, K. H. G., 33,37,50 Verstuyt, J. G., 267,299 Venvoerd, T. C., 195,202 Vetter, W., 48 Vidal, J. C., 58,60,63,66,69,70,80, 85,86 Vigneault, N., 54.84 Vinio, P., 344,358,369 Virtanan, J. A., 77,80,86, 344, 358, 369 Vita, G. M., 308,369 Voelker, T. A., 18,50 Vogel, T., 267,268,269,27 1,298,299 Vogelberg, K. H., 277,279,301 Volwerk, J. J., 54,60,63,66, 77,84,85, 87 von Eckardstein, A., 357,369 von Heijne, G., 31 1,369 Voorma, H. O., 376,390,413 Vorum, H., 186,197 Voshol, H., 377,414

448

AUTHOR INDEX

Voshol, J., 402,404,414 Vu Dac, N., 356,367

Weinstock. J., 156,202 Weintraub, H., 373, 398,401, 413, 414, 415

Weintraub, M. S.,282,301 Weisgraber, K. H., 213,248, 250, 251, Wade, D. N., 179, 181, 182,202 Wagener. R.,208,248 Wahli, W., 156, 198 Waite, M., 53,58,87, 250,266,280,301 Wal, J.-M., 183,203 Wald, J., 345,352, 355,356, 360,366 Waldmann, T. A., 154,202 Waldmann-Meyer, H., 180,202 Walker, J. E., 156, 160, 169, 183, 192, 200,202

Wall, R.,310,365 Wallace, B. A., 313,365 Wallis, S. C., 208, 209,210,211,213, 232, 244,245,246,247

Walsh, A., 267,299 Walsh, M. T., 113, 123, 142, 147,149 Walter, M., 357,369 Wang, C.-S., 40,45,47,50 Wang, K., 41,42,47 Wang, S., 57,58,82 Wang, X.,209,247,248 Wang, Z., 401,414 Wardell, M. R.,2 13,248, 269,279, 284, 285,286,287,288,289,290,301, 302,341,344,345,351,369,378,415 Warmke, G. L., 245 Warrick, M. W., 64,87 Warshel, A., 11,50 Watanabe, Y . , 267,302 Watkins, S., 162, 163,196,200 Watson, H. C., 212,247,310,313,367 Weare, J. H., 166,197 Weber, P., C.,167, 197 Weber, W., 255,275,277,295,300,301 Weckesser, J., 99,151 Weech, P. D., 232,247 Weers, P.M. M., 396,414 Wegmann, T. G., 156, 160,200 Weichel, E. J., 166,200 Weigand, K., 154,201 Weigand, R. C., 45,49 Weil, D., 207,246 Weinberg, R. B., 357,358,369 Weinstein, D. B., 262,299 Weinstein, J. N.,352,354,369

252,253,256,257,258, 259,260, 262,264,265,266,267,268,269, 270,271,272,274,276,277, 278, 279, 280,282,283,284,285, 286, 287,288,289,290,291,292,293, 294,295,296,297,298,299, 300, 301,302, 341,343,344,345,35 1, 358,359,366,368,369,378,415 Weisgraber, K., 359,369 Weisgraber, R. H., 257,258, 259,302 Weiss, M.S., 99,151 Weiss, R. M., 3 10, 3 11, 345,365 Weissfloch, A. N. E.,19,49 Weitkamp, L. R.,163,202 Welches, W., 57,85 Wells, J. A., 74, 75,81 Wells, M. A., 2,50, 93, 104, 113, 121, 122, 131, 132, 133,148,151, 340, 341, 342,357,364,368, 372, 373,374, 375,376,377,378,379, 381, 382, 383, 385,389,390, 391, 394, 395, 396,397,398,399,402, 404,405, 406,410,411,412,413,414,415 Welte, W., 99, 151 Weng, S.. 210, 21 1,245 Weng, S.-A., 212, 215,248 Weng, S.-Z., 207, 21 1,226,248 Wernstedt, G., 93, 104,148 Wery, J.-P., 64,87 Wes, J., 404,414 Wesenberg, G., 341, 342,357,364, 377, 378,379,383,410 Wessler, A. N., 398, 413 Wesson, R. M., 313,365 West, J. B., 2,50 Westerlund, B., 64,87 Westerlund, J . A., 292,293, 294,302 Westerlund, J. R.,280,297 Westphal, U., 177,202 Wetterau, J. R.,257,258,259,260, 262, 264,285,295,302 Wettesten, M., 240,245 Wharton, C. W., 72,86 Wheeler, C. H.,373, 378,402,404,411, 415

449

AUTHOR INDEX

Whitaker, D. R.,74,83 White, A. J., 72,86 White, A., 240,245 White, R.,269,296 White, S. P., 54,59,60,61,62,63,65,67, 68.69, 70, 71,72,73, 76,77, 78,79, 85,86, 87 Whitlam, J. B., 179,202 Whitlock, M. E.,262,296 Whitted, B. E., 252,299 Wierenga, R., 99,150 Wilcox, R. W., 250,266,301 Wiliamson, R.,207,246 Will, G., 166,201 Williams, D. L., 210,246, 250,300 Williams, R.R.,269,296 Williams, S., 264,300 Williamson, M. P., 58.87 Wilson, C., 213,248,285,286,287,288, 289,290,301,302, 341,344,345, 350,369,378,415 Wilson, D., 282,297 Wilson, P. W. F., 307,369 Wilting, J. B., 174,202 Wiman, K., 137,150 Winard, J., 57,83 Windler, E. E. T., 307,366 Windler, E., 266,302 Winkler, F. K., 3,4,9, 10, 16, 18,20,26, 37,42,50 Winkler, F., 98,151 Winter, N. S., 93, 104, 110, 121, 122, 134, 135,149,151 Winter, W. P., 163,202 Wion, K. L., 156,199 Wisniewski, T., 250,302 Witiak, D. T., 178,200 Witling,J., 166, 176,196, 198 Witzturn,J. L., 208,248, 250, 252,253, 296 Wlodawer, A., 352,354,369 Wokstrom, P., 38,50 Woldike, H., 47 Wolf, S. I., 163,198 Wolfer, H., 48 Wollert, U., 155, 178, 184,198,200 Wong, C. K., 45,49 Wong, C.-H., 2,50 Wong, H., 41,42,47,50, 275,296 Wong, J. S., 275,297

Wood, P. C., 195,201 Woodrow,J. R., 195,201 Wootan, k.G., 93, 104, 110, 122, 134, 135, 147,151 Wootton, R.,307,366 Worchester, D.,216,246 Woreell, A. C., 18,50 Wosilait, W. D., 178, 179, 198 Wright, A. K., 161,202 Wright, C. E.,92,93, 140, 148 WU,C.4. C., 257,259,260,264,295, 320, ?69 WU,H.-L., 178,202 Wu, L. L., 269,296 WU,M.-J., 208,247,248 Wu, S. H., 55,87 Wu,S.,3,4,6,7,8, 18,22,50 Wyatt, G. R.,373,377,379,410,411 Wynne, H. J. A., 399,414

X Xie, Z.-F., 19,50 Xiong, W., 209,244 Xu, C. F., 209,245 Xu, C., 209,248 XU,C.-F., 269,299 Xu, Z., 93, 104, 110, 115, 116, 120, 126, 128, 129, 130,142,144,145,149,151 Xuong, N.-H., 75,86,98,150

Y Yagi, K., 308,366 Yakata, M., 162,202 Yamada, J. H., 93,104,149 Yamada, N., 250,267,300,302 Yamada, T., 162,202 Yamaguchi, S., 3, 18,39,40,50 Yamamoto, A., 257,259, 263, 264,269, 291,296,300,302 Yamamoto, K., 267,300 Yamamoto, T., 271,302 Yamamura, T., 269,296,300,302 Yamashita, S., 262,302 Yang, A. S., 16,50 Yang, C. H., 6,39,49 Yang, C. Y., 210,211,212,245 Yang, C.-C., 76,87

450

AUTHOR INDEX

Yang, C.-Y., 207,210,211,212,213, 214, 215,226,245,248,252,298 Yang, F., 156. 160,202,203 Yang, J. T., 320,369 Yang, M., 156,199.201, 212,215,248, 252,298 Yang, N.-C.C., 120, 121, 134, 136, 149 Yao, Z., 235,236,237,248 Yap, W.T., 180,201 Yarranton, G. T., 43,47 Yates, F. E.,177,203 Yates, M., 226,246,411 Yazaki, Y., 267,300 Yazawa, M., 377,378,410 Yeadon, J. E., 38,49 Yeagle, P. L., 215,248, 339,365, 389, 415 Yeh, P., 195,203 Yip, P., 93, 104,149 Yokoyama, S., 257.259, 263,264,291, 296,302, 346, 347, 348,365, 398, 413 Yoshida, M., 76,87 Yoshida, N., 39,47 Yoshimura, A., 262,297 Yoshinura, K., 90, 120, 126, 127,151 Yoshizumi, K., 58,88 Yotsuyanagi, T., 182,200 Young, S . G., 208, 209,232, 235, 236, 237,248,252,253,296 Yu, 58.83 Yu, B.-Z., 71, 76,78,81,83 Yu, Z., 90,93, 104,150 Yuan, W., 54,60,65,67,68.69,72,76, 77, 78, 79,85, 88

Yuregir, G. T., 163,198 Yvon, M., 183,203

z Zaccai, G., 313,365 Zannis, V. I., 209,210,245, 251, 255, 256,257,261,277,302 Zanotti, G., 92,93,98, 104, 130, 138, 139, 150,151 Zappini, M.,162, 198 Zaroslinski,J. F., 179,199 Zasloff, M.,313,320,368,369 Zech, L.A., 267,282,297,300 Zeigler, R., 374, 377,402,413 Zemlin, F.. 69,82 Zepponi, M.,163,200 Zhang, H., 208,248 Zhang, S. H.,267,302 Zhingnesse, M.,269,299 Zhu, S., 207, 226,245 Ziegler, R., 373, 377,402,403,413, 414, 415 Zilversmit, D. B., 308,367, 388, 415 Zimmerman, S. C., 15,50, 75,81,88 Zini, R., 178, 179,202 Zinsmeister, A. T., 308,366 Zipper, P., 385,411 Zoellner, C. D., 250,295 Zollman, S., 207,247 Zorich, N., 250,302 Zuber, H., 140,149 Zurakowski, R., Jr., 176,203 Zurcher-Neely, H., 58.86 Zvelebil, M.J. J. M.,169,203 Zwiers, H., 320,369

SUBJECT INDEX

A

physical measurements, 164 structure early concepts, 161-166 evolution, 190-194 primary, 155-161 X-ray crystallography, 167-169 conformational flexibility, 172- 176 disulfide pairings, 17 1 domain structure, 169-171 history, 166-167 surface charge distribution, 171-172 Alpha helices, amphipathic amino acid side chains charged, 322-336 hydrophobic, 336 antiviral/antiinflammatory activities,

Acet ylcholinesterase crystallographic database, 5 Torpedo calijornica, alp hydrolase fold, 36-37

Adipocyte lipid-binding protein apo and holo forms, 114-1 15 binding affinity and specificity biochemical studies, 119-122 crystallographic studies, 122-126 Ca coordinates, root-mean-square difference, 102 conserved amino acids, 106- 109 crystallographic data, 92-93 function, 127-130 gap regions between pD and BE,

36 1-363

116-119

apolipophorin X-ray structure, 340-344 characterization, 3 10-3 12 classes correlation with lipid associations,

ligand-binding properties, 120 ligand entry into binding cavity, 115-1 16

structural motif, 94-98 structure, 127-130 Adsorption, interfacial, secretory phospholipases A2 catalytic efficiency and, 78-80 surfaces, 76-77 Albumin amino acids conservedhnvariant, 158-159 genetic variants, 162-163 sequence homologies, 160- 161 sequence references, 156 ligand binding, 176-181 binding constants, 177-180 long-chain fatty acids, 186- 187 metals, 188- 190 sites, 195 small organic compounds, 181-185 loop-link-loop pattern, 157

357-358

groups, 313-321 conformational analysis, 344-346 cross-sectional shape, 337-340 evolutionary origin, 322 HDL receptor activity, 360-361 lecithin-cholesterol acyltransferase activation, 359-360 length, 336-337 lipoprotein lipase activation, 358 role in receptor recognition, 358-359 structure-function studies with a% nouo-designed synthetic peptide analogs, 346-349 with intact apolipoproteins, 352-357 with native apolipoprotein sequences, 349-351

X-ray structure determination, 340-344 45 1

452

SUBJECT INDEX

Amino acids albumin conserved, 158-159 evolutionary aspects, 190- 194 genetic variants, 162-163 apolipoprotein E, genetic variants, 269 conserved albumin, 158-159 ApoE, 253 intracellular LBPs, 106- 109 secretory phospholipases A2, 56-57 Amino acid sequence albumin, 155-161 apolipophorin 111,379-381 apolipoprotein B, 2 10-2 13 apolipoprotein E, 25 1-255 intracellular LBPs, 103-105 secretory phospholipase As, 54-58 Amino terminus, secretory phospholipases A2, 60-63 Antibodies, monoclonal, see Monoclonal antibodies Antiinflammatory activity, amphipathic helices, 36 1-363 Antiviral activity, amphipathic helices, 361-363 Apolipophorin 111 amino acid sequence, 379-381 binding to lipophorin, 382-383 distribution among insects, 377 evolutionary aspects, 383-384 function, 378 molecular structure, 378-379 X-ray structure, 340-344 Apolipoprotein A-I amino acid sequence, derived properties, 327 amphipathic helices conformational analysis, 344-346 native sequence studies, 350 properties, 334 function, 306 hydrophobic and Pro residues, HELNET analysis, 338 molecular weight, 306 Apolipoprotein A-I1 amino acid sequence, derived properties, 326 amphipathic helices, properties, 334 function, 306 molecular weight, 306

Apolipoprotein A-IV amino acid sequence, derived properties, 327 amphipathic helices native sequence studies, 35 1-352 properties, 334 function, 306 molecular weight, 306 Apolipoprotein B cotranslational assembly in rough endoplasmic reticulum, 240-243 gene for, 207-209 mapping on LDL surfaces by LDL core circumference related to ApoB fragment size, 235-240 by lipid extraction after LDL attachment to EM grids, 227 with monoclonal antibodies, 227-235 mRNA processing, 209-2 10 polypeptide features, 206 primary sequence, 210-2 13 receptor binding site, formation, 207 Apolipoprotein C-I amino acid sequence, derived properties, 326 amphipathic helices, properties, 334 function, 306 molecular weight, 306 Apolipoprotein C-11 amino acid sequence, derived properties, 326 amphipathic helices, properties, 334 function, 306 molecular weight, 306 Apolipoprotein C-111 amphipathic helices native sequence studies, 350 properties, 334 function, 306 molecular weight, 306 Apolipoprotein E amino acids genetic variants, 269 sequences, derived properties, 327 amphipathic helices Conformational analysis, 344-346 native sequence studies, 351 properties, 334 as chaperone protein, 250 cysteinyl residues, 284-285

SUBJECT INDEX

defective form, effects on lipoprotein clearance, 280-281 discovery, 25 1 domain interactions, 282-284 domain structure, 257-260 function, 306 heparin interaction, 274-277,359 heterogeneity characterization, 255-257 effect on plasma lipid concentrations, 281-282 hydrophobic and Pro residues, HELNET analysis, 338 and hyperlipoproteinemia type 111, 261-262,277-281 lipid binding carboxyl-terminal regions for, 292-294 effects on structure, 290-292 in lipoprotein metabolism, 265-267 molecular weight, 306 multireceptor binding ability, 262-265 nomenclature, 256-257 physical properties, 257-260 in plasma lipoproteins, 260-262 primary structure, 251-255 receptor-binding region, 267-274 22-kDa fragment, 3D structure, 285-290 Apolipoproteins amphipathic helices, see Alpha helices, amphipathic -lipid complexes, amphipathic helices, 352-355 in lipophorins, 389-390

B Beta strands, see also Protein folding in extracellular LBPs, 98 in immunoglobulin fold, 98-99 in intracellular LBPs characteristics, 94-98 gaps between BD and BE, 116- 119 Bile salt-activated lipase, role in lipidllipoprotein metabolism, 44-45 Bilin-binding protein crystallographic data, 92-93 function, 140 structure, 140

453

C Calcium, secretory phospholipase A2 binding loop, 63-64 primary ion, 65 Carboxyl terminus ApoE, lipid-binding regions, 292-294 secretory phospholipases A2, class 11, 66 Catalytic triads, in lipases, stereochemistry, 10-16 Cellular retinoic acid-binding protein amino acid sequence, 103 ligand-binding properties, 120 Cellular retinol-binding protein apo and holo forms, 114-1 15 binding affinity and specificity biochemical studies, 119-122 crystallographic studies, 122-126 binding cavity, 110-1 13 Ca coordinates, root-mean-square difference, 102 conformational similarity to other iLBPs, 99-101 conserved amino acids, 106-109 crystallographic data, 92-93 function, 134-135 gap regions between fiD and BE, 116-1 19 ligand-binding properties, 120 site-directed mutagenesis, 141-142 structure, 134- 135 Cellular retinol-binding protein I1 binding affinity and specificity, biochemical studies, 119-122 binding cavity, 110-1 13 Ca coordinates, root-mean-square difference, 102 conserved amino acids, 106-109 crystallographic data, 92-93 function, 135-136 ligand-binding properties, 121 structure, 135-136 Chaperone proteins, apoE function, 250 Charge distribution, in amphipathic helical domains, 322-336 Cholesterol esterase, see Bile salt-activated lipase Cofactors, binding by human lipoprotein lipase and pancreatic lipase, 41-42

454

SUBJECT INDEX

Colipase interaction with pancreatic lipase, 9 role in interfacial activation, 26-27 COMBO program amphipathic helix classes analyzed by,

Disulfides, pairings in albumin, 171 DNA, human ApoB100, cDNA sequence, 210-213

E

314-319

analysis of potential amphipathic helices, 330 description, 312 COMNET program amphipathic helix classes analyzed by, 314-319

description, 3 12 Conformation albumin, pH effects, 172-176 amphipathic a helices, 344-346 cellular retinol-binding protein, 99- 101 heart fatty acid-binding protein, 97 human pancreatic lipase, 25-26 intestinal fatty acid-binding protein, 99-101

Electron microscopy, lipophorins, 386-388

Electrostatics, in interfacial catalysis of secretory phospholipases A2, 77-78 Emulsion particle model, for low-density lipoproteins, 2 15-2 17 Endoplasmic reticulum, rough, cotranslational assembly of lipoprotein, 240-243 Ester bond, hydrolysis by lipases, 10-18

F Fat body, lipid transport to and from,

lipases, interfacial activation-related changes, 22-29 liver fatty acid-binding protein 11, 99-101

Consensus pentapeptide, in serine proteinases and lipases, 30-33 CONSENSUS program amphipathic helix classes analyzed by, 3 14-319

analysis of amphipathic helices, 332 description, 3 12 Copper(II), binding by albumin, 188-190 Cotranslational assembly, lipoproteins in rough endoplasmic reticulum, 240-243

Cutinase FIIsanum sohni, crystallographic database, 4 alp hydrolase fold, 36-37 Cysteinyl residues, effects on ApoE function, 284-285

D Development, changes in lipophorin metabolism during, 405 Diacylglycerols, in lipophorins, 390-39 1 Dienelactone hydrolase crystallographic database, 5 alp hydrolase fold, 36-37

398-401

Fatty acid-binding proteins heart p barrel conformation, 97 binding affinity and specificity, crystallographic studies, 122- 126 Ca coordinates, root-mean-square difference, 102 crystallographic data, 92-93 function, 130 ligand-binding properties, 120 structure, 130 intestine apo and holo forms, 1 14- 1 15 binding affinity and specificity, crystallographic studies, 122-126

Ca coordinates, root-mean-square difference, 102 conformational similarities, 99-10 1 conserved amino acids, 106- 109 crystallographic data, 92-93 function, 133-134 gap regions between BD and PE, 116-1 19

ligand-binding properties, 12 1 ligand entry into binding cavity, 115-1 16

protein folding, 144- 145 structure, 133-134

455

SUBJECT INDEX

liver crystallographic data, 92-93 type 11, conformation, 99- 10 1 M a n d w a sexta binding affinity and specificity, 122-126 binding cavity, 110-1 13 conserved amino acids, 106-109 crystallographic data, 92-93 function, I3 1- 133 structure, 131-133 Fatty acids, long-chain, binding by albumin, 186-187 Folding, proteins, see Protein folding Fungal lipases, see a h specific lipases activity and structure, 39-40

G Gastric lipase, human, role in lipidllipoprotein metabolism, 43-44 Genes apolipoprotein B, 207-209 lipase, 40-43 Geotrichum canddum lipase crystallographic database, 4 a l p hydrolase fold, 36-37 oxyanion hole, 18 p-cSer-a motif, 32-33 substrate specificity, 18-19 3D structure, 7-8 a2-Globulin, urinary crystallographic data, 92-93 function, 140-141 structure, 140-141 Gold(1). binding by albumin, 188-190

H Haloalkane dehalogenase crystallographic database, 5 alp hydrolase fold, 36-37 HDL receptors, see Lipoprotein receptors, high-density Heart, fatty acid-binding proteins p barrel conformation, 97 binding affinity and specificity, crystallographic studies, 122-126 crystallographic data, 92-93 function, 130 ligand-binding properties, 120

root-mean-square difference in Ca coordinates, 102 structure, 130 Helical net representations, 310-31 1 Helices alpha, amphipathic, see Alpha helices, amphipathic amino-terminal, secretory phospholipases AP,60-63 antiparallel, secretory phospholipases A2,64 HELNET program apoE and apoA-I hydrophobic residues, 338 description, 3 12 examples of amphipathic helices, 328 Heparin, ApoE interaction, 274-277, 359 Hepatic lipase, human cofactors required by, 41-42 role in lipoprotein metabolism, 40-43 Hinged-domain hypothesis, evidence for, 355,357 Hormone-sensitive lipase, role in lipidllipoprotein metabolism, 45-46 Humicola lanugn'nosa lipase, crystallographic database, 4 alp Hydrolase fold, in lipases, 33-37 Hydrophobicity ApoB, 212 nonpolar face, amphipathic helices, 336 Hydrophobic moment analysis, amphipathic helices, 310-31 1 Hyperlipoproteinemia, type 111, apolipoprotein E role, 261-262, 277-281

I iLBPs, see Lipid-binding proteins, intracellular Interfacial activation, lipases colipase role, 26-27 conformation changes, 22-29 lids, 20-22 site, 20-22 Interfacial catalysis, by secretory phospholipase A2 efficiency, 78-80 electrostatics, 77-78 hopping mode, 75-76

456

SUBJECT INDEX

interfacial adsorption surfaces for, 76-77 scooting mode, 75-76 Intestine, fatty acid-binding proteins apo and holo forms, 114- 115 binding affinity and specificity, crystallographic studies, 122- 126 Ca coordinates, root-mean-square difference, 102 conformational similarities, 99- 101 conserved amino acids, 106-109 crystallographic data, 92-93 function, 133-134 gap regions between /IDand PE, 116-1 19 ligand-binding properties, 121 ligand entry into binding cavity, 115-1 16 protein folding. 144-145 structure, 133-134

L 8-Lactoglobulin crystallographic data, 92-93 function, 139-140 structure, 139- 140 LBPs, see Lipid-binding proteins LDLs, see Low-density lipoproteins Lecithin-cholesterol acyltransferase, amphipathic helix-enhanced activation by apoA-I, 359-360 Lids, in lipases, 20-22 Ligand binding, albumin, 176- 181 binding constants, 177- 180 long-chain fatty acids, 186- 187 metals, 188-190 sites, 195 small organic compounds, 181- 185 Ligands, entry in iLBP binding cavity, 115-1 16 Lingual lipase, rat, role in lipid/lipoprotein metabolism, 43-44 Lipases, see aLto spec@ lipases catalytic triad stereochemistry, 10- 16 crystallographic database, 3-6 ester bond hydrolysis, 10-18 evolutionary relationships, 29,37-38 gene family, 40-43

interfacial activation conformational changes, 22-29 lids, 20-22 site, 20-22 in lipid/lipoprotein metabolism, 40-46 oxyanion hole, 16- 18 preduodenal, 43-44 procolipase-lipase complex crystal structure, 27-28 prokaryotic enzymes, 39 and proteinases, relationships, 37-38 8-rSer-a motif, 30-33 substrate specificity, 18-20 tertiary folds, 33-37 Lipid-binding proteins adipocyte, see Adipocyte lipid-binding protein beta strands, gaps between PD and PE, 116-1 19 extracellular members, 93 structural motif, 98-99 intracellular amino acid sequence comparisons, 103-105 apo and holo structures, 114-1 15 binding affinity and specificity biochemical studies, 119-122 crystallographic studies, 122-1 26 ligand binding properties, 120- 121 binding cavity characterization, 110-1 13 homology of residues, 114 ligand entry, 115-1 16 Ca coordinates, root-mean-square difference, 102 characterization, 110-1 13 conformational similarity, 99- 101 homologies, 101- 106 members, 93 need for, 146-147 physiological functions, 145- 148 structural motif, 94-98 protein folding, 92 site-directed mutagenesis, 141-142 stereo diagram, 95 structure, general features, 92 Lipid metabolism bile salt-activated lipase role, 44-45 hormone-sensitive lipase role, 40-46

457

SUBJECT INDEX

human gastric lipase role, 43-44 lysosomal acid lipase role, 44 plasma concentrations, ApoE heterogeneity effects, 28 1-282 rat lingual lipase role, 43-44 Lipids -apolipoprotein complexes, amphipathic helices, 352-355 binding by iLBPs site-directed mutagenesis studies, 141-142

thermodynamics, 142-144 Lipid transport, by lipophorins to developing oocytes, 404-405 from fat body to flight muscle, 401 -404

from midgut to fat body, 398-400 Lipophorin-111,amphipathic helical domains, X-ray structure, 340-344 Lipophorins apolipoproteins in, 389-390 biosynthesis, 394-397 diacylglycerols in, 390-39 1 exchangeable proteins, 377-384 heterogeneity, 384-388 hydrocarbons in, 390 lipid composition, 373-375 lipid transfer particle function, 397-398

lipid transport to developing oocytes, 404-405 from fat body to flight muscle, 401-404

from midgut to fat body, 398-400 metabolism, changes during development, 405 models, 39 1-393 molecular weight, 384-388 phospholipids in, 390 physiological roles, 397-405 shape, 384-388 size, 384-388 structural proteins, 375-377 structure, metabolic implications,

Lipoprotein receptors, high-density, mediation by amphipathic helix, 360-36 1

Lipoproteins assembly cotranslational, 240-243 triglyceride-rich, two-step model, 243 classification,304 composition, 304 insect, see Lipophorin low-density, see Low-density lipoproteins metabolism apolipoprotein E role, 265-267 bile salt-activated lipase role, 44-45 hormone-sensitive lipase role, 40-46 human gastric lipase role, 43-44 lysosomal acid lipase role, 44 rat lingual lipase role, 43-44 plasma, ApoE in, 260-262 Liver, fatty acid-binding proteins crystallographic data, 92-93 type 11, conformation, 99-101 Low-density lipoproteins ApoB mapping by LDL core circumference related to ApoB fragment size, 235-240 by lipid extraction after LDL attachment to EM grids, 227 with monoclonal antibodies, 227-235

composition, 214 definition, 2 14 emulsion particle model, 2 15-2 17, 220-22 1

molecular weight, 2 14 physical properties and composition emulsion particle model, 220-221 experimental values, 223 quantitative molecular model, 2 17-226 structure, 213-215 Lysosomal acid lipase, role in lipid/lipoprotein metabolism, 44

405-408

Lipoprotein lipase, human amphipathic helix-enhanced activation by apoC-II,358 cofactors required by, 41-42 role in lipoprotein metabolism, 40-43

Major urinary protein crystallographic data, 92-93 function, 140-141 structure, 140-141

458

SUBJECT INDEX

Manduca sex& bilin-binding protein, crystallographic data, 92-93 fatty acid-binding protein binding affinity and specificity, 122-126 binding cavity, 110-1 13 conserved amino acids, 106- 109 crystallographic data, 92-93 function, 13 1-133 structure, 131-133 Mercury(II), binding by albumin, 188-190 Metals, binding by albumin, 188- 190 Monoclonal antibodies, ApoB mapping on LDL surfaces with, 227-235 Muscles, flight, lipid transport by lipophorins, 401 Mutagenesis, site-directed, iLBPs, 141-142 Mutations, point, in human serum albumin, 172 Myelin P2 protein binding affinity and specificity, crystallographic studies, 122- 126 binding cavity, 110-1 13 Ca coordinates, root-mean-square difference, 102 crystallographic data, 92-93 function, 126-127 ligand-binding properties, 120 structure. 126-127

Nickel(II), binding by albumin, 188-190

0 Oocytes, lipophorin lipid transport to, 404-405 Organic compounds, long-chain, binding by albumin, 181-185 Oxyanion hole, lipases, 16- 18

P Pancreatic lipase, human conformational changes related to interfacial activation, 25-26

hydrogen bonding in catalytic triads, 11-13 alp hydrolase fold, 36-37 lid features, 20-22 molecular structure, 9-10 oxyanion hole, 18 p-cSer-a motif, 30-33 substrate specificity, 19 tertiary folds, 33-37 Penicillium camemberti lipase, crystallographic database, 4 Peptide analogs, for studies of apolipoprotein amphipathic helices, 346-349 pH, effects on albumin conformation, 172- 176 Phosphatidylcholine, in lipophorin, 374-375 Phosphatidylethanolamine, in lipophorin, 374-375 Phospholipase Ar, secretory amino-terminal helix, 60-63 amino terminus, 60.63 beta wing, 66 calcium-binding loop, 63-64 calcium ion, primary, 65 carboxyl-terminal extension, 66 catalytic networks, 60-61 classes, 55 classification, 54-58 class I loop, 64-66 conservedlinvariant amino acid residues, 56-57 conserved substructures, 60-66 hydrophobic channel, 60-62 interfacial catalysis electrostatics, 77-78 hopping mode, 75-76 interfacial adsorption catalytic efficiency and, 78-80 surfaces, 76-77 scooting mode, 75-76 primary structure, 54-58 related proteins, 57-58 secondary structure, 58-60 substrate binding, 67-71 tertiary structure, 58-60 tetrahedral intermediates, 7 1-73 Phospholipids, in lipophorin, 373-375, 390

459

SUBJECT INDEX

Polymorphisms ApoB gene, 208-209 APE, 255-257 Polypeptides, amphipathic helices hormones, 313,317,320 lytic polypeptides, 313,316,320 Portal hypothesis, ligand entry into iLBP binding cavity, 115-1 16 Preduodenal lipases, 43-44 Procolipase, lipase-procolipase crystal structure, 27-28 Prokaryotic enzymes, 39 Proteinases, evolutionary relationships with lipases, 37-38 Protein folding, see also Beta strands intestinal fatty acid-binding protein, 144-145 intracellular LBPs, 144-145 lipid-binding proteins, 92 tertiary, in human pancreatic lipase, 33-37 Protein kinases, calmodulin-associated, amphipathic helices, 3 13 Proteins, amphipathic helix classes, 313-321 Protein stability, intracellular LBPs, 144-145

R Retinol-binding protein crystallographic data, 92-93 function, 137-139 structure, 137-139 Rhiwmucm miehei lipase conformational changes, interfacial activation-related, 22-24 crystallographic database, 4 hydrogen bonding in catalytic triads, 11-13 alp hydrolase fold, 36-37 lid features, 20-22 oxyanion hole, 17- 18 b-cSer-a motif, 30-33 substrate specificity, 19 3D structure, 6-7 RNA, messenger, ApoB processing, 209-2 10

Root-mean-square difference, in Ca coordinates for iLBPs, 102

S Schiffer-Edmundson helical wheel diagrams, 310-31 1 Serine carboxypeptidase, wheat crystallographic database, 5 alp hydrolase fold, 36-37 8-eSer-a motif, in lipases, 30-33 Serum albumin, see Albumin Silver(I1).binding by albumin, 188- 190 Stereochemistry, catalytic triad in lipases, 10-16 Substrate binding, productive-mode, secretory phospholipases A*, 67-71 Substrate specificity, lipases, 18-20 Surface charge, human serum albumin, 171-172

T Transition state, tetrahedral, secretory phospholipase AP,71-73

W WHEEL program description, 3 11 examples of amphipathic helices, 328

X X-ray crystallography albumin, 167-169 conformational flexibility, 172-176 disulfide pairings, 17 1 domain structure, 169-1 7 1 history, 166-167 surface charge distribution, 171-172 apolipophorin amphipathic helical domains, 340-344 intracellular LBPs binding affinity and specificity, 122- 126 lipases, database, 3-6 lipid-binding proteins, 92-93 X-ray scattering, small-angle, lipophorins, 304-306

I S B N 0-12-034245-6

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