No title

Current Topics in Membranes and Transport VOLUME 29

Membrane Structure and Function

Advlsory Board

G . Blobel

E. Carafoli J . S. Cook D.Louvard

Current Topics in Membranes and Transport

VOLUME 29

Membrane Structure and Function

1987

ACADEMIC PRESS,INC. Harcourt Brace Jovanovich, Publishers

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Contents

Preface, ix Yale Membrane Transport Processes Volumes, xi

Current Views of Membrane Structure ALAN M. KLEINFELD 1. 11. 111. IV. V. VI.

Introduction, I Lipid Structure, 3 Lipid-Lipid Interactions: Single Lipid Species. 4 Lipid Mixtures and Domains, 10 Membrane Protein Structure. 12 Interactions between Lipids and Proteins, I6 References. 20

Ultrastructural Studies of the Molecular Assembly in Biomembranes: Diversity and Slmilarity SEK-WEN HUI 1. 11.

Ill. IV. V.

VI.

Similarity and Diversity of Biological Membranes. 30 Is the Lipid Bilayer a Passive, Homogeneous "Solvenl"'!. 35 Is the Bilayer the Only Form of Membrane Lipids?, 43 Do Membrane Proteins Distribute Randomly in the Bilayer'?, SO What Determines the Conformation of Intrinsic Protein Molecules in Membranes'?, 57 A Look at the Future through the Electron Microscope, 62 References. 65

vi

CONTENTS

The Thermodynamics of Cell Adhesion MICAH DEMBO AND GEORGE 1. BELL I. 11.

Ill. IV. V. V1. VII.

Introduction, 71 The Model, 71 The Chemical Potentials, 74 The Repulsive Potential, 76 Minimizing the Free Energy, 78 Results, 82 Discussion, 87 References, 89

Rotational and Lateral Diffusion of Membrane Proteins and Lipids: Phenomena and Function MICHAEL EDIDIN Introduction, 91 Probes for Diffusion, 92 111. Rotational Diffusion, 93 IV. Lateral Diffusion, 102 V. A Concluding Remark, I I9 References, I I9 1.

11.

Biosynthesis and Distribution of Lipids KENNETH J . LONGMUIR Introduction, 129 The Lipid Composition of Subcellular Membranes, 130 Desaturation and Elongation of Fatty Acids, 141 IV. Phospholipid Biosynthesis, 144 V. Membrane Lipid Tmnsport and Differentiation of Membranes, 163 References, 167 I.

11. 111.

Lipid Exchange: Transmembrane Movement, Spontaneous Movement, and Protein-Mediated Transfer of Lipids and Cholesterol ELlEZAR A. DAWIDOWICZ Introduction, 175 Spontaneous Exchange of Lipids between Membranes, 176 Protein-Mediated Lipid Transfer between Membranes, 186 IV. Transmembrane Movement of Lipids. 189 References. 193

1. 11. 111.

CONTENTS

Membrane Fusion ROBERT BLUM ENTH AL I. Introduction. 203 11. How We Observe Membrane Fusion. 205

I l l . 'The Triggering Event in Membrane Fusion. 213 1v. Movement into Apposition, 216 V. The Recognition Event, 219 VI. Steric Constraints. 22 I VII. Motion of Phospholipids, 223 VIII. Interactions between Bilayer Membranes, 227 I X . Membrane Fusion: Fact. Hypothesis, or Theory'?, 240 References. 245

The Control of Membrane Traffic on the Endocytic Pathway I K A MELLMAN, CHRISTINE HOWE. AND AKI HELENIUS 1. Introduction, 255 11. Endocytosis and Membrane Recycling, 2.56 111. Keceptor-Mediated Endocytosis. 258 IV. Mechanisms and Functions of Endocytic Organelles. 260 V. The Exocytic Pathway. 279

References. 281

Index, 289 Contents of Recent Volumes, 299

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Preface The science of membrane structure and function has seen significant progress over the past 15 years. Needless to say, we are only beginning to understand the intricacies involved in maintaining the compartmentalization that is needed to restrict and regulate metabolic processes within the living cell and to permit an adequate, controlled dialogue with the extracellular environment. It has become increasingly clear that membranes themselves are highly dynamic structures, from a physical as well as a chemical point of view. The study of their structure and function requires sound hypotheses to work from as well as the application of a great variety of biophysical, biochemical, and morphologic techniques. Concepts derived from detailed investigations into t h e properties of artificial model membranes are being utilized more and more to gain fundamental insights into the physiology of biomembranes. Volume 29 of Ciirrerit Topics in Mrmhrtrnc~sand Transport is meant to highlight some of the paths in the large field of membrane biology along which recent progress has been realized and to delineate objectives toward which future investigations might be aimed. Many articles in this volunie carry elements of both a review and an essay in an effort to invite the reader to regard them as an introduction to current thinking rather than only a s a review. In the first article, a broad overview is given of our current knowledge of membrane structure in general. A number of morphological backups for these concepts as well as further, new observations on membrane ultrastructure are given in the second article. Communication between cells and between cells and their environment is mediated by membranes and membrane-based molecules. Theoretical models can be devised which may point to specific molecular mechanisms in such communication. The third article presents an example of building a model for cell adhesion in which density and mobility of membrane surface receptors appear as important parameters. The subsequent fourth article explores many theoretical and experimental aspects of the dynamics of proteins and lipids in the mcmbl-me. In the fifth and sixth articles, topology and metabolism of membrane lipids along with implications for the transmembrane distribution of lipids and the compositional diversity of cellular membranes are discussed. The study of fusion processes in artificial and biological membranes has been ix

X

PREFACE

greatly facilitated by the development, over the past 6 years, of a variety of assays. The seventh article provides a survey of membrane fusion studies, the hypotheses behind them, and the models arising from them. Recent, new understanding of the process of endocytosis and its underlying membrane traffic has given a boost to the investigation of such complicated matters as intracellular routing and sorting in which the precise role(s) of membranes and membrane constituents is still largely unknown. The final article describes the developments in this field and offers a number of leads to further experimental approaches. Virtually any volume on progress in the study of membrane structure and function is bound to be far from complete. This volume is no exception in this respect. As our knowledge of membrane physiology increases, essays and reviews on the subject will render an increasingly integrated picture. JOS V A N RENSWOUDE CHRISTOPH KEMPF RICHARD D. KLAUSNER

Yale Mem bra ne Transport Processes Volumes Joseph F. Hoffman (ed.). (1978). “Membrane Transport Processes,” Vol. I . Raven, New York. Daniel C . Tosteson, Y u . A. Ovchinnikov, and Ramon Latorre (eds.). (1978). “Membrane Transport Proccsses,” Vol. 2. Raven, New York. Charles F. Stevens and Richard W. Tsien (eds.). (1979). “Membrane Transport Processes,” Vol. 3 : Ion Permeation through Membrane Channels. Raven, New York. Emile L. Boulpaep (ed.). (1980). “Cellular Mechanisms of Renal Tubular Ion Transport”: Volume I3 of Crirrctil Topics in Mcr?ihmnes rind Trrinsport (F. Bronner and A. Kleinzeller, eds.). Academic Press, New York. William H. Miller (ed.). (198 I ) . “Molecular Mechanisms of Photoreceptor Transduction”: Volume 15 of Cirrrcnt Topics in MonbrLines mid Trruisport (F. Bronner and A. Kleinzeller, eds.). Academic Press, New York. Clifford L. Slayman (ed.). (1982). “Electrogenic Ion Pumps”: Volume 16 of C‘irrrrnt Topics in Mrmbrrinrs rind Trtinsport ( A . Kleinzeller and F. Bronner, eds.). Academic Press, New York. Joseph F. Hoffman and Bliss Forbush 111 (eds.). (1983). “Structure, Mechanism, and Function of the Na/K Pump”: Volume 19 of Crrrrcnt Topics in Mrnihrrincs cind Trrinsporf (F. Bronner and A. Kleinzeller, eds. ). Academic Press, New York. James B. Wade and Simon A. Lewis (eds.). (1984). “Molecular Approaches to Epithelial Transport”: Volume 20 of Crrrrenf topic..^ in Mombrrinc>s mid Trrrtisport (A. Kleinzeller and F. Bronner, eds.). Academic Prcss, New York. Edward A. Adelberg and Carolyn W. Slayman (eds.). (198s). “Genes and Membranes: Transport Proteins and Receptors”: Volume 23 of Crrrrcnt Topics in Memhrtines und Trrrnsport (F. Bronner and A. Kleinzeller, eds.). Academic Press, Orlando. Peter S . Aronson and Walter F. Boron (eds.). (1986). ‘“a’- H’ Exchange, Intracellular pH, and Cell Function”: Volume 26 of Cirrrenf Topics in Mcmhrcrnc~sand Trrrnsport ( A . Kleinzeller and F. Bronner, eds.). Academic Press, Orlando. Gerhard Giebisch (ed.). (1987). “Potassium Transport: Physiology and Pathophysiology”: Volume 28 of Cirrrent Topics in Mctnhrrrncs tind 7’rrrnsport (F. Bronner and A. Kleinzeller, eds. 1. Academic Press, Orlando.

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CLIRRENT 1 0 P I C S IN MEMBRANES AND TRANSPORT. VOLLIME 29

Current Views of Membrane Structure ALAN

M. KLEINFELD

1.

INTRODUCTION

The current view of membrane structure is similar in many respects to the general outline of membrane structure developed more than a decade ago by Singer and Nicolson (1972). The essential features of the model, consisting of transmembrane and peripheral proteins in association with a lipid bilayer, remain intact. In the intervening decade a more profound understanding has been achieved for lipid and protein structure, the interactions between these membrane constituents, and the dynamics of these interactions. This article presents an overview of recent developments in the structure of membrane components and certain of their interactions. Subsequent articles will further explore the structural and dynamic aspects of these interactions. The rationale for studying the structure of membrane components is that such knowledge will allow one to predict the form of the composite membrane and ultimately to understand membrane function at the molecular level. Current understanding of the constituent structures and their interactions leaves us far from this goal. Where the relationships between molecular structure and function are well understood, as for example in the cases of lysozyme (Phillips, 1967) and hemoglobin (Perutz et d., I%%), physiological function appears to be associated with structural changes that are on the order of I A, suggesting that this resolution is necessary to predict and understand function. Furthermore, because of the dynamic nature of the components forming the membrane, appreciable temporal resolution is also necessary. Thus the molecular coordinates determined from lipid and protein crystals must be modulated by molecular dynamics 1 Copyrighi
2

ALAN M. KLEINFELD

in order to understand membrane function at the molecular level (McCammon and Karplus. 1979; Welch et a l . , 1982). X-Ray diffraction and nuclear magnetic resonance (NMR) studies of lipid crystals have successfully determined lipid structure at high resolution. The structures determined from such studies have had a significant impact on the detailed understanding of lipid bilayer structure and formation (Hauser et ul., 1981). Understanding of integral or transmembrane protein structure is at a much more primitive level than that of lipid structure. Relatively few primary structures are known for membrane proteins, compared to the several thousand different proteins probably associated with cellular membranes. The best resolution obtained for an integral membrane protein is the photoreaction center of Rhodopseudoinonas viridis which has been solved to 3.0-A resolution by Deisenhofer et ul. (1983, using X-ray diffraction of three-dimensional crystals (about 2 A is necessary to identify individual side chains). The next best level of resolution is 7 A and this has only been achieved for a single protein, bacteriorhodopsin (Henderson and Unwin, 1975). Interactions between lipids have been studied extensively in model membranes and there is a reasonably clear understanding of many of the essential features of such interactions. Effects of acyl chain length, degree of acyl chain saturation, and the nature of the head group have been correlated with phospholipid packing and mobility, bilayer width, vesicle size, and the physical state of the membrane. Interactions between different lipids, especially binary mixtures of phospholipids, have been shown to lead to phase separation. It is known how the presence of sterols affect various membrane properties. Biological membranes although vastly more complex exhibit many of the properties of model membranes. Thus the study of model membranes is of considerable help in understanding biological membranes. The interaction between lipids and proteins is central to the understanding of membrane function. It is a subject which has received a great deal of experimental and theoretical attention. Membrane lipid must significantly affect protein function, since membrane proteins generally remain tightly bound to the lipid bilayer and it is likely that the protein structure is at least partially determined by its dissolution in the bilayer. The question then is whether this dissolution is passive, so that any medium of appropriate dielectric constant would achieve the same structure, or whether the interaction between the dissolved protein is specific to the nature of the lipid. The past several years have witnessed a considerable advance in our understanding of this issue, and a significant section of this article will be devoted to its discussion.

CURRENT VIEWS OF MEMBRANE STRUCTURE

3

II. LIPID STRUCTURE

Lipid structure is reasonably well understood at the angstrom level of resolution. X-Ray diffraction studies on single crystals and X-ray diffraction and N M R of aqueous lipid dispersions provide a reasonably complete picture of the molecular structure of the major lipids (see review by Hauser e t nl.. 1981). Although it has not yet been possible to use this structural information to derive a complete picture of the bilayer, these results. as discussed below, place severe restrictions on the possible forms of lipid associations. Molecular structures of phosphatidylcholine and phosphatidylet hanolamine obtained from single crystals are shown in Fig. 1 . This figure illustrates some of the general features of lipid structure. The axis formed by the phosphorylcholine (ethanolamine) groups of the a-chain is nearly perpendicular to the y-chain so that the head group lies parallel to the membrane surface. These results obtained by X-ray diffraction and N M R on single crystals (Hauser et a / . , 1981; Griffin ei a / . , 1978; Herzfeld ('2 d,, 1978) have been confirmed by X-ray and neutron diffraction and NMR on fully hydrated dispersions of phospholipids (Franks, 1976; Worcester and Franks, 1976; Seelig and Seelig, 1980). The surface parallel conformation of the head group is not affected by hydration or by the presence of cholesterol (Worcester and Franks, 1976). Thus the stability of the head group orientation is determined by intramolecular interactions and therefore this configuration probably prevails in biological as well as model membranes. Although the orientation of the phosphorylcholine or -ethanolamine axis relative to the plane of the membrane is rigid, the axis itself undergoes rapid rotation (>lO"/sec) about the C-I-C-2 axis and, of course, the molecule as a whole rotates about an axis normal to the surface. According to the crystal structure, a polar region can be defined which extends from the head group to the carbonyls of the glycerol backbone and has a thickness, depending upon head group and acyl chain length, of between 8 and 10 These results are in excellent agreement with the electron density profile obtained by X-ray diffraction on aqueous dispersions (Blaurock. 1982). Thus a significant fraction of the bilayer thickness may form a region of relatively high dielectric constant [also suggested by electron spin resonance (ESR) and fluorescence studies, see below]. The acyl chain orientation is parallcl to the surface normal in the liquid crystalline state and is either parallel or tilted to the normal in the gel state. In either case, however, the phospholipid crystal structure (Hauser et a / . . 1981) as well as N M R studies of model and biological membranes (Seelig and Seelig, 1980) indicate that the first two carbons of the P-chain

A.

4

ALAN M. KLEINFELD

Y-chain h

a-ch

B-chain FIG.I . X-Ray crystallographic structure of phosphatidylcholine. A similar structure was obtained for phosphatidylethanolamine.(Reprinted with permission from Hauser ct a/.. 1981.)

are parallel to the surface and therefore the P-chain does not extend as far into the bilayer as the y-chain. An important consequence of these studies is that structures determined from the solid crystal can be applied to the fully hydrated state. Even the rotational mobility of the head group about the C-14-2 axis can be inferred from the single crystal structures. On the other hand the strong hydrogen bonding between head groups which is observed in the crystal study is probably not as significant in the fully hydrated state, at least at temperatures above the phase transition where the average separation of head groups exceeds hydrogen bonding distances. Thus while the crystal structure provides an essential starting point for using constituent structures to predict the form of the physiologically interesting aggregate, information from other sources will be necessary to completely understand membrane structure. 111.

LIPID-LIPID INTERACTIONS: SINGLE LIPID SPECIES

A. Bilayer Structure

Aqueous dispersions of phospholipid give rise to a number of distinct structures (Luzzati, 1968) including the bilayer, hexagonal tubes, and micelles, as shown in Fig. 2. The nature of the phase depends on lipid species, lipid concentration, temperature, pH, and ionic strength. Attention has focused on the nonbilayer or hexagonal tube phase which has been found to coexist with the bilayer phase for special lipid mixtures in model systems (Cullis and de Kruijff, 1979; Hui ef al., 1981). Since there is little evidence that nonbilayer and bilayer structures coexist in biological membranes the following section will deal primarily with the bilayer phase.

CURRENT VIEWS OF MEMBRANE STRUCTURE

5 C

Molecular Shape Lysophosphdipids Detergents

----

Micellar

-v-

- - .- _ _ .

Inverted Cone

Sphingomyelin ..

Bilayer

'hosphotldytethandamine (unsaturated)

.-

.

..

Cylindrical

---&

- - -

.. .

'hosphatidic acid Hexagonal (H,,:

. .. .

Cone

FIG.2. Phospholipid phases in aqueous dispersions. The structure of each phase is shown in H, examples of phospholipids which form such phases are given in A. and the overall shape o f the phospholipid molecule is shown in C. (Reprinted with permission from Cullis and de Kruijff, 1979.)

Specific conditions are required for bilayer formation. Phosphatidylcholines with acyl chain lengths greater than 12 carbons, for example, are capable of forming bilayer phases over a wide range of conditions which include the ones relevant to the physiological state. Bilayers of pure phosphatidylethanolamine, however, form only within a restricted range of temperatures and at pH greater than 1 I . Structural studies on single lipid crystals provide insight into conditions necessary for bilayer formation (Hauser et al., 1981). These results indicate that the phosphatidylethanolamine head group occupies an area of about 38 A? in both the anhydrous state and in the presence of excess water. The phosphatidylcholine head group. on the other hand, occupies an area of about 50 A' in the anhydrous state and about 60-70 A in the presence of excess water. Below the transition temperature the sum of the acyl chain cross-sectional areas (for

6

ALAN M. KLEINFELD

saturated chains) is about 20 A’, small enough for these chains to pack either perpendicular or tilted to the surface of both phosphatidylethanolamine and phosphatidylcholine bilayers. As the temperature is raised above the transition temperature the acyl chain area expands to 50 A2. The acyl chains will therefore curve outward from the head group, forming the inverted micelle shown in Fig. 2. For phosphatidylethanolamine, where head groups maintain tight packing even in excess water, acyl chain-head group packing constraints are not consistent with a bilayer phase. The micelles that are formed under these conditions are not stable in isolation and combine to form the hexagonal tube phase (Fig. 2). At elevated pH, phosphatidylethanolamine acquires a net charge and the resulting electrostatic repulsion creates a sufficiently large head group area to accommodate a bilayer phase. In contrast, the head groups of hydrated phosphatidylcholine occupy a sufficiently large area for acyl chain expansion to occur within a bilayer phase. In the liquid crystalline state, in fact, the phosphatidylcholine head groups occupy a considerably greater area than is necessary to accomodate the acyl chains. Over a wide range of conditions, therefore, phosphatidylcholine is able to form bilayers. X-Ray diffraction studies on dispersions of phosphatidylcholines demonstrate that the thickness of the glycerohydrocarbon region increases from about 26 A for 12-carbon acyl chains to about 30 for the 18-carbon lipid (Cornell and Separovic, 1983). Since this increase is less than expected for fully extended acyl chains and since for the same variation in acyl chain length, the area per phospholipid is found to increase from 60 to 75 appreciable folding is suggested for the central portion of the acyl chains. The head to head width of biological membranes is between 40 and 45 A, according to X-ray diffraction studies on multilayer preparations (Blaurock, 1982). This value is consistent with the 30 A width of the glycerohydrocarbon region of an 18-carbon lipid and a head group to glycerol backbone thickness of between 5 and 7 A, from the lipid single crystal results (Hauser r f mf., 1981). The studies of the phosphatidylcholine bilayer thickness indicate that the terminal ends of the acyl chains fold rather than intercalate between the acyl chains of the opposite bilayer leaflet. This would seem to suggest that lipid-mediated coupling of the two bilayer leaflets (a possible transbilayer signaling mechanism) is not a significant factor in phosphatidylcholine bilayers. X-Ray diffraction and electron microscopy of phosphatidylcholines with mixed 10- and 18-carbon acyl chains suggest, however, that interdigitation, and therefore possible bilayer coupling, occurs when the bilayer is in the gel state (Mclntosh el nl., 1984; Hui ef nl., I 984).

A

A.

CURRENT VIEWS OF MEMBRANE STRUCTURE

7

B. Properties of the Bilayer Normal to the Surface The interior of the membrane is not well approximated by a uniform isotropic hydrocarbon fluid, as witnessed by several important properties of the bilayer that exhibit significant variation in the direction normal to the surface. The degree of order as a function of bilayer depth has been studied by 'H or "F NMK using lipids selectively labeled along the acyl chain (Seelig and Seelig, 1980; MacDonald e t a / . , 1984). The reflects the average orientation of each molecular order parameter S,,,,,, segment along the acyl chain. having a value of zero for the completely disordered isotropic chain and a value of unity for a chain in the alltrans state. Seelig and his co-workers have found that, except for a slight dip at the 2 position, Smo,is relatively constant between carbons I and 9 and then falls rapidly for positions deeper in the bilayer, reaching a minimum at the center. The value at the minimum is about 3-fold smaller than the maximum value at the surface of the bilayer. Similar profiles were obtained in the "F studies of MacDonald et a / . (1984). In the liquid crystalline state the profiles are relatively unaffected by fatty acyl chain substitutions. For Acho/c.p/nsma lcridlrrivii B membranes in the gel state. however, enrichment in palmitate. but not unsaturated fatty acids, abolishes the order profile gradient (MacDonald er a / . . 1984). The behavior observed in the 'H and '"F studies is characteristic of model and biological membranes. The order parameter profile has also been investigated by measuring the fluorescence decay anisotropy of the 11-(9-anthroyloxy)fatty acids (Vincent cr d . , 1982; Kutchai et d . , 1983; Storch and Schacter, 1985; A . M . Kleinfeld, unpublished observations). These fatty acids labeled at one of eight different positions along the chain are added exogenously to the membranes and can therefore be used to study a wide variety of membranes. In Fig. 3 the order parameter profiles determined by fluorescence anisotropy are shown for red cell ghosts, liposomes, and paraffin oil. These profiles are in good agreement with those obtained by NMR. The larger S values for ghosts presumably reflect the high cholesterol content of these membranes while, as expected, the isotropic solvent paraffin oil exhibits an almost flat profile. The order parameter reflects the ensembled averaged orientation and may not be simply related to the rotational rate or segmental mobility. Information on the acyl chain mobility has been obtained from "C-NMR and fluorescence decay anisotropy measurements (Lee rt N / . , 1974; Vincent ct a / . , 1982; Kutchai et a/., 1983: A . M . Kleinfeld, unpublished observations). The results of these studies indicate that the mobility profile does, in fact. parallel the order parameter variation.

ALAN

8

0

K

W

!

.42

-

v V

%a ~

M. KLEINFELD

.20

-

w

P I 0

0

+ +

3.6

v

V

t

t

0

v

t

v

t

V

4Y

I

I

I

J

7.2

10.8

14.4

10

ACYL CHAIN

POSITION

PIG. 3. Order parameter variation along fatty acyl chains a s determined by the decay anisotropy of the anthroyloxy fatty acids. Fluorescence decay anisotropies of the n-(9-anthroyloxy) fatty acids ( n between 2 and 16) were measured in red cell ghosts (U). small unilamellar vesicles of egg phosphatidylcholine (A). and parafin oil ( 1. Differential polarized phase lifetimes were measured at an excitation wavelength of 383 nm, and the results were analyzed according to Kutchai et N I . (1983). The order parameter was calculated as S = (r,/r,J"', where r , is the measured value at 383 nm and r, = 0.3. (From A. M. Kleinfeld. unpublished observations.)

+

The polarity of the bilayer displays appreciable variation with position along the acyl chain and probably reflects the degree of water penetration into the bilayer. The first measurements of the polarity profile were performed using the n-doxylstearic spin-labeled fatty acids (Griffith et al., 1974). These electron spin resonance (ESR) measurements, in fully hydrated membranes, demonstrated that the polarity at the C-5 position was greater than ethanol while the polarity between the C-12 and C-16 positions was intermediate between mineral oil and ethanol. In dehydrated membranes, however, all probes sensed a similar polarity, suggesting that the observed gradient was due to water penetration. Confirmation of the hydrated membrane profile has been obtained by fluorescence lifetime studies of the n-(9-anthroyloxy) probes (Chalpin and Kleinfeld, 1983). The degree of water penetration appears to be modulated by lipid composition (Simon et al., 1982). In membranes composed of 1 : I phosphatidylethanolamine: cholesterol the depth to which a significant amount of water can penetrate

CURRENT VIEWS OF MEMBRANE STRUCTURE

9

appears to decrease by about 3.5 A . compared to pure phosphatidylet hanolamine.

C. Phase Transitions

Phase transitions in lipid bilayers have provided an important tool for the investigation of membrane structure. The phenomenon is a collective one and is thought to hold the key to many important cellular events. It is readily observed in model bilayers and its occurrence (in the form of lateral phase separation, discussed below) has been observed in some cellular membranes (Chapman. 197s; Nagel, 1980). Lipid bilayer phase transitions involve changes primarily in the acyl chains from a less to a more disordered state. There are a number of bilayer phases through which the transitions can proceed, and the occurrence of these phases depends primarily on the head group. Using the nomenclature of Ranck r t trl. (1974). the phases below the main phase transition are designated with a p and those above with an a. The main transition is accompanied by a significant enthalphy change (-9 kcal/mol) and is therefore considered to be first order although, since it occurs with a tinite width, it is not rigorously a phase transition (Nagel, 1980). In phosphatidylcholine with saturated acyl chains there is a state in which the chains are all trans, tilted to the bilayer, and in a monoclinic form; this state is designated L,.. In addition. there is another state (the P,. slate) in which the chains are also all trans and tilted but in addition a long range ripple is superimposed upon the monoclinic gel state. This state is more disordered than the L,. and therefore occurs at higher temperature (and is sometimes called the pretransition state). It has been observed (Chen et a / . , 1980; Ruocco and Shipley, 1982; Fuldner. 1981) that in dipalmitoylphosphatidylcholine the gel state is not an equilibrium state and that after prolonged incubation at 0°C the gel state slowly (>3.5 days) transforms into a more ordered state. Upon heating, the transition from this state (crystalline Lp,) to L,. is accompanied by a net enthalpy change. Thus a full description of the transitions in dipalmitoylphosphatidylcholine is L,$.(crystalline) -+ L,. (gel) P,,. + L,,. Only the main transition is observed in phosphatidylethanolamine, and this occurs at approximately 20°C above the corresponding phosphatidylcholine transition (Nagel, 1980). In addition to changes in the acyl chain order, the bilayer width decreases and the area per lipid increases for ;I L,, to L, transition. The increase in lipid area is also associated with an increase in hydration from 1 I to 15 molecules water per dipalmitoylphosphatid ylcholine below the main transition, to about 27 above.

-

10

ALAN M. KLEINFELD

IV. LIPID MIXTURES AND DOMAINS It was implicit in the original formulation of the fluid mosaic model (Singer and Nicolson, 1972) that the membrane constituents formed a wellmixed fluid. Numerous studies now indicate that membrane components exhibit varying degrees of immiscibility. Rates of protein diffusion (as discussed in Edidin, this volume) are generally inconsistent with movement restricted solely by a homogeneous lipid viscosity and in some instances may be consistent with zero mobility. Studies on the lipid composition of biological membranes generally demonstrate compositional asymmetry between the inner and outer bilayer leaflets and therefore an immiscibility of the lipids across the bilayer (see Dawidowicz, this volume). Our focus here is on lateral immiscibility of lipids. Lateral immiscibility in the form of lateral phase separation has been amply demonstrated in model systems (Shimshick and McConnell, 1973; Grant et al., 1974; Luna and McConnell, 1977). Phase separation has also been demonstrated in specialized biological membranes or those perturbed by enrichment or depletion of a particular component (fatty acid supplementation or sterol depletion). A more difficult issue, and one which has received considerable attention recently, is whether lipid immiscibility is a general phenomenon in biological membranes, resulting in small and consequently difficult to detect regions of lipid phase separation. Since these issues have been the subject of reviews elsewhere only a brief discussion of this subject will be presented here (Bach et al., 1977; Jain and White, 1978; Karnovsky et al., 1982a,b; Klausner and Kleinfeld, 1984). A. Model Membranes

Phase behavior has been studied in model membranes composed primarily of binary mixtures. Investigations have focused on mixtures of phosphatidylcholine, using virtually every biophysical method as well as many biochemical techniques. To a lesser degree phosphatidylethanolamine, sterols, sphingolipids, and charged phospholipids have also been studied, It is generally accepted that there are four different phases or phase mixtures in binary systems. (1) For lipids with nearly identical structure and/or phase behavior, bilayers exhibit nearly ideal mixing, so that above some critical temperature the composite system is entirely fluid, while below this temperature the acyl chains are all trans. This is virtually indistinguishable from a single species phase transition discussed above. True ideal mixing, therefore, implies near homogeneity in composition. (2) Binary mixtures of phosphatidylcholine, such as dimyristoylcholinedipalmitoylphosphatidylcholine and phosphatidylcholine-sphingomylin

CURRENT VIEWS OF MEMBRANE STRUCTURE

11

(Lentz rt d., 1976a,b, 1981), in which the components are similar, exhibit nearly ideal mixing but differ somewhat in their phase transition temperature. Thus above a critical temperature the lipids form an ideally mixed liquid phase. As the temperature is reduced, a value is reached where the presence of the higher melting point species causes solid domains to form in equilibrium with the fluid state lipid. The solid domains thus formed are not homogeneous in composition, but have a greater proportion of the higher melting component at higher temperatures. When the temperature is lowered sufficiently the entire phase is a well-mixed solid phase in which both components cocrystallize. (3) For lipids sufficiently different from one another the liquid phase will approach ideality. Lowering the temperature yields a coexisting fluid-solid phase. Finally, as the temperature is reduced further two solid phases will form in which the lower melting point component crystallizes separately from a phase composed primarily of the higher melting point component. (4) In a few special instances liquid-liquid immiscibility is exhibited. This behavior appears to depend on the nature of the lipids (Wu and McConnell, 1975; Galla and Sackmann, 1975) or the radius of curvature of the bilayer (Lentz et ( i l . , I976a,b). The nature of the liquid-solid phase separation has been investigated by combining freeze-fracture electron microscopy, electron diffraction, and ESR (Shimshick and McConnell, 1973; Grant pt ol., 1974). In all cases examined thus far phase separation appears to occur laterally within the plane of a single hemileaflet. Thus the separation does not correspond to separation into distinct membranes (fission), nor does it appear that the phases of the two leaflets of the bilayer are correlated. Lateral phase separations can be induced isothermally, as for example in the case of cation (Ca'+ ) applied to phosphatidylserine-phosphatidylcholine o r phosphatidylcholine-phosphatidylethanolamine mixtures. The detailed topology of the phase separation is difficult to assess although some evidence suggests patches with dimensions of the order of 0.2-0.5 Fm (Hui, 1981). 6. Biological Membranes

Biological membranes are obviously more complicated than the binary model membranes discussed above. The increased heterogeneity manifests itself both in head group and in acyl chain diversity; a considerable fraction of the acyl chains are cis-unsaturated and would therefore be expected to undergo phase transitions at very low temperatures compared to 37°C. In spite of these factors, membranes obtained from cells specifically enriched in a particular fatty acid or depleted in sterols, exhibit thermotropic behavior similar to binary model membranes (Melchior, 1982; Klausncr

12

ALAN M. KLEINFELD

and Kleinfeld, 1984). In addition, there are a number of cells which exhibit macroscopic lateral phase separation (Bearer and Friend, 1980; Wolf, 1983; Wolf and Voglmayr, 1984). These large-scale segregations of certain lipid classes are probably a result of lipid-lipid interactions as well as lipid interactions with other cellular constituents. In addition to these relatively large effects, there appears to be a class of phenomena, suggesting domains or clusters of lipids on a microscopic scale, for which the evidence is more indirect. The evidence for such domains has been obtained from calorimetry, photobleaching, and spectroscopy. Many of these effects have been discovered at temperatures appreciably below the physiologically important ones. Clearly a central issue is whether membrane lipid domains are significant structural factors at physiological temperatures. Several studies do in fact indicate domain formation under physiological conditions (Brasitus et al., 1980; Mutsch rt al., 1983). A number of cellular phenomena have been interpreted in terms of lipid domains. These phenomena generally involve the observation that altering fatty acid composition, either by changing the lipid acyl chain composition (Horwitz et al., 1974; Hatten rt al., 1978) or adding exogenous fatty acids (Karnovsky et al., 1982a,b; Hill et al., 1983; Klausner and Kleinfeld, 1984). alters a specific cellular function although the fatty acids remain unesterified. Modulation of cellular function by sterol alteration also suggests domains in lymphocytes (Hoover et al., 1983). These observations suggest a number of physiological roles which domains might play in cellular function. For example, depending upon their function, proteins may be located in regions where the lipids are in a solid phase and partitioning of lipophilic molecules (fatty acids, drugs, hormones) into this region may serve as a trigger for a particular function or as a larger region to collect a substrate for the protein. There are also a number of studies which suggest that domain formation itself may occur in response to physiological stimulus or growth (Curtain et al., 1980; de Latt et ml., 1979; Packard et al., 1984). V.

MEMBRANE PROTEIN STRUCTURE

Complete understanding of cellular function and the effects of lipids on this function must ultimately be based on the molecular structure of membrane proteins. Although atomic detail of membrane protein structure is not yet available, considerable advances have occurred in the past decade. A number of reviews have surveyed recent results (Senior, 1983; Benga and Holmes, 1984; Eisenberg, 1984). In this section the salient features of membrane protein structure will be reviewed, with emphasis on the

CURRENT VIEWS OF MEMBRANE STRUCTURE

13

portion of the protein associated with lipid. This latter qualifier is necessary since there are a number of membrane proteins that have large cxtramembranous domains which can be cleaved from the membrane-bound portion, crystallized, and studied by conventional X-ray diffraction (Mathews et ul., 1979: Wilson et ul., 1981). In contrast, the structure of proteins that are primarily lipid associated is at a much less refined level. Purification and crystallization for these proteins, for example, present much greater difficulties than for water-soluble proteins. The seminal study of membrane protein structure was performed on the photosynthetic reaction center using X-ray diffraction on the threedimensional crystals of this complex (Deisenhofer et ul., 1985). The naturally occurring two-dimensional crystals of bacteriorhodopsin formed in the purple membrane of Hdohacteriurn lirrlohirrm have been studied to partially solve the structure of this highly integral membrane protein (Henderson and Unwin, 1975). Using electron diffraction and microscopy an image of the protein was obtained at a resolution of 7 A parallel and 14 A perpendicular to the plane of the membrane. More recently this resolution has been improved to 6.5 and I!: A, respectively (Liefer and Henderson, 1983). Although at this resolution the location and orientation of individual amino acid residues cannot be discerned, a general outline of the protein shape and secondary structure can be inferred. The model of Fig. 4 indicates that the protein is composed of seven rods, roughly arranged as a cylinder of radius 15 A . Approximately 70% of the cylinder, whose length is about 40 is contained within the membrane. Based upon the analysis of the electron density, measurements of the circular dichroism (CD), the primary sequence, and theoretical arguments. it has generally been concluded that the seven rods are a-helicies and that the segments connecting the loops are extramembranous and nonhelical 1979; (Henderson and Unwin, 1975; Long r t 01.. 1977; Ovchinnikov et d., Khorana et ul., 1979; Engelman el ul., 1980; Senior, 1983). Recent diffraction studies and CD measurements suggest, however, that several of the rods may be partially @-sheet(Jap et id., 1983). Plausable models based upon the a-helical configuration have been proposed for the spatial arrangement of the primary sequence through the membrane (Engelman r t ul., 1982; Ovchinnikov et ul., 1979; Stoeckenius and Bogomolni, 1982). These models are consistent with the electron diffraction results and with labeling and proteolytic digestion studies which define the topology of the extramembranous regions (Engelman c’t rrl., 1982; Agard and Stroud, 1982; Huang et nl., 1982). Outlines of the membrane-associated portions of several other proteins have been obtained at lower resolution. These include gap junctions (Makowski et a / . , 1984; Unwin and Zampighi, I980), ~ibiquinol:cytochrome c reductase (Leonard et ul.. 1981), cytochrome oxidase (Deatherage rt

A,

14

ALAN M. KLEINFELD *.Q

@,",

.L.

11.

@'" "' "'

ALA C L I

**O@,mo

1.1.

*LA

@

@

ALA

PA)cl

scn 1*1-1.0

GLl

l C1

Leu-

-7""-

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OL1

PLI LCU

.f*

ILC LCU

VAL

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I"".,

100

1.f aL. CLU)

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FIG.4. A model for the structure of bacteriorhodopsin. This proposed arrangement of the primary sequence through seven &-helices is based on electron diffraction, energy min-

imization. and biochemical evidence. (Reprinted with permission from Engelman 1982.)

ei

d..

al., 1982). and the acetylcholine receptor (Changeux et al., 1984). These

studies are generally consistent with a cylindrical shape for the membranespanning portion of the protein. In addition, high-resolution studies have been conducted on small membrane-associated peptides such as alamethicin (Fox and Richards, 1982) and melittin (Terwilliger et d., 1982). These studies have solved the structures at resolutions between 1.5 and 2.5 A,sufficient to determine the location of the amino acid side chains. The results are important for indicating how side chains interact with lipids and should be useful for analyzing larger proteins. In particular, the study on melittin suggests that hydrophobic residues associate with the lipid and hydrophilic residues orient toward the aqueous phase. A wide variety of experimental methods have been used to determine the location of particular portions of membrane-bound proteins and to relate these locations to the membrane architecture. These approaches complement diffraction methods that are as yet unable to distinguish individual residues. They also offer the possibility of monitoring at least

CURRENT VIEWS OF MEMBRANE STRUCTURE

15

part of the protein structure irr sitrr and possibly in different physiological states. Thus chemical labeling and proteolysis with selected proteases have been used to determine the number of times the protein loops across the membrane as well as the location and general topology of the extramembranous segments (band 3, Brock ct t i / . , 1983; bacteriorhodopsin, Engleman et a / . , 1980; histocompatibility antigen, Cardoza cr t i / . , 1984). Regions within the bilayer have been mapped out using feiritin labeling (Henderson et ( I / . , 19781, photoactivatable reagents attached to fatty acids (Robson cf a/., 1982; Takagaki et t i / . , 1983), and fluorescence quenching of tryptophan with spin-labeled fatty acids (London and Feigenson, 198 1a.b). Resonance energy transfer between tryptophan and lipid-associated acceptors has been used successfully to locate individual tryptophan residues and may allow the determination of the distribution of multiple tryptophan residues (Fleming ef d . , 1979; Kleinfeld and Lukacovic, 198.5; Kleinfeld, 198.5). Secondary structure of a number of membrane proteins has been studied by circular dichroism. Many of these studies suggest that the major portion 1977; of the hydrophobic region is a-helical (Guidotti, 1977; Long or d., Senior, 1983; Wallace ef id., 1986). Theoretical arguments have been presented suggesting that the a-helix is the energetically preferred configuration (Engelman and Stietz, 1981 ; Senior, 1983). Several circular dichroism and X-ray diffraction studies. however, have found appreciable p structure (Makowski e t a / . , 1982; Jap c t t i / . , 1983; Wallace er (11.. 1986). In fact. predictions of secondary structure based on the primary structure usually favor p-strand in the hydrophobic stretches of membrane proteins (Senior, 1983). p-Turns or -bends within the membrane have been thought to be energetically unfavorable, yet evcn these structures have been found in at least two cases (Dailey and Strittmatter, 1981; Cordoza ct a l . , 1984). Thus it is likely that the internal structure of membrane proteins is composed of both a-helix and p-sheet (Wallace ct t i l . , 1986). Since membrane protein structural information is so limited it is important to make use of theoretical as well as experimental information a s a guide in further analysis. Amino acid composition and primary structures have been examined in the hope that systematic behavior would emerge that would make it possible to characterize a protein as membrane or water soluble. The average hydrophobicity of the total amino acid composition does not appear to be a reliable predictor. Membrane proteins, however, i n contrast to water-soluble proteins, frequently possess stretches of hydrophobic residues that are long enough (>20 residues) to span the lipid bilayer. The correlation between hydrophobicity and the hydrophobic moment has been shown to be a reasonably successful prcdictor of protein classification (Eisenberg, 1984). On a plot ot of the hydrophobic moment versus the hydrophobicity per residue, proteins separate into surface (large moments and therefore asymmetric and relatively

16

ALAN M. KLEINFELD

small hydrophobicities), globular (intermediate moments and hydrophobicities), and transmembrane (small moments and large hydrophobicities). Hydrophobicity alone does not predict whether a residue will be located within the bilayer region since the most plausible models of bacteriorhodopsin, for example, suggest that a number of charged residues are located within the membrane (Engleman et al., 1980). To account for the considerable energy cost in burying a charged residue (20 kcal/mol) it has been suggested that the charged residues are distributed near the central axis of the cylinder formed by the seven rods and that hydrophobic residues are located at the periphery, in contact with the lipid hydrocarbon chains (Engelman and Zacci, 1980). VI. INTERACTIONS BETWEEN LIPIDS AND PROTEINS

Interactions between lipids and proteins have been studied by observing the effect of lipid on protein and the effect of protein on lipid structure. Proteins have been found to alter the order and mobility of the lipid acyl chain, the translational mobility of the whole lipid, and the lateral organization of the bulk lipid. The effect of lipid on protein structure and dynamics is less well characterized. A number of studies do, however, suggest alterations in specific protein segments in response to changes in lipid structure. In this section these two effects are discussed separately since they are studied by different methods and they potentially represent different physiological functions. A. Effects of Lipids on Proteins

Lipid-protein interactions involve the alteration in secondary, tertiary, or quaternary protein structure induced by changes in the composition or physical state of the lipid. Several studies in reconstituted systems using magnetic resonance have demonstrated that the mobility of individual amino acid side chains can be altered by changes in lipid phase (Hagan et al., 1978; Stollery et al., 1980; Boggs et ul., 1980). Secondary structural changes in response to alterations in lipid composition and lipid to protein ratio have been observed in reconstituted glycosyltransferase (Beadling and Rothfield, 1978) and fd coat protein (Dunker et al., 1982). Evidence also exists which demonstrates that fatty acid perturbation in plasma membranes may affect the conformation of individual amino acid residues (Esfahani and Devlin, 1982; Pjura et al., 1982). Interactions of small peptides with membrane indicate that binding occurs with a specific orientation relative to the membrane surface (Eisenberg, 1984) but that the actual

CURRENT VIEWS OF MEMBRANE STRUCTURE

17

conformational changes are quite subtle (Deber and Benham, 1984). A large body of indirect evidence for lipid-protein interactions has been obtained from functional changes observed in response to lipid and fatty acid alterations (Melchior and Steim, 1976; Criado et ul., 1984; Stubbs and Smith, 1984, Benga and Holmes, 1984). B. Effects of Proteins on Lipids

Protein-lipid interactions have attracted a great deal of attention in the past decade. This interest has been encouraged by the availability of a rich variety of lipid probes and by increased understanding of reconstitution of proteins into lipid bilayers. The central issue in these investigations has been whether and how the presence of protein in membranes affects lipid structure and dynamics. Proteins may affect the local properties of individual lipid molecules, such as the acyl chain order or mobility, or more global properties, such as the lateral and transmembrane distribution and mobility. Transmembrane effects will be discussed in Dembo and Bell (this volume) and lateral mobility in Edidin (this volume). In this section we will not deal with interactions affecting the transmembrane lipid properties and will only briefly touch upon lateral dynamics. The issue which has generated the most interest (and controversy) is whether a tightly bound lipid boundary is formed around transmembrane proteins. Tightness of binding is directly proportional to the residence time of a particular lipid in the protein-lipid interface. There are two important time scales with which to evaluate this residence time: the time characteristic of the protein’s function and the bulk lipid residence time. Since times which are important for protein function are probably greater than lo-‘ sec the lipid residence time should be greater than this in order for the binding to be functionally important. The bulk lipid residence time, the average time one lipid spends near another, is about lO-’sec (Marsh et al., 1982). This, therefore, is the shortest residence time in which any specific lipid can play a structural role. In this section we will pay particular attention to the issue of the lipid boundary, using these time limits to gauge the significance of protein-lipid interactions. The first direct physical evidence that lipid at the protein-lipid interface might exhibit special properties was obtained using cytochrome c oxidase reconstituted into liposomes composed of mitochondria1 lipid and a spinlabeled fatty acid (Jost et al.. 1973). The ESR spectra from such preparations suggest two environments, one in which the spin label exhibits the nearly isotropic tumbling that it does in pure lipid liposomes (only a single environment is detected in pure lipid) and one in which the probe is motionally restricted. Variation of the lipid to protein ratio demonstrated

18

ALAN M. KLEINFELD

that over a broad range of values the immobilized component was proportional to the protein concentration. At sufficiently high concentrations virtually all the probe was immobilized. From the observed saturation ratio it was estimated that the boundary lipid constituted one lipid layer or annulus around the protein. It was suggested that in normal membranes an annulus formed around integral membrane proteins and that the lipids in this annulus were tightly bound. Subsequent studies using ESR, fluorescence, and raman spectroscopy on several other reconstituted and plasma membrane preparations have confirmed the existence of immobilized lipid related to the presence of protein (Hesketh et al., 1976; Cable and Powell, 1980; Marsh and Watts, 1982; Boggs et al., 1982; Thomas et NI., 1982; Wolber and Hudson, 1982; Taraschi and Mendelsohn, 1980; Levin ec NI., 1982). The observation of an immobilized lipid fraction by these techniques does not, however, constitute evidence for immobilized lipid on the time scale of protein function or the bulk lipid exchange rate. These techniques have time windows which range from lo-* to sec. In the case of ESR, for example, it is only necessary for the spin label to exchange between the annulus and the bulk lipid at rates slower than about 108/sec for an immobilized component to appear in the ESR spectrum. NMR, which is sensitive to much slower events (exchange rates smaller than IO’lsec) can be used in the same way as ESR to detect regions of immobilized lipid. In most NMR studies no evidence of an immobile lipid component is observed, and therefore these studies establish a lower limit to the exchange rate of IO’/sec (Oldfield et al., 1978; Seelig et al., 1981, 1982). The constraints set by the ESR and NMR results therefore require that if bound lipid exists its mean residence time in the annulus is between IO-’and IO-’sec. Indeed, there is some evidence obtained from the analysis of the position and line shape of the ESR spectra that the fatty acid spin label exchange rate is about 107/sec(Marsh et al., 1982). This value is about the same as the bulk lipid exchange rate, suggesting that the exchange of lipid in the annulus with bulk lipid is unhindered and, therefore, that the annular lipid is not tightly bound to the protein. Spectroscopic techniques generally reveal a component which is rotationally immobile for times longer than the temporal resolution of the method and, therefore, only indirectly reflect lipid to protein binding affinities. Measurements of the effect of protein on lipid lateral mobility and the lipid-protein association constants represent more direct approaches to the question of how tightly lipid is bound to the protein surface. Several investigations using fluorescence photobleaching recovery to measure the lipid lateral mobility in the presence and absence of protein have been carried out in reconstituted and native membranes (Golan et al., 1984).

CURRENT VIEWS OF MEMBRANE STRUCTURE

19

Results in red cells indicate that the rate of lipid diffusion is slower by a factor of 4 in whole membranes a s compared to liposomes formed from extracted lipid (Golan el NI., 1984). This decrease, however, is entirely explicable in terms of lipid collisions with proteins acting as simple-hard surfaces, suggesting, therefore, that the protein-lipid interaction does not ex hi bi t a significant attractive interact ion. The association constant for the binding of lipid at the protein-lipid interface may be determined from the partition of lipid between the annulus and bulk regions, using spectroscopic markers to indicate the fraction in each region. In addition to the ESR method (Griffith c’t [ I / . , 1982). spinlabeled fatty acids and brominated phospholipids have been used as shortranged quenchers of intrinsic protein fluorescence, to determine the fraction of these probes in close apposition to the protein surface (London and Feigenson, 1981a,b; East and Lee, 1982). The results of these studies indicate that the partition of neutral lipid between the boundary and bulk phases is not significantly different from unity. Fluorescence energy transfer between tryptophan and fluorescent fatty acids suggests that the fatty acid is excluded from a region in immediate apposition to the protein. This is consistent with a preferential association of the phospholipid with the protein as compared to the fatty acid and with a tightly bound lipid annulus (Fleming ef ( I / . , 1979; Lee et ( I / . , 1982; Kleinfeld and Lukacovic, 198s). I t is possible, however, that this represents not tight binding of [he lipids to the protein surface but rather the decreased partition of the lipid probes into a region in which the acyl chain free energy is reduced by restrictions to rotation or by electrostatic repulsion between the protein and the charged probes. Although there is little evidence for specific binding of neutral lipid to the lipid-protein boundary, a number of studies suggest that tight binding may be exhibited by charged lipids and will, therefore, probably be protein specific. Proteins from myelin, studied by a number of techniques. appear to induce formation of domains, enriched in charged lipids, from mixtures 1982). The lipid in these domains of neutral and charged lipids (Roggs et d., as well as in the bulk phase retain their unmixed phase behavior suggesting that the segregation occurs without significantly affecting the acyl chain properties. Studies on cytochrome c’ oxidase and the matrix protein of vesicular stomatitis virus also indicate the segregation of charged lipids and demonstrate an approximately 2-fold increase in partition preference of the charged lipids for the boundary. in comparison to neutral lipid (Cable and Powell, 1980: Griffith ct NI., 1982; Wiener c’t ( I / . , 1983). The Ca” ATPase from sarcoplasniic reticulum, on the other hand, displays no binding specificity for charged lipids (East and Lee, 1982). Although proteins may not retard lateral diffusion of lipids other than

20

ALAN M. KLEINFELD

by providing simple barriers, a number of studies have demonstrated an effect on acyl chain order and mobility (the effect of proteins on transbilayer movement is dealt with in the article by Dawidowicz in this volume, where it is suggested that proteins tend to enhance rather than retard such movement). Deuterium-NMR studies in reconstituted systems of several proteins demonstrate that the degree of acyl chain order decreases in response to protein incorporation (Oldfield el al., 1978; Seelig et al., 1982). This may reflect the influence of the uneven, relatively convoluted surface of the protein on the packing of the neighboring lipid acyl chains (Seelig ef a!. , 1982). Measurements of the fluorescence decay anisotropy of the fatty acid analog paranaric acid confirm the decrease in order but also indicate that the rotational mobility of the lipid acyl chains is decreased by protein (Wolber and Hudson, 1982). The decrease in rotational mobility is consistent with the protein side chains acting to retard the gauche-trans rotation of the lipid methylene groups. Thus it appears that virtually all measured protein interactions are consistent with the protein acting to retard movement by virtue of the slow diffusion of the protein mass as compared to the lipid. These results can in fact be understood in terms of an “infinite” barrier presented by the protein. A lipid molecule adjacent to a protein differs from one in the bulk lipid phase since its possible jump positions are limited by the protein. As the relative protein concentration increases some of the lipids will also be trapped within aggregates of protein and display further immobility. Although these results suggest that on average there is no specific protein-neutral lipid interaction it should be emphasized that only a small fraction of membrane proteins has thus far been studied. ACKNOWLEDGMENTS

I would like to thank Dr. Judith Storch for informative discussions about membrane structure and for her wise counsel concerning the manuscript. I would also like to t h a n k Michael Toon and Sean Condon for their careful reading of the manuscript. This work was supported by a Grant-in-Aid from the American Heart Association and with funds contributed in part by the Massachusetts affiliate (83-789) and a grant from the National Science Foundation (PCM-830268). This work was done during the tenure of an Established Investigatorship of the American Heart Association and with funds contributed in part by the Massachusetts affiliate (82-174). REFERENCES Agard, D. A,, and Stroud, R. M. (1982). Linking regions between helices in bacteriorhodopsin revealed. Biophys. J . 37, 589-602. Bach, D., Bursuker, I., and Goldman, R. (1977). Differential scanning calorimetry and enzyme

CURRENT VIEWS OF MEMBRANE STRUCTURE

21

activity of rat liver microsomes in the presence and absence of I-tetrahydrocannabinol. Biocliim. Bio~phys.Acrci. 469, I7 1-1 79. Beadling. L.. and Rothfield. I,. I. (1978). Modulation of the conformation of a membrane glycosyltransferase by specific lipids. Proc. Nurl. Accid. Sc,i. U.S.A. 75, 3669-3672. Bearer, E. L.. and Freind, D. S . (1980). Anionic lipid domains: Correlation with functional topography in a mammalian cell membrane. Proc.. Ntrtl. Ac,tid. Sci. U . S . A . 77. 66016605. Benga, C . , and Holmes, R . P. (19841. Interactions between components in biological membranes and their implications for membrane function. I'rog. Biophvs. 43, 19.5-257. Blaurock. A . E . ( 1982). Evinence of bilayer structure and of membrane interactions from X-ray diffraction analysis. Bioc,/iirn. Biophy.~.Acrcr 650, 167-207. Boggs, J . M.. Stollery, J . G . , and Moscarello. M. A . (1980).Effect of lipid environment on (he motion of a spin-label covalently bound to myelin basic protein. Bio~hemi,sri:v19, 1226-1 234. Boggs. J . M., Moscarello. M . A,. and Papahadjopoulos. D. (1982). Structural organization of niyelin-Role of lipid-protein interactions determined in model systems. I n "LipidProtein Interactions" (P. C. Jost and 0. H. Griffith. eds.), Vol. 2 , p. I . Wiley. New York. Brasitus, T. A,. Tall, A. R.. and Schachter. I). ( 1980). Thermotropic transitions in rat intestinal plasma membranes studied by differential scanning calorimetry and fluorescence polarization. Bioc~lrc~rnistr?, 19. 1256-1261 . Brock, C. J . . Tanner. M. J. A,. and Kempf. C. (1983).The human erythrocyte anion-transpot1 protein. Bioc,hc.m. J. 213. 577-586. Cable. M. B . . and Powell. C. I,. ( 1980). Spin-labeled cardiolipin: Preferential segregation in the boundary layer of cytochrome c oxidase. Bioc~hrrni.sriy19, 5679-5686. Cardoza. J . I).. Kleinfeld, A. M.. Stallcup. K. C . . and Mescher, M. F. (19x4). Hairpin tr:~ configuration of H-2Kh in liposomes formed by detergent dialysis. B i o c ~ h c ~ ~ i r i s23, 440 1-4409. Chalpin. D. B . . and Kleinfeld. A . M. (1983). Interaction of fluorescent quenchers with the . 731, 4 6 5 4 7 4 . n-(9-anthroyloxy) fatty acid membrane probes. Bioc./iirn. B i o p l i y . ~Acfcr Changeux, J . , Devillers-Thiery, A,. and Cheniouilli, P. ( 1984). Acetylcholine receptor: an allosteric protein. ScYcnct, 225, 1335-1345. Chapman, D. (1975). Phase transitions and tluidity characteristics of lipids and cell membranes. Q . Rci.. Biophys. 8. 185-235. Chen. S . C.. Sturtevant, J . M . , and Gaffney. B. J . (1980). Scanning calorimetric evidence for a third phase transition in phosphatidylchoiine bilayers. Proc.. N i t / / .Ac,trif. S(.i. 1J.S.A. 77, 5060-SO63. Cornell. A , . and Separovic, F. (1983). Membrane thickness and acyl chain length. B i ( ~ ~ / i i i ~ i . Biopliys . A c'rtr 733, 189- 193. Criado. M., Eibl, H., and Barrantes. F. J . (1984). Functional properties o f t h e acetylcholine receptor incorporated in model lipid membranes. J . Biol. Chem. 259. 9188-9198. Cullis, P. R . . and de Kruijff. B. ( 1979). Lipid polymorphism and the functional roles of lipids in biological membranes. Biochim. Bioplrvs. Acrtr 559, 399420. Curtain. C . C.. Looney. F. D.. and Smelstorius. J . A. (1980). Lipid domain formation and ligand-induced lymphocyte membrane changes. Biocliirn. Biophps. Acrtr 5Y6, 43-56. Dailey. H. A , . and Strittmatter. P. (1978). Structural and functional properties o f t h e membrane-binding segment of cytochrome h,. J . Biol. Clrern. 253, 8203-8209. Dailey. H . A,. and Strittmatter. P. (1981). Orientation of the carboxyl and NH-. termini of the membrane-binding segment of cytochrome h, on the same side of phospholipid bilayers. J . Biol. C l r m . 256, 395 1-3955.

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Deatherage, J. F., Henderson, R., and Capaldi, R. A. (1982). Relationship between membrane and cytoplasmic domains in cytochrome c oxidase by electron microscopy in media of different density. J . M d . B i d . 158, 501-514. Deisenhoffer. J., Epp, O., Miki. K., Huber, R., and Michel. H. (1985). Structure of the protein subunits in the photosynthetic reaction centre of Rliodopsertdo/nonus viridis at 3A resolution. N o t ~ r e(London) 318, 618-624. Deber. C. M.. and Benham. B. A. (1984). Role of membrane lipids in peptide hormone function: Binding of enkephalins to micelles. Proc. N u t / . Acud. Sci. U . S . A . 81, 61-65, De Laat, S . W., Van Der Saag. P. T., Elson, E. L., and Schlessinger, J. (1979). Lateral diffusion of membrane lipids and proteins is increased specifically in neurites of differentiating neuroblastoma cells. Biockim. Biophys. Acta 558, 247-250. Dunker, A. K., Fodor, S . P. A,, and Williams, R. W. (1982). Lipid-dependent structural changes of an amphomorphic membrane protein. Biophys. J . 37, 201-203. East, J . M., and Lee, A. G. (1982). Lipid selectivity of the calcium and magnesium ion dependent adenosinetriphosphatase, studied d with fluorescence quenching by a brominated phospholipid. Biochemistry 21, 4144-4151. Eisenberg, D. (1984). Three-dimensional structure of membrane and surface proteins. Annrc. Rev. Biocliem. 53, 595-623. Engelman, D. M., and Steitz, T. A . (1981). The spontaneous insertion of proteins into and across membranes: The helical hairpin hypothesis. Cell 23, 41 1-422. Engelman, D. M., and Zacci, G. (1980). Bacteriorhodopsin is an inside-out protein. Proc. Nrrtl. A w d . Sci. U . S . A . 77, 5894-5898. Engelman. D. M., Henderson, R., McLachlan, A. D., and Wallace, B. A. (1980). Path of the polypeptide in bacteriorhodopsin. Proc. N u t / . Acud. Sci. U . S . A . 77, 2023-2027. Engelman. D. M., Goldman, A.. and Steitz, T. A. (1982). The identification of helical segments in the polypeptide chain of bacteriorhodopsin. Methods Enzyrnol. 88, 81-88. Esfahani, M., and Devlin, T. M . (1982). Effects of lipid fluidity on quenching characteristics of tryptophan fluorescence in yeast plasma membrane. J . B i d . C h e m . 257, 99199921.

Fleming, P. J., Koppel, D. E., Lan, A. L. Y.,and Strittmatter, P. (1979). lntramembrane position of the fluorescent tryptophanyl residue in membrane-bound cytochrome b,. Biocliemistry 18, 5458-5464. Fox, R. O., Jr., and Richards. F. M. (1982). A voltage-gated ion channel model inferred from the crystal structure of alamethicin at I 5-A Resolution. Nature (London) 300, 325-330.

Franks. N . P. (1976). Structural analysis of hydrated egg lecithin and cholesterol bilayers. I. X-Ray diffraction. J . Mol. B i d . 100, 345-358. Fuldner. H. H. (1981). Characterization of a third lipid transition in multilamellar dipalmitoyllecithin liposomes. Biochemistry 20, 5707-57 10. Galla, H. J . , and Sackmann, E. (1975). Chemically induced lipid phase separation in model membranes containing charged lipids: A spin label study. Bioclzim. Biophvs. Acts 401, 509-529. Golan. D. E., Alecio, M. R., Veatch, W . R. and Rando, R. R. (1984). Lateral mobility of

phospholipid and cholesterol in the human erythrocyte membrane: Effects of proteinlipid interactions. Biochemistry 23, 332-339. Grant, C. W. M.. W u , S. H.-W.. and McConnell, H. M. (1974). Lateral phase separations in binary lipid mixtures: Correlation between spin label and freeze-fracture electron microscope studies. Biochim. Biophys. Acia 363, 151-158. Griffin. R. G . . Powers, L., and Pershan. P. S . (1978). Head-group conformation in phospholipids: A phosphorus-3 I nuclear magnetic resonance study of oriented monodomain dipalmitoylphosphatidylcholinebilayers. Biochemistry 17, 2718-2722.

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23

Griffith. 0 . H.. Ikhlinger. P. J.. and Van. S. 1’. 11974). Shape o f the hydrophobic barrier o f phospholipid bilayers (evidence for water penetration in hiological membranes). J . M<’l?lhl’.BiO/. 15, 159-192. Griffith. 0. H.. Hrotherus. J. R.. and Jost. P. C’.11982). Equilibrium constants and number o f hinding sites for lipid-protein interaction\ in membranes. f r i “Lipid-Protein Interactions” (P. C. Jost and 0. H. Criftith. eds.). Vol. ?, p. 225. Wiley. New York. Cuidotti. G. ( 1977). The structure o f intrinsic membrane proteins. J . .Sitpr(/mol. .Y/ruc,!. 7, 489-497. Hagen. D. S.. Weiner, J . H.. and Sykes, B . I). (197X). Fluorotyrosine MI3 coat protein: Fluorine-19 nuclear magnetic rewnance s ~ u d yo f the motional properties o f an integral membrane protein in phospholipid vesicles. Rioc.lrrrtiisrry 17, 3860-3866, Hatten. M . E., Scandella. C. J . , Horwitz. A. F.. and Burger. M. M. 11978). Similarities in the membrane fluidity of 313 and SVIOI-37’3 cells and its relation to concaniiviilin A and wheat germ agglutinin-induced agglutination. J . Riol. Clie/n. 253, 1972-1977. Hauser. H., Pascher, I., Pearson. R. H.. and Sundell. S. (19x1). Preferred conformation and mo1ecul;tr packing o f phosphatitlylethiinolamine and phosphatidylcholine. fjioc/ii/ti.

Biophps.

A ( , f t i 650,

21-51.

Henderson. K . , and Unwin. P. N . T. 11975). Three-dimensional model o f purple membrane obtained by electron microscopy. Nci//rrc, ILoriclorr) 257, 23-32. Henderson, R., Jubb, J. S.. and Whytock. S. (1978). Specific labelling o f the protein and lipid on the extracellular surface of purple membrane. J . Mol. Biol. 123, 259-274. HerLfeld. J.. Griffin, R. G.. and Haberkorn. K . A . 11978). Phosphorus-31 chemical shift tensors i n barium diethyl phosphate and urea-phosphoric acid: Model compounds for phospholipid head-group studies. Bioc.lic~rnis/ry17, 27 I 1-27 18. Hesketh, T . R.. Smith, G . A.. Houslay, M . D., McGill. K. A , , Birdsall. N. J . M.. Metcalle. J . C.. and Warren. G. B . (1976). Annular lipids determine the ATPase activity o f a 15. 4 145calcium transport protein complexed with dip~ilmitoyllecithin.Bioc~hr~rrii,s/r~ 4151. Hill. D. I . . Dawidowicz. E. A., Andrews. M . L... and Karnovsky. M. J. (1983). Modulation o f microsomal glucose-6-phosphate translocase activity by free fatty acids: Implications for lipid domain structure in niicrosonial membranes, J . Crll. Phpsiol. 11.5, 1-8. Hoover, R. L.. Dawidowicz, E. A., Robinson. J . M.. and Karnovsky. M. J . (19x3). Role of cholesterol in the capping of surface immunoglobulin receptors on murine lymphocytes. J . Cell. B i d . 97. 73-80. Horwitr. A. F.. Hatten. M. E., and Rerger. M. M . (1974). Membrane fatty acid replacements and their effect on growth and lectin-induced agglulinability. Proc. Ntc/l. A c ~ i t l .S(,i.

U . S . A . 71, 3115-3119. Huang. K.-S.. Liao, M.-J.. Gupta. C. M.. Royal, N . . Beimann. K.. and Khorana. H. G . (1982). The site o f attachment o f retinal i n hacteriorhodopsin. J . Eiol. Chcrii. 257, X.596-

8599. Hui. S . W. (1981). Geometry o f phase-separated doninins i n phospholipid hilayers hy diffraction-contrast electron microscopy. Bioplip.s. J . 34, 383-395. Hui. S. W.. Stewart. T. P.. Yeagle. P. L., and Albert. A . D. 11981). Bilayer to non-bilayer transition in mixtures o f phosphatidylethanolamine and phosphatidylcholine: Implications for membrane properties. Arch. Bioclicrrr. Eiopliy.\. 207. 227-240. Hui. S. W.. Mason. J. T.. and Huang, C. 11984). Acyl chain interdigitation in saturated mixed-chain phosphatidylcholine bilayer dispersions. Bioc,/icrriisrry 23, 5570-5577. Jain, M. K.. and White, H. B . (1978). Long-range order i n biomembranes. A h . Lipid R r s . 15. 1-60. Jap, B . K.. Maestre. M. F . . Hayward. S. B., and Glaeser, R. M. 11983). Peptidc-chain secondary structure o f bacteriorhodopsin. Bioplr,vs. J . 43, X 1-89,

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Jost, P. C., Grifith, 0.H., Capaldi, R . A., and Vanderkooi, G. (1973). Evidence for boundary lipid in membranes. Proc,. Nut/. Accid. Sci. U . S . A . 70, 480-484. Karnovsky. M. J., Kleinfeld, A. M., Hoover, R. L., Dawidowicz, E. A,, Maclntyre, D. E., Salzman, E. A., and Klausner, R. D. (1982a). Lipid domains in membranes. Ann. N . Y . Acc/d. Sci. 401, 61-75. Karnovsky, M. J., Kleinfeld. A. M., Hoover, R. L., and Klausner, R. D. (1982b). The concept of lipid domains in membranes. J . Cell. Biol. 49, 1-6. Khorana, H . G., Gerber, G . E., Herlihy, W. C.. Gray, C. P., Anderegg, R . J., Nihei. K., and Biemann, K. (1979). Amino acid sequence of bacteriorhodopsin. Proc. NLIII.Actid. Sci. U . S . A . 76, 5046-5050. Klausner, R. D., and Kleinfeld, A. M. (1984). Lipid domains in membranes. In “Cell Surface Dynamics” (A. S. Perelson, C. H. DeLisi, and F. W. Wiegel, eds.), pp. 23-58. Dekker, New York. Kleinfeld. A. M. ( 1985). Tryptophan imaging of membrane proteins. Biochemi.7rr.y 24, 18741882. Kleinfeld. A. M., and Lukacovic. M. I,. (1985). Energy transfer study of cytochrome h, using the anthroyloxy fatty acid membrane probes. Biocliernistry 24, 1883-1890. Kutchai. H., Chandler, L. H., and Zavoico, G . B . (1983). Effects of cholesterol on acyl chain dynamics in multilamellar vesicles of various phosphatidylcholines. Bioclriwz. Biclph.V.7. Act
CURRENT VIEWS OF MEMBRANE STRUCTURE

25

membranes: Purple membrane o f f-ltrlohtrc,/erirrr,i / i d o h i w i . Eioc,/icrrr. E i o p / i j , s . K c ~ s . Co//lln/f/l.75, 725-73 I. Luna, E. J . , and McConnell, H. M . (1977). I-literal phase separation:, in hinary mixfurch of phospholipids having different charges and different crystalline structures. Bioc./rirrr. BiOphy,S. A ( . / < 470, / 303-3 16. L u n a t i . V. (1968). X-Kay diffraction studies o f lipid-water systems. f r i "Biological Membranes" (1). Chapman, ed.). p. 71-123. Academic Press. London. McCammon, J . A., and Karplus, M. (1979). 1)ynamics of activated procesyes i n globular proteins. Proc. N o / / . ActrJ. S c . i . U.S.A. 76, 3585-3589. MacDonald, P. M.. Sykes. B . I).. and McElhaney, K . N. (1984). Fatty acyl chain \ti-tictiire. orientational order, and the lipid phase transition in Af.holc/,/rr.sr,itr / t r i d / t r ) i i i H niembranes. A review o f recent '"F nuclear magnetic resonance studies. Ctrtr. J . Bioc./rc~rrr. 62, 1134-1 150. Mclntosh. T. J . . Simon. S . A,. Ellington. Jr.. J . C.. and Porter. N. A . (1984). New structural model for mixed-chain phosphatidylcholine bilayers. Biochc.mi.srry 23, 40384044. Makowski, L . , Caspar. D. L. I).,Goodenough. D . A.. and Phillips. W. C. (1982). Gap junction structures. 111. The effect of variations in the isolation procedure. Eiop/ry.\. J . 37, 189-191. Makowski, LA.. Caspai-. D. L. D.. Phillips. W. C., Raker. T. S.. and Goodenough, I). A . ( 1984). Gap junction structures. VI. Variation and conservation in connexon conformation and packing. Eiopliy.\. J . 45, 208-2IX. Marsh. I).. and Watts. A. ( 1982). Spin labeling and lipid-protein interactions in membranes. I t i "Lipid-Protein Interactions" (P. C'. Jost and 0. H. Griffith. edh.). Vol. 1. p. 53126. Wiley, New York. Marsh. 0.. Watts, A , . Pates. D.. Uhl. R.. Knowles. P. F., and Esmann, M. (1982). ESK spin-label studies of lipid-protein interactions in membranes. Biophys. J . 37, 165-

214. Mathews. F. S.. Czerwiski. E. W . . and Argos. P. (lY7Y). The X-ray crystallographic s t ~ ~ ~ c t t ~ r e of calf liver cytochrome h,. Irr "The Porphyrins" (I). Dolphin. 4.). Vol. 3. p. 107-145, Academic Press, New York. Melchior. D. L. (1982). Lipid phase transition., and regulation of membrane fluidity in prokaryotes. Crrrr. Top. Mcruhr. Trtrnsp. 17, 263-3 16. Melchior. D. L.. and Steim. J . M. ( 1976). Thermotropic transitions in bionienibranes. Atirrrr. R ~ I ,Eic>p/iys. . Biocrig. 5 , 705-138. Mufsch. B . , Gains. N . . and Hauser.. H. (1983). Order-disorder phase transition and lipid dynamics in rabbit small intestinal bruhh horder membranes. Effect of proteins. E i o c ~ / l c ~ t / l i s / r : \22, .

63366333.

Nagle. J. F. (1980). Theory of the main lipid bilayer phase transition. Anrirr. RoI'. !>/I?.\. C/IC/?I. 31, 157-195. Oldfield. E . . Gilniore. R., Glaser. M., Gutow\ky. H . S.. Hshung. J . C.. Kang. S. Y.. King. T. E.. and Meadows, M. (1978). Deuterium nuclear magnetic resonance investigation of the effects o f proteins and polypeptide\ on hydrocarbon chain order in model membrane systems. Proc.. N o / / . Actrd. .Sf,i. U..S.A.75, 46574660. Ovchinnikov. Yu. A,. Abdulaev. N. G.. Feigina. M. Yu.. Kiselev, A. V . . and Lobanov. N, A. ( 1979). The structural basis ofthe functioning o f bacteriorhodopsin: A n overview. FEES f,t,i/. 100. 219-224. Packard. B. S., Saxton, M . J . , Bissell, M. J . . and Klein. M. P. (1984). Plasma membrane reorganization induced by tumor promoters i n an epithelial cell line. Proc,. Ntr//.Acocl. sc.;. U.S.A.81, 449-452. Perutz. M. F . . Muirhead. H.. Cox. J. M . . and Goanian, L. C. G. (1968). Three-diniensionnl

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fourier synthesis of horse oxyhaemoglobin at 2.8 A resolution: The atomic model. Nutrrre (Loridon) 219, 131-139. Phillips, D. C. (1967). The hen egg-white lysozyme molecule. Proc. N u t / . Acud. Sci. U . S . A . 57, 484-495. Pjura, W. J.. Kleinfeld, A . M., Klausner, R. D., and Karnovsky, M. J. (1982). Fatty acid perturbation of a membrane protein-lipid interaction: A Tb fluorescence study. Bioplrvs. J . 37, 69-71. Ranck, J. L., Mateu, L . . Sadler, D. M., Tardieu, A., Gulik-Krzywicki, T.. and Luzzati, V . J. ( 1974). Orderdisorder conformational transitions of the hydrocarbon chains of lipids. Mol. Biol. 85, 249-277. Robson, R. J . , Radhakrishnan, R., Ross, A . H., Takagaki, Y., and Khorana. H . G . (1982). Photochemical cross-linking in studies of lipid-protein interactions. I n “ Lipid-Protein Interactions” (P. C. Jost and 0. H. Griffith, eds.), Vol. 2, p. 149. Wiley. New York. Ruocco. M. J., and Shipley, G. G . (1982). Characterization of the sub-transition of hydrated dipalmitoylphosphatidylcholine bilayers. Biocltem. Biophys. Ai,/ti 684, 59-66. Seelig, J . , and Seelig, A. (1980). Lipid conformation in model membranes and biological membranes. Q . R e v . Bioplrys. 13, 19-61. Seelig, J., Tamm, L.. Hymel, L., and Fleischer, S. (1981). Deuterium and phosphorus nuclear magnetic resonance and fluorescence depolarization studies of functional reconstituted sarcoplasmic reticulum membrane vesicles. Biot~hc,mi.vr.v20, 3922-3932. Seelig. J . , Seelig. A.. and Tamm, L. (1982). Nuclear magnetic resonance and lipid-protein interactions. I n “Lipid-Protein Interactions” (P. C. Jost and 0. H. Griffth, eds.). Vol. 2, p. 127-148. Wiley. New York. Senior. A. E. (1983). Secondary and tertiary structure of membrane proteins involved in proton translocation. Biochinr. Biopl7.v~.Acrir 726, 81-95. Shimshick, E. J.. and McConnell, H. M. (1973). Lateral phase separation in phospholipid membranes. Bioche~rii.sfry12, 235 1-2360. Simon. S. A., Mclntosh, T . J.. and Latorre, R. (1982). lnfluence of cholesterol on water penetration into bilayers. Science 216, 65-67. Singer. S . J., and Nicolson, G. L. (1972). The fluid mosaic model of the structure of cell membranes. Sciet?i.r 175, 720-731. Stoeckenius, W., and Bogomolni, R. A . (1982). Bacteriorhodopsin and related pigments of halobacteria. Annie. R e v . Bioihem. 52, 587-616. Stollery, J . G.. Boggs, J. M., Moscarello, M. A . , and Deber, C. M. (1980). Direct observation by carbon- 13 nuclear magnetic resonance of membrane-bound human myelin basic protein. Bioi,/irmi.strv 19, 2391-2396. Storch, J . . and Schacter, D. (1985). Calcium alters the acyl chain composition and lipid fluidity of rat hepatocyte plasma membranes in v i t m . Biochivi. Bioplrys. Acrrr 812, 473484. Stubbs, C. D., and Smith. A. D. (1984). The modification of mammalian membrane polyunsaturated fatty acid composition in relation to membrane fluidity and function. Biodiinr. B~OP/I.VS. A(./tr 779, 89-137. Takagaki. Y., Radhakrishnan. R., Gupta. C. M . , and Khorana, G. (1983). The membraneembedded segment of cytochrome / I , as studied by cross-linking with photoactivatable . 9128-9135. phospholipids. J . B i d . C I I P ~258, Taraschi. T.. and Mendelsohn. R. ( 1980). Lipid-protein interaction in the glycophorin-dipalmitoylphosphatidylcholinesystem: Raman spectroscopic investigation. Proi,. Natl. Actid. Sci. U . S . A . 77, 2362-2366. Terwilliger, T. C., Weissman. L., and Eisenberg. D. (1982). The structure of melittin in the form I crystals and its implication for nielittin’s lytic and surface activities. Bioipl~vs. J . 37, 353-361.

CURRENT VIEWS OF MEMBRANE STRUCTURE

27

Thonras, D. D., Bigelow, D. J.. Squier. T. C.. and Hildago. C. (1982). Rotational dynamics of protein and boundary lipid in sarcoplasmic reticulum membrane. Biopliys. J . 37,2 17225. Unwin. P. T.. and Zampighi. G . (1980). Structure of the junction between communicating cells. Nrrtrrr~(Lotidon) 283, S45-549. Vincent. M.. Foresta, B., Gallay. J.. and Alfsen. A. (1982). Nanosecond fluorescence decays of r1-(9-anthroyloxy) Fatty acids in dipalmitoylphosphatidylcholine vesicles with regard to isotropic solvents. Nioc./ierni.v/r:v 21, 708-7 16. Wallace. €3. A , . Cascio. M.. and Mielke, D. 1,. (19X6). Evaluation of prediction method\ for membrane protein secondary structui-es. Proc,. N o / / . A i d . Sci. U . S . A . . in press. Welch, G. R.. Somogyi. B.. and Damjanovich. S. (1982). l h e role of protein fluctuations in enzyme action: A review. f‘ro,q. Biopltys. M o l . B i d . 39, IW-146. Wiener. J . R., Pal. R..Harenholz. Y.. and Wagner. R . K. (1983).Influence o f t h e periphcriil matrix protein of vesicular stomatitis virus on the membrane dynamics of mixed phospholipid vesicles: Fluorescence studies. Bioc./ic~~tri.\/ry 22, 2 162-2 170. Wilson, 1. A,, Skehel, J. J.. and Wiley. D. C. (1981). Structure o f t h e haemagglutin membrane glycoprotein of influenza virus at 3A resolution. ~11tro.eondo don^ 289, 3 6 3 7 3 . Wolber. P. K.. and Hudson. B . S. (1982). Bilayrr acyl chain dynamics and lipid-protein interaction. The effect of the M 13 bacteriophage coat protein on the decay of the tluorescence anisotropy of paraniiric acid. Hioplrys. J . 37, 253-262. Wolf. D. E. (1983). The plasma membrane in early embryogenesis. I n “Development of Mammals’’ (M. H. Johnson. ed.), Vol. 5. Elsevier. New York. Wolf. D. E., and Voglmayr. J . K. (1984).Diffusion and regionaliziition in membi-anes of miituring rat spermatozoa. J . C ’ d l Biol. 98, lh7X-lhX4. Worcester, D.L..and Franks, N. P. (1976).Slructural analysis of hydrated egg lecithin and cholesterol bilayers. 11. Neutron diffraction. J . M d . Biol. 100, 359-378. Wu. S. H.. and McConnell. H. M. (1975). Phase separations in phospholipid membranes. Bioc/ioni,s/rv 14, 847-854.

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C'LIKKENT TOPICS IN MkIMBKANES A N I ) I KANSP(.)KI. V O L U M E 19

Ultrastructural Studies of the Molecular Assembly in Biomembranes: Diversity and Similarity SEK- WEN HUI

A great creation may seem defective. yet its u\efulness is unhthomahle. Lao Tse. sixth century I $ . ( .

Biological membranes have many forms and functions, but they share several common features: all membranes are sheetlike, semipermeable, and have approximately the same thickness. It seems logical to draw a general molecular model that not only represents these common features but also satisfies the physicochemical properties of membrane components. Historically a number of membrane models have been proposed. Two of these models have stood the test of time. The bilayer model originally proposed by Danielli and Davson ( 1935) predicts lipid molecules organizing in a bilayer sheet. A refinement ofthe bilayer model. i.e., the tluid mosaic model proposed by Singer and Nicolson ( 1972), depicts membrane proteins randomly scattered in a sea of fluid lipids, with some protein molecules protruding from one or both sides of the bilayer. These models are still regarded as fair representations of the general structure of biological membranes. Although various amendments have been proposed and added to these two models. their basic concepts remain unchanged. In this article I shall discuss an accurate way to characterize the general structure of biological membranes based on recent structural studies of membrane components. 1 shall first consider the structural similarity and diversity of biological membranes as revealed by electron microscopic and by electron, X-ray, and neutron diffraction studies. I shall then discuss 29 Copyright IYX7 hv Academic I'rc\\. Inc. All tights 01 icprodiiciion in a n y tiwin rcsrrvcd

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SEK-WEN HUI

some fundamental questions concerning the fidelity of the fluid mosaic model. Finally, I shall mention some areas in ultrastructural research that may lead to a better understanding of molecular organization in biomembranes. 1 shall limit myself, however, to consideration of structural studies only. The dynamic aspects of biomembranes will be discussed in a separate article by Edidin in this volume. An excellent review of biomembrane structure research has been given by Robertson (1981).

1.

SIMILARITY AND DIVERSITY OF BIOLOGICAL MEMBRANES

In cross-section views through the electron microscope, biological membranes look strikingly similar. This was noted by Robertson (1959) whose trilaminar picture is commonly seen in today’s literature. As examples, 1 have selected four different membranes to show the similarity and the diversity of the structure of biological membranes. These four membranes, namely, the myelin from neural tissues, the photoreceptor disc membrane from retina rods, the plasma membrane of erythrocytes, and the sarcoplasmic reticulum (SR) from muscle cells, have been studied extensively by various ultrastructural techniques. These membranes have different compositions, cellular locations, and functions, and yet their cross-sectional appearances are very similar. The cross section of myelin in a single layer branched from the multilayer in Fig. la has a symmetric trilaminar profile. The photoreceptor disc membrane shown in Fig. Ib also has a trilaminar appearance, although the separation between the two electron-dense lines is smaller, and the density profile is symmetrical. The erythrocyte plasma membrane shown in Fig. Ic has a wider separation between the two electron-dense lines, and the density profile is asymmetric. The SR membrane in Fig. Id is extremely asymmetric, as revealed by the tannic acid staining procedure which shows the glycocalyx. All four membranes have a common trilaminar structure, with variations in the thickness of each layer and in the relative density of the two darker layers. Based on image profiles one can draw some conclusions about the thickness and asymmetry of the membrane structure. However, in interpreting thin section electron micrographs one has to be cautious about artifacts. For instance, the thickness of the laminae can vary according to the methods of fixation and staining (Jones, 1974). The asymmetry can easily be destroyed by using a different fixation and staining procedure (Saito et al., 1978). Since each sample takes up stain differently, and since the sectioning and photographing steps vary from sample to sample, there is no direct relation between the photographic density and the molecular

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

31

FIG.I . Electron micrographs of cross sections of ( a ) myelin from feline corpus cullowm lHilderbrand and Muller. 1974). (b) photoreceptor disc membrane from frog retina (Jone5. 1974). ( c ) human erythrocyte membrane (Seeman. 1974). and ( d ) isolated sarcoplasmic reticulum vesicles from rabbit skeletal muscle (Saito t i d..1978). Bar. IOU nm.

organization of the membrane. It is not practical to compare the molecular organization of one membrane with another based on cross-sectional appearance. A better way to compare the cross-sectional structures of membranes is by small angle X-ray diffraction, since natural (unprocessed) membrane samples can be used in this method and absolute electron densities can be measured. In order to obtain a structural profile in terms of electron density across the membrane, individual membranes are usually stacked to form a one-dimensional paracrystal so that discrete diffraction intensities can be obtained for crystallographic analysis. In theory unstacked membranes (single unit or only a few repeating units) can also be studied by X-ray scattering. using continuous or semicontinuous scattering intensities (Blaurock and Wilkins, 1972; Hosemann and Bagchi, 1962). However,

32

SEK-WEN HUI

1

0

lOnm

Fici. 2. Electron density profiles derived from small angle X-ray diffraction analysis: ( A ) rabbit optic nerve myelin (Caspar and Kirschner. 1971). ( B ) frog retina disc membrane (Charbre, 1979, (C) human erythrocyte ghost membrane (Pape GIN/.. 1977). and (D) isolated 1981). Electron density sacoplasmic reticulum from rabbit skeletal muscle (Herbette ef d,, is in relative scale.

data collection in this type of experiment is more difficult to accomplish and less accurate. All four membrane examples shown in Fig. I have been studied by small angle X-ray diffraction. Myelin and photoreceptor disc membranes form natural one-dimensional paracrystals. Their structural profiles have been mapped to a considerable resolution. Erythrocyte membranes and SR have to be artificially stacked to form one-dimensional paracrystals. After centrifugation, vesicles of these membranes collapse to form somewhat regular arrays of repeating membrane units. Figure 2 shows the results of small angle diffraction studies of these membranes. The electron density profile of myelin (Caspar and Kirschner, 1971) in Fig. 2A is fairly symmetrical, with a separation of about 5 nm between the two high density peaks. The profile of disc membrane (Charbre, 1975) is very similar to that of myelin. The profile of erythrocyte membrane (Pape et d..1977; McCaughan and Krirnm, 1980) is quite different. The separation of the two high density regions is wider and its density profile is very asymmetric. The cytoplasmic leaflet has a higher electron density than the exoplasniic leaflet. The SR membrane (Herbette et id., 1981) has a thickness similar to that of the disc membrane. The inner half of the vesicular membrane has a higher electron density, but the electron dense band of the outer half is wider. The four membranes have a common characteristic: a low electron den-

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

33

sity region in the center of the membrane. This region is commonly interpreted to represent the hydrocarbon interior of the membrane. Even with a high protein to lipid ratio, such as in the S R membrane, the low density hydrocarbon region of the lipid is still dominant. The effect of the location of membrane proteins on the electron density profile is of secondary importance. The high electron density regions of the profile represent the head groups of the phospholipids rather than the membrane proteins. The 5-nm separations between the phosphate head groups in myelin and photoreceptor disc membranes agree quite well with the cross-section measurements obtained by electron microscopy, which show a separation of 4 nm between the two electron-dense lines. Since the electron-dense lines in cross-section electron micrographs are more representative of the location of unsaturated bonds in the hydrocarbon chains than of the head groups of the phospholipids, the interlaminar spacings in electron micrographs cannot be equated to the separations between the peaks in the electron density profiles measured by X-ray diffraction. The agreement between the wider separation of peaks in the erythrocyte membrane profile shown in Fig. 2C and the wider separation between the two electrondense lines in Fig. l c is not universal. although it is often the case. The asymmetric nature of the electron density profile of SR shown in Fig. 2D also reflects the feature shown in Fig. Id. The results of electron microscopy and X-ray diffraction show the common feature of biological membranes: In spite of their vastly different compositions and functions, the bilayer nature is predominant. This finding supports the essential correctness of the bilayer model proposed by Danielli and Davson SO years ago. However, the differences among membranes in term of bilayer thickness and asymmetry are appreciable, indicating the diversity of the structure of biological membranes. Freeze-fracture electron microscopy offers a new look at the ultrastructure of biological membranes. Freeze-fracture electron micrographs of the same four membrane examples of Figs. I and 2 are shown in Fig. 3. I n all these membranes, intramembranous particles (IMPS) are seen on the fracture planes. There is only one major intrinsic membrane prolein in myelin, photoreceptor disc membrane, and SR (respectively, lipophilin, rhodopsin, and calcium ATPase), and only one intrinsic membrane protein (band 3) is visible as an IMP in erythrocyte membranes (Marchesi et ( I / . , 1973). The densities of lMPs have been directly correlated with the quantities of glucose transporter or calcium ATPase in respective reconstituted membranes using these proteins (Sase et d.,1982; Wang rt d.,1979). We can regard the IMPS seen in Fig. 3 as representing the major integral proteins in these membranes. The densities of IMPS on the four membranes reflect the protein to lipid ratios in these membranes (Table 1). Myelin

34

SEK-WEN HUI

FIG.3 . Freeze-fracture electron micrographs of (a) myelin (Benitez r i d., 1977). (b) photoreceptor disc membrane (Andrews and Cohen, 1979), (c) human erythrocyte membrane (S.-W. Hui, unpublished), and (d) isolated sarcoplasrnic reticulum vesicles (Saito ei a / . . 1978). E and P fracture faces are indicated. Asterisk indicates convex face. Bar. 100 nm.

has the lowest protein to lipid ratio and also the lowest IMP density, whereas SR has the highest protein to lipid ratio and a high IMP density. However, the density of IMP is not always proportional to the protein/ lipid ratio in general, since not all intrinsic proteins are visible as IMPS and many proteins are visible only as dimers, trimers, tetramers, or higher polymers. In all four membranes there are more IMPS on the P face (adjacent to the cytoplasmic side) then on the E face. The uneven distribution of the IMPS between the P and E faces indicates that the proteins locate asym-

35

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

Membrane Myelin Photoreceptor disc Plasma membrane Sarcaplasmic reticulum

l'rotein/lipid ratio (Wt/Wt)

Reference

Human brain Bovine retina

0.28 1.1

O'Brian and Sampson ( 1965) Borggreven ct NI. ( 1970)

Human erythrocyte Human pectoral muscle

I .3 2. I

Steck (1974) Takagi (1971)

Source

metrically across the membrane. Such uneven distribution has also been seen by negative stained electron microscopy of SR (Saito at id., 1978). However, the asymmetric protein distribution is not apparent in electron density profiles of myelin and photoreceptor disc membrane determined by X-ray diffraction analysis, although it is reflected to some degree in electron density profiles of erythrocyte membrane and SR. In all these four examples, we see no regularity in the planar distribution of IMPS on either the E or P face. The irregular distribution of intrinsic proteins in the bilayer fits the description of the fluid mosaic model. Moreover, in many cases, we can induce a change in the distribution of IMPS by manipulating the composition, temperature, or pH and by crosslinking the membrane and cytoskeletal components. The lateral mobility of proteins further supports the fluid mosaic model. In recent years. however, increasingly sophisticated experiments have led us to picture the membrane as a more complex molecular assembly than that depicted by the fluid mosaic model. 1 shall discuss several aspects of the fluid mosaic model that have been reexamined.

II. IS THE LIPID BILAYER A PASSIVE, HOMOGENEOUS "SOLVENT"? According to the fluid mosaic model, the lipid in the membrane is a homogeneous body. Being a homogeneous medium, the only influence the bilayer has on membrane activity is through its bulk physical properties. However, lipid effects on membrane activity cannot always be explained by bulk physical properties; therefore, many suggestions have bcen made that heterogeneous structures exist in the membrane lipids. Whether these heterogeneous structures differ by composition, by molecular organization such as the packing of the acyl chains, or by their proximity

36

SEK-WEN HUI

to a protein molecule is not certain, nor is the size of these heterogeneous structures or domains known. In the discussion of lipid domains and other heterogeneous lipid structures of membranes, I shall discuss only the transient, spontaneously formed lipid microstructures not confined to any particular sites of the cell. Specialized membrane zones permanently located in given sites of polarized cells, such as sperm heads (Friend and Bearer, 1981; Bearer and Friend, 1980), tight junctions in epithelial cells (Robenek et al., 1982; Kachar and Reese, 1982), bile canaliculi on the surfaces of hepatocytes (Robenek et al., 1982). and neuromuscular junctions (Bridgman and Nakajima, 1981; Perrelet et al., 1982; Pumplin and Bloch, 19831, are known to contain lipids and proteins that are different from those in other parts of the membrane. The formation and maintenance of these specialized membrane zones are important topics in cell biology but are beyond the realm of general lipid heterogeneity. The formation of smaller, less permanent membrane zones, such as coated pits and secretion sites (Elias et al., 1978; Montesano et ul., 1979; Plattner, 1981; Nemeth and Rohlich, 1981), depend more on the dynamic association and dissociation of membrane components. These smaller membrane zones could be related to the proposed, even smaller lipid heterogeneities that have no specific macroscopic functions-the lipid domains. In order to understand more about the physical chemistry that leads to the formation of lipid domains, one would naturally turn to the study of a very simple system, namely the model lipid bilayer. There have been many studies of lipid domains in model bilayers. In a single component system, a domain in a gel or solid phase bilayer can be defined as an area of continuous crystallinity. As the bilayer undergoes a phase transition, the melting of solid domains most likely starts at the domain boundary. For an infinite crystal of identical molecules, the phase transition is first order, namely, the transition temperature range is small. Within the transition temperature range, both solid and fluid domains exist. When the transition is completed, all solid domains melt and the bilayer becomes a homogeneous fluid. If the single lipid component is contaminated, or if the system contains more than one component, the transition temperature range is broadened. Within the transition temperature range both solid and fluid domains coexist according to physical chemical principles (Marsh et al., 1977; Lee, 1977). If the multicomponent system is completely miscible in both solid and fluid phases, both fluid and solid domains contain all components in continuously varying proportions during the phase transition. If the system consists of immiscible components, below the onset of phase transition there will be different solid domains consisting of mainly one or other

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

37

components (or the eutectic composition). Above the phase transition, there are heterogeneous fluid areas, each containing clusters of different composition if the fluid components are also immiscible. In the transition temperature region, both solid and fluid domains have distinct compositions according to the phase diagram. Therefore domains can be defined either by their composition o r by the molecular packing (phase). The definitions of miscibility and phase separation are sometimes used in the literature indiscriminately. Domains in the solid phase are similar to “grains” in a polycrystalline metal. Between domains of a given orientation there must be boundaries of mismatching areas. In a metallic sample of alloy, such domains and domain boundaries are visible because its crystalline domains reflect light differently due t o different molecular alignments. In a lipid bilayer, the visualization of such domains is not so simple, because the domains are much smaller and less stable and optical techniques cannot be easily applied. To visualize the molecular packing domains, the most logical choice of optical tool would be X-ray optics. The molecular spacing between acyl chains of phospholipids are typically 0.4 to 0.5 nm, which is slightly larger than the X-ray wavelengths used in diflraction experiments. Unfortunately, the technology of X-ray imaging has not developed to its wavelength limit. Therefore such imaging by X-ray techniques is not yet possible. The lipid packing domains should also be visible by electron optics, which can easily image objects to a resolution of 0.3 to 0.5 nm. However, imaging by conventional electron microscopy requires the sample to be processed so that it can resist the damaging effects of electron radiation and the vacuum environment. Lipid bilayers do not lend themselves to such treatment. However, by putting a lipid bilayer in an environmental chamber to protect it from the vacuum and by using a low dose technique, 1 was able to apply the diffraction contrast method to visualize lipid domains by electron optics. The contrast between domains was created by selecting diffracted electron beams from domains of different molecular orientations and reconstructing the image in the electron microscope. Some examples are shown in Fig. 4. The area percentages of solid and fluid domains measured in the transition temperature regions of phospholipid bilayers matched well with that predicted by calorimetry and other physical measurements (Hui, 1981). This technique is applicable only when the domains are large enough so that their diffracted beams are easily separated by in sitcc optical filtering in the electron microscope. In some lipid systems, especially those of saturated phosphatidylcholine (PC) and phosphatidylglycerol (PG), t h e mismatching of the head groups and the acyl chain cross sections favors a special type of chain tilting that

FIG.4. Freeze-fracture and diffraction contrast electron micrographs of mixed dipalmitoylphosphatidylcholine and bovine brain phosphatidylseiine (molar ratio 2: I ) at 34°C. Arrow heads indicate polygonal ring domains. Bar = I pm.

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

39

results in the formation of ripples at a temperature range just below the solid-fluid phase transition (Sackmann, 1978).The periods of the ripples, typically 12-15 nm, are very large in comparison to molecular dimensions. These ripples can be distinguished from smoother domains of different molecular packings by freeze-fracture electron microscopy. Visualizations of solid domains by this method have been reported by many research groups (van Dijck et ul., 1978; Luna and McConnell, 1978, Stewart et nl., 1979).The ripple phase domains measured by this method are comparable to those measured by diffraction contrast (Fig. 4). Interestingly. whenever the ripple phase is involved, the shape of the ripple phase domain is dictated by the orientation of the ripples, which usually have straight edges and tend to form polygons (Figs. 4c and 4d) to eliminate the singularities at the end of the rippled ridges (Sackmann, 1978; Schneider ef a / . , 1983). The boundaries of rippled domains can sometimes be seen as unzipping line by line during melting, corresponding to the one-dimensional melting theory proposed by Jain (1983). I t has been reported that some tluorescent dyes partition differently into the solid and tluid lipid domains. These dyes include trans- and cisparanaric acid (Sklar et d..1977: Kleinfeld, this volume), diI dye (dioctadecylindocarbocyanine iodide) (Wolff ef c d . , 1977). and merocyanine dye (Williamson et al,, 1981). These dyes can be used to label domains of different molecular packings for fluorescence microscopy. Some lipid zones have been visualized by this method (Williamson r t al., 19811, but application to visualization of microdomains has been limited. Weis and McConnell ( 1985) were able to visualize lipid domains using fluorescent lipid probes. In the study of the characteristics of membrane lipids, steroids and other neutral lipids must be taken into account. Cholesterol constitutes up to 50% of the total lipid in plasma membranes of animal cells. The effects of cholesterol on the formation of lipid domains are best illustrated by studying mixtures of cholesterol and one or two phospholipids of defined composition. There are many physical studies of mixed cholesterol and phospholipid bilayers, using calorimetry, N M R , ESR, fluorescence, and X-ray and electron diffraction. Although the miscibility of cholesterol and phospholipids is still an open question, most of the studies indicate that below 20 mot 96 of cholesterol there are not enough cholesterol molecules to interact with every phospholipid molecule; therefore there are "domains" of pure phospholipids. It is a challenge to electron microscopists to visualize the phospholipid domains. By the freeze-fracture method, long ridges have been visualized in mixtures of PC and cholesterol at low cholesterol content. The relative widths of ridges and valleys change as the cholesterol to phospholipid ratio increases. The long stripe pattern indicates that the cholesterol-rich and cholesterol-poor domains interlayer in a stripc fashion (Copeland and

40

SEK-WEN HUI

McConnell, 1980; Lentz et al., 1980). The stripe domains also correspond to the unidirectional lateral diffusion measurements by the fluorescence recovery after photobleaching (FRAP) method. At mole percentages of cholesterol greater than 20%, domains are no longer visible by freezefracture although one-dimensional molecular alignment can still be detected by electron diffraction (Hui and He, 1983). At higher percentages, cholesterol molecules tend to segregate eventually into ribbonlike domains, as visualized by the diffraction contrast method (Hui and Parsons, 1975). In 1976, Kleeman and McConnell reported a significant observation. When they incorporated proteins from human erythrocyte membranes into a lipid mixture which contained both solid and fluid domains, they found that the proteins preferentially partitioned in the fluid lipid domains. The proteins can be observed as intramembranous particles (IMPS)by freezefracture electron microscopy. The unequal partition of proteins in the solid and fluid lipid domains enables electron microscopists to observe the domain size and shape by the lateral distribution of IMPs. Since most biological membranes contain protein, this method can be widely adopted to observe lipid phase separation, especially when the domains cannot be recognized by the diffraction contrast method because the diffracted beams are too weak. Unfortunately, differential partitioning of proteins in fluid and solid domains is not the only cause of nonrandom distribution of IMPs. Other factors that cause IMPs to redistribute will be discussed in Section IV. Therefore, unless experimental conditions are specified, one cannot make a direct association between the distribution of IMPs and the lipid phase separation phenomenon. Using IMP distribution as a criterion, thermotropically induced lipid domains have been visualized in several bacteria (Verkleij and van Golde, 1975; Halvarson et ul., 1978). Escherichiu coli is a particularly popular specimen for observing membrane domains because the membrane lipids in some metabolically defective strains can be manipulated through feeding. Observations of the IMP distribution clearly show that domains exist in the membrane of these E. coli strains, and the occurrence of lipid phase separation is related to the function of the bacteria (Bayer e t ul., 1977; Legendre er a / . , 1980; Arancia et al.. 1980a). Membrane domains are also observable in Achokeplusmu luidluwii, (Verkleij et a/., 1972), in the nuclear 1979), membrane of Trtruhymenu (Wunderlich et ul., 1974; Giese et d,, and in Salrnonellu (Cottam et ul., 1986). Figure 5 shows the appearance of phase separated domains as visualized by the IMP-denuded areas in Salmonella membranes as the temperature was lowered below the threshold of lipid phase separation. When the temperature was raised over the threshold, the IMP distribution was once again random. The redistribution of IMPs and the lipid phase change are reversible, and the microorganism

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

41

Fic;. 5 , Freex-fracture electron micrographa of ;in unsaturated fatty acid auxotroph of S d w w / / ( i / , v p / i i o i / w i / mcultured with oleic acid hiipplernent and freeze-quenched a t ( a ) 20°C and (b)37°C‘. The fracture plane shown I S the P face o f the inner membrane. Bar. 100 nm. ( I n collaboration with C. Hi).)

can survive such changes. Membrane domains as defined by the distribution of IMPS were also observed in several types of algae (0-iakianand Satir, 1974; Armoud and Staehelin. 1979) and fungi (Puccinelli ct d . , 1977: R . W . Miller, 1980). The vital question to ask is. “Do lipid domains exist in membranes of‘ higher animal cells’?” Much of the earlier evidence showing gel phase lipids in membranes of higher animal cells were shown to be artifactual. Most freeze-fracture electron microscopic observations of the plasma membranes of higher animal cells show that IMPS are “randomly” distributed. Perhaps the high cholesterol content in most plasma membranes of higher animals prevents a large-scale lipid phase separation. In 1980. Hui ct trl. partially removed cholesterol from human erythrocyte membranes. B y lowering the temperature. they were able to observe IMP-

42

SEK-WEN HUI

denuded areas arising from the phase separation of membrane lipids. This observation agreed with their electron diffraction results. Gerritsen et ul. (1979) also observed the redistribution of IMPs in human erythrocyte membranes as a function of temperature when most of the cytoskeletal proteins were removed from the membrane. These observations support the proposition that the cytoskeletal network and the cholesterol content in the plasma membrane of higher animals cells prevent large-scale phase separation. Under special conditions, IMPs in erythrocyte membranes can be induced to form patches which might also be related to lipid phase separation. Such patching of IMPs has been observed with changes of pH and osmotic pressure (Elgsaeter et al., 19761, with the addition of calcium in the presence of phosphates (Vos et ul., 1976; Vosky and Loyter, 1977), in the presence of Sendai virus (Knutton, 1977; Asano and Sekiguchi, 1978; Zakai er ul., 1977; Pinto da Silva et ul., 1980), as well as in the presence of polyethylene glycol (PEG) (Knutton, 1979; Krahling, 1981; Hui et a l . , 1985). The redistribution of IMPs is not restricted to erythrocyte membranes. For instance, myelin can also be induced to form domains by the addition of calcium (Hollingshead et al., 1976; Melchior et u l . , 1979). Temperature-dependent IMP redistribution has also been observed in crab axon membrane (Gulik-Krzywicki et al., 1981) and in astrocytes (Anders and Brightman, 19821, as well as in L cells in the presence of PEG (Roos et al., 1983). Thermotropic IMP patching in mitochondria1 membranes has been observed (Hackenbrock et al., 1976; Hochli and Hackenbrock, 19761, and the degree of patching can be modulated by cholesterol (Schneider et al., 1982). if the IMP patching and the formation of IMPdenuded areas are indeed caused by lipid phase separation, then largescale phase separation domains do exist in membranes of higher animal cells under special circumstances. Although cholesterol can be responsible for the reduction or even elimination of phase separated domains in the membranes of higher animal cells, it would be interesting to find out whether cholesterol itself forms domains in membranes. The polyene antibiotic filipin forms specific complexes with cholesterol and disrupts the planar bilayer arrangements in membranes containing high local concentrations of cholesterol (see review article by Severs and Robenek, 1983). This specific association can be utilized to visualize local concentrations of cholesterol in membranes. However, whether filipin is an indicator of cholesterol already concentrated in local areas or whether filipin cholesterol concentrates to form complexes in local areas remains unclear. In experiments using filipin as a cholesterol label, careful membrane fixation must be carried out to prevent or at least to reduce the redistribution of cholesterol by the action

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

43

of the drug. The reliability of this technique to determine the asymmetric distribution of cholesterol was discussed by Miller ( 1984). Cholesterol-enriched areas have been observed by the filipin technique in A. /ciid/mii (de Kruijff and Dewel. 19741, in the nuclear membrane of Tetrrrhymencr (Sekiya el id., 19791, in flagellate membrane (Melkonian ct d . , 1979), in human erythrocyte membrane in the presence of Sendai virus (Brown et d . , 19821, in the outer nuclear membrane of a number of cells (Feltkemp and van der Waerden. 1982; Robenek and Greven, 1981; Kim and Okada, 19831, and in T lymphocytes a s a function of external fatty 1981). Assuming that the freeze-fracture acid incorporation (Hoover ef d., observation of cholesterol-filipin complexes truly represent local areas of high cholesterol concentration, then cholesterol domains exist in many membranes of higher animal cells. Some recent fluorescence experiments (Klausner pt d.,1980) have also showed the existence of lipid microdomains in higher animal cell membranes. The spectroscopic evidence (see Kleinfeld. this volume) is beyond the scope of this article on microscopy. The available evidence so far indicates that spontaneous domains do exist in membrane lipids, but whether the lipids in higher animal cell membranes are structurally homogeneous or heterogeneous depends on the environment. and may vary from case to case. 111.

IS THE BILAYER THE ONLY FORM OF MEMBRANE LIPIDS?

Although lipids are known to exist in many different forms in the presence of water (Luzzati, 1968), the bilayer is by far the most common form. In fact, it is the only form observed in biological membranes. which are supposed to form an effective barrier to protect the cell and to partition its interior compartments. There is little doubt that membrane lipids exist only in the bilayer form. Based on geometrical considerations alone, the bilayer is not always thc best molecular arrangement for membrane lipids. The head groups of phospholipids have different sizes and hydration capacities. The hydration capacity of the head groups determines how close they are linked together. This feature is well illustrated in crystallographic studies (Hauser ('1 r r l . , 1981). The space occupied by the acyl chains also varies with the number of trans-gauche transitions, which in turn depends on the degree of unsaturation as well as on thermal excitation. Therefore, a close match between the cross section of the head group and that of the acyl chains, as demanded by the bilayer configuration (Figs. 6a and 6b), is a coincidence rather than a rule.

44

SEK-WEN HUI

....... ,:.::":.

.. . . . . . . . .

.. .. .. .. . e . . . .. .. .. m .

m kbbb. w.,.a

,

~

C

a'

..:::............: ..

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

............. . .:;. .... .

. ,. . ..

...

. . . .

. . , . i;

., . .. .,: .. .

.. ...

~~

"w

...........

d

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

....... . . . . . . ::.. . .

. . . .., ... ' .. . .., .. .. .. .I

. .. .. .

FIG.6 . Schematic drawings of various types of molecular packing in phospholipidlwater systems. The phospholipid molecules are drawn (a) with matching cross sections for head groups and acyl chains: (b) with smaller head groups but large hydration shells to separate the head groups; (c, d) with large head groups that require chain tilting or head group stacking to compensate for cross-section mismatching; (e, fl with unequal acyl chain lengths leading to chain interdigitation; (g) with small head groups and insufficient hydration, resulting in the formation of bilayer disruptions with negative curvature only; (h) with compensiiting shapes to form a curved bilayer; or (i, j) with extreme curvatures to form micelles of opposite senses in a nonpolar or polar environment.

However, nature has many ways of accommodating the mismatching

in the bilayer form. For instance, excess area occupied by the head groups may be accommodated by chain tilting (Fig. 6c). Alternatively, stacking

of the head groups may also reduce their total cross-section area. Such stacking results in surface ripples o r undulations as a modified form of chain tilting (Fig. 6d). If the mismatching becomes too great to be accommodated by chain tilting, chain interdigitation (Figs. 6e and 6 0 may occur (Mclntosh et al., 1984; Hui et al., 1984b). For wedge-shaped molecules a highly curved structure (Figs. 6g and 6h) is sometimes preferred. Structures with positive and negative curvatures are formed depending on the relative areas occupied by the head groups and the acyl chains, and the

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

45

degree of hydration of the head groups. For instance, PE with a low hydration and a small head group cross section, coupled with highly unsaturated acyl chains, would favor a structure with negative curvature at high temperature. A structure having negative curvature only, such as that shown in Fig. 6g, would satisfy the requirement. The structure shown in Fig. 6h requires two types of molecules of opposite senses of area compensation to form both positive and negative curvatures. These types of structures form the basis of the inverted cubic and the inverted hexagonal tubes (Fig. 6i) (Boni and Hui 1983; Hui ct cil., 1983b). PC with a bulky and strongly hydrated head group, when coupled with a single short acyl chain, such as in lyso egg PC, would favor positively curved structures such as micelles (Fig. 6j) or hexagonal 1 tubes. The structural types summarized in Figs. 6g-6j are well known but are not thought to be features in biological membranes. Rather they represent disruptions of bilayers. For instance, the micellar form is the basis for the detergentlike reaction of lysolipids in biomembranes. The disruptive properties of lysolipids are related to the disruption of bilayers during the membrane fusion process (Lucy, 1975). This relation was proposed again later by Cullis and Hope (1978). At that time, bilayer disruption points (lipidic particles) were observed by freeze-fracture electron microscopy in lipid mixtures containing cardiolipin in the presence of calcium (Vail and Stollery, 1979; Verkleij e t al., 1979b). These disruption points of the bilayer were formed to satisfy the requirement for highly negatively curved structures in order to accommodate the mismatching between the cross sections of the acyl chains and the head groups of the cardiolipin-calcium complex. Similar disruption points are also observed in bilayers containing unsaturated PE (Hui r t d . , 1981). These disruption points are seen as connections between bilayers in cross fractures. as shown in Fig. 7. The morphology of these “lipid particles” is distinct from IMPS of protein nature on the fracture plane (Fig. 7). Many such types of nonbilayer structures have been reported, and several molecular arrangement models have been proposed (Verkleij et (11.. 1979a; Hui pt (11.. 1981; R. G . Miller, 1980. Hui r t d.,1983b; Verkleij, 1984; Rand et d . , 1981). Naturally, such bilayer disruptions are thought to be either intermediate stages or by-products of bilayer fusion (Hui et ul.. 1981; Verkleij rt ml., 1982). The molecular mechanism of fusion is discussed by Blumenthal in this volume. Based on the nonbilayer structures detected by the change of ”P N M R line shapes given by the motional averaging of molecules under structural constraints (Thayer and Kohler, 1981; Seelig, 1978). some investigators have suggested that nonbilayer structures also exist in biological membranes (van Venetie and Verkleij, 1982; Stier er id., 1978). Hexagonal I1 structures in biomembranes have indeed been observed with electron mi-

46

SEK-WEN HUI

FIG.7. Freeze-fracture electron micrognph of part of a multilamellar vesicle reconstituted with human erythrocyte membrane proteins (Triton X-100extract). soybean phosphatidylethanolamine, and egg phosphatidylcholine. Large lipidic particles are visible in cross fracture as interlamellar connections (bilayer defects). Small particles of protein origin are seen embedded in the bilayer at all levels. Bar, 100 nm.

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

47

croscopy, but only rarely and under low hydration, high temperature, manganese ion, or other unusual environments (Crowe and Crowe, 1982; Corless and Costello, 1981; Chabot and Leopold. 1982; Gruner et a / . , 1982; Gounaris et uf., 1983a; van Venetie and Verkleij, 1982). Nonbilayer structures have also been observed with electron microscopy in total lipids extracted from mitochondria (Cullis el d . , 1980). photoreceptor discs (Albert rt d., 1984; de Grip rt NI.. 19791, chloroplasts (Gounaris et id., 1983b). and E. cofi plasma membrane (Burnell rt al., 19801, although nonbilayer structures have not been seen in the native forms of these membranes. There have been several reports that the incorporation of proteins into lipids inhibited the formation of nonbilayer structures (Taraschi t>t d . , 1982; Goormaghtigh et al., 19821, although a few proteins, such as gramicidin A , have been reported to promote the formation of nonbilayer structures when incorporated into a mixture of lipids containing acidic lipids (van Echteld et d . , 1982). Our recent study of the plasma membrane of Mycopplasma capric.ulrrm showed that nonbilayer structures could be observed in native membranes only if part of the membrane protein had been digested by trypsin. Apparently the absence of nonbilayer structures in biological membranes may be due partly to the presence of membrane proteins. The nonbilayer structures in M. cupriculirm membranes digested with trypsin. and the absence of these structures in Ca"-free membranes. are shown in Fig. 8. From what is known so far about nonbilayer structures, it seems that without membrane proteins, many membrane lipids would have formed nonbilayer structures under physiological conditions. Yet very few functional biological membranes exist in nonbilayer forms. Instead of asking the question: "Do nonbilayer structures exist in biological membranes?" it would be more appropriate to ask the question: "How important are the nonbilayer-forming lipids in biological membranes'?'' It seems more than coincidental that many energy transducing membranes contain a large amount of nonbilayer-forming lipids. For instance, photoreceptor disc membrane contains a large percentage of highly unsaturated PE which by itself would form hexagonal I1 structures in the presence of calcium (Albert e f 01.. 1984). The high percentage of cardiolipin in the inner membrane of mitochondria in animal cells also favors the formation of nonbilayer structures (van Venetie et 01.. 1982). The lipids of thylakoid membranes contain a high percentage of monogalactosyldiglyceride which also forms nonbilayer structures (Sen et ul., 1982). Table I1 gives the composition of nonbilayer-forming lipids in a variety of membranes. On the other hand, the plasma membrane of animal cells contains a large amount of satiirated PC which favors bilayer formation. In passive membranes such as myelin, saturated PC and sphingomyelin dominate; even with a very low protein

P 0)

FIG.8. Frczze-fracture electron micrographs of isolated membranes from Mycoplnsmn cupriculum cultured with (a) normal medium; (b) with 0.5 mM of calcium. resulting in the formation of hexagonal I1 structure: and (c) with 0.5 mM calcium and 20 pghl of egg phosphatidylcholine. resulting in many bilayer defects in the form of lipidic particles. Bar, 100 nm. (In collaboration with R. Bittman.)

T A B L E I1 L I P I DCOMPOSITION o b BIOLOGIC AI. MEMHKANLS

Membrane Myelin Plasma membrane Plasma membrane Sarcoplawnic reticulum Photoreceptor disc Mitochondria1 outer membrane Mitochondria1 inner membrane

Percentage of pho\pholipid“

Source

Chole\lerol/ phospholipid (mol/mol)

PC

SP

PS

Human hrain

0.80

16.6

32.0

10.6

Human erythrocyte Rat hepatocyte Human pectoral muscle Bovine retina Rat hepatocyte Rar hepatocyte

0.79

28.9

26.9

13.0

PI

PE

-

22.4

1.3

27.2

Un\aturation index”

DPG

Totdl

PE

DPG

Reference

72

113

-

O’Brinn and Simpwn (1965) d. Rower

I27

0.63

30.0

22.1

15.3

6.0

19.3

I43

0.42

65.1

1.7

2.7

8.7

16.9

I36

0.15

40.2

3.6

13.3

2.3

38.6

228

0.06

59.0

4.8

20.3

0.02

39.3

7.5 3.4

6.6

34.4

(25.3) I69 (46.0) 219 (42.3) 21x (36.8)

-

(11

(196X)

-

274 (105.8)

-

129

133

67.5 (4.3)

1x1

(27.0) I 87 (64.3)

13.5 (18.5)

van Hoeven and Emmelot (1973) Takagi (1971)

Anderwn and Maude (1970) Huet (’1 t r l . (1968) Huet c v t i / . (1968)

PC, Phosphatidylcholine: SP. sphingomyelin; PS, phosphatidylserine; PI. phosphatidylinosiiol: PE. phosphatidylethanolamine: DPG. cardiolipin. ” The unsaturation index is the sum of the product of the mole percentage of fatty acid and the numher of double bonds (figures in parentheses normalized to respective mole percentages of phospholipids). I’

50

SEK-WEN HUI

content, the formation of nonbilayer structures in myelin is very unlikely. In general, the metabolically active membranes contain a larger percentage of nonbilayer-forming lipids. Such correlation indicates a special role for the nonbilayer-forming lipids in these membranes. What are the functions of the nonbilayer-forming lipids in metabolically active membranes? Since these lipids by themselves do not form stable bilayers, functional proteins must play a role in stabilizing the bilayer structure. The proteins may make up the geometric mismatching of lipid molecules in order to maintain the bilayer form of the membrane. On the other hand, the structural fluctuation associated with the instability induced by the nonbilayer lipids may be necessary for these proteins to carry out their high metabolic activities. This hypothesis is supported by the reconstitution experiments of Navarro et ul., (1984), Bosterling and Trudell (1982), and us (Cheng rt al., 1986). In our experiment, when the calcium ATPase from sarcoplasmic reticulum was incorporated into liposomes of various mixtures of lipids, the energy coupling efficiency (calcium ions transported per ATP hydrolyzed) was found to be elevated when the lipid mixtures contained a large amount of nonbilayer-forming unsaturated PE. Whether these lipids allow higher protein flexibility in the vacant volume created by geometric mismatching or allow faster protein movement (translational or rotational) through structural defects in the unstable bilayer remains to be determined.

IV.

DO MEMBRANE PROTEINS DISTRIBUTE RANDOMLY IN THE BILAYER?

The fluid mosaic model depicts the intrinsic membrane proteins distributing randomly in the plane of the bilayer. There is no provision for any interaction or organization of intrinsic proteins. This model was partly based on the electron microscopic observation of labeled lectin receptors on plasma membranes. The distribution of these labels was apparently random (Singer and Nicolson, 1972). There were a number of studies of the planar distribution of membrane proteins by labeling techniques. For instance, the glycoprotein in human erythrocyte membrane was labeled by ferritin, which was visualized by freeze-etching electron microscopy. The distribution pattern of the ferritin-labeled glycoprotein and the distribution of IMPS in the fracture plane of the membrane matched (Marchesi et ul., 1973). These studies showed that although membrane proteins formed patches under certain conditions, no regularity could be ascribed to their distribution. Because many protein distribution studies were based on observations

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

51

of surface labels or IMPs, identification of these particles is very important. In membranes consisting of only one major intrinsic protein, the identification of IMPS is simple but by no means trivial. For instance, rhodopsin accounts for 85% of the total protein of the photoreceptor disc membrane. The IMPs in the disc membrane must be associated with rhodopsin molecules, although we do not know whether each particle represents one, two, or several molecules. Myelin contains two major proteins. One is an intrinsic protein which is appropriately named lipophilin. and the other is a more hydrophilic basic protein. The IMPs in rnyelin are most likely lipophilin. I n SR, the IMPs can be attributed with certainty to the major protein (calcium ATPase). The asymmetric distribution of IMPS in the convex and concave fracture faces, as shown in Fig. 3d, has been used to support the asymmetric location of this protein in the bilayer (see Section 11). In erythrocyte plasma membrane, the major intrinsic protein of considerable molecular weight (95.000) is the band 3 protein. However, glycophorin (Hui et d., 1985)and glucose transporter (Sase ct u / . , 1982) also form IMP in reconstituted membranes. Sase et ( I / . 11982) determined the molecular weight of glucose transporter by counting the density of IMPS in reconstituted membranes of known proteinhpid ratio. In membranes containing a number of ma-jor proteins. the identification of surface components or IMPs is not simple. In surface-labeling processes, one can use electron-dense markers conjugated to specific ligands such as lectins or antibodies to selectively reveal t h e location of particular receptors or antigens. Small, electron-dense molecules such as ferritin have been used for this purpose. More recently, colloidal gold has been widely used as an electron-dense marker. Colloidal gold can be conjugated to a variety of molecules, including lectins. antibodies, protein A , biotin, and other small proteins. The conjugated gold microspheres are then used for direct or indirect labeling. One of the advantages of colloidal gold is that it can be made into different sizes for double labeling. It is very visible in both transmission and scanning electron microscopy. It should be noted that in determining the distribution of proteins, the labeling efficiency must be high enough so that most of the proteins are labeled in order that the distribution of labels in the visual field be representative of the true protein distribution. The identification of IMPs is a very tricky technical problem. Since IMPs are visible only when the membrane is frozen and fractured, labeling frozen samples poses a formidable obstacle. The problem has been solved by two different approaches. Pinto da Silva and Torrisi (1982) thawed frozen, fractured specimens, labeled them with either lectin-conjugated or immunoconjugated gold microspheres, and then critical-point-dried the specimens and replicated them in the dry or frozen state. This approach

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permitted identification of the particles on the P phase on the plasma membrane as WGA receptors (Fig. 9). The other approach is to form replicas of freeze-fractured samples in the frozen state, using the conventional freeze-fracture method. The samples are then thawed, labeled, embedded, and sectioned, From the structural features of the serial sections, Rash et af. (1982) were able to recognize the IMPS and identify them by immunolabeling using colloidal gold microspheres. Once the nature of the IMP is known, one may equate the distribution of IMPS to the distribution of certain proteins in more complex membranes. To truly assess the randomness of protein distribution, mathematical criteria must be applied. Anionic membrane proteins can be labeled with cationized ferritin, colloidal iron, or colloidal gold. Figure 10 shows a thin section of a Chinese hamster ovary (CHO) cell labeled with colloidal iron which attaches to the sialic acid moieties of the membrane glycoproteins. This type of one-dimensional distribution of labels was statistically analyzed (Weiss et al., 1972). The analysis showed that the one-dimensional distribution was not strictly random according to Poisson statistics. A number of factors were found in other studies to alter the random distribution. Among these factors are electrostatic replusions by neighboring cells, uneven distribution of electric charges on membrane surfaces of different curvatures, fixation artifacts (Arancia et al., 1980b), glycerination artifacts (Gilula et al., 1975; Pinto da Silva and Martinez-Palomo, 1975), and cytoskeletal restrictions (see below). In several two-dimensional studies of IMPS or labeled surface components, assessments of randomness of the distribution of protein components have been made. One such analysis was reported by Abbas et af. (19751, who studied the distribution of immunoglobulin on the surface of lymphoblasts. They found that the distribution was not exactly random by measuring the density of labels in each subdivision of the visual field and statistically comparing the distribution with that predicted by random models. A truly random distribution does not mean equal numbers of particles in each square subdivision, but numbers corresponding to a statistical model such as the Poisson or the binomial distribution, depending on the average number of particles in each square. Weinstein (1976) studied the distribution of IMPS in the plasma membrane of urinary bladder carcinoma cells. He also compared the density distribution of IMPS to that derived from the Poisson distribution and quantitated the nonrandomness by the coefficient of dispersion. Markovics et al. (1974) applied a different strategy, namely they measured the interlabel distance and applied a radial distribution analysis to study the distribution of nuclear pores. Pearson et al. (1979) studied the distribution

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Fiti. 9. (a. b) Partition of wheat germ agglutinin hinding sites between P and E fr~icture faces of erythrocyte membrane, respectively. as labeled with ovamucoid-coated colloidul gold. (c) Thin section micrograph shows the partition of label5 between complementary membrane halves (Pinto da Silva and Torrisi, 1982). KBC. Red blood cell: BSA, bovine serum albumin. Bar, 100 nm.

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FIG. 10. Thin section electron micrograph of a Chinese hamster ovary (CHO) cell. The cell surface sialic acid was labeled with collidal iron particles. Bar. I pm.

of IMPs in human erythrocyte membranes under different conditions, using both radial distribution and density distribution analyses. They compared the results to those derived from statistical models, taking into account the exclusion areas occupied by the IMPs. By using parameters derived from both radial and density distribution analyses, they devised a pattern recognition routine to classify the distributions of various samples. Figure 11 illustrates the procedures to obtain the radial, angular, and density distributions of IMPs in human erythrocyte membranes. The radial distribution method has been refined and applied to analyze IMP distributions in other studies (Niedermeyer and Wilke, 1982; Pearson et al., 1983; de Laat ef ul., 1983; Gershon el ul., 1979; Finegold, 1976). Hui and Frank (1985) also applied autocorrelation analysis to study membrane protein distribution. Various random distribution models have been developed to allow for the range of protein-protein interactions (softsphere or hardsphere interaction potentials, sticking time, etc.). The aim of the increasingly sophisticated models was to simulate the nature of protein-protein interactions and the randomization forces of the lipid bilayer. In summary, we know so far that protein distribution in most biological membranes is not ordered, but neither is it random. The details of the forces that keep them from being truly random are still not completely understood, and may differ from case to case.

--

'-

n

50 40

5 u)

30 In 6 20

I0

20

rinml

0

2

4

6 8 1 0 1 2 Occupancy. n

I4

FI(j, I I. ( a ) Freeze-fracture electron micrograph ofthe I l k i c e ofhuman erythrocyte t11cI11hranc. Lines adjoining nearest ncighhoring inti.iinienibriinous particles ( I M P S ) are usell to determine the r d i a l distribution (b). In the lower left corner o f i i . ii 40-nm squ:ire grid i s imposed to obtain the density distrihution (c). The dottcd. dashed. m c i solid lines in b ;ind c arc calculated from random models assuming lypical particle diameters t o he 0. 10. Lrnd 8 nni. respectively.

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Cross-linking of surface receptors is one of the earliest known factors to contribute to the redistribution of membrane protein components. The cross-linking of lectin receptors by multivalent ligands and surface antigens by divalent antibodies causes the receptors and antigens to form patches. For some cells the redistributed surface components would eventually cap on one pole of the cell. Cell cytoskeleton is known to play an important role in the patching and capping processes (Yahara and Edelman, 1974; Bougiuignon and Singer, 1977). In some cases, the distribution of IMPs was found to be independent of the distribution of lectin receptors (Yahara and Edelman, 1974; Kuby and Wofsy, 1981), whereas in other cases, the IMPs were found to cocap with lectin receptors (Loor, 1973; Bennett and Condeelis, 1982). Ash ez ul. (1977) observed the relation between the lectin receptors on the cell surface and the underlying microfilament bundles in fibroblasts, and they suggested that the network of microfilament bundles limited the range of the movement of surface components. In selected cases, the linkage between integral membrane proteins and cytoskeletal proteins is known in some detail (Bennett and Branton, 1977). There is no doubt that certain protein components in the membrane are closely linked to the cytoskeleton and that the linkage is controlled by cell activity. This linkage could be the single most important factor in controlling the distribution of membrane proteins. During transit between plasma membrane and cytoplasmic membrane, such as internalization of coated pits to form coated vesicles or insertion of membrane from Golgi vesicles to plasma membrane, controls of selective connection and disconnection of intrinsic proteins from the cytoskeleton are interesting “questions awaiting answers.” The movement of surface components and IMPs in human erythrocyte membranes are impeded by the underlying cytoskeletal network. No electrophoretic movement of either was observed. The relative immobilization of IMPs by the cytoskeletal network is further supported by the observation (Gerritsen ef ul., 1979) that, after the complete removal of actin and spectrin, the IMPs could be induced to form large patches and large IMP-denuded areas when the temperature was lowered. Such redistribution is most likely caused by the formation of phase separated lipid domains. If the cytoskeleton was left more or less intact, the thermally induced phase separated domains were much smaller, indicating the limited mobility of the IMPs. A normal level of cholesterol in the membrane obliterated the phase change, thereby eliminating the thermally induced IMP patching (Hui et ul., 1980). Thermotropically and osmotically induced IMP patching in lymphocyte membranes was also observed by Speth rt cil. (1981). Lectin receptors in several types of cells were found to move in response

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to an external DC electric field (Zygyansky and Jard, 1979; Poo, 1981). The electric field-related movement of the fluorescent labeled lectin receptors is driven by either electrophoretic o r electroosmotic forces (McLaughlin and Poo, 1981). If the receptors can be displaced by an external electric field and are free to diffuse back after the field is turned off, they must be free to move among membrane lipids. IMPs in isolated inner mitochondria1 membrane vesicles were also observed to be driven to one side of the vesicle by the electromotive force (Sowers and Hackenbrock, 1981). The electric field-induced translocation of IMPs in the plasma membrane of fibroblasts was also reported, although no obvious movement was associated with the Con A receptors (Yang cJt (JI., 1984). There is no evidence so far to prove that protein movement is heterogeneous or homogeneous on the microscopic scale. Such information would be very valuable in determining the relative importance of cytoskeletal control and lipid domains in the electromobility of membrane components. A detailed discussion of membrane component mobility is given in Edidin (this volume). The knowledge gained in the past several years on the identification and distribution of proteins in membranes enables us to say more about the fluid mosaic model. In general, the membrane proteins visualized by fluid mosaic models are essentially correct. There is no regular pattern of protein distribution except in special cases. The distribution, however, is not entirely random. The most likely factors that affect the random distribution of membrane proteins are control by the cytoskeleton, formation of microdomains of membrane lipids, and interaction of membrane proteins themselves. V.

WHAT DETERMINES THE CONFORMATION OF INTRINSIC PROTEIN MOLECULES IN MEMBRANES?

When the fluid mosaic model was proposed, protein structure within the bilayer was virtually unknown. Protein molecules were drawn as globular without any details. A question often asked by membrane biochemists and biophysicists is, “How do intrinsic protein molecules structure themselves in the bilayer?” This problem has been approached from many directions. The amino acid sequences of most intrinsic proteins usually contain a segment or segments that are more hydrophobic than average. From the distribution of the hydrophobic residues of the protein, one can guess which part of the protein is likely to be in the bilayer. For example, in the erythrocyte membrane glycoprotein glycophorin, a sequence of 19 amino acids are more hydrophobic on average than the rest of the mol-

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ecule. This segment was predicted to locate within the bilayer (Furthmayr et al., 1978). An oligomer of these molecular segments can be seen as an IMP (Fig. 12a). These 19 residues account for the a-helix that is long enough to transverse the thickness of the lipid bilayer, as depicted in Fig. 13a. A number of peptides have segments of alternating hydrophobic and hydrophilic residues in a cycle of approximately 39'2 residues. If the sequence of this segment is plotted in a polar diagram representing the endon view of an a-helix (Epand et a/., 1983; Keiser and Kezdy. 1984), the hydrophobic residues would concentrate on one side of the a-helix and the hydrophilic ones on the other side. These so-called amphiphilic peptides would bury themselves half in the bilayer interior and half in the polar environment. Glucagon is an example of an amphiphilic peptide. A crystallographic structure of glucagon (Blundell et d., 1976) showed two amphiphilic a-helices separated by three nonperiodic residues. The amphiphilic segments are plotted in the polar form together with the rest of the sequence in Fig. 13b. Other amphiphilic peptides include serum apolipo proteins, melittin, pendorphin, calcitonin, corticotropin release factor, and prothyroid hormones. They usually react strongly with membrane lipids, especially at the phase transition temperatures of the lipids when bilayer defects are at a maximum. The reactions are so strong that in some cases they completely disrupt the bilayer structure to form individual lipid-protein complexes (Pownall et (11.. 1978; Epand et a/., 1981; Hui et al., 1984a; Epand et ul., 1985). An example of glucagon-DMPC complex is shown in Fig. 12b. The multilamellar structure of the lipid is disrupted by individual

FIG.12. Freeze-fracture electron micrographs of (a)glycophorin in reconstituted vesicles with egg PC (in collaboration with P. L. Yeagle); (b) glucagon/DMPC mixture, at 23°C (in collaboration with R. Epdnd): and (c) bacterial rhodopsin in the purple membrane of Htrlohricterium lialohirrrn (from Fisher and Stoeckenius, 1977). Circles indicate repeating units in (c). Bar, 100 nm.

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lipid-protein complexes. These complexes often appear as stacked disks in negatively stained electron microscopy samples, but freeze-fracture and X-ray diffraction studies found little stacking in aqueous suspensions. The interaction between amphiphilic peptides and phospholipids is not always entirely hydrophobic. In the case of calcitonin, for example, t h e electrostatic force and the exclusion volume effect of the molecules also play an important part (Epand ct a / . , 1983; Hui ct d.,1984a). For larger amphiphilic protein molecules, the more complicated structures within the bilayer cannot always be predicted by the amino acid sequence. However. judging by the hydrophobic moments (vector representations of the hydrophobicity of each residue), Eisenberg c f trl. (1986) were able to predict which proteins were likely to be intrinsic or disruptive to the bilayer. The amino acid sequence of bacterial rhodopsin has been mapped (Gerber et ( I / . , 1979; Ovchiunikov et L I / . , 1979). However. without the guide of a molecular conformation study, its structure within the bilayer would be difficult to unravel. For a long time protein structure in the bilayer could not be determined by electron microscopic studies because of limited contrast and radiation sensitivity of membrane proteins to the electron beam. In 1975, the first membrane protein structure, namely, bacterial rhodopsin in the purple membrane of halobacteria, was derived from an electron microscopic study (Unwin and Henderson, 1975). In this study, glucose-embedded samples were used to improve the resolution normally obtainable by the negative staining technique. A low dose image was also utilized to calculate the phase angles of the electron diffraction data, and a medium resolution structure of the rhodopsin molecule was reconstructed. A three-dimensional projection of bacterial rhodopsin was later derived from a tilting series of electron diffraction data (Henderson and Unwin, 1975). The molecular structure in Fig. 13d shows that each molecule consists of seven a-helices transversing through the bilayer. At 0.7nm resolution, individual amino acids cannot be discerned. The identification of each u-helix (Fig. 13c) was obtained later by the careful labeling studies of Englemen P r a/. (1980), using neutron diffraction. The historic success of "solving" this structure is rather fortunate. The purple membrane of Hri/ohacrer.ir~mprovides several favorable factors for study: ( I 1 the rhodopsin molecules are positioned in a regular array its if in a twodimensional crystal (Fig. 12c); (2) the purple membrane is relatively resistant to radiation; and (3) the amino acid sequence of the molecule is known. Since this ground-breaking work, a number Of membrane proteins have been studied. These include the porin in E. c d i membrane (Engel ct d., 1982). the protein of the gap junction or connexon (Zampighi and Unwin,

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1979; Baker et al., 1983), the protein in the membrane of epithelial cells of urinary bladder (Robertson and Vergara, 1980), the pore-forming protein in the outer membrane of mitochondria (Mannella et al., 1984), cytochrome c oxidase (Fuller et al., 1979), NADH:Q oxidoreductase (Boekema et al., 1982), and the light-harvesting chlorophyll-protein complex (Kuhlbrandt, 1984). These few membrane proteins either already form natural two-di-

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FIG. 13. The amino acid sequences of (a) glycophorin (after Furthmayr er u / . , 1978) and (c) bacterial rhodopsin (after Engelman and Zaccai. 1980) are drawn with respect to their relation to the lipid bilayer (shaded area). (b) Part of the amino acid sequence of glucagon is drawn in a polar diagram. (d) The molecular structure of bacterial rhodopsin. derived from electron diffraction, is also shown (Henderson and Unwin, 1975).

mensional crystals or can be induced to form two-dimensional crystals. Although resolution of the structures of these proteins is not as high as that obtainable from conventional X-ray crystallographic studies, these studies nevertheless represent the true structure of the proteins in the membrane environment. It was not possible to determine the conformation of these proteins until the recent advances of the low dose and image reconstruction techniques. Structural studies of membrane proteins, interesting as they may be, are somewhat restricted. The technique so far can be applied only to the study of regular arrays of molecules. The majority of membrane proteins, as predicted by the fluid mosaic model, are excluded from this type of structural study. Moreover, the membranes have to be relatively stable in the electron beam to withstand the rigor of electron optical recording. The crystals have to be sufficiently thin so that the kinematic theory of electron diffraction applies without the complications of the dynamic scattering effect. The two-dimensional crystals must be large enough so

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that a great number of units can be sampled to reduce the electron beam dosage on each unit. Finally, the position of these units must be well ordered and the crystals must be extremely flat so that the mosaic spread of the electron diffraction peaks is sufficiently narrow to give high resolution data. In order to overcome the last obstacle, which commonly exists in membrane protein crystals, image processing techniques have been applied to “correct” such misalignments. One approach is to use the correlation function method developed by Frank (1980) with the aid of a cornputer to artificially form a perfect crystal from a distorted one. With such tools available we would expect more membrane protein structures to be resolved at higher resolution.

VI.

A LOOK AT THE FUTURE THROUGH THE ELECTRON MICROSCOPE

There are many unsolved problems in membrane structure, many of which concern the molecular dimension which is barely touched by modem electron microscopic techniques. Research of these problems may lead us further into unraveling the mystery of the unique molecular assembly of biomembranes. I shall discuss only a few aspects here as my concluding remarks. The dispute over the existence of microdomains of lipids in biological membranes has not yet been settled, Present electron microscopic techniques cannot visualize compositional or structural lipid domains or clusters of less than 100 lipid molecules. Electron diffraction from such small areas is not strong enough to be utilized as a contrast mechanism. If labeled lipids are used, the tendency of these lipids to form microdomains may be altered. Unfortunately, isotope-labeled lipids do not provide sufficient contrast for electron optics. Some chemical analogs may be used to identify lipid domains by electron microscopy. These lipid analogs, such as thiophospholipids (Hui rt al., 1983a; Vasilenko et al., 1982), may be utilized in electron beam microanalysis. The one or two layers of lipid molecules surrounding intrinsic protein molecules, the so-called boundary lipids, are even more difficult to identify by ultrastructural techniques. The “boundary lipid” supposedly exchanges rapidly with the “bulk lipid.” At present, the mapping of such a thin, rapidly exchanging lipid shell by a relatively static electron microscopic method is not yet possible. Recently, single paraffin molecules were imaged by low dose, low temperature electron microscopy (Zemlin er a,!.,1985). The application of this technique to the study of lipid structure in biomembranes would be most interesting.

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The insertion of proteins into lipid bilayers is another interesting problem. In some cases, insertion of protein is at a maximum during the lipid phase transition (Wickner. 1977; Pownall et d., 1978; Epand cr ul., 1981; Prendergast ef a / . , 1982; Terwilliger rr d.,1982; Hui et ul., 1984a). Presumably the hydrophobic protein inserts itself into bilayer defects which express at a maximum during the highly fluctuating state of phase transition. Structural defects also exist below the phase transition along the domain boundaries and at points where the bilayer structure is disrupted 1981). These structural defects (Blaurock and Gamble, 1979; Hui er d., were suggested to be responsible for spontaneous membrane fusion (Larrabee, 1979; Boni er d.,1984) and also to facilitate the incorporation of membrane proteins (Hah et d.. 1983; Cheng er d . , 1986). However, the actual insertion process has never been visualized by electron microscopy as a kinetic process, although the breaking-up of vesicles of DMPC and sphingomyelin by glucagon has been arrested by rapid freezing electron microscopy (Epand et al., 1981). Electron microscopy has long been regarded as a static technique. Only samples that have been fixed at a certain stage of reaction can be studied. The time resolution is determined by the length of time taken to fix a specimen. With chemical fixation, this process takes minutes. With the advance of rapid freezing, the fixation time can be reduced to milliseconds, Heuser and Reese (1981) have improved the fixation time to less than a millisecond in order to catch the dynamic events of synaptic vesicle release in the neuromuscularjunction upon electric stimulation. Figure 14 shows a reconstructed time sequence for the release of synaptic vesicles using the rapid freezing method. Other kinetic events have also been studied by rapid freezing techniques. These include membrane fusion by calcium injection, using a stopflow apparatus (Miller and Dahl, 1982),and the triggering of membrane fusion by an electric pulse (Stenger and Hui, 1986). Such kinetic studies have made electron microscopy a dynamic tool rather than a static one. In a strict sense, only static pictures are obtained, and the dynamic events must be reconstructed from a series of static observations. The true imaging of dynamic processes such as those observed with light microscopes can only be accomplished by using an in sifu environmental stage in an electron microscope (Costa and Hui, 1979) or by using flash X-ray pulses in an X-ray microscope (Penessa-Warren cr id., 1982; Fader r t d.,1985). If the membrane sample is amenable to crystallographic analysis, synchrotron X-ray diffraction offers a possibility for sampling a dynamic process at millisecond intervals. The structural changes in photoreceptor disc membranes upon exposure to light were documented by Charbre (1975). The increasing spatial and temporal resolutions from electron micro-

FIG.14. Montage of six different forms of synaptic vesicle exocytosis selected fromfreezefracture micrographs of the active zone of frog neuromuscular junction stimulated at a given time before rapid freezing. Stimulation-freeze intervals were (a) 3.7 msec, (b, c, d) 5.2 msec, (e) 20 msec, (0 50 msec. Also shown are freeze-substituted sectional views of nerve terminals frozen at rest (g) and frozen 5.2 msec after stimulation (h). Many pockets which resemble collapsed vesicles occur in the stimulated active zone (Heuser and Reese, 1981). Bar, 100 nm.

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scopic and crystallographic studies will dramatically improve our understanding of the actual molecular arrangements in membranes. In combination with spectroscopic studies, we may eventually be able to visualize the membrane structure in the molecular dimension.

AC KNOW LE DG M E NTS

I wish to thank Dr. Arindam Sen for helpful discussions and for reading the first clrnft of the manuscript. Some of the material in this article was organized in preparation for a workshop supported by the Neurosciences Institute. Permissions lo reproduce micrographs from many authors quoted in this article are gratefully acknowledged. Those illustrations not referenced are from my laboratory. Research leading t o these results was supported by research grants from the National Institutes of Health (GM-28120. GM-30969, and RR-01474) and from the American Cancer Society (BC-248).

REFERENCES Abbas. A. K., Auk, K. A.. Karnovsky. M . J . . and Unanuc. E. K. (1975). ftiwirrtio/ogv 114, 1197-1204. Albert. A., Sen, A.. and Yeagle. P. L. (19x4). 5ii)diim. 5 i o p h y s . Ai.tir 771, 28-34. Anders, J . J . . and Brightman, M. W. (1982). J. Nerrrocyiol. 11, 1009-1029. Anderson, R. E.. and Maude, M. B. (1970). Bioc.h~ini.srry9, 3624-3628. Andrews, I,.. and Cohen, A. I. (1979). 1.Cc4 Biol. 81, 215-228. Arancia. G.. Belli. S . , Donelli, G.. and Trovalusci, P. (1980a). J . Giw. M i c ~ ~ h i oIIY, l . ISS164. Arancia. G., Valente. F. R., and Crateri, P. 1’. (1980b). J . Micros. 118, 161-176. Armond. P., and Stdehelin, A. (1979). Proc. Ncrrl. Acrid. Sci. U.S.A. 76, IWI-1905. Asano. A.. and Sekiguchi. K . (1978). J. Sirprwnol. S’rrucr. 9, 44-52. Ash. J. F.. Louvard, D., and Singer. S. J . (1977). P r w . N d . Arcid. Sii. U.S.A. 74, 55845587. Baker. T. S . . Caspar, D. L. D.. Hollingshead, C. J., and Goodenough, D. A. (1983). ./. Cell Biol. 96, 204-216. Bayer. M. E., Dolack, M.. and Houser, E. (1977). J . Btrctrriol. 129, 1.563-1573. Bearer, E. L., and Friend. 0. S. (1980).Proc. N o t / . Accrd. Sii. U.S.A. 77, 66014605. Benitez, C., Caspar, D. L. D., and Kirschner. 0. A. (1977). In “Electron Microscopy” (G. W. Bailey, ed.). pp. 6 ~ I Claiton. . Baton Rouge, Louisiana. Bennett, H., and Condeelis, J. (1982). Biophys. J . 37, 17%. . 252, 2753-2763. Bennett. V., and Branton. D. (19771. J. B i ~ l Clwm. Blaurock, A. E . . and Gamble, R . C. (1979). J . Mrnihr. Lliol. 50, 187-204. Blaurock, A. E., and Wilkins, M. H. F. (1972). Ntrrrrre ( L o n d o n ) 236, 313-314. Blundell. T. C., Dockerill, S.. Sasaki. K., Tickle, 1. J.. and Wood. S. P. (1976). Mctuho/i.vtiz 25 (SUPPI.I ) , 1331-1336. Boekema. E. J.. van Breeman, J. F. L.. Keegstra. W.. van Bruggen, E. F. J.. and Albracht. S. P. J. (1982). Biodiiw~.Biophys. Artti 679, 7-11. Boni, L. T., and Hui. S. W. (1983). Biochitn. Bioplr,w. Actrt 731, 177-1x5.

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SEK-WEN HUI

Boni, L. T., Hah, J. S. , Hui, S. W., Mukhejee, P., Ho, J. T., and Jung, C. Y. (1984). Biochim. Biophys. Actu 775, 409-419. Borggreven, J. M. P. M., Daemen, F. J . M., and Bonting, S. L. (1970). Biochim. Biophys. Acta 202, 374-38 I . Bosterling, B., and Trudell, J. R. (1982). Biochim. Biophys. Actu 689, 155-160. Bougiuignon, L. Y. W., and Singer, S. J. (1977). Proc. Nut/. Acud. Sci. U . S . A . 74, 50315035. Bridgman. P. C., and Nakajima. Y. (1981). Proc. Nut/. Acud. Sci. U.S.A. 78, 1278-1282. Brown, D., Montesan, R., and Orci, L. (1982). J . Hisfoehem. Cytochem. 30, 702-706. Burnell, E., van Alphen, L., Verkleij. A. J., and de Kruijff, B. (1980). Biochim. Biophvs. Actu 597,492-501.

Caspar, D. L. D., and Kirschner, D. A. (1971). Nuture (London) 231, 46-52. Chabot, J. F., and Leopold, A. C. (1982). Am. J . Bot. 69, 623-633. Charbre, M. (1975). Biochim. Biophys. Actu 382, 322-343. Cheng, H. K., Lepock, J . R., Hui, S. W., and Yeagle, P. L. (1986). J . B i d . C h e m . 261, SO8 1-5087. Copeland, B. R., and McConnell, H. M. (1980). Biochim. Biophys. A i m 599, 95-109. Corless, J. M., and Costello, M. J. (1981). Exp. Eye Res. 32, 217. Costa. J. L., and Hui. S. W. (1979). Biophys. J . 19, 307-313. Cottam, P. F., Hui, S . W., He, N. B.. and Ho, C. (1986). Biochern. Biophys. Actu 862, 413-428. Crowe, L. M., and Crowe, J. H. (1982). Arch. Biochem. Biophys. 217, 582-587. Cullis, P. R . . and Hope, M. (1978). Nutrrre (London) 271, 672-674. Cullis, P. R., de Kruijff, B., Hope, M. J.. Nayar, R.,Rietveld, A., and Verkleij, A. J. (1980). Biochirn. Biophys. ACIN 600, 625-635. Danielli, J . F., and Davson. H. (1935). J . Cell. Physiol. 5, 495-508. de Grip, W. J., Drenthe. E. H. S . , van Echteld, C. J. A., de Kruijff, B., and Verkleij, A. J . (1979). Biochirn. Biophys. Acta 558, 330-337. de Kruijff, B . , and Dewel. R . A. (1974). Biochim. Biophys. Actu 339, 57-70. de Laat, S. W., Tertoolen. L. G. J., van er Saag, P. T., and Bluemink, J. G. (1983). J . CelI. Biol. 96, 1047-1055. Eisenberg, D., Wilcox. W., and McLachlan, A. D. (1986). J . Cell Biochem. 31, 11-17. Elgsaeter, A., Shotton, D. M., and Branton, D. (1976). Biochim. Biophys. Actu 426, 101122. Elias, P. M . , Friend, D. S. , and Goerke, J. (1978). J . Cell Biol. 78, 577-596. Engel. A.. Massalski, A., Rosenbusch, J . P.. Dorset, D., and Saxton, 0. W. (1982). Proc,. 10th I n ! . Congr. Electron Microsc. 3, 43-44. Engelman, D. M.. and Zaccai. G. (1980). Proc. Nut/. Acud. Sci. U.S.A. 77, 5894-5898. Epand, R. M., Epand, R. F., Stewart, T. D., and Hui, S. W. (1981). Biochim. Biophys. Acta 649, 608-615. Epand, R. M., Epand, R. F . , Orlowski, R. C., Boni, L. T.. and Hui, S . W . (1983). Biochemistry 22, 5074-5084. Epand, R. M., Epand, R. F., Hui, S. W., He, N. B . , and Rosenblatt, M. (1985). Int. J . Pept. Protein Res. 25, 594-600. Fader, R.,Banton, V., Sayre, D., Costa, J., Baldoni, M., and Kim, B. (1985). Science 227, 63-64. Feltkemp, C . A.. and van der Waerden, A. W. M. (1982). CellBiol. In/. R e p . 6, 137-145. Finegold, L. (1976). Biochim. Biophys. Actu 448, 393-398. Fisher, K. A.. and Stoeckenius, W. (1977). Science 197, 72-74. Frank, J. (1980).-In"Computer Processing of Electron Microscope Images" (P. W. Hawkes, ed.), Springer-Verlag, Berlin and New York.

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

67

Friend. 1). S., and Bearer, E. L. (1981). Hiutochem. J . 13, S3S-546. Fuller, S. D.. Capaldi, R. A,. and Henderson, R. (1979).J. M o l . Biol. 134, 305-327. Furthmayr. H.. Galardy, R. E., Tomita. M.. and Marchesi, V. T . (1978). Art,/?.Bioc.herir. Bioph>’.s. 185, 21-29.

Gerber. G. E.. Anderegg, R. J., Helihy, W. C.. Gray, C. P.. Biemann, K., and Korana, H. G. f 1979). PWC. NU//. Al.rit/. S(.i. U . S . A . 76, 227-23 I . Gerritsen, W. J.. Verkleij. A. J., and van Deenen. L . 1,. M. (1979). Bicwhirn. Biopl7y.s. Actcr 555, 26-4 I . Gershon, N. D.. Demsey, A.. and Stackpole. C. W. (1979). Exp. Cell Res. 122, 11-126. Giese. G., Fromme. I.. and Wunderlich. F. (1979). Eur. J. Biochrm. 95, 275-28s. Gilula. N. B., Eger, R. R.. and Fifkin. D. B. (1975). Proc. N o t / . Acnd. Sci. U . S . A . 72, 3594-3598.

Goormaghtigh, E.. van den Branden, M., Ruysbchaert. J . M., de Kruijff. B. (1982).Bioclrirn. B i o p h y s . A c ~ 685, ~ I 137-143. Gounaris, K., Brain, A. P. R., Quinn. P. J., and Williams, W. P. (1983a). FEBS Ltrr. 153, 47-52.

Gounaris, K.. Sen. A.. Brain. A. P. R.. Quinn. P. J.. and Williams, W. P. ( 1983b). Biochirn. Biophys. Ac/u 728. 129-139.

Gruner-. S. M.. Rothchild. K. J.. and Clark. N . A . (1982). Bioplrv.~.J . 39, 241-245. Gulik-Krzywicki. T., Balerne, M . , Vincent, J. P.. and Lazdunski, M. (19x1). B i i d r i r n . Biop l ~ y .A ~ c. / ( I 643, 101-104.

Hackenbrock, L. R., Hochli. M..and Chau. R. M . (1976). Biochim. Biop/ry.s. A(,/tr 455, 466-484. Hah. J. S.. Hui, S. W.. and Jung, C. Y. (1983). Bioclrenri.s/ry 22, 4763-4769. Halvarson, C. A.. Esser. A. F., and Sonja, K. A. (1978). J . S~cprcrrnol.S/rirc/. 8, 129-138. Hauser, H., Pascher. I., Pearson, R. H., and Sundell, S. (1981). Biochirn. Riopkys. Actci 650, 2 1-5 1. Henderson. R.. and Unwin, P. N . T. (1975). Ntr/irrc (Lorrdon) 257, 28-32. Herhette, L.. Scarpa, A,. Blasic, J . K., Wang. C . T . , Saito. A . , and Fleischer. S. ( l 9 X l ) . Biopky.S. J . 36, 47-72. Heuser. J. E., and Reese. T. S . (19x1). J . roll B i d . 88, 564-580. Hildebrand. C.. and Muller. H. (1974). Nrrrrohiology 4, 7 1 4 1 . Hochli. M.. and Hackenbrock, C. R. (1976). Proc. Nnrl. Acrid. Sci. U . S . A . 73, 1636-1640. Hollingshead. C. J.. Caspar. D. L. I)., Melchior. V., and Kirschner, D. A. (19761. Proc.. Nn//. Aiwd. Sci. U . S . A . 73, 1636-1640. Hoover, R. L.. Davidowicz, P. A.. Robinson. J . M . , and Karnovsky, M. J . (1981) C/W/??.97, 73-80. Hosemann. R.. and Bagchi, S . N . (1962). “Direct Analysis of Diffraction by Matter,” p. 111. North-Holland Publ.. Amsterdam. Huet, C., Levy, M., and Pascaud. M. (1968). Biochiwr. Biophys. Act11 150, 521-524. Hui. S. W. (1981). Biophys. J. 34, 383-394. Hui. S. W.. and Frank. J. (198s). J . Micwsc. 137, 293-303. Hui. S . W.. and He, N . B. (1983). Bioc/rt,mi.srry 22, 1159-1164. Hui. S. W., rind Parsons, D. F. (1975). S ~ ~ i c ~ n190. c e 383-384. Hui. S. W.. Stewart, C. M.. Carpenter. M. P . . and Stewart, T. P. (1980). J . Cc4l B i d . 85, 283-291. Hui. S. W., Stewart. T. P., Boni. L. T.. and Yeagle, P. L. (1981). Science 212, 921-93. Hui. S. W., Boni. L. T., Stewart, T. P.. and Isac. T. (1983a). Biockrmislp 22, 351 1-3516. Hui. S. W., Stewart. T. P.. and Boni. L. T. (1983b). Clrem. Phys. lipid.^ 33, 113-126. Hui. S. W.. Epand, P. M., Dell, F. R.. Epand. R. F.. and Orlowski, R. C. (1984a). Bioclrim. Bio/)hy.v. A(’r11772, 264-272.

68

SEK-WEN HUI

Hui, S . W., Mason, J., and Huang. C. H. (1984b). Biochemistry 23, 5570-5577. Hui, S . W., Isac, T., Boni, L. T., and Sen, A. (1985). J. Membr. Biol. 84, 137-146. Jain, M. K. (1983). In “Membrane Fluidity in Biology: Concepts in Membrane Structure” (R. Aloia, ed.), pp. 1-37. Academic Press, New York. Jones, G. J. (1974). J . Cell Sci. 16, 199-219. Kachar, B., and Reese, T. S. (1982). Nufure (London) 296, 464-465. Keiser, E. T., and Kezdy, F. J. (1984). Science 223, 249-255. Kim, J., and Okada, Y. (1983). Eur. J . Biochem. 29, 244-252. Klausner, R. D., Kleinfeld. A. M., Hoover, R. L., and Karnovsky, M. J. (1980). J. B i d . Chem. 255, 1286-1289. Kleeman, W., and McConnell, H. M. (1976). Biochim. Biophys. Arfrr 419, 206-222. Knutton, S. (1977). J . Ce/l Sci. 28, 189-210. Knutton, S. (1979). J. Cell Sci. 36, 61-72. Krahling, H. (1981). Acta Histochem. Suppl. 23, 219-233. Kuby, J. M., and Wofsy, L . (1981). J . Cell B i d . 88, 591-598. Kuhlbrandt, W. (1984). Nufitre (London) 307, 478-479. Larrabee, A. L. (1979). Biochemistry 18, 3321-3326. Lee, A. G. (1977). Biochim. Biophys. Acta 472, 237-281. Legendre, S., Letellier, L., and Schechter, E. (1980). Biochim. Biophys. Acta 602, 491. Lentz, B. R., Barrow, D. A., and Hochli, M. (1980). Biochemistry 19, 1943-1954. Loor, F. (1973). Eur. J . Immunol. 3, 112-1 16. Lucy, J. A. (1975). In “Cell Membrane, Biochemistry, Cell Biology and Pathology” (G. Weissman and R. Claiborne, eds.), pp. 75-83. H P Publishing, New York. Luna, E. J., and McConnell, H. M. (1978). Biochim. Biophys. Acta 509,462-473. Luzzati, W. (1968). In “Biological Membranes: Physical Facts and Functions” (D. Chapman, ed.), pp. 71-123. Academic Press, New York. McCaughen, L., and Krimm, S. (1980). Science 207, 1481-1482. McIntosh. T. J., Simon, S. A., Ellington, J. C., and Porter, N. A. (1984). Biochemistry 23, 4038-4438.

McLaughlin, S., and Poo, M. M. (1981). Biophys. J . 34, 85. Mannella, C. A., Colombini, M., and Frank. J. (1984). Proc. Not/. Acad. Sci. U.S.A. 80, 2243-2247.

Mdrchesi, V. T., Jackson, R. L.,Segrest, J. P., and Kahane. I. (1973). Fed. Proc., F f d . Am. Sac. Exp. B i d . 32, 1833-1837. Markovics, J.. Glass, L.. and Maul, G. G. (1974). Exp. Cell Res. 85, 443-451. Marsh, D. A., Watts, A.. and Knowles, P. F. (1977). Biochim. Eiophys. Actu 465, 500-514. Melchior, V., Hollingshead, C. J., and Caspar, D. L. D. (1979). Biochim. Biophys. Acta 554, 204-226.

Melkonian, M., Robenek, H., and Steup, M. (1979). Photop/asmu 109, 349-358. Miller, D. C., and Dahl, G. P. (1982). Biochim. Biophys. Actn 689, 165-169. Miller, R. G. (1980). Nature (London) 287, 166-167. Miller, R. G. (1984). Biochim. Biophys. Acta 774, 151-157. Miller, R. W. (1980). Can. J . Biochem. 58, 1138-1 143. Montesano, R., Perrelet, A., Vessalli, P., and Orci. L. (1979). Proc. Nut/.Acrid. Sci. U . S . A . 76, 6391-6395.

Navarro, J., Toivio-Kinnucan, M., and Rocker, E. (1984). Biochemistry 23, 130-135. Nemeth, A.. and Rohlich, P. (1981). Acta Hisrochem. Suppl. 23, 181-187. Niedermeyer, W., and Wilke, H. (1982). J . Microsc. 126, 259-273. O’Brian, J., and Sampson. E. L. (1965). J . LipidRes. 6, 537-551. Ojakian. G. K., and Satir, P. (1974). Proc. Natl. Acad. Sci. U.S.A. 71, 2052-2056.

THE MOLECULAR ASSEMBLY IN BIOMEMBRANES

69

Ovchiunikov. Yu, Abdulaev, N.. Fergira. M.. Kiselev. A,. and Lobanov, N. (1979). FEBS Lerr. 100, 219-224.

Pape. E. H., Klott. K.. and Kreulz, W. (1977). Biophys. J. 19, 141-161. Pearson. R. P., Hui, S. W.,and Stewart, T. P. (1979). Biochitn. Biophys. Actcr 557, 265282.

Pearson, L. T., Chen, S. I., Lewis, B. A.. and Engelman. D. M. (1983). Biophvs. J. 43, 167- 174.

Penessa-Warren, B. J., McCorkle, K. A.. Hoffman. P.. Warren, J. B., and Colman, G. (

1981). U//rrirtzicroscc,p?,6, 139-148.

Perrelel. A,, Garcia-Segura. L. M., Singh. A . . and Orci. L. (1982). Proc. N i t / / . Actid. Sci. U . S . A . 79, 2598-2602.

Pinto da Silva. P.. and Martinez-Palomo. A. (1975). Proc. NNII.Accid. S t i . U.S.A. 72, 572576.

Pinto da Silva, P., and Torrisi, M . K. (1982). J . Cell Biol. 93, 463-469. Pinto d a Silva. P.. Shiniizu, K., and Parkinson. C. (1980).J . Cell Sci. 43, 419-432. Plattner. H. (1981). Cell B i d . f n t . Rep. 5, 435459. Poo. M. M. (1981). Annrr. Rev. Biophys. Bioeng. 10, 245. Pownall, H. J., Massey, J. B., Kussero. S. K.. and Gotto, A. M. (1978). Biockemistn. 17. 1183-1 189.

Prendergast. F. G.. Lu, J.. Wei, G. J.. and Bloomfield, V. A. (1982). Biocliemisiry 21,6963697 1.

Puccinelli, V . , Innocenli. M., Lasagni, A., and Caputo. K. (1977). Br. J . Derrrrriiol. 96. 515519.

Pumplin. D. W., and Bloch. R. J. (1983). J . Bit>/.Clirrn. 97, 1043-1054. Rand. R . P., Reese. T. S., and Miller, R. G. (19x1). Nrirrrrc, (LotidonJ293, 237-239. Rash, J . E.. Johnson, T. J. A,. Hudem, C. S . . and Giddings. F. D. (1982). J . M i c ~ s c .128, . I2 1-128. Robrnek, H.. and Greven, H. (1981). Etrr. J. Bioc.hc,rn. 25, 131-135. Robenek. H.. Jung, W.. and Ciabhardt. R. (1982). J . Ulrrtrsrr. Rcs. 78, 91-106. Robertson, J. D. (1959). Biodretn. S y m p . 16, 3 4 3 . Robertson. J . D. (1981). J . Cell B i d . 91, 1895-2045. Robertson. J. D.. and Vergara, J . (1980). J . Cell B i d . 86, 514-528. Roos, D. S.. Robinson. J. M., and Davidson, R. L. (1983). J . Cell B i d . 97, 909-917. Rouser. G.. Nelson, G. J.. Fleischer, S.. and Simon. G . (1968). In “Biological Membranes” (D. Chapman, ed.), pp. 5-69. Academic Press, New York. Sackmnnn, E. (1978). Bcr. Biinscwge.7, Phys. Chem. 82, 891-909. Sailo. A.. Wang. C. T., and Fleischer. S. (197x1. J. Cell Biol. 79, 601-616. Sase. S . . Anraku. Y.. Nagano, M.. Osuni. M., and Kasahara. M. (1982). J . Biol. CltcJrii. 257, I 1100-1 1105. Schneider. H.. Hochli, M.. and Hackenbrock. C. (1982). J. Cell Biol. 94, 387-393. Schneider, M., Chan, W. K.. and Webb. W. W. (1983). Bioplivs. J . 43, 1.57-165. Seelig, J . (1978). Biochirn. Bioph,vs. Actcr 515, 105-140. Sekiya, T.. Kitajima, Y .. and Nozawa, Y . (1979). Biochini. Biopliys. Ac,/tr 550, 269-778. Sen. A.. Williams. W. P., Brain, A. P. R.. and Quinn. P. J. (1982). Bicwhitrt. B i o p l i w . Actcr 685. 295-306.

Severs. N. J.. and Robenek. H. (1983). Bioc.kitti. Biopliys. Actu 737, 3 7 3 4 0 4 . Singer, S. J.. and Nicolson. G. (1972). Scicvicr 173, 7 2 e 7 3 1 . Sklar, 1.. A., Hudson, H., and Simoni. R. I). (1977). Biotherni.sr/.v 16, 819-828. Sowers. A . E., and Hackenbrock. C . P. (19x1). /‘roc.. Nor/. Actrcl. .S(,i. U.S.A. 78, 6?46. Speth, V . . Bruer. H. L.. and Brunner. G. (1981). Bioc,/rirn.B i o p h y . ~A. m 649, 113-120.

70

SEK-WEN HUI

Steck, T. L. (1974). J . Cell Biol. 62, 1-19. Stenger, D., and Hui, S. W. (1986). J . Membr. Biol. 93, 43-50. Stewart, T. P., Hui, S. W., Portis, A. R., and Papahddjopoulos, D. (1979). Biochim. Biophys. A c / 556, ~ 1-16. Stier, A., Finch, S. A. E., and Bosterling, B. (1978). FEES Lett. 91, 109-1 12. Tdkagi, A. (1971). Biochim. Biophys. Actu 248, 12-20. Taraschi, T. F., van der Steen, A. T. M.. de Kruijff, B., Tellier. C., and Verkleij, A. J. (1982). Biot~hemi.stry21, 5756-5764. Terwilliger, T., Weissman, L., and Eisenberg, D. (1982). Biophys. J . 37, 353-361. Thayer, A. M., and Kohler, S. J. (1981). Biochemistry 20, 6831-6835. Unwin. P. N . T., and Henderson, R. (1975). J . Mol. B i d . 94, 425-450. Vail, W. J., and Stollery, J . G. (1979). Biochim. Biophys. Actrr 551, 74-84. van Dijck, P. W. M., de Kruijff, B., Verkleij, A. J., and van Deenen, L. L . M. (1978). Biochirn. Biophys. Actcc 512, 84-96. van Echteld. C. J. A.. de Kruijff. B., Verkleij, A. J., Leunissen-Bijelt, J., and de Gier, J. (1982). Biochitn. Biophys. Actu 692, 126-138. van Hoeven, R. P., and Emmelot, P. (1973). In "Tumor Lipids" ( R . Wood, ed.). pp. 126138. Am. Oil Chem. SOC.,Champaign,Illinois. van Venette, R., and Verkleij, A. J. (1982). Biochitn. Biophys. Acru 692, 397-405. Vasilanko, I . A., de Kruijff. B.. and Verkleij. A. I. (1982). Biochim. Biophys. Acra 658, 144- 152. Verkleij, A. J . (1984). Biochirn. Biophys. Actu 779, 43-63. Verkleij, A . J., and van Golde. L. M. G. (1975). J . Bucreriol. 124, 1522-1528. Verkleij. A. J., Ververgaert, P. H. S., van Deenen, L. L. M., and Elbers, F. (1972). Biochitn. Biophys. Actu 288, 326-332. Verkleij. A. J . , Mombers, C., Gerritsen, L., Leunissen-Bijvelt. L., and Cullis. P. R . (1979a). Biochitn. Biophys. Acra 555, 358-65. Verkleij, A. J.. Mombers, C., Leunissen-Bijvelt, J., and Ververgaert, P. H. S. (1979b). Nutitre (London) 279, 162-163. Vos. J., Ahkong, Q. F.. Bothan, G. M., Quick, S. J . , and Lucy, J. A. (1976). Biochem. J . 158, 651-653. Vosky, D., and Loyter, A. (1977). Biochitn. Biophys. A m 471, 243-259. Wang. C. T., Saito. A., and Fleischer, S. (1979). J . B i d . Chem. 249, 302-309. Weinstein, R. S. (1976). Cuncer Res. 36, 2518-2524. Weis. R. M., and McConnell, H. M. (1985). J . Phys. Chem. 89, 4453-4459. Weiss, L., Jung, 0. S., and Ziegel, R. (1972). In/. J . Cuncer 9, 48-56. Wickner, W. T. (1977). Biochemistry 16, 254-258. Williamson, P. L., Massey, W. A,, Phelps, B. M., and Schlegel, R. A. (1981). Mol. Cell. B i d . 1, 128-135. Wolff, D. E.. Schlessinger, J., Elson, E. L., Webb, W. W., Blumenthal, R.. and Henkart. P. (1977). Biochemisrry 16, 3476-3483. Wunderlich, F., Batz. W.. Speth. V., and Wallach, D. F. H. (1974). J. Cell Biol. 61, 633. Yahara, I.. and Edelman, G. M. (1974). Ann. N . Y . Acud. Sci. 253, 455. Yang, W. P., Onuma. E.. and Hui. S. W. (1984). Exp. Cell Res. 155, 92-104. Zakai, N . , Kulka, R. G . . and Loyter, A. (1977). Proc. N u t / . A c a d . Sci. U.S.A. 74, 2417242 I . Zampighi, G . , and Unwin, P. N. T. (1979). J . Mol. B i d . 135, 541-564. Zemlin, F., Reuber, E., Beckmann, B., Zeitler, E., and Dorset. D. L. (1985). Science 229, 46 1-462. Zygyansky. Y. A., and Jard, S. (1979). Nuture (London) 280, 591-593.

CUKKF.NT‘ropicsI N M E M B K A N E S A N D .IKANSPOKT.

V O L U M E ?Y

The Thermodynamics of Cell Adhesion MICAH DEMBO A N D GEORGE I. BELL lhrorcticcil Division Lo.\ A1trmo.s Nirrionul Lriborirrorv Los Altrtnos. N e w Me.ti1.o 87545

1.

INTRODUCTION

Equilibrium thermodynamics as a rigorous discipline is restricted to the description of completely inanimate or ”conservative” systems. Nevertheless, even in biological systems that are more or less far from equilibrium, thermodynamic models can give us some idea of the natural end point towards which the system would tend if active metabolism were neglected. In the case of cell-cell and cell-surface adhesion the study of thermodynamic models is also important because these models can provide a language for the clarification and integration of a variety of subtle and vaguely defined notions.

II. THEMODEL

In order to illustrate the principles involved in taking a thermodynamic approach to modeling cell adhesion, we will present the detailed formulation and analysis of an elementary example: adhesion of a cell to a large flat surface, herein called the “substrate.” To start with, consider that the cell has “receptors” that can bind, in a lock and key fashion. to complementary ‘‘sites” on the substrate. We will regard the sites as fixed and immobile in the plane of the substrate. In contrast, we will allow the receptors to be freely mobile in the entire plane of the plasma membrane.’

‘ A detailed discussion of a different special case. adhesion of a cell to another cell where receptors on both cells are free to diffuse laterally, appears elsewhere (Bell (’I ( / I . , 1984). 71

Copyright nl 19x7 by Aciidemic Pres*. Inc. All rights of‘ reproduction in a n y form re\erved

72

MICAH DEMBO AND GEORGE I. BELL

In sum, we will view cell adhesion as resulting from the formation of specific receptor-site bonds; the net effect of all other forces between cell and substrate is taken to be repulsive. Given this rather elementary formulation, the next problem is to define precisely what is meant by “contact” between the cell and the substrate (see Fig. 1). To accomplish this, let the surfaces of the cell and the substrate be represented by two sets of points, {A,} and {As},respectively. We now propose that interactions between the cell and the substrate can only occur within a definite subset of “contact points,” {A}. Naturally, inside the set {A} the cell surface and the substrate must be in relatively close proximity, with the average separation distance S. We will regard points on the remainder of the cell surface as being separated from the substrate by distances very much greater than S. Let us denote the total surface density of receptors and sites at a position x by n,(x) and n,,(x), respectively. Since the substrate is a uniform material, and since sites are immobile, it is clear that n,,(x) = n,, must be constant for all x E {As}.Furthermore, because receptors and sites can only be in one of two states, free or attached, we surmise that both total surface densities can be broken into two terms

n,t = n,(x) -t nh(x)

X

E {A,}

(2)

where n,(x) and n,(x) are the surface densities of unattached receptors and sites and &(x) is the local surface density of cell-to-substrate bridges. If we consider the distribution of bridges over the plasma membrane, it is obvious that nh(x) must be zero outside the contact region. On the other hand, since entropy must be maximized near thermal equilibrium, the equivalence of all contact points implies that the density of cell-substrate bridges will be a positive constant for points inside the contact set. Therefore, if A is the area of the set {A}, and N h represents the absolute number of bridges between the cell and the substrate, then near equilibrium the density of bridges on the plasma membrane is a discontinuous function nh(X) = nh

Nb/A

x E {A}

and nh(X) = 0

x E {A,} - {A>

(3a)

Furthermore, inserting this expression into Eq. (2), it is easily seen that the density of free sites on the substrate is also discontinuous

73

THE THERMODYNAMICS OF CELL ADHESION

~I,(x) = n,,

-

x E {A}

n,,

and n,(x)

x E {A,}

= n,,

-

{A)

(3b)

In contrast to sites on the substrate and cell-substrate bridges, unattached receptors are freely mobile. Thus, since free receptors can exist with equal probability at all points on the cell surface, both inside and outside the contact region, we see that close to thermal equilibrium n , ( x ) must have the same density both inside and outside the contact area. Thus, if A, is the area of the set {A,} (i.e., A, is the total cell surface area). then IZ,(X)

= n, =

(A',, - N,)/A,

x E {Ac}

(3C)

We are now in a position to consider the Gibbs free energy of the closed system formed by a cell in contact with a substrate. To this end, let the repulsive force between membrane and substrate, at a separation distance S , be F(S) per unit area of contact. Furthermore, let p,[n,(x)], p,[n,(x)l, and pb[nh(x),S]represent the chemical potentials of receptors, sites, and bridges at an arbitrary position x. Given this notation, the free energy will

Unattached Receptors

Unattached Sites

f

Cell - Substrate Bridges

FIG. I . The model of cell adhesion. A cell is assumed lo possess a fixed number of freely diffusing receptors. whereas the substrate is covered by a constant density of imrnohile sites. Binding between receptors and sites can occur only in a "contact region'' where the separation distance between the two surfaces is small.

74

MICAH DEMBO AND GEORGE I. BELL

be of the form

(4a)

+ constant terms In order of appearance the various terms in Eq. (4a) represent (1) the free energy of unattached receptors on the cell surface; (2) the free energy of unoccupied sites on the substrate; (3) the free energy of cell-substrate bridges; (4) the work that must be done against nonspecific forces in order to bring the cell and substrate to a separation distance of S over an area A; and (5) constant terms representing the free energy of those parts of the system that are unperturbed by the occurrence of contact. Using Eqs. (3a), (3b), and (3c) we can quite simply compute the various integrals in Eq. (4a). When this is done, we obtain the simpler expression G(Nd,S)

=

(Nrt

-Nb)F,[(N,t - Nt,)/A,I

i(Anst -

NtJPJn,, -(NdA)I

+ (A, - A)nst~Jn,t1

(4b)

+ Nbpb(Nb/A,S)+ A r ( S ) + constant terms where A, is the total area of the substrate and z

us) = .I- m d Z S

is the repulsive potential per unit area of contact. When written in the form of Eq. (4b), it is apparent that the free energy can be regarded as a function of only three independent variables: &, A, and S. Thus, by finding the minima of Eq. (4b) with respect to these three variables, we can determine the number of distinct stable states of the cell-substrate system as well as the most probable equilibrium properties of these states. However, in order to proceed with an analysis of the free energy function, we must first obtain explicit expressions for the chemical potentials and for the potential energy of nonspecific repulsion.

111.

THE CHEMICAL POTENTIALS

In the “ideal solution” limit, the chemical potentials of the free receptors and free sites can be related to the corresponding concentrations according to the usual expressions

75

THE THERMODYNAMICS OF CELL ADHESION

and

pr(n,)= p:!

+ kT t‘n(n,)

(Sa)

k,(n,) = ky

+ k7’ tn(n,)

(Sb)

Notice that although the receptors are freely mobile and the sites are fixed in space, this makes no difference as far as the form of the expressions for the chemical potential is concerned. The chemical potential of a cellsubstrate bridge is only slightly more complicated: pLh(nh,S) = pE(S) + k’l’ h(n,,)

(5c)

In Eqs. (5aL (5bL and (5c), p:, p:‘, kE(S) represent the chemical potentials of receptors, sites, and bridges at a “standard” concentration (e.g., one molecule per pm’). k: and p! are simply constants because the internal energy of unattached states is not affected by the amount of cell-substrate separation. In contrast, pE(S) depends quite critically on the cell-substrate separation because changes in this distance result in stretching or compression of the bridges (see Fig. 2). If we think of a bridge as having a certain amount of elasticity, then we can characterize the S dependence of pE(S) in terms of the “resting” or unstressed length of the bridge, L ,

0

s =L

1

FIG.2. The chemical potential of cell-substrate bridges is dependent on the separation distance. I f S is too small. as in A, then the chemical potential is large because the bridges are compressed. If S is too large. as in C . then the chemical potential is large because the bridges are stretched. There is thus a certain value of the separation distance. .Y = 1.. such that the chemical potential of cell-substmte bridges is minimal ( B ) .

76

MICAH DEMBO AND GEORGE I. BELL

and a force constant for stretching of the bridge, K. In terms of these parameters, the chemical potential of a unit concentration of bridges is

Cl”,(s)=

&(L)

+ i K ( S - L)’

-k

...

(6)

In subsequent developments, it will be useful to define the idea of an equilibrium constant for cell-substrate bridging at a given separation distance. By direct analogy with the classical equilibrium constant of solution phase reactions, we will therefore introduce K ( S ) = exp [

( ~ y+

p! - pt(S)

+ kT)/kT]

(7a)

Inserting Eq. (6) into (7a) we see that to a good approximation K ( S ) = K , exp [ - ~ K ( S- L)’/kT]

(7b)

where KL = K f L ) represents the equilibrium constant for formation of an unstressed bridge. IV. THE REPULSIVE POTENTIAL

The repulsive potential, T(S),is defined as the mechanical work that must be done against nonspecific forces in order to bring a unit area of membrane from an infinite separation to a separation of S. If the repulsive force between unit areas of membrane and substrate is F(S), then F(S)

=

-rl(s)

or equivalently

r(s)= 7 m ) d z S

According to recent estimates (Bongrand and Bell, 1984), the forces resisting adhesion arise mainly from a combination of two effects: (1) electrostatic repulsion between negative chafges associated with the surfaces, and (2) the so-called steric stabilization effect.* The steric stabilization force is well known in the theory of colloid suspensions (Napper, 1977); it arises because cell membranes are coated by hydrated layers of long chain polymer molecules (i.e., the glycocalyx). In many systems the substrate could also be coated by a layer of absorbed polymers (e.g., serum proteins); this might enhance repulsive effects somewhat, but it is not an essential factor for the steric stabilization force. As the cell approaches *Since cell surfaces are generally negatively charged, it is possible to obtain an attractive electrostatic force to a positively charged substrate, e.g., one coated with polylysine. However, in this article we assume the net force to be repulsive.

77

THE THERMODYNAMICS OF CELL ADHESION

A

0

s< T FIG.3. The repulsive force between cell and substrate. If the separation distance is much larger than the average thickness of the glycocalyx ( A ) . then the repulsive force will be negligible. If the separation distance is less than the average thickness of the glycocalyx (B), then several factors will lead t o repulsive forces. There are ( 1 ) the osmotic tendency of solvent to return into a region of high polymer concentration. ( 2 ) the steric compression of the polymer chains, and ( 3 ) electrostatic repulsion between fixed charges on the substrate (if they are negative) and the glycocalyx.

the substrate, the polymer molecules of the glycocalyx are compressed between rigid surfaces, and in addition, water of hydration is squeezed out of the cell-substrate gap (see Fig. 3). The repulsive force results partly from the steric compression of the polymers and partly from the osmotic pressure caused by the tendency of solvent to return into the gap. An exact description of the complex interactions that go into determining the detailed form of electrostatic and steric stabilization forces are beyond the scope of this article. Instead, we will adopt a simple phenomenological expression for US) that has the correct qualitative behavior,

U S ) = -YS exp ( - S I T )

(8)

From a descriptive point of view, the parameter y measures the stiffness of the glycocalyx; the larger the value of y the harder it is to compress the polymers. The parameter T measures the thickness of the glycocalyx. If S is much greater than T , then there is a negligible compression of the glycocalyx; if S is much less than T , then the glycocalyx is compressed to the point where it fills the entire cell-substrate gap uniformly (i.e., the thickness of the gap is much smaller than the thickness of the glycocalyx). Typical values of T for cell-substrate interactions are in the range of SI S nrn.

78

MICAH DEMBO AND GEORGE I. BELL

Theoretical estimates of y can be derived by considering the statistical mechanics of chain molecules anchored to rigid surfaces. For typical cell parameters such estimates yield values of y on the order of lo-‘ to lo-’ dynes (Bongrand and Bell, 1984). The contribution of electrostatic repulsions to the value of y can also be estimated on the basis of theoretical argument. However, it seems that electrostatic effects do not significantly add to the value of y unless the glycocalyx is very sparse or the charge density is unusually high (Bongrand and Bell, 1984).

V.

MINIMIZING THE FREE ENERGY

Returning to the process of finding the minimum value of the free energy with respect to N b , A, and S, we should first note that none of these variables has a completely unrestricted range. For example, since the separation distance is obviously strictly positive, we must require that (94

OSSSCO

In a similar way, it is obvious that the number of cell-substrate bridges cannot increase if there are no free receptors left on the cell or if all the sites on the substrate are already occupied. Thus the number of bridges must fall in the range 0 s N , s min [N,,An,J

(9b)

Finally, the area of cell-substrate contact cannot increase if the cell has become completely flattened out to the point where there are no further reserves of “slack” membrane. As indicated by Fig. 4, if the surface-tovolume ratio of the cell is small, then the maximum possible amount of spreading will be small; if the surface-to-volume ratio is large, then the maximum contact area will approach ‘LAc. In any event, the contact area will, in general, be subject to a constraint of the form

0
s

S

A,,,

S

AAc

(9C)

Let N h , A,and be the equilibrium values of the three variables N h , A , and S. These values will correspond to the minimum value of G subject to the construint conditions of Eqs. (9u)-(9cJ. Because the constraint conditions are important, we are forced to approach the problem of computing N,,A, and 3 in two stages. In the first stage, we find values of N,, A , and S that minimize G without regard to the constraint conditions. If A,. A , and are values of the variables that correspond to such an unconstrained or “absolute” minimum, then in stage two we check to see whether or not these values are consistent with all the constraints [Eqs.

79

THE THERMODYNAMICS OF CELL ADHESION

C

FIG.4. Because of obvious geometrical conhtraints. the area of contact between a cell and a substrate cannot exceed a definite upper limit. If the surface area of the cell i b very large compared to the volume of cytoplasm ( A ) . then thih upper limit will be approximately one-half of the total cell surface area. As the ratio between surface area and cytoplasmic volume decreases (B), the maximum contact area a l s o decreases. As the ratio o f volume to surface area approaches the minimum value, namely. that of a sphere (C), the maximum contact area shrinks to zero.

(9a)-(9c)]. If the values are consistent, then we need proceed no further since the unconstrained minimum is also a permissible minimum. On the other hand, if the absolute minimum is not permissible, then, depending on which constraints are violated, we must set one or more of the variables equal to its constraint limit and attempt to minimize G only with respect to the remaining unknowns. Since G is a continuously differentiable function, f i b , A , and Lq must satisfy the three simultaneous equations dNbG= d,G = d,G

=

0

Inserting Eqs. (5a)-(5C) into Eq. (4b), and carrying out the appropriate differentiations, these three equations become d,,G

d,G

=

=

Nl,)/ALl- kT

-

/.LoI - kT Yn[(N,,

-

/.L:) -

kT Yn[(n,, -NJ/AL] - kT

+

&S)

+ kTYn(N,/A) +

n,,/.~:+ n,,XT tn[(n,,

-

-

X 7 = 0

N,,)/A] + k7N,,/A

80

MICAH DEMBO AND GEORGE I. BELL

Nh = n,,A

N,

(+I -

( 1 Ib)

?)y exp (-S/T)]/S2TK(S- L )

[A(S

=

exp [ - y exp ( - S l ~ ) / n , , k T S l

( I Ic)

To solve these three equations, we first insert Eq. ( I Ib) into Eq. ( I Ic) obtaining a single transcendental equation in S, 1 - exp

(

1

[ - y exp ( - S/.r)]/n,,kTS

=

[(s+ 7)y eXp ( - s / T ) ] / s ’ T K ( s

-

L)n,, (12)

This expression does not lend itself to exact solution; however, it can be shown that a permissible solution, 3, in the range L < 3 < m exists. Furthermore, this solution can easily be computed by standard numerical techniques (e.g., Newton’s iteration). It is also possible to obtain various approximate analytic expressions for 3. For example, if the repulsive potential is not too strong, i.e., if r(S)ln,,kT s l , then Eq. (12) becomes simply

s2- (L + kT/KT)S

-

(kT/K) = 0

Solving this quadratic we then conclude that

3

4 [ ( L + kT/K‘T)

+ v ( L + k T / K T ) 2 + 4kT/K]

Regardless of how one proceeds to solve Eq. (12), once the value of known, it can be inserted into Eqs. (1 la) and ( 1 1b) to obtain

A

= [Nrt

fib= N,, - e(3) - S($I “(3) + Ac/K(3)1/n&(3)

3 is

( I3a)

(13b)

where

is a nondimensional parameter that measures the ratio of repulsion to binding. Eqs. (13a) and (l3b) immediately show that the values of & and A will both be permissible if, and only if, two conditions are satisfied; first,

“3) N,,

(Ida)

and second,

( + V[N,.,

k(S) 3 4 N , -

Amnxnst - A,/K(S

1

- Amaxnst - A,/K(S)]’

(14b)

+ 4A,Nr,/K(~)]

THE THERMODYNAMICS OF CELL ADHESION

81

A thermodynamic system is said to exhibit a phase transition if the fundamental equations governing the behavior of the system are different for different ranges of the parameter values. According to this definition, plays Eqs. ( 14a) and ( 14b) indicate that the nondiniensional parameter t(,!?) a critical role in determining the phase of the thermodynamic system formed by a cell and a substrate. This is illustrated geometrically by considering the typical structure of a phase diagram in the plane defined by and N,., (see Fig. 5). As indicated by Fig. 5 . inequalities coordinates t(,!?) (144 and (14b) lead to division of the phase diagram into three regions. In region I inequality (l4a) is violated and (14b) is satisfied; in region 11 both inequalities are satisfied; and in region 111 inequality (14b) is violated but (14a) is satisfied. It is impossible to have a region where both of the inequalities are violated simultaneously. I n the first region of phase space, Eqs. (13a) and (13b) predict negative values of A and A,. In this case simple inspection shows that G is an increasing function of both A and N , , so that the permissible minimum must occur when N, = A = 0. Inspection also shows that the minimum

W

FIG. 5. Phase diagram of cell-to-substrate adhe\ion. The nondimensional parameter €(.TI [see Eq. ( 13c)] meawres the relative strength ofthe repulsive potential and bridge formarion 1i.e.. Inrge viilues o f < ( S ) correspond to large repulsive forces and/or weak binding]. For ii fixed value of <($, there are three possible phases o f cell-substrate attachment depending on the number of receptors per cell, N c , . If N , , i s too small. then stable binding will not occur (phase I behavior). Next. there i s a range o f N , , values (phase 11) for which binding i s st;ible. but the contact area i s submaximal ( A r' A",,,,). Finally. at large values of N , , , the conlact iirea will be fixed at its upper limit (phase Ill).

82

MICAH DEMBO AND GEORGE I. BELL

value of G is independent of S, so that is indeterminate in the region 0 s 3 s m. Obviously from the nature of this class of solutions, region 1 is the portion of phase space in which stable cell-substrate adhesion cannot occur. In the second region of phase space the values of f i b . A, and j. are all consistent with the constraint conditions. In this case the absolute minimum of G and the permissible minimum of G coincide, so that N b = f i b , = A, and 3 = 3, where f i b , A , and are given by Eqs. (13a), (13b), and ( I 2), respectively. In region 111 of phase space Eq. (13b) predicts a value of A greater than A,,,. In this region G can be minimized by using the constraint condition = A,,, in place of Eq. ( I Ib) and then solving Eqs. ( 1 la) and ( I Ic) simultaneously to determine fib and 3. Once again, it is not possible to obtain explicit expressions for x b and 3 although it can be shown that a unique solution exists and that it is quite easy to compute this solution by standard numerical methods. VI.

RESULTS

Table I presents a list of the various physical parameters that characterize cell-substrate adhesion according to our model. This table also presents some preliminary estimates of the parameters; these estimates are

TABLE I CONIKOL PARAMETERS Parameter Number of receptors per cell Density of sites on the substrate Total surface area of cell Maximum possible contact area Equilibrium constant for formation of unstressed bridge Length of unstressed bridge Force constant for stretching of cell-substrate bridges Compressibility coefficient of the glycocalyx Thickness coefficient of the glycocalyx

Symbol

Estimated value"

2 x lo-' pm dynes/pm

Y 7

10 6dynes lo-' p m

"In general, the numbers given are appropriate for a small cell such as a lymphocyte and for noncovalent bonds of the type produced by antigen-antibody interactions. See Bell et a/. (1984) for a detailed discussion of the arguments and measurements that lead to these estimates.

83

THE THERMODYNAMICS OF CELL ADHESION

subject to considerable uncertainty, but they are useful for purposes of orientation when carrying out illustrative computations. At least in principle, all the parameters in Table I are fixed by nature or tire under the control of a sufficiently omnipotent experimentalist. For any choice of such “control parameters,” the most probable configuration of a cell adhering to a substrate then follows from the laws of thermodynamics and can be computed using the method outlined in the previous section. The variables that characterize the detailed configuration of a cell interacting with a substrate include the area of contact, the cell-substrate separation distance, the total number of cell-substrate bridges, the density of bridges in the contact area, the density of unattached receptors outside the contact area, the density of unattached sites in (or out) of the contact area, etc. In general one can regard any quantity that changes during the interaction of a cell and a substrate as a “configuration variable” of the system. Suppose that we make observations of a configuration variable (e.g., the area of contact A) at various values of one of the control parameters (all other control parameters being fixed). In such an experiment our model predicts that there will be certain critical values of the control parameter at which the configuration variable will undergo a sharp threshold type transition due to the crossing of a phase boundary. Between such critical points, there will be regions where the phase remains unchanged, and there is smooth variation of the configuration variable. To illustrate these general considerations Fig. 6 shows the predicted behavior of the three configuration variables N,,, and as the density of sites on the substrate, n,,,is increased. At low values of I ] , , , bridge formation between cell and substrate is not sufficient to overcome the repulsive interaction. Thus for this range of parameter values, stable binding cannot occur, and the system is in phase I (i.e., = N,, = 0 and s = s).As n,, continues to increase, ultimately a critical point is reached at which the system leaves region 1 and enters region II of phase space (see the phase diagram, Fig. 5 ) . As soon as the system enters phase 11, the cell-substrate separation distance suddenly drops to a finite value and the contact area and number of cell-substrate bridges start to increase. A moderately surprising aspect of Fig. 6 is that once the system enters region 11, the increase in contact area is very rapid compared to the rate of increase in the number of cell-substrate bridges. Thus after only a small percentage of the cell’s receptors are involved in bridging, the cell has already become completely spread out into its maximum contact configuration. Of course, the point at which the cell can no longer continue to spread corresponds to the phase It-phase 111 boundary. Accordingly, al-

x,

04

MICAH DEMBO AND GEORGE I. BELL

0.5 -

4

0. 10-1

I

,

, 1 1 1 4 1

10’ n,t (103 sites/Fm2

I00

I

I

I

4

I 1 I I I

I02

FIG.6. Variation of configurationvariables as the density of adhesive sites on the substrate, n,,, is increased. Control variables other than n,, were fixed at the values given in Table 1. For convenience of representation on a single scale, the configuration variables are expressed in nondimensional form: N, = Nb,Nr,,A AIA,,.,,, and SIL. According to our model, there are three critical values for n,, in the sort of computation shown. The first represents the onset of stable adhesion, the second represents the onset of maximum contact, and the third represents the loss of maximum contact at the start of the “rounding up” phase. See the text for a more detailed discussion of the physical significance of these phase transitions.

most as soon as the system has entered phase I1 from phase I, it reaches a new critical point and leaves phase 11, entering phase 111. Once the system has entered phase 111, further increases in n,, gradually cause formation of more cell-substrate bridges, which in turn pull the cell closer to the substrate. Finally, as n,, gets larger, all the receptors on the cell become incorporated into bridges (N,,/N,, 1). At this point, one might think that the story would be pretty much over; however, if we continue to raise the density of sites on the surface still more, a very surprising thing happens. At very high levels of nst, a third critical point is reached at which the system leaves phase 111 and reenters phase 11. In other words, an increase in the number of adhesive sites on the surface causes the cell to lose its maximally spread configuration and to begin a process of “rounding up”! After a rapid initial decline in contact area, the rounding up process continues at a slower and slower pace as additional increases in a,, proceed.

85

THE THERMODYNAMICS OF CELL ADHESION

Ultimately the contact area approaches an asymptotic limit at a value 5060% less than A,,,,. In addition to changes in the contact area, the rounding up process is characterized by a monotone decline in the cell-substrate separation distance. As with the decline in contact area, the decline in gradually slows and approaches a limiting value as n,, + =. Finally, in contrast to the behavior of and 3. the number of cell-substrate bridges remains at a saturating level throughout the rounding up phase (i.e., all receptors are incorporated into bridges). Note that for very large values of n,,, Eqs. (13c) and (13b) predict 5 + A,r/n,,(kT)and A + N,,kT/I’. The unexpected third critical point in Fig. 6 demolishes the naive notion that changes in cell-substrate contact are more or less proportional to changes in cell-substrate binding. Clearly, a more sophisticated assessment of the balance of forces governing adhesion is needed. To provide such an explanation, we first note that the cell is torn between the need to minimize repulsive interactions (i.e., rounding up) and the need to maximize attractive interactions (i-e., bridge formation). If the site de sity on the surface is low, then many new bridges will be formed for e. ch new increment in contact area. Thus there is a large driving force for increasing contact, and the cell spreads to a maximal extent. In contrast, if the site density on the substrate is sufficiently large, then all the receptors on the cell can be saturated without the need for maximum contact. At the margin, this means that the driving force for spreading is decreased, and the cell starts to round up under the influence of the repulsive forces. The rounding up process slows down and stops at very high levels of n,, because the free energy decrease caused by reducing repulsive interactions is balanced by the free energy increase required to compress the cell-substrate bridges into a small contact area. In addition to variations in the number of sites on the substrate as in Fig. 6, we could equally well consider the changes in configuration variables caused by such things as the number of receptors on the cell ( N , , ) , the compressibility coefficient of the glycocalyx (y), or the binding constant for unstrained bridge formation (KL). In each of these cases, however, the results of computation are in line with what one would expect on the basis of the phase diagram in Fig. 5; i.e., one passes in orderly progression from one phase to the next as the parameter is varied, only two critical points occur, and there are no additional surprises. As an illustration of this kind of computation, Fig. 7 shows the changes in configuration caused by increasing the repulsion between the cell and substrate (i.e., variation of Y). If it is possible to manipulate two or more of the control variables in Table 1 simultaneously, then we can achieve an additional level of complexity in the analysis of adhesion data. To illustrate, suppose that the

s

f

86

MICAH DEMBO AND GEORGE 1. BELL

y (microdynes)

FIG.7. Variation of configuration variables as the compressibility coefficient of the glycocalyx, y . is increased. The configuration variables are expressed in the same nondimensional form as in Fig. 6. In contrast to the computation shown in Fig. 6, only two critical points occur as the value of y is increased. These correspond to the transition from phase I11 to phase I1 at low values of y (see the expanded scale insert on the right-hand side of the figure) and to the transition from phase I1 to phase I at the point where cell-substrate contact is finally broken.

simulation in Fig. 6 is carried out at several different values of N,, (the number of receptors per cell). For each experiment we then determine the three critical values of n,, and plot their values versus the value of Nrt.The result of this procedure is nothing other than the phase diagram of adhesion in the N& - n,, plane (see Fig. 8). A computation such as that shown in Fig. 6 is equivalent to traveling upward along a path parallel to the y axis in Fig. 8. Figure 8 shows that if the value of N,, is greater than about I .2 x lo5 receptors per cell, then such a vertical path will lead in stages from region I to region I1 to region 111 and then back to region I1 (this is the behavior observed in Fig. 6). If N,, is less than 1.2 x 10' receptors per cell, then a vertical line will not intersect region 111 of the phase diagram and maximum contact will never be achieved; an example of such a computation is shown in Fig. 9. Finally, if N , is larger than about 7 x 10' receptors per cell (see the insert in Fig. 8), then vertical lines in the phase diagram will not intersect the upper boundary of region 111. This means that if N , is sufficiently large, a decline in contact area as n,, will no longer occur.

-

87

THE THERMODYNAMICS OF CELL ADHESION

4

3

-

N

E

i c0) m"

-

2

Phase

0

U

A < Amox

c UI

/

/

c

Phose ffi

I

i.0

0 N,, (105receptors/cetl)

FK;.8 . Phase diagram of cell-to-wbstrate adhesion in the plane formed by control parameters n,, and Nr,. The various regions of the phase diagram have the same significance as in Fig. 5. The insert shows the phase 11-phase 111 separation line for large values of the variables.

VII.

DISCUSSION

The simple model of cell-substrate adhesion tha- we have presented and analyzed in this paper can be viewed as a special case of a very much larger class of related models [see Bell et a / . (1984) for a more complete discussion]. Nevertheless this model demonstrates the power of thermodynamics in its being able to integrate a large number of complicated variables into an effective computational and predictive tool. The model we have discussed has had to incorporate notions about the receptors in the cell (laterally mobile, fixed number per cell) and the sites on the substrate (laterally immobile, fixed surface density); notions about the cell-

88

MICAH DEMBO AND GEORGE I. BELL

1.0 -

0.5

1

0 10-1

10‘ ns+(103sites/Fm2)

100

I

I

I

, 1 1 1 1

I02

FIG.9. Variation of configuration variables as the density of adhesive sites on the substrate, is increased. Conditions are identical to Fig. 6 except that N,, is reduced by a factor of 2. As in Fig. 6, a decline in contact area still occurs as n,, -+ m. However, in contrast to Fig. 6 , even at the optimum value of n,,, the contact area never reaches A,,,.

n,,,

substrate bridges (length, spring constant, binding constant); notions about the repulsive forces between the cell and substrate (compressibility of the glycocalyx, thickness of the glycocalyx); and finally, notions about the purely geometrical parameters of adhesion (uniform separation distance in the contact area, maximum contact constraint). In order to achieve the power and generality of a thermodynamic approach, a price must be paid in terms of certain definite limitations. For example, our model cannot account for the possibility that cells might actively “grip” the substrate or that there might be rapid temporal synthesis and degradation of receptors. In addition, thermodynamic models say nothing about the kinetic process involved in reaching adhesive equilibrium. However much one wishes to make of these “nonequilibrium” factors, they certainly do not detract from the need to treat the basic thermodynamics as well. The most striking single prediction of our model of adhesion is the existence of various critical points characterizing such things as the onset of stable adhesion as the density of sites on the substrate increases (see

THE THERMODYNAMICS OF CELL ADHESION

89

Fig. 8). Solid experimental confirmation of the existence of such critical points is not easy to find although threshold phenomena have been reported in a few systems (Weigel et al.. 1979). Perhaps now that a theoretical basis for such phenomena is known, experimentalists will be alerted to look for their existence and study their behavior in a systematic way.

ACKNOWLEDGMENTS Work was performed under the auspices of the U.S. Department of Energy. Work was also supported by NIH Grants K04-AKM566and ROI-AI21002.

REFERENCES Bell, G. I., Dembo, M., and Bongrand, P. (1984). Cell adhesion: Competition between nonspecific repulsion and specific bonding. Eioplrys. J . 45, 1051-1064. Bongrand, P., and Bell, G.1. (1984). Cell-cell adhesion: Parameters and possible mechanisms. In “Cell Surface Dynamics: Concepts and Models” (A. Perelson, C. DeLisi, and F. Wiegel eds.), Chapter 14. Dekker, New York. Napper. D. H. (1977). Steric stabilization. J . Colloid Interfuce Sci. 58, 390407. Weigel, P. H., Schnaar, R. L., Kahleuschmidt. M. S., Schmell. E.. Lee, R. T., Lee, Y. C.. and Roseman, S. (1979). Adhesion of hepatocytes to immobilized sugars: A threshold phenomenon. J . Eiol. Chem. 254, 10830-10838.

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('IJRKEN'I TOI'I('S IN MI(MI3RANhS A N I ) 'I'HANSPOKI'. V O I LIME ?Y

Rotational and Lateral Diffusion of Membrane Proteins and Lipids: Phenomena and Function MICHAEL EDIDIN

I. INTRODUCTION Motion of cell membrane components occurs over a wide range of timc and distance scales. This article covers the rotational and translational diffusion of membrane lipids and proteins. motions over a part of the time and distance scales. Small scale motions, in the range of tens of angstroms and with characteristic times of 10" to 10" sec-I will not be discussed, nor will very slow motions, usually directed and metabolically driven, on distance scales of hundreds of micrometers and time scales that may range up to days. The short-range and rapid motions, especially of membrane lipids. are summed up in the terms "viscosity" or "fluidity," and this subject has been well reviewed by Quinn (1981). The slow motions, over long times and distances, are the province of work on cell locomotion, membrane flow, and recycling. Here again, reviews of a very large literature, and one outside our present scope (Abcrcrombie, 1980; Steinman ef a / . , 1983; Brown et ul., 19831, can only be mentioned. We will be concerned with rotational motions in the range of IO'to 10' sec-l and with lateral movements in the range of 10-l' to lo-' cm' sec I. Methods for measuring diffusion will be described and then some of the results obtained with these methods will be covered. Finally, an attempt will be made to discuss biological functions of diffusion in membranes. The first two approaches might be said to be those of the physicist, to the extent that they describe some properties of membrane components without ascribing function to them. The last approach is very much that ~

91 C'upyright '1 I Y X 7 hy Academic PIC,\. Inc. All right3 n1 rcproduution in any form rvwrved

92

MICHAEL EDlDlN

of the biologist who, from the time of Aristotle, has enquired about final causes of things seen in nature. II. PROBES FOR DIFFUSION

All the techniques for measuring diffusion depend upon some aspects of the spectra of selected probes. These probes may be roughly divided into optical probes (mainly fluorescent labels, but also endogenous chromophores with characteristic absorption spectra) and magnetic resonance probes [labels such as 'H (deuterium) for nuclear magnetic resonance (NMR) and stabilized paramagnetic labels, notably nitroxides, for electron spin resonance (ESR)]. The two sets of techniques cover all the range of time scales summarized above. They differ in their sensitivities and in the degree to which they perturb the membrane being measured. Fluorescence measurements are the most sensitive, NMR measurements the least. NMR is the least perturbing, depending upon the molecule labeled, ESR or fluorescence may be the most perturbing. Spectroscopic methods for membranes may be compared in a published collection of reviews (Grell, 1981). Magnetic resonance methods always probe populations of cells or synthetic membranes. They give averages for the population. Fluorescence methods may be used to probe populations, but, with care, they can be adapted for microscopy and hence can be used to analyze single cells, or even particular regions of cells, for example, the brush border of epithelial cells or the ruffled leading edge of moving fibroblasts. Signals from fluorescence probes localized to portions of a cell membrane may then be associated with structural or functional properties of that region of the membrane. We also note in a general way that both magnetic resonance and optical probes may be used to measure motions over a range of time scales. However, the ease of applying a particular class of probes to a particular diffusion measurement varies considerably. For example, while the lifetime of the excited state of fluorescence probes is long enough to be affected by rotation of lipid and lipid-like molecules, it is too short to effectively measure rotation of membrane proteins. Instead, spectral properties of the longer lived triplet state must be used, with consequent loss in sensitivity. Spin labels also report on times shorter than those for rotational diffusion of membrane proteins or lipids. However, ESR techniques for measuring protein rotational diffusion are now available. They involve measurement of details in spectra obtained under unusual conditions in which spectral intensities are not linearly dependent upon intensity of the incident microwave field (Hyde, 1981).

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

111.

93

ROTATIONAL DIFFUSION

A. Methods for Measuring Rotational Diffuslon

I . OPTICALMETHODS The general approach to measurement of rotational diffusion in a population of molecules requires selection of molecules with a definite orientation relative to the coordinates of the experimental system and measurement of the time taken for molecules of the selected population to reorient and randomize. In general this is readily done by defining molecules with a flash of plane-polarized light and measuring the anisotropy of the system in terms of polarization of fluorescence or in terms of anisotropy of absorption. Measurements of anisotropy versus time give values for rotational relaxation, which are proportional to the rotational diffusion coefficient. Cherry (1979) discusses the various constants of proportionality relating rotational relaxation and rotational correlation times, d, or T ~ to , D,,,for isotropic and anisotropic rotation. Rotation of proteins and lipids around a membrane normal is anisotropic, and Cherry The deviation of initial suggests the appropriate relationship is d, = l/D,.c,t. anisotropy from the theoretical maximum gives some indication of motions of the probe or of intramolecular motions which reorient selected molecules during the time of the selecting flash. Residual, persistent anisotropy gives some measure of restriction to rotational diffusion, for example, due to formation of large aggregates of molecules. Work on rotation of probes has branched into work on small, lipidsoluble molecules whose moderately long lifetimes, tens of nanoseconds, make them useful as probes of lipid order and “microviscosity” (reviewed by Quinn, 1981; Edidin, 1981). The theory for behavior of the most studied of these probes, the rod-shaped molecule diphenylhexatriene, has been thoroughly developed by Hare (1983), whose paper includes an extensive bibliography of work on this probe, and on polarization of fluorescence in general. As noted above, optical techniques for measuring the relatively slow anisotropic rotation of membrane proteins (microsecond time scale) require long-lived probes. These may be intrinsic chromophores, for example, intermediates in the bleaching of rhodopsin or heme groups of cytochromes, or exogenous chromophores which can be excited to yield longlived triplets. Rotation of labeled molecules is usually followed in terms of dichroism (Cherry, 1978; Kawato and Kinosita, 1983) or of polarization of luminescence or fluorescence (Jovin rt al., 1981). Both these approaches are quite insensitive compared to fluorescence methods. Methods of ground state depletion for measuring rotational diffusion have been developed. In one method fluorescence is bleached with polarized light and

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MICHAEL EDlDlN

rotation of labeled probes is followed in terms of fluorescence recovery (observed with an attenuated beam with polarization parallel to the bleaching beam) or fluorescence depletion (polarization of measuring beam perpendicular to that of the bleaching beam) (Smith et af., 1981). This approach has been used only to measure rotation of lipid-soluble dyes, but it could be readily applied to measurement of slower rotations of membrane proteins. Measurements of lipid-soluble dyes in synthetic bilayers gave excellent agreement with theory and provide a firm foundation for work on native membranes. The polarized fluorescence photobleaching method is also more readily applied to intact cells than methods depending upon generation of triplet states, since photobleaching measurements may be made in the presence of oxygen. A second, similar, approach, depending on the long lifetime of the triplet state has been applied to membrane proteins covalently labeled with eosine or other triplet-formers (Johnson and Garland, 198 I). After determination of steady-state fluorescence levels, about 20-30% of the probe molecules are excited to the triplet state by a microsecond flash of polarized laser light. The laser is then attenuated, and recovery of fluorescence intensity is measured with the polarized beam. When the ground state has been fully repopulated, excitation to the triplet state is repeated, with the polarization of the exciting beam rotated by 90". Anisotropy as a function of time, r, is calculated in the usual way as Y,

=

( I 11 -

m z + ~2 1 , )~

(1)

The recovery of polarized fluorescence depletion itself is affected by rotational diffusion of triplet state labels, by the return of triplets to the ground state, and by lateral diffusion of ground state molecules into the spot excited by the laser beam. Return to ground state and lateral diffusion should make the same contribution to both Ill and I,. Hence, r, ought to be solely a function of the rotation of triplet-labeled molecules. Tradeoffs between fluorescence yield and ease of excitation to the triplet state determine the sensitivity in a particular application, but the method is up to lo9 times more sensitive than luminescence or absorbance methods. Probes useful for the method are discussed by Johnson and Garland (1982). Though not yet widely used, the fluorescence depletion method promises to make rotational diffusion measurements available from any laserlmicroscope configured for the determination of lateral diffusion coefficients by fluorescence photobleaching and recovery (see Section IV,B,2,c). 2. SATURATION-TRANSFER ESR (S-T ESR) Electron spin resonance methods for membrane components are generally used to determine the amplitudes and angles of displacement of

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

95

probes from a membrane normal. The spectra obtained by conventional ESR techniques depend upon motional averaging which is sensitive to molecular movements in the range lo-' to 10-l' sec, that is to motions that occur within the time scale for return of spin states to equilibrium in the x-y plane of the nitroxide label: this is defined by a relaxation time T.. However, if saturating levels of microwavc power are used, then relaxation of the nonequilibrium population of spins occurs by exchange of energy between the population of spins and other degrees of freedom, that is by so-called spin-lattice relaxation, characterized by a time TI.The value of TI is approximately 300 times less than 7?.TI is affected by slower molecular motions than affect T2 and is particularly affected by rotation of molecules around the membrane normal. Thus S-T ESR spectra can be used to detect slow rotations, with correlation times in the range of hundreds of microseconds (for reviews see Marsh, 1981; Hyde, 1981). The method was developed for isotropic rotation, and the first applications to anisotropic rotation of membrane proteins used isotropically tumbling models to aid in interpretation of the spectra. However, there has been progress in the analysis of anisotropic models (Gaffney, 1979; Delmelle et a / . , 1980) in the use of 3s-GHz microwave probes (Q-band) which yield higher resolution spectra than do thc usual 9.5-GHz (X-band) probes (Johnson and Hyde, 1981; Johnson ut d.,1982).

B. Rotational Diffusion of Membrane Proteins Rotational diffusion of a few very well-characterized and abundant membrane proteins has been studied with a variety of techniques. Rotation of erythrocyte band 3, bacterio- and vertebrate rhodopsin, calcium ATPase of sarcoplasmic reticulum, cytochromes and cytochrome oxidases, and acetylcholine receptor has been measured in native and synthetic membranes. These measurements yielded information on protein-protein association as a function of membrane protein concentration, on coupling of integral membrane proteins with the cell cytoskeleton. and on coupling of enzymatically interacting components, for example, in redox reactions. I will review measurements of rotational diffusion for only three proteins. Rotational diffusion of other membrane proteins is summarized in Table I1 (see Section 111,B,4).

I.

ERYTHROCYTE

BAND 3

Cherry and co-workers ( 1976) prepared eosine-labeled erythrocyte ghosts in which band 3 was the predominant labeled species. The anisotropy decay indicated the existence of two populations of band 3 molecules, one moving so slowly that its rotation could not be detected in

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MICHAEL EDlDlN

the time of the experiment (-4 msec). Improved techniques led to an as 160 psec for the rapidly rotating component and beestimate of tween 3400 and 6460 psec for the slowly rotating form. The rapidly rotating form is a dimer, since chemical cross-linking of band 3 to form dimers did not change the rotational correlation time (Nigg and Cherry, 1979). Slowly diffusing oligomers of band 3 form as a function of temperature, and the highly aggregated forms of the protein predominate below 25°C. Association of band 3 molecules as a function of temperature probably reflects temperature-dependent changes in the organization of membrane lipids. Membranes whose cholesterol content is varied by incubation with liposomes contain varying proportions of dimers and higher oligomers (Muhlebach and Cherry 1982). However, this effect is seen only in membranes digested with trypsin, which cleaves the C-terminal portion of band 3 and hence stops its interactions with cytoskeletal proteins. Despite this modification, the model may form useful basis for consideration of the effects of temperature on protein-protein association in native membranes. The phosphorescence anisotropy of labeled band 3 reports a rotational correlation time of 300 Ksec (Austin et al., 1979), fairly close to that reported by Cherry and co-workers. A slowly rotating labeled component is also resolved in these experiments, with relaxation times ranging from 2.2 msec at 4°C to 0.4 msec at 30°C. Here too, the proportion of slowly rotating oligomer increases with lowered temperature. Rotational diffusion of the band 3 dimer is probably restricted by the interaction of its C-terminal cytoplasmic portion with erythrocyte cytoskeletal proteins. Nigg and Cherry (1980) found that trypsin digestion of band 3 to cleave its cytoplasmic portion, or salt extraction of membranes to remove cytoskeletal proteins, enhanced the decay of absorbance anisotropy. As in earlier investigations, the decay curve gave evidence for two populations of band 3 molecules, one rotating with a correlation time of about 100-200 psec (over a temperature range from 45 t o 12°C) and the other rotating with a correlation time in the range 2-9 msec. Trypsinization or salt extraction did not significantly affect the rotational correlation times, but it did change the proportion of slowly rotating and rapidly rotating molecules. In untreated membranes 24% of band 3 molecules were rapidly rotating and 52% could be assigned to the slowly rotating population. These proportions are reversed in lightly trypsinized ghosts and are 40 and 35% in salt-extracted ghosts. The data are interpreted as showing an effect of the cytoskeleton on band 3 rotation, since viscous drag on the cytoplasmic tail of band 3 should be insignificant compared to the drag on the intrabilayer portion of the molecule. This interpretation is reinforced by the results with salt extraction, though the effects of high plus low salt extraction are not as marked as the effects of trypsin digestion.

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The stoichiometry of linkage is complex. Only 20% of band 3 dimers can be linked directly to ankyrin (since t h e ratio of ankyrin to band 3 dimer is about 1 5 ) . but 405%of the total band 3 may be mobilized by trypsinization or salt extraction. Higher oligomers of band 3 may be linked to single ankyrins, or perhaps other peripheral membrane proteins, for example. bands 4.1 or 4.2 which are linked directly to band 3. Glycophorin may also associate with band 3 complexes. Antibodies to glycophorin inhibit rotational diffusion of band 3 (Nigg L’t a / . , 1980). Band 3 rotation in synthetic [dimyristoylphosphatidylcholine(DML)] membranes has been measured with S-T ESR by Sakaki and co-workers (1982). They detected a single species of labeled band 3 with D,,,, =4 X lo4 sec-’, which gives a correlation time, l/D,,,,,of about 25 Fsec, in contrast to the value of about I50 Fsec found by Cherry and co-workers. Addition of a cocktail of cytoskeletal proteins increased the rotational correlation time of band 3 about 2-fold. Some of the discrepancies between these results and those of Nigg and Cherry is probably due to differences in the assumptions made in evaluating and interpreting the experimental spectra. Differences between the viscosity of red cell membrane lipids and the viscosity of DML vesicles may also give rise to the observed differences in rotational correlation times. Sakaki et al. (1982) estimate the viscosity of DML vesicles, above the phase transition, as about 2 poise (P).This value gives good agreement between the value predicted for lateral diffusion of band 3 in DML, 2.1 x lo-’ cm’ sec-’ (Saffman and Delbruck, 19751, and the value measured, 1.6 x lo-’ cm’ sec-’ (Chang et al., 1981). However, it is lower than the viscosity estimated for DML from the rotational diffusion of bacteriorhodopsin in DML, 3.7 k I .3 P, at a ratio of lipid to protein of 168:I (Cherry and Godfrey. 1981; Heyn et al., 1981). Either value for viscosity of DML implies red cell membrane viscosities of about 10 P, a high, but not unreasonably high value. Even this high value must be postulated only if there are no other sources for the discrepancies between flash photolysis and S-T ESR measurements. This is unlikely. 2.

VERTEBRATE AND BACTEKIORHODOPSIN

The rotation of rhodopsin was first observed by Cone (1972) who bleached the intrinsic chromophore of vertebrate rhodopsin with a flash of polarized light. The rotational relaxation time, 20 psec, and the estimate of membrane lipid viscosity, 0.7-6.0 P, remain benchmarks for other work on diffusion in bilayers. The theory and methodology of these measurements have been reviewed (Ah1 and Cone, 1982). Rhodopsin of two other visual systems appears to be organized differ-

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MICHAEL EDlDlN

ently than vertebrate rhodopsin. The rotational relaxation time for squid rhodopsin is around 200 psec, suggesting that the protein exists in the photoreceptor membrane as oligomers, rather than as monomers or dimers (Pasternak et al., 1981). No decay of flash-induced dichroism of rhodopsin is seen in 0.75%formaldehyde-fixed crayfish rhabdomeres (Goldsmith and Wehner, 1977) though rotation of vertebrate rhodopsin is not affected by fixation with as much as 5% formaldehyde (Cone, 1972; Brown, 1972). Bacteriorhodopsin forms a crystalline lattice and is immobile in native membranes (for review see Henderson, 1977), but rotates freely, with a rotational relaxation time close to that of vertebrate rhodopsin monomers, when reconstituted into phospholipid vesicles at 1ipid:protein ratios similar to those in native disc membranes (around 0.5:l.O w/w) (Heyn et al., 1977; Cherry and Godfrey, 1981; Wey et a / . , 1979) Some aggregates form at high 1ipid:protein ratios and these are detected in deviations of the experimental anisotropy decay curve from the theoretical curve for a single diffusing species. At low 1ipid:protein ratios it is suggested that the solute (protein) concentration affects membrane viscosity (Cherry et al., 1977; Peters and Cherry, 1982). Thus at an L:P ratio of 168: I the apparent viscosity of DML above its phase transition is 3.5 P. At an L:P ratio of 1 1 1: 1 almost all of the bacteriorhodopsin is still present as monomer, but it reports an apparent viscosity of 8.6 P. Extensive S-T ESR studies of phospholipid vesicles containing bacteriorhodopsin give similar effects which are interpreted in terms of aggregate formation, rather than protein effects on lipid viscosity. Bacteriorhodopsin in DML vesicles reconstituted with an L:P ratio of 240:l rotated with correlation times ranging from 10 to 60 psec (Kusumi et al., 1980). Pure phospholipid vesicles of other chain lengths also showed some effect of protein concentration on rotational correlation time (Kusumi and Hyde, 1982). These data are summarized in Table 1. All the values given are for phospholipids above the phase transition. The temperature of this transition is lowered by the added protein. Apparently, protein-rich fluid regions segregate from the rest of the phospholipids. Kusumi and Hyde also measured rotational diffusion of vertebrate rhodopsin in intact discs. They find a rotational correlation time of 20 psec, in agreement with the value from Cone’s (1972) photodichroism experiment. Hence, they argue that the value of approximately 10 psec. obtained as the limit when diluting bacteriorhodopsin into lipid vesicles, must represent the true value for a rhodopsin monomer and that rhodopsin in discs is aggregated, presumably into dimers. This argument neglects possible effects of protein concentration on the apparent viscosity of lipid bilayers, which Cherry and co-workers believe to be important

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99

TABLE I B.4CI'ERIOKHOI)OI'SIN AS A FLINCI'ION OV LIPOSOMI.. COMPOSITION A N D LIPID '1-1) PROTEIN ( w l w ) R,vrio"

~0'l'AI'ION.Al. DIFFUSION OF

Phospholipidh DLPC (C,?PC) DMPC (C,, PC) DPPC tC,, PC) DSPC (C,"PC)

Rotational correlalion time ( psec) L:P 15O:l L:P 5 0 : I 27 I0 8 I2

30 17 17 24

"From Kusumi and Hyde (1982).

hDLPC, dilinoleoylphosphatidycholine: IIMPC. dimyiistoylphosphatidylcholine: DPPC. dipalmitoylphosphatidylcholine: DSPC. distearoylpho.lphatidylcholine.

in membranes [this view is summarized by Cherry (1979) and by Peters and Cherry ( 198211. 3. CYTOCHROMES P-450

The degree of aggregation and the molecular associations of microsomal and mitochondria1 cytochromes and their associated enzymes have been estimated from measurements of rotational diffusion. The most extensive measurements have been made on species of cytochrorne P-450 of the microsomal rnonooxygenase system. This is a terminal electron acceptor receiving electrons from NADPH-cytochrome P-450 reductase and from cytochrome h, (for review, see White and Coon, 1980). There are 20-30 cytochrome P-450 molecules per molecule of reductase, and while P-450 appears to be deeply embedded in the lipid bilayer (De Pierre and Ernster, 1977). the reductase is anchored to the bilayer only by a hydrophobic tail. The stoichiometry of P-450 associations and its interaction with substrate may be inferred from measurements of rotational diffusion which are readily accomplished by several methods, notably flash photolysis of the cytochrome heme group. The results, though restricted to only a few species and to only a few types of drug-induced cytochromes P-450, give a general indication of the molecules' functional associations in microsomal membranes. Flash photolysis measurements were first performed on intact microsomes and indicated slow rotation of P-450 (Richter et al., 1979). From one-half to two-thirds of the total cytochrome P-450 appeared to be immobile in rat or rabbit liver microsomes, and the rotational correlation time for the mobile fraction was estimated at 100-300 Fsec. Later work refined these observations (Kawato pt d.,1982a). Rotational correlation

100

MICHAEL EDlDlN

times for the mobile population of molecules were estimated as 63 2 6 psec at 37°C and 120 36 psec at 20°C. Lowering the temperature increased the fraction of molecules immobile on a millisecond time scale from 47% at 37°C to 60% at 20°C to 95% at 5°C. This suggests that cytochrome P-450 molecules aggregate, even at physiological temperatures, and that this aggregation can be enhanced at lower temperatures. Other factors may also cause aggregation, since cytochrome P-450 was 100% immobile in some freshly isolated microsome preparations. The weight ratio of lipid to protein in microsomes is about 0.4. Behavior of purified cytochrome P-450 reconstituted into mixed phospholipid liposomes was studied in liposomes containing from 30:l to 1:1 lipid to protein (w/w). The fraction of immobile cytochrome P-450 increased with increasing protein content of the liposomes. It also increased when the liposomes were cooled. The results with liposomes bear out the observations on intact microsomes and indicate that cytochrome P-450 molecules are aggregated in functional microsomes. A study of exogenously labeled cytochrome P-450 in liposomes also finds about half the molecules immobile on a millisecond time scale, and suggests that the molecules free to rotate are aggregated as hexamers (Greinert et al., 1982a). A detailed analysis of this system led to a model in which each P-450 hexamer interacts with one P-450 reductase molecule and in which substrate affects the depth at which the hexamer lies in the lipid bilayer (Greinert et id., 1982b). The aggregation of the cytochrome molecules is relieved by reconstitution with equimolar amounts of NADPH-cytochrome P-450 reductase. Under these conditions all P-450 molecules are mobile, with a rotational correlation time of about 40 psec. On the other hand, in vesicles with a P-450: reductase ratio of 5: I there is little effect on the immobile fraction of P-450. Thus the enzyme appears to interact with the cytochrome in a 1: I complex. Aggregated cytochromes are mobilized when this interaction occurs (Gut et al., 1982). Antibody to the reductase immobilizes cytochrome P-450 in vesicles with high protein content (L:P I : I ) containing equimolar amounts of cytochrome and reductase (Gut et al., 1983). This result then reinforces the view that the cytochrome and the reductase form I : 1 complexes. It implies that electron transfer to cytochrome P450 occurs by two different mechanisms: complex formation and lateral diffusion. Saturation-transfer ESR studies of spin-labeled cytochrome P-450 in liposomes (Schwartz et al., 1982a) and in microsomes (Schwartz et d., 1982b) report slower correlation times than found by flash photolysis. In liposomes with an L:P ratio around 12: I P-450 appears to be in clusters of 8-12 molecules, while flash photolysis experiment (Kawato et al., 1982a) suggest that there are no more than dimers formed in liposomes of similar

*

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

101

composition. The S-T ESR data suggest that similar size complexes form in native microsomes as well, and there are no free P-450molecules, again in contrast to the flash photolysis data. Despite these differences we oughtto stress that both sets of measurements indicate clustering of the cytochrome molecules in native microsomes.

TABLE I I ROTATIONAL DIFFLISION OF SOME MEMBRANE PROTEINS"

Protein H-2 antigens in

plasma membrane Acetylcholine receptor from 7orperlo Epidermal growth factor receptors Cytochrome b, in DML vesicles N A I)H-c y t oc hrome c oxidoreductase in synthetic membranes Complex I I I only Cytochrome oxidase in native mitochondria1 inncr membranes ADP/ATP translocator in native mitochondrial inner membranes Ferredoxin:NADP ' oxidoreductase in thylakoid membranes Ca" ATPase in sarcoplasrnic reticulum

Method Phosphorescence anisotropy S-T ESR

Phosphorescence anisotropy Luminescence an i sotropy S-T ESR

Rotational correlation time ( p s e c )

References"

17-22

I

> 10o)o (Native membranes) 5ObIOO ( p H I I treated) 15-90 (at 4°C) 0.4. 9.0

2, 3

40-80 (L./P 0 . 5 )

h

4

5

900

IL/P 0.2) 20 700- 1400 ( U P 0.2) (60% immobile)

7

Ahsorhance anisotropy

99 2 20

X

Absorbance anisotropy

0.25 ( N o ferredoxin) 44) (Ferredoxin added) 200 30 50

Absorbance anisotropy

S-T ESR S-T ESR Absorhance mistropy

(37°C) 9

I0

II 12

" Most values quoted are for temperatures at or near physiological. Cooling of membranes has significant effects on rotational diffusion of many of the proteins listed above. " ( 1 ) Ddmjanovich cr r d . (19x3): (2) Rousselet PI trl. (19x1 1: (3) Rousselet et ( I / . (1982); (4) Zidovctzki e l t i / . (1981); ( 5 ) Vaz el a / . (1979): ( 6 ) Poore ( ' I r i l . (1982): (71 Kawato C I ( t l . (198%); (8) Muller ('I d . (1982); (9) Wagner pi ril. ( 19x2): ( 10) Kirino PI rrl. ( 1978): ( I I ) 'rhomils and Hidalgo ( 1978); (12) Hoffman el id. ( 1979).

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MICHAEL EDlDlN

S U M M A R Y REMARK O N MEASUREMENTSOF ROTATIONAL DIFFUSION

4. A

The detailed discussions of the preceding sections as well as the papers summarized in Table I1 show that rotational diffusion measurements may tell us something about the functional states of membrane proteins. However, it will also be noted that different methods for measuring rotational diffusion may give conflicting values for rotational correlation times, or for the fraction of a protein population that is immobile during an experiment and, presumably, aggregated. Both optical and magnetic resonance methods still depend upon a number of assumptions in arriving at numerical estimates of rotation, and both are limited by the quality of spectra obtainable with a given label. Indeed, in a number of instances data are quoted as approximations, o r even as order of magnitude estimates. However, even such estimates have proved useful in understanding the associations between membrane proteins and the effects of temperature or other perturbations upon this association. IV.

LATERAL DIFFUSION

A. Lateral Diffusion in Cell Membranes The lateral diffusion of membrane lipids and proteins has been demonstrated many times. Despite these repeated demonstrations, some authors still confuse thermally driven motions, lateral diffusion, and metabolically driven motions, patching and capping, and many more fail to distinguish between lateral diffusion on a small scale, over fractions of a micrometer, and lateral diffusion over micrometers and tens of micrometers. The field is further made troublesome by some authors’ refusal to accept results obtained by the most widely used method for measuring lateral diffusion, fluorescence photobleaching. Finally, though most measurements have been made on plasma membranes, it may be that lateral diffusion is most important in the functioning of endomembranes, for example, those of mitochondria. I hope that the material in this section will clarify some of this confusion and stimulate thinking about the function of lateral diffusion in coupling reactions in membranes.

B. Methods for Measuring Lateral Diffusion in Membranes

All methods for lateral diffusion take one of two approaches: measurement of frequencies of molecular encounter, or measurement of the time required for marked molecules to fill a defined area of membrane. The second approach has proved by far the more versatile.

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1. ESTIMATES OF LATERAL DIFFUSIONF R O M FREQUENCIES OF

MOLECULARCOLLISIONS If the spectrum of a probe molecule contains features due to collisions or other interactions between molecules, then the magnitude of these features may be used to determine the frequency of collisions and from this the lateral diffusion rate of the colliding molecules. This approach was applied to lateral diffusion of spin-labeled lipids, where the broadening of nitroxide spin-label line widths gave a good measure of the interaction between probe molecules (Devaux and McConnell, 1972; Sackman and Trauble. 1972). Though this work is complicated and limited in application by the high concentrations of spin labels required, similar results have recently been achieved with spin saturation ESR measurements (ELDOR and saturation recovery) which are sensitive to quite low concentrations of spin labels (Popp and Hyde, 1982). All the ESR measurements estimate lateral diffusion of lipid labels (fatty acids and phospholipids) in synthetic bilayers in the range lo-' to lo-' cm2 sec-'. The newer approach is potentially applicable to measurements in native membranes. It might allow estimation of rotational and translation diffusion of a given probe in a single preparation and in a single instrument. The features of fluorescence spectra that report on excimer formation offer another indicator of molecular encounters by lateral diffusion. Vanderkooi and Callis (1974) first used this method to measure diffusion of pyrene in liposomes. It has also been applied to erythrocyte ghosts (Galla et a / . , 1979). Diffusion may also be important in features of fluorescence energy transfer spectra since energy transfer between two fluorophores may be limited by diffusion (Thomas cJt a / . , 1978; Stryer ef ( i l . , 1982). Note that this approach to lateral diffusion measures diffusion on the scale of angstroms and tens of angstroms. 2. ESTIMATES OF LATERAL DIFFUSIONF R O M LARGE-SCALE REDISrRIBUTION OF LABELS Estimates of lateral diffusion on a micrometer scale are made by adding a patch of unlabeled membrane to an otherwise uniformly labeled membrane. A lateral diffusion coefficient is estimated from the kinetics of filling of the unlabeled region with label, appropriate controls being done to rule out mechanisms other than diffusion, for example, metabolically driven flow, which could drive label into the measured area. Three techniques are available for making unlabeled regions in labeled membrane: ( 1 ) formation of heterokaryons by fusion of cells with different surface marker proteins, (2) electrophoresis of membrane proteins in sitr, to create protein-poor regions, and ( 3 ) bleaching of spots or bands in an otherwise uniformly labeled membrane. The first and second techniques

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MICHAEL EDlDlN

follow redistribution of surface markers over a hemisphere nearly as large as a cell: the third technique follows entry of markers into a spot that is usually significantly smaller than one cell diameter. a . Lateral Diffusion in Fused Cells. Lateral diffusion of membrane proteins was first shown in experiments in which the intermixing of surface antigens of mouse/human heterokaryons was followed, using fluorescent antibodies to visualize the progress of marker (histocompatibility)antigens from the “mouse” hemisphere to the “human” hemisphere of the heterokaryons (Frye and Edidin, 1970). The heterokaryons are created almost synchronously in a few minutes, and the population is sampled over a period o f a few hours. The marker antigens are visualized after the population is sampled, that is they diffuse in the absence of added antibodies. Samples are characterized in terms of the fraction of all heterokaryons with intermixed surface antigens. A number of controls indicate that intermixing is not energy dependent; rather, it appears to be due to Brownian motion. N o diffusion coefficient was obtained from the first data of this sort, but it was clear that diffusion was rapid; a time on the order of 10 min was required for redistribution of the antigens of half the heterokaryons. A few years later, the diffusion equation for heterokaryons (all diffusing species concentrated in one hemisphere at time zero) was solved (Huang, 1973) as tliz =

[r,,1’/2D

(2)

where I”’ is the time for 50% reduction in antigen concentration in the source hemisphere, I-,) the heterokaryon radius, and D the diffusion coefficient of the marker antigen. The equation could be used, with some assumptions, to yield order of magnitude estimates of diffusion coefficients and to measure the effects of drugs and other agents on these coefficients (Edidin and Wei, 1977a,b; Edidin and Wei, 1982a). This approach has also been used to estimate the diffusion of band 3 molecules in erythrocytes (Fowler and Branton, 1977; Fowler and Bennett, 1978). However, even though the method can be used quantitatively, it is cumbersome, and at present it is mainly used to check results obtained by other methods (Edidin and Wei, 1982b; Koppel and Sheetz, 1981). 6 . Relaxation after Electrophoresis in Situ. Electrophoresis of mobile membrane components in the plane of the membrane was first suggested by Jaffe (1977) as a mechanism for the effects of electric fields on cell development and differentiation. Jaffe (1977) and Po0 et a / . (1979) developed the equations describing the distribution of molecules in a steady extracellular electric field. This steady-state distribution is a function of electrophoretic mobility and back diffusion of molecules. Experimental

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

105

systems in which cells are placed in a chamber between two electrodes may be characterized in terms of an asymmetry index, A , A(t) =

[C(180",t) - C(O".t)]/[C(I8O0,I) + C(O",t)]

(3)

where C(180")and C(0") are the surface density of molecules facing the anode and cathode, respectively. While A is a function of lateral diffusion, D , and electrophoretic mobility, D can be estimated from the decay of A with time after turning off the current as A ( t ) = B exp ( -2Dt/r')

(4)

where B is a constant characteristic of the system and r is the cell radius. Po0 and Robinson (1977) first showed experimental electrophoresis irr sitir and back diffusion of concanavalin A receptors in Xenopils myoblasts in a field with a potential drop of about 12 mV across a cell width. Diffusion coefficients for concanavalin A-binding proteins are variously estimated from decay of A as 6 x lo-''' cm' seccl (Po0 et u l . , 1978) or 6 x lo-' cm' sec-' (Po0 et d., 1979) in amphibian cell membranes and as I x 10- 'I to 3 x 10-"'cm2 sec-I in cultured rodent cells (Zagyansky and Jard, 1979). It should be emphasized that here, as in heterokaryons, the membrane molecules of interest are not visualized until after electrophoresis or back diffusion are underway or completed. Back diffusion is not observed if Con A binding sites are held in an electric field for 14-24 hr (Po0 et al., 1978; Orida and Poo, 1978)and some aggregation of sites is found after electrophoresis for 30 min (Poo, 1981). Another membrane protein, acetylcholine receptor, aggregates irreversibly after brief electrophoresis. The aggregates themselves are electrophoretically mobile, but they remain stable to lateral diffusion for up to 2 days after electrophoresis for 10 min [cited in Po0 (1981)l. The aggregates grow at the expense of unaggregated, diffusely distributed acetylcholine receptors, and they may mimic the way in which patches of receptors form in mature myocytes. There are some indications that the very large-scale (tens of micrometers) mobility observed in the relaxation is regulated differently than lateral diffusion over a few micrometers. Zagyansky and Jard (1979) aggregated lectin binding sites by electrophoresis in sitii so that these sites would not redistribute back to their original positions. Despite this, they could show appreciable local lateral diffusion, on a scale of a few micrometers, estimating D at 2 x lo-"' cm' sec-l from photobleaching. It would be interesting and useful to develop the instrumentation (scanning spot or quantitiative intensified video microscopy) and labels necessary to follow electrophoresis and back diffusion on membranes in real time on single cells.

106

MICHAEL EDlDlN

Relaxation after electrophoresis has also been used to show lateral diffusion of mitochondria1 membrane proteins. Electric fields applied to mitochondrial suspensions caused aggregation of intramembrane particles in the membranes and once the field was removed. The aggregates appeared to disperse by diffusion at -8 x lo-’” cm’ sec-’ (Sowers and Hackenbrock, 1981). The theory, methodology, and experimental results for electrophoresis in situ have been reviewed (Poo, 1981). c . Phorobleaching Methods. The first determination of a coefficient for lateral diffusion of a membrane protein was made by Po0 and Cone (1974). They bleached the rhodopsin on one side of a rod outer segment, and, by measuring absorbance of unbleached rhodopsin alternately on each side of the rod, they were able to follow both return of unbleached pigment to the bleached side and loss of unbleached pigment from the unbleached side. Controls ruled out alternatives to diffusion as the mechanism for rhodopsin redistribution. A diffusion coefficient for vertebrate rhodopsin, D -3.5 ? 1.5 x cm’ sec-I, was calculated for frog and rat rhodopsins. A similar result was obtained by Liebman and Entine (1974). Both the numerical value obtained and the method are important for all later work on lateral diffusion. A D value of 3 x to 4 x lo-’ cm’ sec-’ is expected from the classical work on diffusion (Einstein, 1905) and from the estimates for membrane viscosity obtained from rotational diffusion measurements (Cone, 1972). While some other proteins appear to diffuse this fast in native membranes, diffusion coefficients for integral proteins typically are an order of magnitude smaller than those measured by Po0 and Cone. It will be seen that considerable effort has been expended on investigating the theoretical and physical bases for this discrepancy. The photobleaching approach to measuring lateral diffusion was limited by the low sensitivity of absorbance measurements (the full scale for changes in rhodopsin absorbance seen by Po0 and Cone was 0.05 O.D.) and by the absence of endogenous labels in most cell membranes. Fluorescent labeling of cell surfaces clearly offered a route to generalizing a photobleaching experiment, and this route was taken by Peters and coworkers, who attempted to measure lateral diffusion of band 3 molecules in erythrocyte ghost membranes labeled with fluorescein isothiocyanate (Peters er nl., 1974). N o recovery of fluorescence was observed after bleaching one-half of a labeled membrane, implying D < 3 x lo-” cm’ sec-’. Despite the failure to actually observe diffusion in the plane of the ghost membrane, this experiment is important as the first in which elements of later photobleaching experiments were combined. Three laboratories further developed photobleaching techniques, appropriately called fluorescence recovery after photobleaching (FRAP) or

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

107

fluorescence photobleaching and recovery (FPR), and obtained the first lateral diffusion coefficients for membrane lipids and proteins (Edidin Pt a / . , 1976; Zagyansky and Edidin, 1976: Jacobson et d., 1976; Schlessinger et (11.. 1976). All the groups used laser light for bleaching or measuring fluorescence in a selected spot on a cell surface. The light, typically from a cw argon ion laser, is attenuated about 1000-fold and focused through a microscope onto the labeled membrane. Spots with a I/e’ diameter of 2-3 k m are readily obtained. and these can be used to measure lateral diffusion on cells or organelles as small as 5-7 pm in diameter, using a wide range of fluorescent probes and labels. After measuring the steadystate intensity of fluorescence, a fraction of the fluorophores is bleached by a brief pulse, typically tens of milliseconds, of the unattenuated beam. The laser beam is attenuated once again and used to measure the recovery of fluorescence in the bleached spot. Diffusion coefficients are estimated from the time dependence of fluorescence intensity in the spot. The level to which fluorescence finally recovers is a measure of the fraction of molecules free to diffuse during the time of the experiment. An idealized experiment is shown in Fig. 1. The recovery curve is not a simple exponential, but it may be fit by nonlinear least squares routines to yield half-times for recovery of fluorescence, f I l 2 , which in turn are related to D,;,, as D,,, = w214t1,2~

(5)

where y, a function of the fraction of molecules bleached, ranges between I .O and 2.2. Values of f I l 3 may also be estimated from the recovery curves themselves, as can the level of fluorescence recovery (Axelrod et d., 197ha,b). The recovery curve shape is sensitive to flow, and in principle components of recovery due to flow, rather than to lateral diffusion. can be detected from this. In practice it is difficult if not impossible to decide if a curve is due to a mixture of flow and diffusion or if it is a composite of two different diffusion coefficients. However, a new linearization procedure for FPR data is sensitive to flow or to multiple diffusion coefficients, and allows fast data fitting on quite small computers as well (van Zoelen el d., 1983). This promising approach should greatly improve the analysis of FPR data. d. Other FPR Methods. Variations on the basic FPR technique have evolved in several laboratories. Peters and co-workers ( 198I ) observed the rate of continuous bleaching in a spot rather than the rate of return of fluorescence to a highly bleached spot. Koppel rt t i / . (1980) used a spot scanning method to measure changes in fluorescence intensity in and out of the bleached spot. This demonstrates both gain of fluorescence in the spot and loss from the population of unbleached molecules outside the

108

MICHAEL EDlDlN INITIAL CONDITION

AFTER RECOVERY

AFTER PHOTOBLEACHING

....... .:.: .".

6 .

0

z W V

8 a 3 J LL

I

I

!

0

'

I

%

TIME

-

FIG.I . An idealized photobleaching experiment. Fluorescent, unbleached molecules on the cell membrane are shown as dots, and bleached molecules are shown as crosses. The area of bleaching and measurement is defined by a circle. F,, Initial fluorescence level in the measurement region; F,, level of maximum recovery of fluorescence after photobleaching: tl12,half-time for recovery of fluorescence. (Reproduced from Jacobson et a/.. with permission of Fed. Prcic.. Fed. Am. SOC.Exp. B i d . )

spot, giving a more reliable evaluation of lateral diffusion than measurements of fluorescence in the spot alone, Several laboratories have sacrificed information on single spots in a membrane for the accuracy, speed, and sensitivity gained by imposing a pattern of bleached and unbleached regions across an entire cell (Smith and McConnell, 1978; Davoust et al., 1982). Beside being more sensitive than spot photobleaching, the method also detects anisotropic lateral diffusion, and its most notable result is that membrane glycoproteins diffuse perpendicular to an oriented cytoskeleton at least an order of magnitude more slowly than they diffuse parallel to the cytoskeleton (Smith er al., 1979). Progress in instrumentation and analysis and newer results with FPR are summarized in a workshop report (Jacobson et al., 1983). C. Accuracy of Lateral Diffusion Coefficients by FPR

All three laboratories that first published on FPR obtained "typical" values for lateral diffusion of membrane glycoproteins of around 2 x lo-'"

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

109

cm' sec- I . Fluorescent dyes, notably dioctadecylindocarbocyanine iodide (diI), were also used to probe lateral diffusion in cell membrane lipids. Typical diffusion coefficients for diI labels in cells were 0.5 x 10- to 2 x cm' sec-', about the same as observed for the dyes in synthetic bilayers. Many other laboratories since have found comparable values in FPR measurements. They are tabulated in several reviews (Edidin. 1981; Cherry, 1979; Peters, 1981), and representative results are shown in Tables I11 and IV (see below). The results for membrane lipid probes in cell membranes are generally in good agreement with results in synthetic membranes and with values obtained by other techniques, for example ESR. The diffusion coefficients for lipid probes are those predicted from the viscosity of phospholipid bilayers estimated from rotational diffusion of membrane proteins. In contrast, many results for plasma membrane proteins are not at all those expected from the results of Po0 and Cone (1974) nor from calculations of lateral diffusion from rotational diffusion coefficients or from protein diameter and membrane viscosity (Saffman and Delbruck, 1975).The following section will show that this discrepancy is largely due to cytoskeleton effects on lateral diffusion, and perhaps also to effects of membrane protein concentration on effective membrane viscosity. The unexpectedly small lateral diffusion coefficients found in the first FPR experiments and the experimental conditions in which an intense pulse of light is focused on a small spot have led to suggestions that lateral diffusion measured by FPR is artifactually low due to membrane damage and photoinduced cross-linking of labeled proteins. These views have been vigorously pressed, especially by Bretscher (1980), whose theory of cell locomotion and membrane flow will not accommodate diffusion coefficients for membrane proteins as small as IO-l('cm' sec-l (Bretscher, 1976). A calculation by Axelrod (1977) showed clearly that local heating is unlikely to occur in a typical FPR experiment. Some work on bleaching of tluorescent molecules in cuvettes does suggest that photoinduced crosslinking is likely to occur (Lepock er a / . . 1978; Sheetz and Koppel, 1979). However, these measurements were made with light intensities lower than those used in an FPR experiment, and higher light fluxes induced less cross-linking than lower fluxes. It appears then that the rute of bleaching, not the total number of photons irradiating the sample, is important in cross-linking. Other indications that FPR might give falsely low diffusion coefficients come from an electrophysiological measurement of acetylcholine receptor diffusion (Poo, 1982) and from an observation that the rapid clustering of rhodamine-bungarotoxin-labeled acetylcholine receptors is inhibited by light fluxes of the magnitude usually used in fluorescence microscopy (Olek et d.,1983). The effects of low light fluxes on lateral movements are

110

MICHAEL EDlDlN

probably comparable to the observations by Lepock et al. (1978) and by Sheetz and Koppel (1979). However, the electrophysiological estimate of D,,, for unlabeled acetylcholine receptors of about 2 X cm’ sec..’ (Poo, 1982) is 100-fold greater than that estimated from FPR measurements of rhodamine-bungarotoxin-labeledacetylcholine receptors (Axelrod ef ul., 1978). The discrepancy may be due to some effect of the bungarotoxin label required to mark the receptors for FPR rather than to the effects of light on the receptor-label complex. Covalently labeled acetylcholine receptor from Torpedo gave D = 2 x lo-* to 3 x lo-’ cm’ sec-’ when reconstituted in liposomes (Criado et al., 1982), and even receptor labeled cm2 sec-’ when difwith fluorescent bungarotoxin gave D = 2.0 x fusion was measured on cell blebs largely free of cytoskeleton (Tank el ul., 1982). Perhaps the binding of bungarotoxin affects the associations between acetylcholine receptor molecules and the cytoskeleton. A number of control experiments have now been performed which tell strongly against the possibility that diffusion coefficients measured by FPR are artifactually low. Wolf and co-workers (1980) labeled a single class of receptors with a mixture of two different fluorochromes, fluorescein and rhodamine, and showed that the diffusion coefficients measured were not affected by previous measurements at a given spot. This control itself depends upon FPR, and though arguing against photodamage it cannot rule it out. Wey and co-workers (1981) were able to covalently label rhodopsin in intact disc membranes and to repeat the measurements of Po0 and Cone (1974), using FPR rather than absorbance measurements. They found the same diffusion coefficient by FPR that had previously been cm’ sec-’, has found by flash photolysis. This value, around 3 x also been measured by a method independent of any bleaching, in which diffusion is calculated from birefringence changes in the presence of a rapidly reversing electric field (dynamic Kerr effect) (Takezoe and Yu, 1981).

Koppel and Sheetz (1981) covalently labeled band 3 of erythrocytes with fluorescein and then fused mixtures of labeled and unlabeled cells. Lateral diffusion could be evaluated in fused pairs of labeled and unlabeled cells, in terms of the redistribution of fluorescence from labeled to unlabeled partner, using a spot scanning method (Koppel, 1979) to improve on the original design of this experiment (Fowler and Branton, 1977). Lateral diffusion was estimated in fused pairs of labeled cells in the same population by photobleaching one-half of each pair and estimating the diffusion coefficient by following the return of fluorescence to the bleached spot, that is, by FPR. Both methods gave D = 3.8 x l o - ” cm’ sec-’. Here FPR gives the same diffusion coefficient as a method which does not depend on rapid photoselection of molecules. A comparison between

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

111

diffusion coefficients for histocompatibility-2 antigens measured in heterokaryons (Edidin and Wei, 1977a) or by FPR (Edidin and Wei, 1982b) also gives no indication that values from FPR are artifactually low. Indeed, if anything, the FPR measurements missed some very slowly diffusing molecules which could be detected in late time samples of heterokaryons. In summary, control experiments fail to show that diffusion coefficients obtained by FPR are artifacts of the photobleaching method. Rather, there appear to be constraints to lateral diffusion in many, though not all, cell membranes. If these constraints are removed, then FPR measurements show that molecules such as band 3 may difuse 50 times faster than they do in native membranes.

D. Constraints to Lateral Diffusion in the Plasma Membrane Lateral diffusion may be constrained by interactions of membrane proteins with other molecules within the membrane bilayer or on either side of the membrane. Limited data on interaction of external peripheral proteins, notably fibronectin, indicate that external protein coats do not affect lateral diffusion (Schlessinger et NI., 1977). Membrane protein concentrations do affect dffusion coefficients to some extent. Diffusion of a number of proteins in synthetic bilayers is faster than observed in native membranes in the absence of cytoskeletal interactions (see Table IV below). and diffusion of a given protein reconstituted into a defined lipid bilayer depends on the protein to lipid ratio in the reconstituted membrane (Peters and Cherry, 1982). There is also some evidence that lateral diffusion is 10 times faster than usual in native membranes with very low protein concentrations (Small et al., 1982). However, the magnitude of these effects (less than 10-fold reductions in lateral diffusion) is significantly less than that of the effect of residence in a cell membrane on protein lateral diffusion (reduction of 10- to 1000-fold from the rate in synthetic or cytoskeleton-free bilayers). Thus it seems that molecular interactions within the plasma membrane bilayer do not in general regulate lateral diffusion rates. Protein concentrations could dominate lateral difusion in endomem branes. We are left with membrane-cytoskeleton interactions as the locus for constraints to lateral diffusion in plasma membranes. Experiments on capping implicate the cytoskeleton (Taylor et d.,1971; for reviews see Edelman, 1976; Nicolson, 1976a.b; Schreiner and Unanue, 1976; Edidin, 1981)as do the effects of cytochalasin B on lateral diffusion of membrane proteins (Edidin and Wei, 1977a; Schlessinger r t al., 1976). Cytochalasin B blocks capping but immobilizes membrane proteins, rather than releasing constraints to their lateral diffusion. Though unexpected, the results at

112

MICHAEL EDlDlN

least suggest that something involving actin filaments is implicated in the control of lateral diffusion. Study of erythrocyte band 3 molecules has given the most abundant evidence for cytoskeletal control of lateral diffusion. The diffusion of band 3 measured by label mixing in fused cells or by FPR in ghost membranes is around 3 x lo-'' to 4 x lo-'' cmz sec-'; values in a particular system depend on temperature and on the ionic strength of the buffer used to suspend the ghosts (Fowler and Branton, 1977; Koppel et a/., 1981; Schindler et al., 1980; Sheetz et al., 1980; Golan and Veatch, 1980). Lateral diffusion is affected by treatments which alter the cytoskeleton: aging of red cells before measurement (Fowler and Branton, 1977), removal of spectrin by addition of solubilized ankyrin (Fowler and Bennett, 1978), removal of cytoskeletal proteins by increasing temperature and lowering the ionic strength of incubating buffers (Golan and Veatch, 1980), or covalent cross-linking of spectrin (Smith and Palek, 1982). Lateral diffusion of band 3 in ghost spherocytic cells from mice congenitally lacking spectrin is 50-100 times faster than in normal mouse erythrocyte ghosts (Sheetz et a / . , 1980; Koppel er a/., 1981). These data are summarized, together with results on lateral diffusion of membrane lipids, in Table 111. The results are not unique to band 3 and erythrocytes. Webb and coworkers were able to create F-actin-free blebs on nucleated cells (myoblasts and lymphocytes). They found that lateral diffusion of membrane proteins within these blebbed regions lacking the most abundant cytoskeletal protein was around 100-fold faster than in adjacent regions of a given cell which had remained attached to the cytoplasm (Tank, et al., 1982; Wu et al., 1982). Still higher rates of lateral diffusion may be observed for membrane proteins reconstituted into liposomes. Protein concentration affects diffusion coefficients to some extent. Coefficients in the range of native and synthetic membranes are summarized and compared in Table IV. The interactions of membrane integral proteins with the cytoskeleton are still undefined. These interactions could involve specific coupling proteins and coupling sites or could be due to nonspecific steric hindrance of diffusion by the polymer matrix of the cytoskeleton. These alternative models are discussed by Koppel et al. (1981), whose data on lateral diffusion of band 3 are consistent with a matrix control model for regulation of lateral diffusion. On the other hand, there are several reports of the coisolation of membrane proteins and cytoskeletal proteins. Koch and coworkers (Koch and Smith, 1978; Flanagan and Koch, 1978) found actin associated with capped and shed membrane immunoglobulins and with shed H-2 antigens. Later, several groups isolated membrane Ig-actin complexes also containing a protein of molecular weight -56,000 which

LATIXN. DIFILISION

'TABLE 111 M E M B H A N LlPll)S E

01: EKYTHROCY'I'A

AND

PHOI'ICINS

I)

Preparation Human cells I 2'C 25°C 37°C Mouse cells 25 "C 37°C Human ghosts 12°C 25°C 37°C Mouse ghosts 23°C Human ghosts

y'c

Labeled molecule(s)

Method

x 10"')

FPR

References" 1

31 82 210 130 300 I 72'8

7x 190 NBD-PE-Lipid label

7 I 50

Band 3

Cell fusion

-. 1

30°C 37°C 30°C iankyrin Human ghosts 30°C 30°C crosslinked spectrin Human ghosts Human ghosts 5 mM salt 21°C 30°C I3 mM salt 21°C 30°C 37°C 26mM salt 21°C 30°C 37°C Mouse ghosts Normnl Spectrin deficient

(

2 0 .Oh' 0.30 0.40 2 x over

3

buffer alone Band 3

Cell fusion

4

0.30 0 . Ih

Band 3 Band 3

Cell fusion FPR

5 h 0.S 1.1

0.7" I .4 6.5

0.7 0.7 1.4

Band 3

7 0.45

25

" References: ( 1 ) Bloom and Wehh (1983): ( 2 ) Fowler and Branton (1977): 13) Fowler and Bennett (1978); (4)Smith and Palek (1982): ( 5 ) Koppel and Sheetz (1981); ( 6 ) Golan and Veatch (1980); (7) Koppel e ~ d(1981). . * Significantly lower than the D value in intact cells at the sanie temperature. ' Lower limit in cells from fresh blood. " Mobile fractions significantly higher at 30 and 37°C than at 21°C.

114

MICHAEL EDlDlN

TABLE IV LATERAL DIFFUSION OF SOMEMEMBRANE PROTEINS IN CELLSAND I N SYNTHETIC MEMBRANES D in native membranes Protein Vertebrate rhodopsin Bacteriorhodopsin L:P 21O:l L:P 140:l L:P 9o:l Acetylcholine receptor Calcium ATPase of sarcoplasmic reticulum Histocompatibility antigens

(

x 10"')

30 Immobile

Immobile

- looh 1-40

D in synthetic membranes (X

10"')

References"

160-300

I, 2

340 230 I30 200-300 200

3

I00

2, 4 2

5-9

References: ( 1 ) Po0 and Cone (1974);(2) Vaz et a / . (1982);( 3 ) Peters and Cherry (1982); (4) Criado ef c d . (1982); ( 5 ) Cartwright et a / . (1982); (6) Smith P r a / . (1982); ( 7 ) Henis and Gutman (1983); (8) Damjanovich el a / . (1983); (9) Edidin and Wei (1982a). Estimated from rotational diffusion. 'I

may serve to link the membrane protein to actin (Petrini et al., 1983). There are numerous other indications of specific membrane-cytoskeleton interactions, mainly from fluorescence studies of cell adhesion plaques and from analysis of the membrane proteins that remain associated with Triton X-100-insoluble cell cytoskeletons (for review see Geiger, 1983). Though extraction experiments suggest specific associations they do not give definite evidence for such associations and against the matrix model of Koppel et al. (1981). The cytoskeletal constraints to lateral diffusion are not static. Diffusion coefficients and the fraction of mobile molecules change significantly through the cell cycle (de Laat ef af., 1979) and also may be altered by treatment of cells with the calcium ionophore A23187 or with metabolic inhibitors (Axelrod et al., 1978; Edidin and Wei, 1982a,b). These treatments may alter either the rate of lateral diffusion or the fraction of molecules free to diffuse, or both. The distinction between changes in rate of lateral mobility and changes from immobile to mobile was first explicitly noted by Golan and Veatch (1980), who modified lateral diffusion in erythrocyte ghosts by manipulating temperature and ionic strength of the cell bathing medium.

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

115

E. The Limits to Lateral Diffusion: Surface Membranes of Cells in Tissues Demonstrations of long-range lateral diffusion lead us to the paradox that cells in tissues often have surfaces that are highly organized with particular sets of enzymes, pumps, and receptors consistently localized to particular regions of membranes. If the diffusion coefficients measured for free-living cells like lymphocytes or for cells of cultured cell lines also held for membranes of tissue cells, then one might expect that such cells would use most of their metabolic resources to synthesize new membrane proteins in order to maintain gradients of protein concentration on their surfaces. This is not the case. Rather it appears that diffusion is greatly restricted in cells in tissues. This assumption is supported by the FPR measurements of Gall and Edelman (1981) comparing diffusion in membranes of cells in tissue slices and in membranes of single cells dissociated from tissues. They found the same diffusion coefficients for labeled antigens in both cases, but while membrane proteins were mobile in more than 95% of isolated liver cells these proteins were immobile in two-thirds of liver cells in tissue slices (Dla,< 5 x 10P” cm’ s e c P 1 ,mobile fraction < 10%). Cultured epithelial cells (MDCK) form oriented, morphologically polarized monolayers which, like epithelial cells in v i w , are water tight and transporting. Cells in these monolayers have organized and segregated their surfaces into regions of different protein (Louvard, 1980; Cereijido et a / . , 1980) and lipid (van Meer and Simons, 1982). Dragsten and coworkers (1981, 1982) studied the redistribution of lipid-soluble dyes from apical to basal surfaces of these cells. They concluded that the tight junctions between cells form the major barrier to interdiffusion of basal and apical surfaces but that this barrier could be crossed by some dyes. In a test cell system, not MDCK, the dyes that could redistribute were incompletely quenched by water-soluble quenchers external to cells while dyes that could not redistribute were completely quenched. Dragsten ct a / . (1982) interpret this result as showing that dyes which partition only into the outer bilayer leaflet cannot pass the junctional complex while dyes in the inner leaflet can pass the complex. This interpretation implies that all of the membrane lipid asymmetry observed in MDCK and other epithelial cells, for example, intestinal epithelia, is restricted to phospholipids in the outer bilayer leaflet. Even though the phospholipid composition of the two halves differs, this difference is not absolute and 1 would expect the eventual redistribution of all membrane lipids because of partitioning between outer and inner bilayer leaflets. However, the tight

116

MICHAEL EDlDlN

junctional band around each cell in an epithelium does seem a plausible barrier to lipid diffusion. The restrictions on protein lateral diffusion in epithelial cells are not known. One might expect tight junctions to segregate proteins as well as lipids into apical and basolateral membrane domains. On the other hand, it is also plausible that the same cytoskeletal mechanisms that operate to impede lateral diffusion in membranes of isolated cells function more efficiently and to a greater extent in epithelia. It is known that apical membrane proteins and glycoproteins redistribute when bladder (Pisam and Ripoche, 1976) or intestinal epithelial cells (Ziomek et ul., 1980) are dissociated, and we have observed redistribution of H-2 glycoproteins from basolateral to apical surfaces of dissociated intestinal epithelial cells (F. Weaver and M. Edidin, unpublished). Redistribution of alkaline phosphatase or leucine aminopeptidase from apical to basal surfaces was not inhibited by fixation in as much as 3% paraformaldehyde or by uncouplers. Hence, the redistribution process is probably diffusion, not metabolically driven flow. Diffusion coefficients for the enzymes were estimated as 5 x 10-"' to 9 x lo-'" cm' sec-' at 37°C. No redistribution was seen in cells in strips one cell thick by three to five cells long; their cells therefore could not bear intact bands of tight junctions since they lacked neighbors on two sides. Treatment of single cells with the uncoupler CCCP enhanced redistribution rates up to 2-fold, suggesting metabolic requirements for constraints to redistribution. It may be noted that treatment of HeLa cells with uncouplers disrupts microtubules and intermediate filaments (though not actin filaments) (Maro and Bornens 1982). Single cultured epithelial cells may polarize in the absence of tight junction formation. MDCK cultured kidney epithelial cells are polarized for budding of enveloped viruses as well as for morphological and enzymatic membrane markers. Influenza, Sendai, and SV5 viruses all bud from the apical surfaces of MDCK monolayers, while vesicular stomatitis virus buds from the basal surface (Rodriguez-Boulan and Sabatini, 1978). This polarity is lost if the cells are dissociated and maintained in suspension culture, but is regained by cells that form small aggregates in culture, or by single cells that are allowed to attach to a collagen substrate (Rodriguez-Boulan et ul., 1983). This behavior can be compared to the diffusion of markers on strips of intestinal cells. If the tight junctions do serve as barriers to lateral diffusion of proteins as well as lipids, then membrane proteins of epithelial cells ought to be mobile within the apical or basal regions. On the other hand, if restrictions to diffusion are cytoskeletal, then all markers should be immobile, wherever they are examined.

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

117

F. Lateral Diffusion and Membrane Function: Diffusion-Coupled Reactions The data reviewed above clearly show lateral diffusion of membrane components. Is this diffusion functional? There are far fewer experiments on the role of lateral diffusion in membrane function than measurements of lateral diffusion. However, even the limited data indicate functions for lateral diffusion in coupling reactions between cell surface and interior and between proteins of organelle membranes in mitochondria. All discussions of diffusional coupling of reactions in membranes turn on analyses of the rates of encounters by diffusion as a function of the dimensionality of diffusion. This problem was first discussed by Adam and Delbruck (l967), who considered the problem of encounter between a molecule and a target of size a in a space of size b, h >> U . They showed that the time for encounter between diffusing molecule and target was proportional to a tracking factor, .f(blcr). The factor was linear in hltr for three-dimensional diffusion, was proportional to log hlu in two dimensions, and was independent of hln in one dimension. They argued that two-dimensional diffusion or combinations of three- and two-dimensional diffusion would result in significantly shorter "catch" times than would threedimensional diffusion alone. The biological example used was the catch of pheromones on the waxy cuticle surrounding specific pheromone sensors of moths. This evaluation was carried further by Hardt (19791, who showed how the concentration of interacting species affects reaction rates, and showed that the sensitivity of reaction rates to concentrations of reactants increases with decreasing dimensionality. The first general suggestions that diffusion might couple reactants came from work on activation of adenylate cyclase by agonist-receptor complexes. The stoichiometry of activation indicated that a single pool of enzyme was activated by many different receptor-agonist complexes, and this was best accommodated by a model in which receptors encountered cyclase at random by diffusion (Cuatrecasas, 1974; De Haen, 1976). It was later shown that receptors implanted in membranes by cell fusion or by partion from detergent extracts could functionally activate adenylate cyclase of the recipient cells (Orly and Schramm, 1976; Eimerl of d., 1980). further implying that receptor and enzyme interacted by random collision. Hanski et a/.(1979) were able to show that reducing membrane viscosity by incorporation of cis unsaturated fatty acids increased the rate of interaction about 2-fold, and they derived a plausible diffusion coeficient for the receptor and/or the cyclase: D -4 x lo-" cm' sec-'. However, the model is complicated by the demonstration of a coupling protein, the GTP-regulatory protein, that mediates the interaction of receptors with

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adenylate cyclase (for review see Levitski, 1981). The coupling protein need not be integral to the membrane bilayer and may diffuse on the inner surface of the bilayer. Thus, while the evidence for lateral diffusion coupling of all reactants remains strong, the diffusion coefficient derived for the reaction needs to be reevaluated. One other interaction between components of the plasma membrane appears likely to be mediated by lateral diffusion: the entry of low density lipoprotein (LDL) receptors into coated pits. Goldstein et af. (1981) used published data on the lateral diffusion rate of LDL receptors, -lo-" cm' sec-', and on the surface density and metabolism of coated pits and receptors to derive a plausible model for the entry of receptors into coated pits by lateral diffusion. This analysis might be generalized to all receptors which internalize through coated pits. Two studies suggest that components of electron transfer systems in endoplasmic reticulum interact by diffusion and random collision (Yang, 1977; Strittmatter and Rogers, 1975). However, these studies are based on interactions in reconstituted model membranes. The most thorough and convincing series of papers on diffusion coupling of reactions comes from Hackenbrock and co-workers, who have been able to relate lateral diffusion and coupling in electron transport chains of mitochondria1 inner membranes. The work involved the demonstration of lateral diffusion of membrane particles quoted earlier (Sowers and Hackenbrock, 198 I), the creation of lipid-enriched mitochondria which allowed study of the dependence of redox reactions on the density of donors and acceptors, and the use of FPR and a variety of specific fluorescent labels to study lateral diffusion of cytochrome b-c,, cytochrome oxidase, cytochrome c, and ubiquinones in inner membranes of giant mitochondria (Gupte et a/., 1984). The integral membrane proteins cytochrome b-c, and cytochrome oxidase diffused at around 4 x lo-"' cm' sec-', somewhat more slowly than estimated for typical membrane proteins by Sowers and Hackenbrock (1981), while a ubiquinone analog and a lipid-soluble dye diffused 6 and 10 times faster, respectively. Cytochrome c , a peripheral protein, diffused at a rate dependent on ionic strength, being almost immobile in low ionic strength medium and diffusing at around 2 x cm' sec-' in 25 mM saline buffer. Recovery of cytochrome L' fluorescence after bleaching appeared to be due to a mixture of two- and three-dimensional diffusion. Recoveries of the other labeled components were around 90%. Thus recoveries are as high as those found for proteins in synthetic or cytoskeleton-free membranes, while the diffusion coefficients, even for the lipid probes, are about an order of magnitude slower than seen in synthetic membranes. Much of this difference must be due to the high protein concentration in the inner membrane. Effects of membrane protein concentration on lateral diffusion have been discussed above.

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The diffusion coefficients estimated for complexes I-IV from these measurements were used to calculate collision frequencies for redox components, and these were found to be greater than the maximum turnover numbers observed for all redox components. Thus lateral diffusion rates are sufficient to couple all components in mitochondrial electron transport, which then must be randomly arranged in the mitochondrial membrane. Here it seems we have found a membrane in which lateral diffusion serves effectively to organize function.

V.

A CONCLUDING REMARK

I have tried to use a limited number of examples to cover the methodology and results for rotational and lateral diffusion of membrane proteins. I hope that this article has shown some directions in which the measurements of diffusion may be taken to shed light on membrane function.

REFERENCES Abercrombie. M. A. (1980). The crawling movement of metazoan cells. P r w . R. S o c . Lom/oti B 207, 129-147. Adam, C., and Delbruck. M. (1967). Reduction of dimensionality in biological diffusion processes. Itr "Structural Chemistry and Molecular Biology" (A. Rich and R. Davidson. eds.), pp. 198-2 15. Freeman. San Francisco. Ahl. P. L..and Cone, K. A. (1982). Photodichroism and the rotational motions of rhodopsin and bacteriorhodopsin. M t ~ / / i dBi:.vu~o/. .~ 88, 74 1-750. Austin, R . H.. Chan, S. S., and Jovin, T. M. (1979). Rotational diffusion of cell surface components by time-resolved phosphorescence anisotropy. P r o f . N d . A w d . .Sc,i. U.S.A. 76, 5650-5654. Axelrod. D. ( 1977). Cell surface heating during fluorescence photobleaching recovery experiments. Biopliys. J . 18, 119-131. Axelrod, D., Kavdin. P., Koppel, D. E., Schlessinger. J., Webb, W. W., Elson, E. L.. and Podleski, T. R. (1976a). Lateral diffusion of fluorescently labeled acetylcholine receptors in membranes of developing muscle fibers. Pr~ic,.N d . A w d . Sci. U.S.A. 73, 45944598. Axelrod. D., Koppel, D. E.. Schlessinger. J.. Elson, E. L., and Webb. W. W. (1976h). Mobility measurements by analysis of tluorescence photobleaching recovery kinetics. Biciphys. J . 16, 1055-1069. Axelrod. D.. Ravdin, P.. and Podleski, T. R. (1978). Control of acetylcholine receptor mobilify and distribution in cultured muscle membranes. Biochini. Bioplivs. Aciu 51 I, 23-38. Bloom, J. A,. and Webb. W. W. (1983). Lipid diffusibility in the intact eryfhrocyte membrane. Bioph.vs. J . 42, 295-305. Bretscher. M. (1976). Directed lipid flow in cell membranes. Natitre ( f . o f i d f i ~260, J 21-23, Bretscher. M. (19x0). Lateral diffusion in eukaryotic cell membranes. T w m k Bioc/irnr. S c i . 5, R b R 7 .

120

MICHAEL EDlDlN

Brown, P. K. (1972). Rhodopsin rotates in the visual receptor membrane. Nutiire (New) Biol.) 236, 35-37. Brown, M., Anderson, R., and Goldstein, J . (1983). Recycling of Receptors: The roundtrip itinerary of migrant membrane proteins. Cell 32, 663-667. Cartwright, G. S ., Smith. L. M., Heinzelmann, E., Ruebush. M. J., Parce, J. W., and McConnell, H. M. (1982). H-2Kk and vesicular stomatitis virus G proteins are not extensively associated in reconstituted membranes recognized by T cells. Proc. Nutl. Acad. Sci. U.S.A. 79, 1506-1510. Cereijido. M., Ehrenfeld, J., Meza, I., and Martinez-Palomo, A. (1980). Structural and functional membrane polarity in cultured monolayers of MDCK cells. J . Mrmhr. Biol. 52, 147- 159. Chang. C.-h., Takeuchi, H., Ito, T., Machida, K.. and Ohnishi, S. (1981). Lateral mobility of erythrocyte membrane proteins studied by the fluorescence photobleaching recovery technique. J . Biochem. 90, 997-1004. Cherry, R. (1978). Measurement of protein rotational diffusion in membranes by flash photolysis. Methods Enzymol. 5 4 , 4 7 4 . Cherry, R. J. (1979). Rotational and lateral diffusion of membrane proteins. Biochim. Biophys. Acrn 559, 289-321. Cherry, R. J.. and Godfrey, R. (1981). Anisotropic rotation of bacteriorhodopsin in lipid membranes. Comparison of theory and experiment. Biophys. J. 36, 257-276. Cherry, R. J., Buerkli. A.. Busslinger, M., Schneider. G., and Parrish, G. A. (1976). Rotational diffusion of band 3 proteins in the human erythrocyte membrane. Ncifitre(London) 263, 389-393. Cone, R. A. (1972). Rotational diffusion of rhodopsin in the visual receptor membrane. Nutitre (New B i d . ) 236, 39-43. Criado, M., Vaz, W. C. L., Barrantes, F. J., and Jovin, T. M. (1982). Translational diffusion of acetylcholine receptor (monomeric and dimeric forms) of Torpedo mcirmornlrr reconstituted into phospholipid bilayers studied by fluorescence recovery after photobleaching. Biochemistry 21, 5750-5755. Cuatrecasas, P. (1974). Membrane receptors. Annu. Rev. Biochem. 43, 169-214. Damjanovich, S.. Tron, L.. Szollosi. J., Ziodvetzki, R., Vaz, W. L. C.. Regateiro, F.. AmdtJovin, D., and Jovin, T. M. (1983). Distribution and mobility of murine histocompatibility H-2Kk antigen in the cytoplasmic membrane. Proc. Nut/. Acnd. Sci. U.S.A. 80, 59855989. Davoust, J.. Devaux. P. F., and Leger, L. (1982). Fringe pattern photobleaching, a new method for the measurement of transport coefficients of biological macromolecules. EMBO J . 1, 1233-1238. De Haen, C. (1976). The non-stoichiometric floating receptor model for hormone-sensitive adenyl cyclase. J . Theor. B i d . 58, 383-400. de Laat. S. W., van der Saag, P. T.. Elson. E. L., and Schlessinger, J . (1979). Lateral diffusion of membrane lipids and proteins during the cell cycle of neuroblastoma cells. Biochim. Biophys. Actu 558, 247-250. Delmelle, M., Butler, K. W., and Smith, I. C. P. (1980). Electron spin resonance spectroscopy as a probe of anisotropic motion in model membrane systems. Biochrniistry 19, 698704. DePierre. J. W., and Ernster, L. (1977). Enzyme topology of intracellular membranes. Annrc. Rev. Biochem. 46, 210-262. Devaux, P.. and McConnell, H. M. (1972). Lateral diffusion in spin-labeled phosphatidylcholine multilayers. J . A m . Chern. Soc. 94,4475-4481. Dragsten. P., Blumenthal, R., and Handler, J . (1981). Membrane asymmetry in epithelia: Is the tight junction a barrier to diffusion in the plasma membrane? Natirre (London) 294, 718-722.

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

121

Dragsten, P. R., Handler. J . S . . and Blumenthal, R. (1982). Asymmetry in epithelial cells: Is the tight junction a barrier to lateral diffusion in the plasma membrane'? In "Membranes in Growth and Development" (C. F. Fox, ed.), pp. 525-536. Alan R. Liss. New York. Edelman. G. M. (1976). Surface modulation in cell recognition and cell growth. Science 192, 218-226. Edidin, M. (1981). Molecular motions and membrane organization and function. In "Membrane Structure and Function" (J. B. Finean and R. H. Michell, eds.) New Comprehensive Biochemistry, Vol. I ), pp. 37-82. Elsevier, Amsterdam. Edidin, M., and Wei. T. (1977a). Diffusion rates of cell surface antigens of mouse-human heterokaryons. I. Analysis of the population. J . Cell B i d . 75, 475-482. Edidin, M., and Wei. T. (1977b). Diffusion rates of cell surface antigens of mouse-human heterokaryons. 11. Effects of membrane potential on lateral diffusion. J. Ce//B i d . 75, 483-489. Edidin. M.. and Wei. T. (1982a). Diffusion rates of cell surface antigens of mouse-human heterokaryons. 111. Regulation of lateral diffusion rates by calcium ions J. Cell B i d . 95,453457. Edidin. M.. and Wei, T. (1982b). Lateral diffusion of H-2 antigens o n mouse fibroblasts. J . Cell B i d . 95, 458-462. Edidin, M., Zagyansky. Y.,and Lardner, T. J. (1976). Measurement of membrane protein lateral diffusion in single cells. Science 191, 466-468. Eimerl. S.,Neufeld, G., Korner. M . . and Schramm. M. (1980). Functional implantation of a solubilized p-adrenergic receptor in the membrane of a cell. Proc. Nail. A w d . Sci. U.S.A. 77, 760-764. Einstein. A. (1905). In "Investigations on the Theory of the Brownian Motion" (R.Furth. ed.). pp. 1-35. Dover. New York (reprint). Flanagan, J., and Koch. G. L. E. (1978). Cross-linked surface Ig attaches to actin. Nu/iire (London)273, 278. Fowler, V.. and Bennett. V. (1978). Association of spectrin with its membrane attachment site restricts lateral mobility of human erythrocyte integral membrane proteins. J . S i i protnol. Sirrici. 8, 21.5-22 I . Fowler. V.. and Branton. D. (1977). Lateral mobility of human erythrocyte integral membrane proteins. Nutiire (London) 268, 23-26. Frye. L. D., and Edidin. M. (1970). The rapid intermixing of cell surface antigens after formation of mouse-human heterokaryons. J. Cell Sc.i. 7, 3 19-335. Gaffney. B. J. (1979). Spin label-thiourea adducts. Model for saturation transfer EPR studies of slow anisotropic rotation. J. Phvs. Chrm. 83, 3445-3449. Gall. W. E., and Edelman, G . M. (1981). Lateral diffusion of surface molecules in animal cells and tissues. Science 213, 903-905. Galla. H.J.. Hartman. W., Theilen, V.. and Sackman. E. (1979). Two-dimensional passive random walk in lipid bilayers and fluid pathways in biomembranes. J . Memhr.. B i d . 48, 21.5-236. Geiger. B . (1983). Membrane-cytoskeleton interactions. Biochi,n. Biop/iy.v.Acttr 737, 30.5341. Golan. D. E . , and Veatch, W. (1980). Lateral mobility of band 3 in the human erythrocyte membrane studied by fluorescence photobleaching recovery: Evidence for control by cytoskeletal interactions. Pro(,. N u / / .Acud. Sci. U.S.A. 77, 2.537-2.541. Goldsmith, T. H.. and Wehner, R. (1977). Restrictions on rotational and translational diffusion of pigment in the membranes of a rhabdomeric photoreceptor. 1. Gen. Physiol. 70, 453490. Goldstein. B.. Wofsy, C., and Bell, G. (1981). Interactions of low density lipoprotein receptors with coated pits on human fibroblasts: Estimate of the forward rate constant and comparison with the diffusion limits. Proc. N u t / . A c i d . Sci. U . S . A . 78, 5695-5698.

122

MICHAEL EDlDlN

Greinert, R., Finch, S. A. E., and Stier, A. (1982a). Conformation and rotational diffusion of cytochrome P-450 changed by substrate binding. Biosci. Rep. 2, 991-994. Greinert, R., Finch, S. A. E., and Stier, A. (1982b). Cytochrome P-450 rotamers control mixed-function oxygenation in reconstituted membranes. Rotational diffusion studied by delayed fluorescence depolarization. Xenohiotica 12, 7 17-726. Grell, E. (ed.) (1981). “Membrane Spectroscopy.” Springer-Verlag. Berlin, Heidelberg, and New York. Gupte, S., Wu, E.-s., Hoechli, L., Hoechli, M., Jacobson, K., Sowers, A., and Hackenbrock, C. R. (1984). Relationship between lateral diffusion, collision frequency and electron transfer of mitochondria1 inner membrane oxidation-reduction components. Proc. Nutl. Acad. Sci. U.S.A. 81, 2606-2610. Gut, J.. Richter, C., Cherry, R. J., Winterhalter, K., and Kawato. S. (1982). Rotation of cytochrome P-450. 11. Specific interactions of cytochrome P-450 with NADPH-cytochrome P-450 reductase. J. Biol. Chem. 257, 7030-7036. Gut, J., Richter, C., Cherry, R. J., Winterhalter, K.. and Kawato, S. (1983). Rotation of cytochrome P-450. Complex formation of cytochrome P-450with NADPH-cytochrome P-450 reductase in liposomes demonstrated by combining protein rotation with antibodyinduced cross-linking. J. B i d . Chem. 258, 8588-8594. Hanski, E., Rimon, G., and Levitzki, A. (1979). Adenylate cyclase activation by the padrenergic receptors as a diffusion-controlled process. Biochemistry 18, 846-853. Hardt, S. L. (1979). Rates of diffusion controlled reactions in one, two and three dimensions. Biophys. Chem. 10, 239-243. Hare, F. (1983). Simplified derivation of angular order and dynamics of rodlike fluorophores in models and membranes. Simultaneous estimation of the order and fluidity parameters for diphenylhexatriene by only coupling steady-state illumination polarization and lifetime of fluorescence. Biophys. J . 42, 205-218. Henderson, R. (1977). The purple membrane from Hulohucteriurn halohiitm. Annu. Rev. Biophys. Bioeng. 6 , 87. Henis. Y., and Gutman, 0. (1983). Lateral diffusion and patch formation of H-2Kk antigens on mouse spleen lymphocytes. Biochim. Biophys. Acta 762, 281-288. Heyn, M . , Cherry, R. J . , and Dencher, N. (1981). Lipid-protein interactions in bacteriorhodopsin-DML vesicles. Biochemistry 20, 840-849. Hoffman, W., Sarzala. G . , and Chapman, D. (1979). Rotational motion and evidence for oligomeric structures of sarcoplasmic reticulum Ca” -activated ATPase. Proc. Null. Acud. Sci. U.S.A. 76, 3860-3864. Huang, H. W. (1973). Mobility and diffusion in the plane of the cell membrane. J. Thcor. B i d . 40, 11-17. Hyde. J. S. (1981). Saturation-transfer spectroscopy. Methods Enzymol. 49,480-51 1. Jacobson, K., Wu, E.-s., and Poste, G. (1976). Measurement of translational diffusion of concanavalin A in glycerol-saline solutions and in the cell surface by fluorescence recovery after photobleaching. Biochim. Biophys. Acra 433, 215-222. Jacobson, K., Elson, E., Koppel, D., and Webb, W. (1983).International workshop on the application of fluorescence photobleaching techniques to problems in cell biology. Fed. Proc.. Fed. Am. Soc. Exp. B i d . 42, 72-79. Jaffe, L. F. (1977). Electrophoresis along cell membranes. Nature (London) 265, 600-602. Johnson, P., and Garland, P. (1981). Depolarization of fluorescence depletion: A microscopic method for measuring rotational diffusion on the surface of a single cell. FEBS Lett. 132, 252-256. Johnson, P., and Garland, P. B. (1982). Fluorescent triplet probes for measuring the rotational diffusion of membrane proteins. Biochem. J . 203, 313-321.

DIFFUSION O F MEMBRANE PROTEINS AND LIPIDS

123

Johnson. M. E.. and Hyde, J. S. (1981). 35 G H z (()-band) saturation transfer electron paramagnetic resonance studies of rotational diffusion. Biochrttrisrn 20, 2875-2XXO. Johnson. M. E., Lee, L.. and Fung, L. W.-M. (1982).Modelsfor \low anisotropic rotational diffusion in saturation transfer electron paramagnetic resoniince at 9 and 35 G H r . Bioc~/wr?tistr:v 21, 4459-4466. Jovin. T. M.. Bartholdi, M.. Vaz. W . I-. C., and Austin, K . H. (1981). Rotational diffusion o f hiological macromolecules by time-resolved delayed luminescence (phosphorescence, fluorescence) anisotropy. Anrr. N . Y . A<.rrt/.St,;. 366, 176196. Kawato. S., and Kinosita. K. ( 1983). Time-dependent absorption anisotropy and rotational diffusion o f proteins in membranes. Bioph?.~.J . 36, 777-296. Kawato. S . , Gut. J.. Cherry. K. J.. Winterhalter. K . H . . and Kichter, C. (1982a). Kotation o f cytochrome P-450. i. Investigation5 o f protein-protein interactions of cytoJ I . 7077chrome P-450 in phospholipid vesicles and liver microsomes. J . B i d . C / I C . ~257.

7029. Kawato. S.,Lehner, C . . Muller. M.. and Cherry. K. J. (1982b). Protein-protein interactions . 6470o f cylochrome oxidase i n inner mitochondria1 membranes. 1.Aiol. C ' h ~ r n 257, 6476. Kirino. Y..Ohkuma, T., and Shimizu. H. (1978). Saturation transfer electron spin resonance study on the rotational diffusion o f calcium- and magnesium-dependent adenosine triphosphatase in sarcoplasmic reticulum membranes. J . Bioclterri. 84, I 11-1 15. Koch. G . L. E., and Smith. J. J. (1978). A n association between actin and the ma.jor histocompatibility complex antigen H-2. Nutro~,( L o t i t l o t i ) 273, 274. Koppel. I>. E. (1979). Fluorescence redistribution after photobleaching: a new multipoint analysis o f membrane translational diffusion. Biophw. J . 28, 281-291. Koppel. D. E . . and Sheetz. M. P. (1981). Fluorescence photobleaching does not alter the lateral mobility of erythrocyte membrane glycoproleins. Nrrrrire Ilontlow) 293, 159-161, Koppel. D. E., Sheetz. M. P.. and Schindler. M . (1980). Lateral diffusion i n biological niembranes: a normal-mode analysis of diffusion on a spherical surface. Biophys. J . 30, 187-

192. Koppel. D. E., Sheetz. M. P . . and Schindler. M. (1981). Matrix control o f protein diffusion in biological membranes, Proc,. Ntirl. Actrtl. .Sc.i. U . S . A . 78, 3576-3580. Kusumi. A,. and Hyde, J. ( 19x2). Spin label saturation-transfer electron spin resonance detection of transient association o f rhodopsin in reconstituted membranes. Bioc~/irrtri.s/r? 21, 5978-5983. Kusunii. A,. Sakaki. T.. Yoshizawa. T., and Ohnishi. S. ( 1980). Protein-lipid interaction in rhodopsin recombinant membranes as studied by protein rotational mobility and lipid 83, 108-1 I I . alkyl chain flexibility measurements. J . Eioc~hrr~t. Lepock, J. K..Thompson, J. E., Kruuv, J.. and Wallach. 1). F. H. (1978). Photainduced ~t. cross-linking o f membrane proteins by fluorescein isothiocyanate. B i r ~ h ~ rBiophy.~. Hcs. Conirnrin. 85, 344-350. Levitski, A. (1981). The p-adrenergic receptor and it\ mode o f coupling to adenylnte cyclase. CRC' C r i / . Reis. Bioc.lrern. 81-1 12. Liebman. P. A,. and Entine. G. (1974). Lateral diffusion o f visual pigment i n photoreceptor disc membranes. Sc.icrwr 185, 4.57-459. Louvard. D. (1980). Apical membrane aniinopeptidase appears at site of cell-cell contact in cultured epithelial cells. Proc-. Nrifl. Actrtl. .Sci. U.S.A. 77, 4132-4136. Maro, B . . and Aornens. M. (1982). Reorganization of He1.a cell cytoskeleton induced by an uncoupler o f oxidative phosphorylation. N(rtrirr2fLorrdort) 295, 334-336. Marsh, D. (1981). Electron spin resonance: Spin labels. Irr "Membrane Spectroscopy" ( E . Grell. ed.), pp. 51-14?, Springer-Verlag. Berlin.

124

MICHAEL EDlDlN

Muhlebach, T., and Cherry, R. J. (1982). Influence of cholesterol on the rotation and selfassociation of band 3 in the human erythrocyte membrane. Biochmtisfry 21,4225-4228. Muller, M., Krebs, J. J. R., Cherry, R. J., and Kawato, S. (1982). Selective labeling and rotational diffusion of the ADP/ATP translocator in the inner mitochondria1 membrane. J . Biol. Chem. 257, 117-120. Nicolson, G. L. (1976a). Transmembrane control of the receptors on normal and tumor cells. I. Cytoplasmic influence over cell surface components. Biochim. Biophys. Actu 458, 57-108. Nicolson, G. L. (1976b). Transmembrane control of the receptors on normal and tumor cells 11. Surface changes associated with transformation and malignancy. Biochirn. Biophys. A c f a 458, 1-72. Nigg, E. A., and Cherry, R. J. (1979). Dirneric association of band 3 in the erythrocyte membrane demonstrated by protein diffusion measurements. Nature (London) 277, 493494. Nigg, E. A., and Cherry, R. J. (1980). Anchorage of a band 3 population at the erythrocyte cytoplasmic membrane surface: Protein rotational diffusion measurements. Proc. Nutl. Acad. Sci. U.S.A. 77, 4702-4706. Nigg, E. A., Bron, C., Giradet, M., and Cherry, R. (1980). Band 3-glycophorin A association in erythrocyte membranes demonstrated by combining protein diffusion measurements with antibody-induced cross-linking. Biochemistry 19, 1887-1893. Olek, A. J., Pudimat, P. A., and Daniels, M. P. (1983). Direct observation of the rapid aggregation of acetylcholine receptors on identified cultured myotubes after exposure to embryonic brain extract. Cell 34, 255-264. Orida, N., and Poo, M.-m. (1978). Electrophoretic movement and localisation of acetylcholine receptors in the embryonic muscle cell membrane. Nufure (London) 275, 31-35. Orly, J., and Schramm, M. (1976). Coupling of catecholamine receptor from one cell with adenylate cyclase from another cell by cell fusion. Proc. Natl. Acad. Sci. U . S . A . 73, 44 10-44 14. Pasternak, C., Corda, S. H., and Shinitsky, M. (1981). Rotational diffusion of rhodopsin in membranes as determined by steady-state polarized photochromism. Ann. N . Y. Acud. Sci. 366, 265-273. Peters, R. (1981). Translation diffusion in the plasma membrane of single cells as studied by fluorescence microphotolysis. Cell Biol. I n f . Rep. 5, 733-760. Peters, R.. and Cherry, R. J. (1982). Lateral and rotational diffusion of bacteriorhodopsin in lipid bilayers: Experimental test of the Saffman-Delbruck equations. Proc. Natl. Acnd. Sci. U . S . A . 79, 4317-4321. Peters. R., Peters, J., Tews, K. H., and Bahr, W. (1974). A microfluorimetric study of translation diffusion in erythrocyte membranes. Biochirn. Biophys. Acru 367, 282-294. Peters, R., Brunger, A., and Schulten, K. (1981). Continuous fluorescence microphotolysis: A sensitive method for study of diffusion processes in single cells. Pro(.. Nutl. Acad. Sci. U.S.A. 78, 962-966. Petrini, M., Emerson, D., and Galbraith, R. M. (1983). Linkage between surface immunoglobulin and cytoskeleton of B lymphocytes may involve G , protein. Nature (London) 306, 73-74. Pisam, M., and Ripoche, P. (1976). Redistribution of surface macromolecules in dissociated epithelial cells. 1.Cell Biol. 71, 907-920. Poo, M.-m. (1981). In s i f u electrophoresis of membrane components. Annu. R e v . Bioppliys. Bioeng. 10, 245-276. Poo. M.-m. (1982). Rapid lateral diffusion of functional ACh receptors in embryonic muscle cell membranes. Nuture (London) 295, 332-334.

DIFFUSION OF MEMBRANE PROTEINS AND LIPIDS

125

Poo. M.-m., and Cone, R. A. (1974). Lateral diffusion of rhodopsin in the photoreceptor membrane. Nuture (London) 247, 438441. Poo. M.-m., and Robinson, R. R. (1977). Electrophoresis ofconcanavalin A receptors along embryonic muscle cell membranes. Nuictri, (London) 265, 602-605. Poo, M.-m.. Poo. W.-j.. and Lam. J. (1978). Lateral electrophoresis and diffusion of concanavalin A receptors in membranes of embryonic muscle cells. J . Cell B i d . 76, 483501.

Poo. M.-m., Lam, W.-j.. H., Orida, N ., and Chao. A. W. (1979).Electrophoresis and diffusion in the plane of the cell membrane. Biophys. J . 26, 1-21. Poore, V . M.. Fitzsimons. J. T. R., and Ragan. C. I. (1982). The effects of lipid fluidity on rotational diffusion of complex I and complex I1 in reconstituted NADH-cytochrome i’ oxidoreductase. Biochim. Biophvs. Acia 693, 113-124. Popp. C. A., and Hyde. J. S. (1982). Electron-electron double resonance and saturationrecovery studies of nitroxide electron and nuclear spin-lattice relaxation times and Heisenberg exchange rates: Lateral diffusion in dimyristoyl phosphatidylcholine. Pror.. Null. A c i d . Sci. U . S . A . 79, 2559-2563. Quinn. P. (1981). The fluidity of cell membranes and its regulation. Prog. Biophvs. Mol. B i d . 38, 1-104, 198. Richter, C.. Winterhalter, K. H., and Cherry, R. J. (1979). Rotational diffusion of cytochrome P-450 in rat liver microsomes. FEBS Lerr. 102, 151-154. Rodriguez-Boulan, E., and Sabatini, D. D. (1978). Asymmetric budding of viruses in epithelial monolayers: A model system for study of epithelial polarity. Pror. Nail. A i d . Sci. U . S . A . 75, 5071-5075. Rodriguez-Boulan. E., Paskiet, K.. and Sabatini, I>. D.(1983). Assembly of enveloped viruses in Madin-Darby canine kidney cells: Polarized budding from single attached cells and from clusters of cells in suspension. J. CelI Biol. 96, 866874. Rousselet. A., Cartaud, J.. and Devaux. P. F. (1981). Effects of temperature, lipid modification and pH on the mobility of the major proteins of the receptor-rich membranes from Torpedo murmoruiu. Biochirn. Bioplivs. Acru 648, 169-185. Rousselet, A., Cartaud. Jr., Devaux, P.F.. and Changeuax, J.-P. (1982). The rotational diffusion of the acetylcholine receptor in Torpado murmoruiu membrane fragments studied with a eosin-labeled a-toxin: Importance of the 43,000 proteinb). EMBO J. I, 439445. Torpcvfo murmoruiu membrane fragments studied with a rhodopsin-labeled a-toxin: Importance of the 43,000 protein(s). EMBO J. I , 439445. Sackman, E., and Trauble. H. (1972).Studies of crystalline-liquid crystalline phase transitions of lipid model membranes. I . Use of spin labels and optical probes as indicators of phase transitions. J . Am. Chim. Soc. 94, 4482-449 I. Saffman. P. G., and Delbruck, M. (1975). Brownian motion in biological membranes. Proc. Null. A c ~ d Sci. . U . S . A . 72, 3111-3113. Sakaki, T., Tsuji. A,, Chang, C.-H., and Ohnishi. S. (1982).Rotational mobility of an erythrocyte membrane integral protein, band 3, in dimyristoyl phosphatidylcholine reconstituted vesicles and effect of binding cytoskeletal peripheral proteins. Bioclic~mi.sirv21, 2366-237 I. Schindler. M., Koppel, D. E., and Sheetz, M. P. (1980).Modulation of membrane protein lateral mobility by polyphosphates and polyamines. Proc. N u t / . Actrd. Sci. U.S.A. 77, 1457-1461. Schlessinger. J.. Koppel, D. E.. Axelrod, D., Jacobson, K.. Webb, W. W.. and Elson, E. 1,. (1976). Lateral transport on cell membranes: Mobility of concanavalin A receptors on myoblasts. Proc. Null. Acud. Sri. U . S . A . 73, 2409-2413.

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Schlessinger, J . , Barak. L. S . , Hammes, G . G . . Yamada, R. M., Pastan, I . , Webb, W. W., and Elson, E. L. (1977). Mobility and distribution of a cell-surface glycoprotein and its interaction with other membrane components. Proc. Nmfl. Acud. Sci. U . S . A . 74,29092913. Schreiner, G. F.. and Unanue. E. R. (1979). Membrane and cytoplasmic changes in Blymphocytes induced by ligand-surface immunoglobulin interaction. A d v . Immrtnol. 24, 37-165. Schwarz, D., Pirrwitz, J.. Coon, M . J., and Ruckpaul. K. (1982a). Mobility and clusterlike organization of liposomal cytochrome P-450 LM2: Saturation transfer EPR studies. Artu Biol. Med. Germ. 41, 425-430. Schwarz, D., Pirrwitz, J., and Ruckpaul, K. (1982b). Rotational diffusion of cytochrome P450 in the microsomal membrane-Evidence for a clusterlike organization from saturation transfer electron paramagnetic resonance spectroscopy. Arch. Biochem. Biophys. 216, 322-328. Sheetz. M.. and Koppel, D. E. (1979). Membrane damage caused by irradiation of fluorescent concanavalin A. Proc. Nuil. Acad. Sci. U . S . A . 76, 3314. Sheetz, M. P., Schindler, M., and Koppel, D. E. (1980). Lateral mobility of integral membrane proteins is increased in spherocytic erythrocytes. Nature (London) 285, 510-5 12. Small, R., Blank, M., and Pfenninger, K. H . (1982). Diffusion of protein complexes in the plasma membrane of growing axons. J . Cell B i d . 95, 248a. Smith, B.. and McConnell, H . M. (1978). Determination of molecular motion in membranes using periodic pattern photobleaching. Proc. N a f l . Acad. Sci. U . S . A . 75, 2759-2763. Smith, D. K., and Palek, J . (1982). Modulation of lateral mobility of band 3 in the red cell membrane by oxidative cross-linking of spectrin. Nafure (London) 297, 424-425. Smith, B . , Clark, W. R., and McConnell. H . M. (1979). Anisotropic molecular motion on cell surfaces. Proc. Null. Acnd. Sri. U . S . A . 76, 5641-5644. Smith, L. M., Weis, R. M., and McConnell, H. M. (1981). Measurement of rotational motion in membranes using fluorescence recovery after photobleaching. Biophys. J . 36, 7391. Smith, L. M., Petty, H. R., Parham, P., and McConnell, H. M . (1982). Cell surface properties of HLA antigens on Epstein-Barr virus-transformed cell lines. Proc. N a f l . Acud. Sci. U . S . A . 79, 608-612. Sowers, A . E., and Hackenbrock, C. R. (1981). Rate of lateral diffusion of intramembrane particles: Measurement by electrophoretic displacement and rerandomization. Proc. Nail. Acad. Sci. U . S . A . 78, 6246-6250. Steinman, R., Mell.man, 1.. Muller, W., and Cohn, Z. (1983). Endocytosis and the recycling of plasma membranes. J. Cell B i d . 96, 1-27. Strittmatter, P., and Rogers, M. J . (1975). Apparent dependence of interactions between cytochrome b, and cytochrome bSreductase upon translational diffusion in dimyristoyl lecithin liposomes. Proc. Nail. Acad. Sci. U . S . A . 72, 2658-2661. Stryer, L., Thomas. D. D., and Meares, C. F. (1982). Diffusion-enhanced fluorescence energy transfer. Annu. Rev. Biophys. Bioeng. 11, 203-222. Takezoe, H., and Yu, H. (1981). Lateral diffusion of photopigments in photoreceptor disc membranes by dynamic Kerr effect. Biochemisfry 21, 5275-5281. Tank, D. W., Wu, E-S., and Webb, W. W. (1982). Enhanced molecular diffusibility in muscle membrane blebs: release of lateral constraints. J . Cell B i d . 92, 207-212. Taylor, R. B.. Duffus. P. H., Raff, M. C.. and dePetris, S. (1971). Redistribution and pinocytosis of lymphocyte surface immunoglobulin molecules induced by anti-immunoglobulin antibody. Nafure (New Biol.) 233, 225-229. Thomas, D. D., and Hidalgo, C. (1978). Rotational motion of sarcoplasmic reticulum C a t + ATPase. Proc. Nutl. Acad. Sci. U . S . A . 75, 5488-5492.

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Thomas. D. D.. Carlsen. W . F . , and Stryer. L. (1978). Fluorescence energy transfer in the rapid diffusion limit. Proc. N o r / . A(,crd. .Sc.i. O . S . A . 75, 5746-5750. Vanderkooi. J . M.. and Callis, J. B . (19741. Pyrene. A probe o f lateral diffusion in the hydrophobic region of membranes. Biochc~vii.yti:\.13, 4(K)O. van Meer. G . , and Simons, K. (1982). Viruses budding from either the apical or the basolateral plasma membrane domain of MDCK cells have unique phospholipid composttionc. E M U 0 J . I , 847-852. van Zoelen. E . J . J.. Tertoolen. L. G . J.. and de Laat. S . W. (1983). Simple computer method for evaluation of lateral diffusion coefficients from fluorescence photobleaching recovery kinetics. Biopliys. J . 42, 103-108. Vaz. W., Austin. R. H.. and Vogel. H . (1979). The rotational diffusion of cytochrorne h, in lipid bilayer membranes. B i o p h ? ~J. . 26, 415426. V u . W. L. C.. Criado, M., Madeira. V. M. C.. Schoellmann. G . . and Jovin. T. M. (1982). Size dependence of the translational diffusion of large integral membrane proteins in liquid-crystalline phase lipid bilayers. A study using fluorescence recovery after photohleaching. Bioc./imiisrn 21, 5608-561 2. Wagner. R.. Carrillo. N., Junge, W . . and Vallejos. R. H . (1982). On the conformation of reconstituted ferredoxin:NADP' oxidoreductase in the thylakoid membrane. Studies via triplet lifetime and rotational diffusion with eosin isothiocyanate label. L i i o c , / i i r t t . Riop/rys. Ac/tr 680, 3 17-330. Wey. C.-l.. Ahl. P.. Cone, R. A , , and Ciaffney. B . J . (l979j. Membrane viscosity-ii small lipid probe reports the same viscosity iis two integral membrane proteins. Riophys. J . 25, 169a. Wey. C-I,, R. A,. Cone. and Edidin. M. A . (1981). Lateral diffusion of rhodopsin in photoreceptor cells measured by fluorescence photobleaching and recovery. Biopfty.y. J . 33, 225-232. White, R. E.. and Coon. M. J . (1980). Oxygen activation by cytochrome P-450. Annrr. HPI.. Bioc~//cm.49, 3 15-3.56. Wolf. D. E . . Edidin. M. A,. and Drngsten. P. K. (1980). Effect of bleaching light on nieasurements of lateral diffusion in cell membranes by the fluorescence photobleaching recovery method. Proc. Ntirl. Acntl. S c i . U . S . A . 77, 2043-2045. Wu. E.-S.. Tank, D. W . . and Webb, W. W. (1982). Unconstrained lateral diffusion of Con A receptors on bulbous lymphocytes. Pro(.. Nctrl. A(,cid. .%i. U . S . A . 79, 4962-4966. Yang. C. S. ( 1977). Organization and interaction o f monooxygenase enzymes in the microsoma1 membrane. k f i , Sci. 21, 1047-10.57. Zagyansky. Y . , and Edidin, M . (19761. Lateral diffusion ofconcanavalin A receptors in the plasma membrane in mouse fibroblasts. B i o ( ~ / i i v iBiop/iys. . Actci 433, 209-214. Zagyansky, Y ., and Jard. S. (1979). Does lectin-receptor complex formation produce Lone\ of restricted mobility within the membrane? Ntrtirrc f L o n t h r i ) 280, 591-593. ZidovetLki. R . , Yarden, Y.. Schlessinger, J . . and Jovin, T. M. (19x1). Rotational diffusion of epidermal growth factor complexed to cell surface receptors reflect\ rapid niicroaggregation and endocytosis of' occupied receptors. Proc.. Nor/. Accrd. Sci. U . S . A . 78, 698 1-6985.

Ziomek. C. A , . Schulman. S., and Edidin. M. (1980). Redistribution of membrane proteins in imlated mouse intestinal epithelial cells. J . Cell B i d . 86, 849-857,

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C U R R h N T IOPICS IN MkMBRANLS A N D I R A N S P O R T . VOLUME 29

Biosynthesis and Distribution of Lipids K E N N E T H J . LONGMUIR Di,pcirtrnent of Physiology (rnd BiopliysicJ Ctrlifornia College of Medicine Llnit~ersitvof California. Irvine Irvine. Cdifornirr 92717

1.

INTRODUCTION

Membrane biogenesis in eukaryotic cells is an important topic of research in contemporary cell biology. This problem is being successfully investigated largely by studies which explore the intracellular transport of newly synthesized, membrane-bound protein (Sabatini et n l . , 1982). Current models suggest that polypeptides, synthesized on ribosomes in the endoplasmic reticulum, move through the endoplasmic reticulum and Golgi apparatus, where they are sorted and targeted to other regions of the cell such as the plasma membrane, lysosomes, or secretory organelles. A less conclusive picture is available for the synthesis, transport, and targeting of membrane lipids in eukaryotic cells. This lack of information is due partially to the experimental difficulties encountered when attempting to follow subcellular synthesis and transport of lipid. Whereas antibodies and lectins are successfully used to determine the location of intracellular proteins, these methods are not generally available to the lipid biochemist. Interpretation of data obtained by subcellular fractionation methods is often limited by contamination of one subcellular membrane fraction by another. Autoradiography of lipids in cells is difficult unless care is given to the retention of lipid during tixation, dehydration, and embedding procedures. Despite the experimental limitations the subcellular sites of lipid synthesis, the intracellular lipid transport processes, and the targeting of lipids to specific membranes are recognizcd as important topics of current interest to lipid researchers. As a result, considerable data have accumulated in recent years which address these subjects. This article provides an overview of lipid metabolism in mammalian 129 Copyrighi 'C' 1987 hy Academic h e $ \ . Inc.

All righlr 01' repr(iduction

in

any forin rcaeivcd.

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KENNETH J. LONGMUIR

cells from the point of view of subcellular synthesis, transport, and targeting of lipid. It discusses the available data which indicate that different subcellular membranes have, to some extent, different lipid compositions. It discusses the various biosynthetic pathways of fatty acid and phospholipid biosynthesis, with emphasis placed on the subcellular locations of the biosynthetic enzymes. Finally, this review discusses the possible mechanisms responsible for the transport of lipid between subcellular membranes and the mechanisms by which cells produce and maintain membranes of different lipid compositions. This article covers a wide range of topics and is intended as an overview for those not familiar with lipid biosynthesis. Some areas are not considered because they are not pertinent to the subject of membrane biogenesis or because excellent reviews have appeared in recent years. For example, fatty acid biosynthesis has been reviewed recently (Wakil et al., 1983), and only fatty acid desaturation and elongation reactions are considered here. Also, space does not permit discussion of glycolipid biosynthesis, nor of the interesting changes in lipid biosynthesis that occur under hormonal or developmental control. Excellent reviews have appeared on nearly all of the topics in this article, and these are cited in the individual sections. II. THE LIPID COMPOSITION OF SUBCELLULAR MEMBRANES A. Nomenclature 1. FATTY ACIDS

Table I contains the names of the principle fatty acids esterified to the lipids of mammalian tissue. The list is limited to those fatty acids which are present in amounts greater than a few percent in most tissues or which are major products of fatty acid biosynthetic pathways. Because fatty acids have long systematic names, a shorthand notation is used to indicate the number of carbon atoms followed by the number of cis (or Z) double bonds. Hence palmitic acid is 16:O and arachidonic acid 20:4. There are two conventions for designating the locations of double bonds. One numbers from the carboxyl end, the other (the o nomenclature) from the terminal methyl group. Linoleic acid, for example, is designated 18:2 (9,12) when numbering from the carboxyl end, and 18:2 w6,9 when numbering from the terminal methyl group. The w nomenclature is useful because of the nature of fatty acid desaturation and elongation reactions in mammalian tissue. Desaturation occurs between an existing double bond and the carboxyl end. Elongation reactions occur at the carboxyl end. As a result, despite the many com-

131

BIOSYNTHESIS AND DISTRIBUTION OF LIPIDS

TABLE 1 PRINCIPAL FATTYACIDS01 M A M M A L I ATISSUE. N Common name

Systematic name

Myristic Palmitic Stearic Palmitoleic Oleic Linoleic Linolenic Arac hidonic

T,etradecanoic Hexadecnnoic Octadecanoic 9-Hexadecenoic 9-Octadecenoic 9.12-Octadecadienoic

-

4.7.10.13.I6.19-Doco\ahexaenoic

“

9,12.15-Octadecatrienoic 5 . 8 , I I , 14-lcosatetraenoic

Shorthand notation 14:O 16:O IX:0 I 6 : I (9)” I8:I (9) 18:2 (9.12)

18:3 (9,12,15) 20:4 (5.8.1 1.14) 22:6 (4,7,10,13.16,19)

All double bonds are cis (or Z ) and are numbered from carboxyl end.

binations of desaturation and elongation reactions that take place, the number of carbon atoms between the terminal methyl group and the nearest double bond remains constant for a given family of fatty acids. Oleic acid and the products of desaturation and elongation of oleic acid are the (119 ) or w9 fatty acids. Palmitoleic acid and its products are (n-7) or w7 fatty acids. Linoleic acid and arachidonic acid are both w6 or (n-6)fatty acids. Linolenic acid and its products are the w3 or the ( n - 3 ) family of fatty acids. [The principal desaturation and elongation pathways of mammalian tissues are listed in Section I11 (Table 1111.1 Double bonds of polyunsaturated fatty acids are separated by one methylene carbon. Using the w nomenclature, it is necessary to specify only the position of the double bond nearest the terminal methyl group. Using this convention, arachidonic acid can be specified as 20:4 06 (which can be written out as 20:4 w6,9,12, IS).

2. GLYCEKOLIPIDS Glycerolipids share the common feature of the glycerol backbone, which in biological systems becomes stereospecific when esterified with phosphate groups, fatty acids, or both. Their structures are best indicated using the stereospecific numbering system (Hirschmann, 1960; IUPAC-IUB, 1978) which has become widely accepted for the naming of glycerol derivatives of biological interest. With a few exceptions (see Section IV,E,S), glycerolipids are products of sn-glycerol 3-phosphate. The phosphate group is found at s n - 3 , while fatty acids are esterified to sn-1 and sn-2 (Fig. I , A). The particular class of glycerophospholipid is determined by the nature of the polar head group (Fig. 1. B).

132

KENNETH J. LONGMUIR

A sn- 1

sn-2 I

I 0 I II HZC-0-P-X

sn-3

I

0B Phosphatidic a c i d

-OH

Phosphatidylcholine

Phosphatidylethanolamine

coo -

Phosphatidylserine

-0-CH

-CH,

/

~

NH3

OH

OH

Phosphatidylinositol

-OQOHOH

Phosphatidylglycerol

-0-CH,-CH-CHI I

I

OH OH

FIG.I . ( A ) Stereochemistry of sn-glycerol3-phosphate and its glycerophospholipid products. X = polar head group. (B)Names and polar head groups of the principal glycerophospholipids derived from phosphatidic acid.

The parent compound of the glycerolipids is phosphatidic acid. If ambiguities arise, the position of the phosphate group on the glycerol backbone may be indicated (e.g., 3-sn-phosphatidic acid) or it may be specified by using the systematic name, 1,2-diacyl-sn-glycerol 3-phosphate. When the fatty acid composition is known, it may be included (e.g., I-stearoyl2-oleoyl-sn-glycerol 3-phosphate). Glycerophospholipids are named most often as derivatives of phosphatidic acid, hence the terms phosphatidylcholine, phosphatidylethan-

BIOSYNTHESIS AND DISTRIBUTION OF LIPIDS

133

olarnine. phosphatidylserine, phosphatidylinositol, and phosphatidylglycerol. The location of the phosphate group is usually not indicated but for the most part is assumed to be srr-3. When necessary, the position of the phosphate may be indicated (e.g., 3-sii-phosphatidylcholine) or the name may be written out to the known degree of specificity (e.g., 1,2diacyl-sn-glycero-3-phosphocholineor I-palmitoyl-2-linoleoyl-sn-glycero3-phosphocholine). Some glycerolipids in mammalian tissue (the “ether lipids”) contain a hydrocarbon chain attached to the glycerol backbone by an ether linkage at position s n - I , with a fatty acid esterified to position sn-2. If a double bond is not present at position I ’ of the hydrocarbon chain, these are named I-alkylglycerolipids. When the alkyl and acyl chains are known, their names may be included, such as I-hexadecyl-2-oleoyl-stl-glycero-3phosphoethanolamine. A second class of ether lipids are those which have a cis (or Z ) double bond between carbons 1’ and 2’ of the chain at position s t i - I . They are called I-( I ‘-alkenyl)-2-acylglyccrolipids, but are usually referred to by their common name, plcrsmrrlogetzs. Choline plasmalogens are I-( I ’-alkenyl)-2acyl-.s~~-glycero-3-phosphocholines, and ethanolamine plasmalogens are I ( I ’-alkenyl)-2-acyl-sn-glycero-3-phosphoet hanolamines. Finally, the designation 0 indicates that the fatty acid or fatty alcohol is linked via the oxygen atom to the glycerol backbone. The proper designation for phosphatidic acid is therefore I -O-acyl-2-O-acyl-sn-glycerol 3-phosphate. It is permissible to omit the 0 designation. as is usually done.

6. Phospholipid Composition This section provides an overview of the phospholipid and fatty acid compositions of the subcellular membranes of mammalian tissue. Studies of phospholipid composition are dependent, of course, upon successful separation of tissue into the various subcellular fractions in vitro. For mammalian tissue, the best data are for the liver, as the methods for obtaining clean subcellular fractions are more advanced for liver than for other tissues. Prominently cited data of the phospholipid compositions of liver subcellular fractions are the reviews of McMurray and Magee (1972), McMurray (1973), and the data tables of White (1973). Except for the lysosomal membrane, the data compiled by McMurray and Magee (1972) have been adapted here in Table 11. Data for the lysosomal membrane are from the work of Bleistein rr ul. (1980). It must be cautioned that one relies upon the combined data of several investigators to obtain an overview of the compositions of subcellular membranes, even for liver tissue.

TABLE I1 PHOSPHOLIPID COMPOSITION OF SUBCELLULAR MEMBRANES FROM RAT LIVER" Mitochondriab Phospholipid Phosphatidylcholine Sphingomyelin Phosphatidylethanolamine Phosphatidylserine Phosphatid ylinositol Phosphatidylgl ycerol Cardiolipin Phosphatidic acid Lysophosphatidylc holine L ysophosphatid ylethanolamine Bis(monoacylg1ycero)phosphate

Inner membrane

Outer membrane

Nuclear membranes'

Rough endoplasmic reticulumd

Golgi membranesd

Plasma membrane'

Lysosomal membranes'

45.4 2.5 25.3 0.9 5.9 2. I 17.4 0.7

49.7 5.0 23.2 2.2 12.6 2.5 3.4 1.3

61.4 3.2 22.7 3.6 8.6

60.9 3.7 18.6 3.3 8.9

45.3 12.3 17.9 4.2 8.7

34.9 17.7 18.5 9.0 7.3 4.8

41.6 9. I 27.3 -

-

1 1.5

5.9

6.3

9.4

-

4.4 3.3

1.9

1.3 4.0

' Except for lysosomal membrane data, table adapted from McMurray and Magee (1972). Reproduced. with permission, from the Annird Review ofBiochernistry. Vol. 41 0 1972 by Annual Reviews Inc. Values are expressed as the percentage of the total phospholipid in a given fraction. Where no value is indicated, the phospholipid was either not reported or below the limit of detection. McMurray and Dawson (1969). ' Kleinig (1970). Keenan and Morre (1970). ' Ray e t a / . (1969). Bleistein et al. (1980). Unidentified phospholipid amounted to 3.2% of the total.

BIOSYNTHESIS AND DISTRIBUTION OF LIPIDS

135

Variations occur with the purity of subcellular fractions and with the separation techniques used. As a result, strict comparisons among the various columns of data in Table I 1 are difficult. Only general trends should be observed. Those trends that in the opinion of this reviewer seem most clearly accepted are discussed below.

I . NITKOGEN-CONTAINING PHOSP~IOI.IPIDS a. P h o s p h c i t i ~ y l c h o l i n e . Phosphatidylcholine is the most abundant glycerophospholipid of mammalian tissue. It is found in the greatest proportion in nuclear membranes and in endoplasmic reticulum. The current view of membrane biogenesis is that membrane formation begins in the endoplasmic reticulum. Membrane material then proceeds through the Golgi apparatus to the plasma membrane (Sabatini ef a l . , 1982). The percentage of phosphatidylcholine decreases as this transition is followed. in in the order nuclear membrane = endoplasmic reticulum > Golgi apparatus > plasma membrane. An intermediate level of phosphatidylcholine is found in the mitochondria. On a percentage basis, there is less in the mitochondria than in the nuclear membranes and the endoplasmic reticulum but more than in the plasma membrane. h. S p h i n g o m y e l i n . Sphingomyelin (Fig. 2 ) is most abundant in the plasma membrane. It is present up to a few percent in the mitochondria, if at all, as some investigators do not detect sphingomyelin in clean mitochondrial fractions from rat liver (Fujiki el NI., 1982). The sphingomyelin content of subcellular membranes follows the progression nuclear membrane = endoplasmic reticulum < Golgi apparatus < plasma membrane. The relative proportions of phosphatidylcholine and sphingomyelin vary from tissue to tissue. However, it is noted that for most tissues the amount of choline-containing phospholipid (phosphatidylcholine + sphingomyelin) is relatively constant at approximately one-half of the total phospholipid (reviewed by Barenholz and Thompson, 1980). c . Phosphutidylethunolarninr. Phosphatidylethanolamine is the second most abundant phospholipid of cell membranes. It is enriched in the mi-

l

c=o I

R

FIG.2 .

Sphingomyelin.

136

KENNETH J. LONGMUIR

tochondria. Otherwise, it appears to be nearly equally distributed among subcellular membranes. d. Phosphafidylserine. Phosphatidylserine is a minor lipid component of cells, with the exception of brain tissue, where it can amount to 15% of the total lipid (Baranska, 1982). In liver, the proportion of phosphatidylserine is lowest in mitochondria, intermediate in endoplasmic reticulum and Golgi, and highest in the plasma membrane. 2. ACIDICPHOSPHOLIPIDS a. Phosphatidylinosifol. Phosphatidylinositol makes up approximately 5-10% of the phospholipid of liver membranes, with the exception of the inner mitochondria1 membrane, where the phosphatidylinositol content is low. Otherwise, there is little indication of a preferential enrichment of phosphatidylinositol in any one subcellular fraction. b. Phosphatidylslycerol. Phosphatidylglycerol is present up to a few percent in mitochondria, where it is a substrate for cardiolipin biosynthesis. By some reports, it is present in the plasma membrane. Phosphatidylglycerol is present in a strikingly large proportion (up to 10% of the total lipid) in pulmonary surfactant, where it is the second most abundant phospholipid (Sanders, 1982). c. Cardiolipin. Cardiolipin (Fig. 3 ) is found in the mitochondria only. It occurs principally in the inner membrane, where it is approximately 15-20% of the total inner membrane phospholipid (reviewed by Ioannou and Golding, 1979). The amount of cardiolipin normally present in the outer mitochondrial membrane is not clear, as available data are quite variable. d . Bis(monoacy1glycerolhosphate. Bis (monoacylglycero) phosphate (Fig. 4, also called 1ysobisphosphatidic acid) is localized exclusively to the lysosome. Detection of bis(monoacylg1ycero)phosphate in mitochondria appears to result from lysosomal contamination (Bleistein et al., 1980).

::

0 I1

R,-C-0-CH,

H2C-O-C-R4

HC-O-C-R3

I

0-

I

OH

b-

FIG. 3. Cardiolipin (diphosphatidylglycerol).

137

BIOSYNTHESIS AND DISTRIBUTION OF LIPIDS

0

0 II

II

R,-C-0-CH,

H,C-O-C-R,

I

I

HO-CH

HC-OH

0Flcj. 4. Bisf monoacylg1ycero)phosphate (lysobi~phosphatidicacid).

3. CHOLESTEROL

Available data clearly indicate differences in cholesterol content among the various subcellular membranes although numerical data are somewhat variable. The numbers that follow are from the work of Colbeau et af. (1971). The inner mitochondrial membrane has the lowest cholesterol content (less than 0.05 kmol cholesterol per pmol lipid phosphorus). The outer mitochondrial membrane and the microsomes have similar cholesterol contents (-0.1 pmol/ynol). However, when microsomes are fractionated to yield smooth and rough endoplasmic reticulum, it is found that smooth endoplasmic reticulum contains more cholesterol (-0.24.3 p.mol/ pmol) than rough endoplasmic reticulum (-0.06 p m o l / ~ m o l ) Plasma . membrane contains by far the most cholesterol (0.76 pmol/p.mol). 4. GENERAL OBSERVATIONS A few remarks should be made about the various subcellular membranes themselves. The inner and outer nuclear membranes have similar lipid compositions (Virtanen et al., 1977). Smooth and rough endoplasmic reticulum have similar lipid compositions (Glaumann and Dallner, 1968). I n contrast, inner and outer mitochondrial membranes differ in several respects. The outer membrane is characterized by amounts of phosphatidylinositol and phosphatidylserine similar to the endoplasmic reticulum (Cobleau et al., 1971). The inner membrane has less phosphatidylinositol and phosphatidylserine than other membranes, a higher content of phosphatidylethanolamine, and it contains most of the cellular cardiolipin. Most of the trends concerning the subcellular distribution of phospholipids in liver hold true for the subcellular membranes of other tissues (White, 1973). However, a few prominent differences should be noted. Brain tissue contains more phosphatidylserine and sphingomyelin than most other tissues due to enrichment in myelin. Brain tissue also contains

138

KENNETH J. LONGMUIR

a substantial portion of ethanolamine phospholipid as ethanolamine plasmalogen (McMurray, 1973). Kidney tissue is rich in ethanolamine plasmalogen (Yeung and Kuksis, 1974). In heart tissue, a substantial portion of both the choline and ethanolamine phospholipids are present as choline and ethanolamine plasmalogen. Heart tissue contains about 2-fold more cardiolipin than other tissues, owing to the abundance of mitochondria in cardiac muscle (McMurray, 1973). The phospholipid compositions of subcellular membranes of some mammalian cells in culture have been studied also. The best data are for the BHK-21 cell line, and the subcellular distribution of lipids in this cell line closely corresponds to the distribution found in mammalian tissue (Brotherus and Renkonen, 1977). In contrast, there are reports that some cell lines exhibit abnormal subcellular phospholipid compositions. Bergelson et al. (1974) have reported that some hepatoma cell lines contain (1) significant cardiolipin in subcellular membranes other than the mitochondria, (2) an abnormally large percentage of sphingomyelin in the intracellular membranes, and (3) a higher ratio of phosphatidylethanolamine to phosphatidylcholine in the microsomal fraction. Other investigators have not substantiated the presence of cardiolipin in extramitochondrial membranes of the hepatorna cell lines. However, they have confirmed the abnormally high intracellular sphingomyelin content and the slightly altered phosphatidylethanolamine to phosphatidylcholine ratio in the rnicrosomal membranes (Hostetler et al., 1976, 1979). C. Fatty Acid Composition

Most phospholipid molecules contain a saturated fatty acid at position sn-1 and an unsaturated fatty acid at position sn-2. Most triacylglycerol molecules also contain a saturated fatty acid at position sn-1 and an unsaturated fatty acid at sn-2. In liver tissue, position sn-3 of triacylglycerol contains principally unsaturated fatty acid (Akesson, 1969).

The profile of fatty acids found at each position depends on the class of lipid. In rat liver, triacylglycerol contains substantial amounts of palmitate (16:O) at sn-l and oleate [18: l (9)] or linoleate [ 18:2 (9,12)] at sn2. Triacylglycerol contains less stearate ( N O ) and arachidonate [20:4 (5,8,11,14)] than is found in the glycerophospholipids (Akesson, 1969). Phosphatidylethanolamineand phosphatidylcholine contain significant amounts of all the major fatty acids. Palmitate (16:O) and stearate (18:O) are abundant at s n - I , whereas oleate [18:1 (9)], linoleate [18:2 (9,12)], ara c h i d o n a t e [20:4 ( 5 , 8 , 1 1 , 1 4 ) ] , and d o c o s a h e x a e n o a t e [22:6 (4,7,10,13,16,19)] are found at sn-2. Phosphatidylinositol and phosphatidylserine have relatively low contents

BIOSYNTHESIS AND DISTRIBUTION OF LIPIDS

139

of palmitate, oleate, and linoleate. Instead, stearate is usually found at sn-1 and arachidonate at sn-2 (Holub and Kuksis, 1978). Cardiolipin is equally unusual. In mammalian tissue. it contains upward of8094 linoleate (loannou and Golding, 1979). Nonrandom pairings of fatty acids have been reported (reviewed by Holub and Kuksis, 1978). In liver, preferen.tial pairings of palmitate with linoleate and stearate with arachidonate are noted, although these pairings are far from exclusive. Pairing of fatty acids is most evident in phosphatidylinositol, where in liver approximately 80% of the total phosphatidylinositol consists of stearate at position s n - I and arachidonate at position SH-2.

Differences in fatty acid composition among subcellular membranes are less evident than differences among phospholipid classes. For liver tissue, no clear differences i n the fatty acid profiles of phosphatidylcholine, phosphatidylethanolamine, phosphatidylinositol, and phosphatidylserine are found between the mitochondria lipids and those of the endoplasmic reticulum (Colbeau et a / . . 1971; Holub and Kuksis, 1978). However, it is generally agreed that the plasma membrane contains a greater percentage of saturated fatty acid than the intracellular membranes such as the endoplasmic reticulum and the mitochondria (Keenan and Morre, 1970: Cobleau r f d., 1971). Golgi apparatus appears to contain a level of saturated fatty acid between that of the endoplasmic reticulum and the plasma membrane. There are two aspects to this enrichment with saturated fatty acid. First, the major phospholipids (phosphatidylcholine, phosphatidylethanolamine, and phosphatidylserine) contain a higher percentage of saturated fatty acid in the plasma membrane than in the intracellular membranes (Keenan and Morre, 1970). Second, plasma membrane is enriched with sphingomyelin. Sphingomyelin has a higher percentage of saturated fatty acid than most of the other phospholipids (White. 1973). Some tissue-specific differences should be noted. Lung surfactant, a lipoprotein material secreted by a lung alveolar cell, is highly enriched with palmitate at both the sn-1 and sn-2 positions of phosphatidylcholine (Sanders, 1982). It is this saturated phosphatidylcholine which gives surfactant its remarkable surface tension lowering properties. The lipid of brain tissue is also enriched with saturated phosphatidylcholine (though not to the extent found in lung) because of its abundance in myelin. In contrast, lipids of reproductive tissue are notably enriched with ii variety of polyunsaturated fatty acids (Kuksis, 1978). Products of the linoleate series (the 0 6 family of fatty acids) include not only arachidonate [20:4 (S,8,11,14)]. but also 20:3 (8,11,14), 22:4 (7,10,13,16), and 22:5 (4,7,10,13,16). Products of the linolenate series (the m3 family) include not only docosahexaenoic acid [22:6 (4,7,10,13,16,19)] but 20:s

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KENNETH J. LONGMUIR

(5,8,11,14,17)and 225 (7,10,13,16,19)as well. Some polyunsaturated fatty acids of the oleate family [e.g., 22:4 (4,7,10,13)] are also found. The reader is referred to the reviews of Holub and Kuksis (1978) and Kuksis (1978) and to the data tables of White (1973) for thorough compilations of the fatty acid profiles of the various phospholipid classes, the various subcellular membranes, and the various mammalian tissues.

D. Asymmetry of Phospholipids in Subcellular Membranes

The concepts of lipid asymmetry in membranes have been thoroughly reviewed (Op den Kamp, 1979; Etemadi, 1980; Krebs, 1982). These reviews discuss the evidence for the asymmetric distribution of lipids and offer critical appraisals of the methods used for determining lipid asymmetry. The absolute asymmetry found for membrane proteins is not found for membrane lipids. Lipids of all classes can be found on both sides of a membrane. The best data for preferential enrichment of lipid on one side of a membrane are for the erythrocyte. This asymmetry was first demonstrated by chemical labeling studies, when it was discovered that phosphatidylethanolamine and phosphatidylserine are preferentially distributed on the intracellular (cytoplasmic) side of the erythrocyte membrane (Bretscher, 1972a,b; Gordesky and Marinetti, 1973). Studies with phospholipases and sphingomyelinase confirmed the enrichment of phosphatidylethanolamine and phosphatidylserine on the intracellular side and indicated that phosphatidylcholine and sphingomyelin are preferentially distributed on the extracellular side (Verkleij et al., 1973; Kahlenberg et al., Renooij el al., 1976). These results have been confirmed recently by the use of nonspecific phospholipid transfer proteins (Crain and Zilversmit, 1980). Lipid asymmetry is less well established for intracellular membranes. However, consistent data have accumulated for the mitochondrial inner membrane (Crain and Marinetti, 1979; Krebs et al., 1979; Harb et a / . , 1981). These studies have used ( I ) chemical probes for labeling the free amino group of phosphatidylethanolamine, (2) phospholipases, and (3) antibodies to cardiolipin. The results of these experiments indicate that phosphatidylcholine is somewhat enriched on the cytoplasmic side of the inner mitochondrial membrane, whereas phosphatidylethanolamine and cardiolipin are located mostly on the matrix side. An asymmetric distribution of lipids across microsomal membranes has not been conclusively demonstrated, even though microsomal preparations can be obtained which are clearly asymmetric with respect to marker enzymes. The lack of conclusive data may be due to the inherent difficulties

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OF

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with the methods used to demonstrate asymmetry. The impermeant reagents needed to modify one side of the membrane may in fact cross to the other side. Alternatively, the lipids of the endoplasmic reticulum may undergo flip-flop across the membrane during the time course of the experiments used to determine asymmetry. Lipids are synthesized on the cytoplasmic face of the endoplasmic reticulum (Section IV,G), yet newly formed lipid must accumulate on both sides ofthe membrane. Hence, flipflop of lipid from the cytoplasmic side to the lumenal side seems physiologically reasonable. Evidence for rapid movement of lipid across microsomal membranes is supported by "P-NMR experiments (de Kruijff et ul., 1978) which suggest that a portion of the microsomal lipid exhibits isotropic motion on the time scale of the N M R experiment. Such N M R data are consistent with the concept that a small portion of the membrane lipid exists in an arrangement other than a lipid bilayer, where rapid motion in all directions can take place to redistribute lipid from one side to the other. These observations suggest that asymmetry of the endoplasmic reticulum in vivo (and microsomal membranes in vifro)as a static phenomenon may be irrelevant. 111.

DESATURATION AND ELONGATION OF FATTY ACIDS

The principal product of dr mvo fatty acid biosynthesis in mammalian tissue is palmitic acid (16:O). De niwo fatty acid biosynthesis takes place in the cytoplasm in a cycle of reactions catalyzed by the multifunctional fatty acid synthetase complex. The reactions leading to the formation of fatty acid from acetyl-CoA and malonyl-CoA are well understood, and the regulation of the fatty acid synthetase complex has been recently reviewed (Wakil el d.,1983). Fatty acids can be modified ( I ) by desaturation reactions which introduce cis double bonds and (2) by elongation reactions which lengthen the fatty acid from the carboxyl end by two carbons per step. Desaturation of fatty acids in mammalian liver tissue can occur at positions nine, six, five, and four from the carboxyl end. These reactions are catalyzed by separate desaturase enzymes, termed the A", Ah, A', and A4 desaturases (James, 1977). Mammalian cells do not possess desaturase activity which will introduce a double bond beyond position 9 from the carboxyl group. Fatty acids with cis double bonds beyond position 9 are obtained by elongation of existing unsaturated fatty acid or by dietary uptake. Linoleic [ 18:2 (9,12)] and linolenic [18:3 (9,12,15)] are the two principal fatty acids supplied by dietary uptake and are called essential fatty acids. They are required for synthesis of polyunsaturated fatty acids and for prostaglandin formation.

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Fatty acid produced by the fatty acid synthetase complex and fatty acid taken up from the circulation must be converted to the fatty acyl-CoA thioester by the enzyme acyl-CoA synthetase before elongation and desaturation can occur. Desaturase enzymes are membrane bound and are located in the microsomal fraction of the cell. The A9 desaturase system (also called the stearoyl-CoA desaturase) is a complex of three proteins: (1) NADH-cytochrome b, reductase, (2) cytochrome b5,and ( 3 ) the terminal desaturase enzyme, often called the cyanide-sensitive protein. The NADH provides reducing power to perform the oxygen-dependent introduction of the double bond,
+ N A D H + H' + 0, +
+ N A D ' + 2 H,O

Elongation of fatty acids takes place in several steps beginning with condensation of an existing fatty acyl-CoA with malonyl-CoA. The resulting p-keto compound is ( I ) reduced to a secondary alchohol, (2) dehydrated to form a trans a,p unsaturated fatty acid, and ( 3 ) reduced to give the elongated fatty acid. Elongation enzymes are microsomal. The initial condensation reaction is rate limiting (Bernert and Sprecher, 1977). Desaturation and elongation generally follow an ordered sequence which can be summarized as follows (Jeffcoat, 1979):

I . When palmitoyl-CoA (16:O) is the substrate, elongation to stearoylCoA (18:O) is usually the first step. 2. For a saturated fatty acid, the first double bond is inserted at position 9 by the A' desaturase. 3. Subsequent double bonds are inserted between the carboxyl group and the double bond nearest the carboxyl group. 4. For polyunsaturated fatty acids, desaturation alternates with elongat ion. These general constraints on fatty acid modification tend to limit the number of possible reactions carried out in mammalian tissue to a few principal pathways outlined in Table 111. Because elongation of palmitic acid usually occurs before desaturation, products of the palmitoleic acid pathway are not found in large amounts in mammalian tissue. Further metabolism of oleic acid to other products of the oleic acid series normally does not take place but will occur as the result of a dietary deficiency of essential fatty acids. The presence of the final product of the series, 5,8,1 I-icosatrienoic acid [20:3 ( 5 3 , I l)], is diagnostic of an essential fatty acid deficiency and is detectable well before the clinical manifestations of the disease appear (Holman, 1977; Laposta et al., 1982). Essential fatty acid deficiencies are also characterized by elevated levels of palmitoleic [16:1 (9)] and oleic [18:1 (9)] acids.

TABLE 111

MAJORPATHWAYS OF FATTY Arm DESATURATION AND ELONGATION" Palmitoleic acid series (the w7 family) 16:O

I9

palmitic acid

c 16:l (9) palmitoleic acid

E

.

18:l ( 1 1 )

cis-vaccenic acid

Oleic acid series (the w9 family) 16:O

palmitic acid 20:2 (8.1 I )

E

"

Linoleic acid serie4 (the w6 family) A6 18:2 (9.12) linoleic acid

c 18:O stearic acid ,20:3 (5.8.1 I )

18:3 (6.9.12) y-linolenic acid

Linolenic acid series (the w3 family) Ah 1 8 3 (9.17.15) c 18:4 (6.9.12.15) a-linolenic acid 2 : 5 (7,l0.l3,16.19)-~ AJ 2 2 6 (4.7.10.13.16.19) "

AY

l8:l (9) oleic acid

.

Ah

D18:2 (6.9)

-

E

20:3 (8.11.14) homo-y-linolenic acid

I'

,20:4 (5.8.11.14) arachidonic acid

E

c 2 0 : 4 (8.11.14.17)

''

D20:5 (5.8.11.14.17)

Reactions are designated as desaturation (4". Ah. 4'.and Ad. catalyzed by t h e respective desaturases) or elongation ( E l

E

E ~

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KENNETH J. LONGMUIR

The linoleic acid series is the pathway followed for the synthesis of homo--y-linoleic acid [20:3 (8,11,14)] and arachidonic acid [20:4 (5,8,11,14)] which are utilized for prostaglandin biosynthesis. The presence of linoleic acid suppresses desaturation and elongation of oleic acid as well as the desaturation of palmitic acid to palmitoleic acid. The mechanisms of these inhibitory processes are not clear. Partial degradation of fatty acids should not be overlooked as a mechanism that regulates the composition of the fatty acid pool available for lipid biosynthesis. As reviewed by James (1977) and Sprecher (1981). longchain fatty acids can be shortened at the carboxyl end by two carbons per step by a process termed “chain shortening” or “retroconversion.” For example, when rats are fed 22:4 (7,10,13,16), this lipid is converted principally to arachidonic acid [20:4 (5,8,1 I ,14)] then incorporated into lipid. IV. PHOSPHOLIPID BIOSYNTHESIS A. Formation of Phosphatidic Acid

1 . ACYLATION OF W-GLYCEROL 3-PHOSPHATE sn-Glycerol3-phosphate is acylated with fatty acyl-CoA by the enzyme glycerophosphate acyltransferase. sn-Glycerol 3-phosphate is derived from NADH-dependent reduction of dihydroxyacetone phosphate (produced by glycolysis). It is also formed by ATP-dependent phosphorylation of glycerol by glycerokinase and by gluconeogenesis (Hers and Hue, 1983). These latter sources appear to be of quantitative importance only in the liver. Fatty acid is obtained by de novo biosynthesis within the cytosol, or by uptake from the circulation. Fatty acid is first esterified with coenzyme A by the enzyme acyl-CoA synthetase, as only the coenzyme A thioesters of fatty acids serve as substrates for acylation reactions in mammalian cells. In liver, acyl-CoA synthetase is found in both microsoma1 and mitochondria1fractions (reviewed by Bell and Coleman, 1980). The acylation of sn-glycerol 3-phosphate occurs at position sn-1 (Yamashita and Numa, 1972; Yamashita et al., 1972, 1973), producing I-acylsn-glycerol3-phosphate. Acylation of 1-acyl-sn-glycerol3-phosphatewith a second fatty acyl-CoA by the enzyme I-acylglycerophosphate acyltransferase produces phosphatidic acid. There is good evidence that the two acylation steps are functions of different enzymes. In rat liver, the two activities can be partially resolved in vitro (Yamashita and Numa, 19721, and they respond differently to a variety of inhibitors. Glycerophosphate acyltransferase activity is found both in microsomal

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145

and mitochondrial compartments. The microsomal and mitochondrial activities appear to arise from separate enzymes on the basis of a number of kinetic observations, including heat stability, pH optima, and sensitivity to detergents, proteases, and organic solvents (Bell and Coleman, 1980). The microsomal activity is sensitive to N-ethylmaleimide, whereas the mitochondrial activity is not (Coleman and Bell, 1980). In liver, glycerophosphate acyltransferase activity is distributed evenly between mitochondrial and microsomal compartments (Haldar et ul., 1979). In other tissues, such as adipose, heart, kidney, brain, and adrenal tissue, and in some cultured cells, the microsomal activity is several times that found in mitochondria (Haldar er al., 1979; Fitzpatrick et al., 1982). suggesting that in general the microsomal acylation reactions have the responsibility for producing physiologically important amounts of phosphatidic acid for glycerolipid biosynthesis. 2. ACYLATION OF DIHYDKOXYACETONE PHOSPHAI'E An alternate pathway for formation of phosphatidic acid begins with the acylation of dihydroxyacetone phosphate to produce I-acyldihydroxyacetone phosphate. The activity of the enzyme which catalyzes this reaction, dihydroxyacetone-phosphate acyltransferase, is found in the microsomal and peroxisomal compartments of the cell. Based upon sensitivity to inhibitors, heating, and proteases. it appears that the enzymes in the two fractions are different. In the microsomal fraction, it is likely that a single enzyme is responsible for acylation of both dihydroxyacetone phosphate and sn-glycerol3-phosphate. This conclusion is based on exhaustive kinetic analyses which find that the two microsomal enzyme activities are equally sensitive to pH, acyl chain length, heating, proteases, detergents, and inhibitors (Schlossman and Bell, 1977). During differentiation of the 3T3-LI preadipocyte cell line, the glycerophosphate and dihydroxyacetone-phosphate acyltransferase activities increase in a coordinate fashion (Coleman and Bell, 1980). In fetal rat, both acyltransferase activities are virtually undetectable, then the activities increase together more than 70-fold postpartum (Coleman and Haynes, 1983). These coordinate changes in both microsomal activities suggest the developmental regulation of a single enzyme. After 1-acyldihydroxyacetone phosphate is formed, it is converted to I-acyl-sn-glycerol 3-phosphate by an NADPH-dependent oxidoreductase. The I-acyl-sn-glycerol 3-phosphate is then acylated to form phosphatidic acid. I-Acylglycerophosphate acyltransferase activity is found in microsomes, but its existence in mitochondrial fractions is unclear (Bell and Coleman, 1980).

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KENNETH J. LONGMUIR

Despite numerous experiments by many investigators, the relative importance of the microsomal dihydroxyacetone phosphate and the sn-glycerol 3-phosphate acylation pathways remains controversial. The experimental approach normally taken to distinguish between the two routes is to incubate tissue slices or cells with ['4C]glycerol plus [2-3H]glyceroland to follow incorporation of label into lipid. Products of the acylation of snglycerol 3-phosphate contain both I4C and 'H label, whereas the 'H label is lost upon oxidation of sn-glycerol 3-phosphate to dihydroxyacetone phosphate. Alternatively, tissues can be incubated with [3H]NADPHand 13H]NADHbecause reduction of dihydroxyacetone phosphate to sn-glycerol 3-phosphate is NADH dependent, whereas reduction of l-acyldihydroxyacetone phosphate requires NADPH (Agranoff and Hajra, 197I). Interpretation of experimental data is complicated by the fact that oxidation of [2-.'H] glycerol exhibits a kinetic isotope effect which discriminates against tritium, hence some investigators have concluded that the dihydroxyacetone phosphate pathway is unimportant. When the isotope effect is taken into account, other investigators have reached the conclusion that acylation via the dihydroxyacetone phosphate pathway in liver (Manning and Brindley, 1972; Rognstad ef al., 1974) and in lung (Mason, 1978) accounts for approximately one-half of the cellular phosphatidic acid biosynthesis. Specificity of the acyltransferases for various acyl-CoA's is discussed in Section 1V.F. B. Formation of the Ether Bond

The committed step in ether lipid biosynthesis is the formation of 1alkyldihydroxyacetone phosphate from 1-acyldihydroxyacetone phosphate. In this reaction, the fatty acid esterified to dihydroxyacetone phosphate is replaced with a fatty alcohol. The enzyme which catalyzes the reaction, alkyldihydroxyacetone-phosphatesynthase, is best characterized in Ehrlich ascites cells (Brown and Snyder, 1983). It apparently catalyzes only the exchange of the acyl group with the fatty alcohol. N o hydrolysis of the acyl group (to give dihydroxyacetone phosphate) is detected when preparations of partially purified enzyme are used. Alkyldihydroxyacetone-phosphate synthase shows little selectivity toward long-chain fatty alcohols. Instead, control of chain specificity probably resides with the oxidoreductase that reduces fatty acyl-CoA thioesters to the corresponding alcohols. In rat brain (Bishop and Hajra, 1981) this enzyme effectively utilizes palmitoyl- (16:0), stearoyl- (18:0), and oleoyl(18: I) CoA's as substrates, and these alkyl chains are the predominant ones found in ether lipids in mammalian tissues. Shorter acyl-CoA's and the polyunsaturated acyl-CoA's are not effective substrates.

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Alkyldihydroxyacetone-phosphatesynthase is a membrane-bound enzyme, but its subcellular location in mammalian cells in not clear (reviewed by Hajra, 1983). For example, in Ehrlich ascites tumor cells and in brain tissue, enzyme activity is found principally in the microsomal fraction. In guinea pig liver, the enzyme is enriched in the mitochondrial fraction. More extensive subcellular fractionation procedures have partially resolved this ambiguity. Hajra and Bishop (1982) have shown that the enzyme in guinea pig liver is largely associated with the peroxisomes. Recent clinical observations support a peroxisomal location for at least one of the enzymes of ether lipid biosynthesis. Infants withZellweger syndrome, a genetic disorder where peroxisomes are found in low levels in cells, have low levels of plasmalogens (Heymans r t n l . , 1983). Alkyldihydroxyacetone phosphate is metabolized by the enzymes of lipid biosynthesis in the same manner as acyldihydroxyacetone phosphate. Following ether bond formation, the I-alkyldihydroxyacetone phosphate is reduced by an NADPH-dependent oxidoreductase. This enzyme can be found in microsomal and mitochondrial fractions, and, as with the other enzymes of ether lipid biosynthesis, it appears to be enriched in peroxisomes from guinea pig liver. The product of the reduction, I-alkyl-snglycerol 3-phosphate, is acylated at position sn-2 to form the I-alkyl analog of phosphatidic acid. The acyltransferase enzyme from rat brain has been studied and appears to preferentially utilize unsaturated acyl-CoA thioesters (Fleming and Hajra, 1977). C. The Dlacylglycerol Pathway 1. HYDROLYSIS OF PHOSPHATIDIC ACTD

The committed step for the synthesis of nitrogen-containing phospholipids and triacylglycerol is the hydrolysis of phosphatidic acid to diacylglycerol by the enzyme phosphatidate phosphohydrolase. This enzyme is specific principally for phosphatidic acid. but it will also hydrolyze I acyl-sn-glycerol3-phosphate(Bleasdale rr d., 19781, as well as phosphate esters of fatty alcohols (Agranoff, 1962). Phosphatidate phosphohydrolase is distinct from other phosphatases such as acid phosphatase. alkaline phosphatase, and phospholipase C. Phosphatidate phosphohydrolase activity is found in virtually every subcellular compartment of the cell. In rat liver, it is reported to have nearly equal specific activity in endoplasmic reticulum. Golgi. plasma membrane, mitochondria, and cytosol (Jelsema and Morre. 19781, and it is present in lysosomes. However, it should be noted that the specific activities of the soluble and particulate fractions vary depending upon the assay conditions (reviewed by Bleasdale and Johnston, 1982).The reason

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KENNETH J. LONGMUIR

for the widespread appearance of the enzyme is not apparent. Conversion of diacylglycerol to end-products of lipid metabolism is carried out by microsomal enzymes. The fates of the diacylglycerols produced by phosphatidate phosphohydrolase in other membranes remain to be established. 2. SYNTHESIS OF PHOSPHATIDYLCHOLINE AND PHOSPHATIDY LETHANOLAMINE u . Choline PhosphatP und Ethunolumine Phosphate. Choline and ethanolamine are phosphorylated to form choline phosphate and ethanolamine phosphate, respectively. These reactions, catalyzed by the enzymes choline kinase and ethanolamine kinase, are dependent upon ATP and Mg” Most but not all investigators report that the choline and ethanolamine kinases are separate enzymes (reviewed by Ulane, 1982) and that there are several isoenzymes of each kinase (Brophy et al., 1977). Both kinase activities are cytosolic. h. CDPcholine arid CDPethunolumine. Choline phosphate and ethanolamine phosphate combine with CTP to form CDPcholine and CDPethanolamine. Although the enzymes are referred to by various names, they are usually termed the choline-phosphate cytidylyltransferase and ethanolamine-phosphate cytidylyltransferase. The two enzymes are separate, as purified choline-phosphate cytidylyltransferase does not contain any ethanolamine-phosphate cytidylyltransferase activity (Choy et ul., 1977) and vice versa (Sundler, 1975). The choline-phosphate cytidylyltransferase exists in two forms: a cytosolic form and an aggregated, particulate form which can be induced by the interaction of the cytosolic form with lipids (Choy and Vance, 1978). It is the aggregated form of the enzyme that has the higher specific activity. The formation of CDPethanolamine and CDPcholine appears to be the rate-limiting step for the synthesis of phosphatidylethanolamine (Sundler and Akesson, 1975) and phosphatidylcholine (Vance and Choy, 1979). Because of its potential role as a regulatory enzyme of lipid biosynthesis, the choline-phosphate cytidylyltransferase in particular has received considerable attention. Current experimental data indicate that a redistribution of the choline-phosphate cytidylyltransferase between cytosolic and membrane-bound compartments is a mechanism by which a cell can regulate the level of phosphatidylcholine biosynthesis. In these experiments, cultured cells are partially depleted of phosphatidylcholine by treatment with phospholipase D (Sleight and Kent, 1983a) or by substitution of choline analogs for choline in the culture medium (Sleight and Kent, 1983~). When these treatments are carried out, phosphatidylcholine biosynthesis is enhanced. This stimulation of phosphatidylcholine biosynthesis appears

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due to a shift in cytidylyltransferdseactivity away from the cytosolic form of the enzyme to the membrane-bound (particulate) form (the more active form of the enzyme) (Sleight and Kent, 1983b). This transition results in an increase in the overall choline-phosphate cytidylyltransferase activity without the need for new protein biosynthesis. The activities of the other enzymes of phosphatidylcholine biosynthesis remain unchanged (Sleight and Kent, 1983b). c . Phosphcitidyk.tiolinc. cincl Phos~~h~iri~~l~~tl~crnolalliin~~. Phosphatidylcholine and phosphatidylethanolamineare formed by the combination of diacylglycerol with CDPcholine and CDPethanolarnine. The enzyme activities which catalyze these reactions, cholinephosphotransferase and ethanolaminephosphotransferase,were believed to be the dual functions of a single enzyme (van den Bosch, 1974). This conclusion appears incorrect. The enzyme activities differ in their specificity for diacylglycerol and in their sensitivity to detergents, phospholipase, protease, and metal ion (reviewed by Bell and Coleman, 1980). Both enzyme activities are exclusively microsomal (Jelsema and Morre, 1978). d . Erhunolclmine Plasmdogen. Ethanolamine plasmalogen is obtained by insertion of a 2 double bond at the I ' position of the alkyl chain of 1alkyl-2-acyl-sn-glycero-3-phosphoethanolamine (Wykle and Schremmer, 1979). The reaction is catalyzed by a microsomal A ' desaturase enzyme complex similar to the desaturase systems that insert double bonds on fatty acyl-CoA's (Section 111). I n vitro studies indicate that A' desaturation of the 1-alkyl group of 1alkyl-2-acyl-sn-glycero-3-phosphoethanolamineis not influenced by the class of acyl chain found at the sn-2 position (Wykle and Schremmer, 1979). Consistent with this observation, it is found that the fatty acid profiles of I-alkyl-2-acyl-sn-glycero-3-phosphoethanolamines and the ethanolamine plasmalogens are similar (Wood and Snyder, 1969). Desaturation of the I-alkyl chain has been shown only for the ethanolamine ether lipids. Hence it is likely that choline, serine, and inositol plasmalogens, as well as I-( 1'-alkenyl)diacylglycerols,are all derived from ethanolamine plasmalogen. Various pathways to form these other classes of plasmalogens can be followed (reviewed by Paltauf, 1983). First, l-(1'alkenyl)-2-acylglycerolcan be formed by action of phospholipase C or by reversal of the reaction catalyzed by et hanolaminephosphotransferase. The resulting ether analog of diacylglycerol can then be utilized for the formation of I-( 1 '-alkenyl)-2,3-diacylglycerolby an acyltransferase reaction or for the production of choline plasmalogen by the action of cholinephosphotransferase. Serine plasmalogen can be formed by base exchange with the ethanolamine plasmalogen (Section IV,D,2). Finally, ethanolamine plasmalogen can be methylated to form choline plasmalogen. This

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KENNETH J. LONGMUIR

methyltransferase activity (Section IV,D,I) has been shown t o exist in brain tissue (Mozzi et al., 1981) and heart tissue (Mogelson and Sobel, 1981).

3. SYNTHESIS OF TRIACYLGLYCEROL The final step of the synthesis of triacylglycerol is the acylation of diacylglycerol with fatty acyl-CoA by the enzyme diacylglycerol acyltransferase. The enzyme is found only in microsomes, and it is clearly distinct from the enzymes glycerophosphdte acyltransferdse and I-acylglycerophosphate acyltransferase (Yamashita et al., 1981). Unlike those acyltransferases which catalyze the formation of phosphatidic acid, the diacylglycerol acyltransferase utilizes a broad range of fatty acyl-CoA substrates, both saturated and unsaturated (Hosaka et al., 1977). D. lnterconversions of the Nitrogen-Containing Phospholipids 1. METHYLATION OF PHOSPHATIDYLETHANOLAMINE

Phosphatidylcholine can be formed by sequential methylation of phosphatidylethanolamine by transfer of the methyl groups from 3 equivalents of S-adenosyl-L-methionine. In mammalian tissues, this pathway is of importance only in liver where, by one report, the phosphatidylethanolamine methyltransferase activity accounts for approximately one-quarter of the total phosphatidylcholine biosynthesis (Sundler and Akesson, 1975). Methylation activity is low in other tissues. The enzyme activities for the three methylation steps in liver are located in the microsomal fraction. It is not known conclusively whether a single enzyme catalyzes all three methylation steps (Audubert and Vance, 1983). The first methylation reaction appears to be rate limiting. Intermediate products of the pathway, phosphatidylmonomethylethanolamineand phosphatidyldimethylethanolamine,are found in trace amounts. However, these two lipids can be introduced in substantial quantities into the membranes of cultured cells which lack methylation activity (Schroeder et al., 1976; Sleight and Kent, 1983~).This is done by substituting monomethylethanolamine and dimethylethanolamine for choline in the culture medium. Incorporation of these analogs into phospholipid occurs via the cytidylyltransferase pathways. 2. SYNTHESIS OF PHOSPHATIDYLSERINE

In eukaryotes, the synthesis of phosphatidylserine occurs predominantly, if not exclusively, by base exchange of serine with phosphatidylethanolamine (reviewed by Baranska, 1982). This base exchange activity

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is energy independent, is Ca” dependent, and, when assayed in vitro, is optimal at a slightly alkaline pH. The enzyme activity responsible for this interconversion is located in the microsomal fraction (Dennis and Kennedy, 1972; van Golde i’t ul., 1974). The enzyme from rat brain has been partially purified (Taki and Kanfer, 19781, and it was found that the most effective acceptors for serine are phosphatidylethanolamine and ethanolamine plasmalogen. Phosphatidylcholine, phosphatidic acid, phosphatidylinositol, and phosphatidylserine are ineffective as acceptors of serine. It should also be noted that serine base-exchange activity is independent of phospholipase D activity. At least in rat brain tissue. the two activities can be separated in vitro (Taki and Kanfer, 1978, 1979). There is a report that phosphatidylserine can be formed in rat liver by the combination of CDPdiacylglycerol with serine (Jelsema and Morrc, 1978). which is the principal route of phosphatidylserine biosynthesis in bacteria (Raetz, 1978). However, the activity is quite low, and it is unlikely that this route is of significance in mammalian tissue. There are also reports (reviewed by Baranska, 1982) o f a mitochondria1 enzyme activity that catalyzes the ATP-dependent, CMP-stimulated incorporation of serine into lipid to form phosphatidylserine. The substrate for this reaction has not been characterized. This activity is also low, and it does not appear that this pathway has physiologic significance. Separate base-exchange enzymes are also present in microsomes that catalyze the exchange of ethanolamine and choline into glycerophospholipid to produce phosphatidylethanolamine and phosphatidylcholine (reviewed by Kanfer, 1980). However, unlike the serine base-exchange enzyme which is solely responsible for phosphatidylserine biosynthesis, it has not been established that the choline and ethanolarnine base-exchange activities have a major role in phosphatidylcholine and phosphatidylethanolamine biosynthesis.

3. DECARBOXYLATION OF PHOSPHA’TIDYLSEKINE Phosphatidylserine is converted to phosphatidylethanolamine by the enzyme phosphatidylserine decarboxylase. In mammalian tissue, the enzyme is located in the mitochondria (Dennis and Kennedy, 19721, specifically in the inner mitochondrial membrane (van Golde et ul., 1974). Newly synthesized phosphatidylserine must therefore be transported from the microsomal compartments to the mitochondria to participate as a substrate for this reaction. Phosphatidylserine decarboxylase is found in bacteria, where it is the major enzyme activity responsible for the formation of phosphatidylethanolamine (Raetz, 1978). Its role in eukaryotic cells is less clear. The presence of the enzyme in mitochondria suggests that it is a source of

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KENNETH J. LONGMUIR

some mitochondria1 phosphatidylethanolamine,but the proportion of phosphatidylethanolamine derived from this pathway remains to be established. 4.

SYNTHESIS OF SPHINGOMYELIN

Until relatively recently, it was concluded that sphingomyelin was formed by the transfer of phosphocholine from CDPcholine to ceramide (Sribney and Kennedy, 1958). Data indicate that this pathway may not be important for sphingomyelin biosynthesis. Instead, most sphingomyelin biosynthesis may result from the enzyme-catalyzed transfer of a phosphocholine group from phosphatidylcholine to ceramide (Bernert and UI1man, 1981; Marggraf et al., 1981; Voelker and Kennedy, 1982). This enzyme activity is energy independent (Voelker and Kennedy, 1982). In rat liver (Voelker and Kennedy, 1982) and in transformed mouse fibroblasts (Marggraf et al., 19811, the enzyme activity that catalyzes this transfer of phosphocholine appears to be localized to those subcellular membranes which have significant 5'-nucleotidase activity (plasma membrane and Golgi apparatus). Less activity is found in the endoplasmic reticulum. The presence of this transferase activity in plasma membranes is consistent with the finding that plasma membranes, compared to other subcellular membranes, are enriched with sphingomyelin and partially depleted of phosphatidylcholine. E. The CDPdiacylglycerol Pathway 1. SYNTHESIS OF CDPDIACYLGLYCEROL

CDPdiacylglycerol (also called CDPdiglyceride or CMPphosphatidate) is formed by the combination of phosphatidic acid with CTP. The enzyme phosphatidate cytidylyltransferase is found both in microsomal and in mitochondrial fractions. The microsomal fraction contains the greater specific activity. The enzymes phosphatidate phosphohydrolase and phosphatidate cytidylyltransferase reside at an important branchpoint in the biosynthesis of glycerolipids in eukaryotic cells. Formation of diacylglycerol is the committed step for triacylglycerol biosynthesis and for the biosynthesis of phospholipids that contain nitrogen. Formation of CDPdiacylglycerol from phosphatidic acid is the first step leading to the formation of the acidic phospholipids phosphatidylinositol, phosphatid ylglycerol, cardiolipin, and bis(monoacylg1ycero)phosphate. The majority of phosphatidic acid formed de novo is no doubt committed to the diacylglycerol pathway, because approximately 10% or less of the

BIOSYNTHESIS AND DISTRIBUTION OF LIPIDS

153

total glycerolipids of the cell are products of the CDPdiacylglycerol pathway. However, the question of the relative proportions of phosphatidic acid that flow into the two pathways is a complicated one. Cells may contain pyrophosphatases that hydrolyze CDPdiacylglycerol. Such enzyme activity has been demonstrated in bacteria (Raetz et a / . , 1976) and in mammalian brain tissue (Rittenhouse et a l. , 1981). Phosphatidylinositol, a major product of the CDPdiacylglcyerol pathway in most cells, undergoes turnover far more rapidly than other end-products of glycerolipid metabolism (Nishizuka, 1984). Cardiolipin biosynthesis requires two molecules of CDPdiacylglycerol to form one molecule of cardiolipin (Section IV,E,4). The relative proportions of phosphatidic acid committed to the two pathways has been studied in microsomes in vitro by generating phosphatidic acid by phospholipase D hydrolysis, then allowing diacylglycerol or CDPdiacylglycerol biosynthesis to take place (van Heusden and van den Bosch, 1978). Over a wide range of microsomal phosphatidic acid concentrations, an approximate 3: 1 ratio of diacylglycerol to CDPdiacylglycerol synthesis is found. However, it should be noted that to assay the maximal phosphatidate cytidylyltransferase activity in vitro, nonphysiologic incubation buffer containing 40 mM Mg’+ is used. As pointed out by Sturton and Brindley (l977), the relative flux of phosphatidic acid into the two pathways is in fact a function of Mg” concentration, at least in in v i m experiments. Most investigators conclude that the enzyme phosphatidate cytidylyltransferaseoperates in vivo well below maximal activity. The flow of phosphatidic acid into the CDPdiacylglycerol pathway can be enhanced by introducing a variety of amphiphilic cations into assay systems of intact cells (Allan and Michell, 1975) or isolated microsomes (Sturton and Brindley, 1977). Examples of such cationic compounds include chlorpromazine, fenfluramine. and propranolol (Sturton and Brindley, 1977). These agents all decrease the rate of biosynthesis of triacylglycerol, phosphatidylcholine, and phosphatidylethanolamine and result in an accumulation of the acidic phospholipids. Using rat liver microsomes, these amphiphilic cations have been shown to inhibit the activity of phosphatidate phosphohydrolase (Brindley and Bowley, 1975). At the same time, they appear to directly stimulate the activity of phosphatidate cytidylyltransferase at physiologic concentrations of magnesium ion (Sturton and Brindley, 1977). 2. BIOSYNTHESIS OF PHOSPHATIDYLINOSITOL Phosphatidylinositol biosynthesis occurs by the reaction of CDPdiacylglycerol with myoinositol to form phosphatidylinositol and CMP. There is little doubt that the enzyme CDPdiacylglycerol-inositol phosphatidyl-

154

KENNETH J. LONGMUIR

transferase resides only in the microsomal fraction (Bell and Coleman, 1980; Jelsema and Morre, 1978). Measurement of the net synthesis of phosphatidylinositol is complicated by other enzyme-catalyzed reactions involving this lipid. The synthesis of phosphatidylinositol itself is a reversible reaction. Addition of CMP to microsomes in v i m will enhance the formation of CDPdiacylglycerol from phosphatidylinositol and CMP (Hokin-Neaverson et al., 1977; Bleasdale et al., 1979). Second, microsomes from several tissues possess a Mg’*dependent enzyme activity that catalyzes the exchange of free n?yo-inositol to phosphatidylinositol (Takenawa and Egawa, 1980; Bleasdale and Wallis, 1981). Finally, phosphatidylinositol participates in a series of reactions, known as the phosphatidylinositol cycle, which involves the metabolism of phosphatidylinositol first to diacylglycerol, then to phosphatidic acid, then back to phosphatidylinositol via CDPdiacylglycerol (Nishizuka, 1984). 3. BIOSYNTHESIS OF PHOSPHATIDYLGLYCEROL Biosynthesis of phosphatidylglycerol from CDPdiacylglycerol takes place in two steps: ( I ) combination of CDPdiacylglycerol with sn-glycerol 3-phosphate to form phosphatidylglycerol3-phosphate,and (2) hydrolysis of phosphatidylglycerol 3-phosphate to form phosphatidylglycerol. The glycerol head group is therefore attached to the phosphate at position snI . In mammalian liver, these activities appear to reside principally in the inner mitochondria1 membrane (Hostetler and van den Bosch, 1972), where phosphatidylglycerol serves as a substrate for cardiolipin biosynthesis. Both enzymes, the glycerophosphate phosphatidyltransferase and the phosphatidylglycerophosphatase, have been partially purified from rat liver mitochondria (McMurray and Jarvis, 1978; MacDonald and McMurray, I 980). The separate cellular locations of phosphatidylglycerol and phosphatidylinositol biosynthesis suggest that these two lipids are formed from separate pools of CDPdiacylglycerol. CDPdiacylglycerol is synthesized in both microsomal and mitochondrial fractions. Phosphatidylinositol biosynthesis is a microsomal function. Phosphatidylglycerol biosynthesis is a mitochondria1 function. However, experimental data are available suggesting that in some systems the enzymes of phosphatidylinositol and phosphatidylglycerol biosynthesis compete for a common pool of CDPdiacylglycerol. In pulmonary surfactant, phosphatidylglycerol is the major lipid product of the CDPdiacylglycerol pathway. Infusion of myo-inositol in vivo (Hallman and Epstein, 1980) or addition of myo-inositol to tissue in culture (Longmuir et al., 1982) effectively suppresses phosphatidylglycerol biosynthesis and enhances phosphatidylinositol biosynthesis. A similar inhibition of phosphatidylglycerol biosynthesis by myo-inositol has

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155

been observed in cells that under various conditions have been made to accumulate phosphatidylglycerol. These include pancreatic islet cells (Freinkel et d.,19751, pineal gland (Eichberg et u/., 19791, and a mutant Chinese hamster ovary cell line defective in myo-inositol biosynthesis (Esko and Raetz, 1980). It has not been resolved whether this apparent competition for the available substrate is the result of an equilibrium bctween microsomal or mitochondria1 CDPdiacylglycerol pools, or whether the microsomal compartments posse\s some phosphatidylglycerol biosynthetic capability. 4. BIOSYNTHESIS OF CARDIOLIPIN Cardiolipin (also called diphosphatidylgl ycerol) is formed by the combination of phosphatidylglycerol with CDPdiacylglycerol. This enzyme activity is exclusively mitochondria1 and is localized to the inner membrane (Hostetler and van den Bosch, 1972). The enzyme has been partially purified from rat liver mitochondria (McMurray and Jarvis, 1980) and is found to have an unusual metal ion requirement in the order Co” > Mn” > Mg’+. It is also found that the enzyme does not show a strong preference for phosphatidylglycerol and CDPdiacylglycerol substrates with particular fatty acid compositions (Hostetler Pt ti/., 1975). Hence the observation that cardiolipin in vivo contains mostly linoleate [18:2 (9,12)] cannot be explained on the basis of the selectivity of the cardiolipin synthase enzyme. 5 . BIOSYNTHESIS OF BlS(M0NOACYLC;LYCEl~O)PHOSPHA‘~E

Bis(monoacy1glycero)phosphate (also called lysobisphosphatidic acid) is an unusual lipid found only in lysosomes. A distinctive feature of this glycerolipid is its unique stereoconfiguration. I t has a backbone of ,sn-Iglycerophospho-sn-I ‘-glycerol rather than the sn-3-glycerophosphate backbone found in other lipids (Joutti or ul., 1976). The biosynthesis of bis(monoacylg1ycero)phosphate takes place in lysosomes. In vitro experiments using lysosomes isolated from rat liver have shown that the lipid can be synthesized from either cardiolipin or phosphatidylglycerol (Poorthuis and Hostetler, 1978). In early studies, it was proposed that bis(monoacylg1ycero)phosphate obtained from phosphatidylglycerol was produced by a deacylation of a fatty acid from phosphatidylglycerol and an acylation of the glycerol head group (Matsuzawa r t ( I / . , 1978). Fatty acyl-CoA is not a substrate for the acylation reaction. Instead, phosphatidylinositol is the most suitable donor of the acyl chain. Other experiments, using liver lysosomes in v i m , have found that the bis(monoacylg1ycero)phosphate product has the same S Y I - I -glycerophospho-sn- 1 ’-glycerol backbone stereoconfiguration that is found irr v i w

156

KENNETH J. LONGMUIR

(Joutti and Renkonen, 1979). Since both the phosphatidylglycerol and cardiolipin substrates have sn-3-glycerophosphate configurations,some rearrangement of the backbone must take place. The individual steps of this rearrangement have not been established.

F. Fatty Acid Specificity and Deacylation-Reacylation Cycles Glycerolipids contain mostly saturated fatty acids at position sn-I and unsaturated fatty acids at position sn-2 (see Section 11,C). It appears that this specificity is introduced into the glycerol backbone at the level of phosphatidic acid biosynthesis. However, after the end-products of lipid metabolism are formed, they can be remodeled to form glycerolipids with fatty acid profiles characteristic of the individual phospholipid class. Most in vitro studies of the acylation of sn-glycerol 3-phosphate indicate that the glycerophosphate acyltransferase (which acylates sn-glycerol 3phosphate at position s n - I ) preferentially utilizes saturated fatty acyl-CoA thioesters. Numa and co-workers (Yamashita and Numa, 1972; Yamashita et al., 1972, 1973) separated rat liver glycerophosphate acyltransferase from other acyltransferases from rat liver and found, by in vitro assay, that palmitoyl-CoA and stearoyl-CoA are the most effective substrates on the basis of relative rates of acylation. Oleoyl-CoA, linoleoyl-CoA, and arachidonoyl-CoA are poor substrates. The enzymes that acylate dihydroxyacetone phosphate show a similar preference for saturated fatty acids (Hajra, 1968). In contrast, l-acylglycerophosphate acyltransferase, which acylates position in-2, exhibits relative rates of acylation on the order oleoyl-CoA > linoleoyl-CoA = palmitoleoyl-CoA > myristoyl-CoA = palmitoyl-CoA. Based on these observations, it can be concluded that position sn-1 of phosphatidic acid, at least in liver tissue, should contain mostly palmitate and stearate (16:O and 18:O) and little unsaturated fatty acid. Position sn-2 should contain mostly monoenoic and dienoic fatty acids (e.g., 18:l and 18:2), with little of either the saturated or the polyunsaturated fatty acids. This fatty acid specificity is indeed what is found for phosphatidic acid in liver tissue (Possmayer et al., 1969). Glycerolipid products of phosphatidic acid can then be remodeled by a deacylation-reacylation cycle. Tissues contain phospholipases A, and A,, capable of deacylating glycerophospholipids, and also contain acyltransferases specific for the individual lysoglycerophospholipids. Deacylation-reacylation activity has been most thoroughly studied for phosphatidylcholine. In liver tissue, there is good evidence that this cycle is important for the introduction of polyunsaturated fatty acids (principally arachidonate) into position sn-2 (reviewed by Holub and Kuksis, 1978). This conclusion is supported by labeling studies which followed the time

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157

course of appearance of various molecular species of phosphatidylcholine. In general, during short incubation periods with radiolabeled substrates, label is found in monoenoic and dienoic species of phosphatidylcholine. As incubations times lengthen, label accumulates in the more polyunsaturated species. These data are consistent with the concept that newly formed phosphatidylcholine has a fatty acid profile similar to the precursor phosphatidic acid and that the phosphatidylcholine is subsequently remodeled to a more unsaturated species. In v i m studies of the lysophosphatidylcholine acyltransferase activity in liver have supported the conclusion that its role is to introduce a polyunsaturated fatty acid into phosphatidylcholine. Yamashita er al. (1981) separated rat liver microsomal lysophosphatidylcholine acyltransferase activity from the other acyltransferase activities. The enzyme is specific for lysophosphatidylcholine and does not acylate sn-glycerol 3-phosphate, 1-acyl-sn-glycerol 3-phosphate. or lysophosphatidylethanolamine. Based upon relative rates of acylation, the enzyme shows a preference for arachidon yl-CoA. Phosphatidylcholine-remodelingenzymes appear to have a different responsibility in lung tissue (reviewed by Batenburg, 1982). The pulmonary type 11 epithelial cell, which synthesizes pulmonary surfactant, produces significant amounts of phosphatidylcholine with palmitate at positions sn1 and sn-2. It is believed that deacylation-reacylation activities are responsible for the introduction of the saturated fatty acid (principally palmitate) at position sn-2.

G. The Topography of Lipid Biosynthesis Figure 5 summarizes the principal pathways of phospholipid biosynthesis discussed in the previous sections. Table IV lists the principal enzymes of phospholipid biosynthesis. Most studies cited in the previous sections that have explored the subcellular locations of the enzymes of lipid biosynthesis have assigned activity to the microsomal, the mitochondrial, or the cytosolic fractions. Microsomes obtained by subcel I ular fractionation consist principal Iy of endoplasmic reticulum but also contain Golgi apparatus, nuclear envelope, peroxisomes, and plasma membrane. More complete subcellular fractionations of liver tissue have been carried out, and these studies appear to reveal further localizations of the enzymes of glycerolipid biosynthesis (Jelsema and Morre, 1978). Lipid biosynthetic enzyme activities which are found in the endoplasmic reticulum are also found in Golgi membranes. In Golgi, the specific activities of these enzymes are anywhere from 25 to 85% of the activities

Dihydroxyacetonephosphate

1

1

Acyldihydroxyacetonephosphate

Glycerol 3-phosphate

1 I -Acylglycerol 3-phosphate

Alkyldihydroxyacetonephosphale

I

b

b

1-Alkyl glycerolipids

1

Phosphatidic acid

Elhanolamine plasmalooen I

i

Other plasmalogens

CDPdiac ylgl ycerol

Discylglycerol

4-1 PhosDhatidyleth.nolamine

Phosphatidylserine

PhosDhatidyIcholine

1

Sphingomyelin

TrlacyIaIyCerol

Phosphatidyllnosltol

P h o s p h a t l d ~ l ~ l ~ c e1 rol

4 - A

Csrdlolipln

FIG.5 . Principal pathways of phospholipid biosynthesis in mammalian cells.

TABLE 1V PRINCIPAL ENLYMES OF PHOSPHOLIPID BIOSYNTHESIS

Recommended name" Plrosphciridate hiosynrhesis

Long-chain-fatty-acid-CoA ligase (acyl-CoA synthelase) Glycerol-3-phosphate acyltransferase

2

I-Acylglycerol-3-phosphate acyltransferase

fJl (D

Glvcerone-phosphate acyltransferase (dihydroxyacetone-phosphate acyltransferase) Alkylglycerone-phosphate synthase (alkyldihydroxyacetonephosphate synthase) Acylglycerone-phosphate reductase (acyldihydroxyacetonephosphate reductase) Diric:\.lglwer.ol ptiilitc~tiy Phohphatidate phosphatase

React ion

Systematic name Acid:CoA ligase (AMP-forming)

Number

Fatty acid + CoA + ATP fatty acyl-CoA + AMP

-

+ sn-glycerol 3-phosphate + I-acyl-sn-glycerol 3-phosphate Acyl-CoA: I-acyl-sn-glycerol- Acyl-CoA + I-acyl-sn-glycerol 3-phosphate 0-acyltransfer3-phosphate + phosphatidate ase Acyl-CoA:glvcerone-phosAcyl-CoA + dihydroxvacetone phate phosphate --t acyldihydroxy0-acyltransferase acetone phosphate Acyl-CoA:sn-glycerol-3-phos-Acyl-CoA

E.C. 6.2.1.3

Microsomes. mitochondria

E.C. 2.3.1.15

Microsomes. mitochondria

E.C. 2.3. I .5 I

Microsomes

E.C. 2.3. I .42

Microwme\. neroxisome\

phate 0-acyltransferase

-

Fatty alcohol + acyldihydroxv- E.C. 2.5. I .26 acetone phosphate 3-phospho-2-oxopropanylalkyldihydroxyacetone transferase phosphate + fatty acid I-Palmitoylglycerol-3-phos- Acyldihydroxyacetone phosphate E.C. I . I .1 . 1 0 1 + NADPHphate:NADP' oxidoreduc lase I-acyl-sn-glycerol 3-phosphate + NADP'

I-Acyl-glycerone-3-phos-

phate:long-chain-alcohol 0-

3-sn-Phosphatidate phosphohydrolase

Phosphatidate

-

diacylglycerol

Principal subcellular location

E.C. 3. I .3.4

Peroxisomes

Microsome\. mitochondria

All whcellular fractions

TABLE IV (continued)

Recommended name" Choline kinase Ethanolamine kinase

a 0)

0

Choline-phosphate cytid ylyltransferase Ethanolamine-phosphate cytidylyltransferase Ethanolaminephosphotransferase

Cholinephosphotransferase

Diacylgly cerol acyltransferase

Systematic name

ATP + cholinecholine phosphate + ADP ATP + ethanolamine + ethanolamine phosphate + ADP CTP + choline phosphate CTP:choline-phosphate 3 . . cytidylyltransferase CDPcholine CTPethanolamine-phosphate CTP + ethanolamine phosphate cytidy Iyltransferase + CDPethanolamine CDPethanolamine:I ,2CDPethanolamine + diacylglycerol diacyglycerol --* ethanolaminephosphosphatidylethanolamine + photransferase CMP CDPcholine: I ,2CDPcholine + diacylglycerol diacylglyc erol phosphatidylcholine + CMP cholinephospho transferase Acyl-CoA: 1.2-diacylglycerol Acyl-CoA + diacylglycerol + 0-ac yltransferase triacylglycerol

-

S-Adenosyl-L-methionine: phosphatidylethanolamine N-methy bansf erase

Phosphatidylethanolamine serinetransferase

Number

ATP:choline phosphotransferase ATP:ethanolamine 0phosphotransferase

Inrerconversions of the nitrogen-containing phospholipids

Phosphatidylethanolamine met hyltransferase

Reaction

S-Adenosyl-L-methionine + phosphatidylethanolamine + phosphatidylcholine Phosphatidylethanolamine + senne --., phosphatidylserine + ethanolamine

Principal subcellular location

E.C. 2.7. I .32

Cytosol

E.C. 2.7.1.82

Cytosol

E.C. 2.7.7.15

E.C. 2.7.8.

Cytosol, microsomes Cytosol, microsomes Microsomes

E.C. 2.7.8.2

Microsomes

E.C. 2.3.1.20

Microsomes

E.C. 2.1.1.17

Microsomes

E.C. 2.7.7. 4

Microsomes

Phosphatidylserine decarbox ylase Sphingomyelin synthase

CDPdiucvlgIycemI purhwuy Phosphatidate CytidylyhrdnSferdSe CDPdiac y Igl ycerol-inositol 3-phosphatid y ltransferase

' 2

CDPdiac ylgl ycerol-gl ycerol3-phosphate 3phosphatidyltransferase (glycerophosphate phosphatidyitrdnsferase) Phosphatidylglycerophosphatase Cardiolipin synthase

Bi s(monoac y lgl ycero)phosphate" synthase

Phosphatid yl-~-serine Phosphatidylserine + phosphdti- E.C. 4.1.l.hS carboxy-1yase dylethanolamine + CO? Phosphatidylcholine:ceramide Phosphatidylcholine + cerdmide phosphocholinetransferase + sphingomyelin + diac y lglycerol CTPphosphatidate cytidyl yltransferase CDPdiacyIglycero1:myoinositol 3-phosphatid yltransferase CDPdiac ylg1ycerol:snglycerol-3-phosphate phosphatidyltransferase

CTP + phosphatidate --9 CDPdiacylglycerol CDPdiacylglycerol + m y o inositol -+ phosphatidylinositol + CMP CDPdiacylglycerol + sn-glycerol 3-phosphate + phosphatid ylgl ycerol 3-phosphate + CMP

Phosphatidylglycerophosphate Phosphatidylglycerol phosphoh ydrolase 3-phosphate + phosphatidy lglycerol CDPdiacylglycerol + phosphatid ylglycerol + cardiolipin + CMP Cardiolipin + bis(monoacyigiycero)phosphdte Phosphatidylglycerol + bis(monoacylg1ycerolphosphate ~~~

"

E.C. 2.7.7.41

Mitochondria Plasma membrane

E.C. 2.7.8. I 1

Microsomes. mitochondria Microsomes

E.C. 2.7.8.5

Mitochondria

E.C. 3.1.3.27

Mitochondria

Mitochondria

Lysosomes

~~

Recommended names established by International Union of Biochemistry ( 1984). Names in parentheses refer to the 1978 recommendations. Synthesis of bis(monoacylglycero)phosphate from cardiolipin or phosphatidylglycerol involves more than one enzyme-catalyzed reaction.

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KENNETH J. LONGMUIR

found in the endoplasmic reticulum. Nuclear membranes and plasma membrane fractions contain some, but not all, of the lipid biosynthetic enzymes found in the endoplasmic reticulum. They have substantial phosphatidate phosphohydrolase activity, phosphatidate cytidylyltransferase activity, and phosphatidylserine exchange activity. The terminal enzymes of phosphatidylcholine, phosphatidylethanolamine, and phosphatidylinositol biosynthesis are virtually absent from both the nuclear membrane and the plasma membrane. It should be noted that the Golgi membranes, the nuclear membranes, and the plasma membrane each contain less than 5 1 0 % of the total membrane protein of the cell. In contrast, the endoplasmic reticulum accounts for approximately 50% of total membrane protein. Hence it can be concluded that on the basis of total enzyme activity, far more lipid biosynthesis can take place in the endoplasmic reticulum than in any other membrane which comprises the rnicrosomal subcellular fraction. Several experiments have also addressed the asymmetrical orientation of the microsomal phospholipid biosynthetic enzymes in mammalian tissue (Vance et af., 1977; Coleman and Bell, 1978; Ballas and Bell, 1980, 1981; reviewed by Bell et al., 1981). These experiments rely on the fact that microsomes from cell homogenates form sealed vesicles with the outside face corresponding to the cytoplasmic side of the endoplasmic reticulum. Integrity of these microsomal vesicles can be assessed by the measurement of mannose-6-phosphatase, an enzyme activity which resides on the lumenal side of the endoplasmic reticulum. If microsomal vesicles are intact, mannose-6-phosphatase activity is low. Activity increases several fold when vesicles are disrupted. The orientation of the enzymes of glycerolipid biosynthesis has been explored by measuring the susceptibility of individual enzyme activities to proteases, such as chymotrypsin, trypsin, and pronase, and to the impermeant inhibitor mercury dextran. Parallel assessment of vesicle integrity was also performed. Inactivation of enzyme activity in an intact vesicle is taken as evidence that at least a portion of the enzyme is exposed to the outside of the microsomal vesicle (which corresponds to the cytoplasmic face of the endoplasmic reticulum). The results show that at least a portion of all rnicrosomal enzymes of lipid biosynthesis are exposed on the cytoplasmic face of the endoplasmic reticulum. They are inhibited by one or more proteases or by mercury dextran. These enzymes include the acyl-CoA synthetase, all enzymes of phosphatidic acid biosynthesis, all enzymes of the diacylglycerol pathway, all enzymes which catalyze the interconversion of nitrogen-containing phospholipids, and the enzymes phosphatidate cytidylyltransferase and CDPdiacylglycerol-inositol phosphatidyltransferase. The case for a cytoplasmic orientation of the enzymes of phospholipid

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163

biosynthesis is further strengthened by observations that many of the substrates of lipid metabolism do not penetrate across endoplasmic reticulum (reviewed by Bell el ul., 1981). These substrates include fatty acyl-CoA, NADPH, CDPcholine, cytidine nucleotides, and ATP, all of which are believed to reside in vivo in the cytosol of the cell or in the cytosolic side of the endoplasmic reticulum.

V. MEMBRANE LIPID TRANSPORT AND DIFFERENTIATIONOF MEMBRANES A. Introduction

As indicated in the previous sections, most of the lipid material of the cell is formed on the endoplasmic reticulum, then transported to the different intracellular membranes. Various cellular processes must move lipid from endoplasmic reticulum to other cell membranes. Also, mechanisms to produce different membranes with different lipid compositions must exist. This section discusses three intracellular processes which may be responsible for the transport and sorting of lipid within the cell: ( I ) synthesis of certain classes of lipid at the membranes in which they are found, (2) transport of lipid by phospholipid transfer proteins, and (3) transport of lipid by a process of membrane flow.

B. Lipid Biosynthesis at Specific lntracellular Sites Although most lipid biosynthetic activity resides in the endoplasmic reticulum, some enzymes of lipid biosynthesis are found in substantial activity in other membranes. When one analyzes the major differences in lipid composition among the subcellular membranes, one finds that some enzymes of lipid biosynthesis colocalize with the lipids that they synthesize. In particular, the following features are noted. I . Mitochondria have a greater proportion of phosphatidylethanolamine and a smaller proportion of phosphatidylserine than other membranes. The enzyme phosphatidylserine decarboxylase (Section IV,D.3), which converts phosphatidylserine to phosphatidylethanolamine,is a mitochondrial enzyme. 2. Mitochondria contain most of the cellular phosphatidylglycerol and cardiolipin. The enzymes of phosphatidylglycerol biosynthesis (glycerophosphate phosphatidyltransferase and phosphatidylglycerophosphatase) and the enzyme of cardiolipin biosynthesis (cardiolipin synthase) are mitochondrial enzymes (Section 1V.E).

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KENNETH J. LONGMUIR

3. The plasma membrane has a greater proportion of sphingomyelin and a smaller proportion of phosphatidylcholine than the intracellular membranes. Recent investigations suggest that the enzyme which synthesizes sphingomyelin from phosphatidylcholine is located principally in the plasma membrane (Section IV,D,4). 4. Bis(monoacylglycero)phosphate, a lysosomal lipid, is synthesized by enzymes which reside mostly, if not exclusively, in the lysosomes (Section IV ,E 3).

Not all enrichments of lipid in various parts of the cell can be accounted for by the localization of certain enzymes of lipid biosynthesis. There are examples which indicate that some lipids must be sorted and transported from the site of synthesis to the region of the cell where they accumulate, as follows. I. Many cells, notably adipocytes, contain cytosplasmic lipid droplets which contain principally triacylglycerol. Triacylglycerol is formed from diacylglycerol by an enzyme which is localized in the microsomal fraction. No diacylglycerol acyltransferase activity is found in the lipid droplets themselves. Hence the triacylglycerol must be sorted out and transported from the endoplasmic reticulum to the lipid droplet. 2. Pulmonary surfactant is principally a lipid material formed by the type I1 alveolar cell of the lung. Its composition is substantially different from the intracellular membranes. It has a high lipid to protein ratio (greater than 10: I w/w), and it contains upward of 80% phosphatidylcholine and 10% phosphatidylglycerol (Sanders, 1982). Other lipids that are substantial components of other membranes, phosphatidylethanolamine, phosphatidylinositol, and phosphatidylserine, are present in only a few percent. Most strikingly, over one-half of the phosphatidylcholine contains only saturated fatty acid at positions sn-I and sn-2. With the possible exception of phosphatidylglycerol, lipids destined for surfactant formation, including the saturated phosphatidylcholine, are produced in the endoplasmic reticulum. They are then transported to a cytoplasmic organelle called the lamellar body, where the lipid material is stored prior to secretion. An intracellular transport and lipid sorting process must be present in order to move lipid to the lamellar body and to enrich the lamellar body material with saturated phosphatidylcholine. C. Phospholipid Transfer Proteins

Since the discovery of phospholipid transfer proteins in 1968, considerable attention has been given to their properties. Phospholipid transfer activity has been found in the cytosolic fractions of all eukaryotic cells

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165

that have been investigated so far. It is now generally accepted that there are three different classes of transfer proteins (reviewed by Wirtz, 1983): I . A transfer protein that is specific for phosphatidylcholine. 2. A transfer protein that shows a preference for phosphatidylinositol but will transfer phosphatidylcholine at roughly one-tenth the rate of phosphatidylinositol. 3. A nonspecific transfer protein that will readily transfer phosphatidylcholine, phosphatidylinositol. phosphatidylethanolamine, phosphatidylglycerol, sphingomyelin, phosphatidic acid, and cholesterol.

The original assays of the phospholipid transfer proteins were carried out in such a manner that exchange of phosphatidylcholine between two classes of membrane was demonstrated. More recently, it has been shown that these proteins are capable, at least in in vim) assays, of carrying out a net transfer of lipid from donor membranes to acceptor membranes. In particular, the phosphatidylcholine-specifictransfer protein can extract a molecule of phosphatidylcholine from a donor membrane without inserting one in return. The phosphatidylcholine-specific transfer protein can also release a molecule of phosphatidylcholine into a membrane that does not contain phosphatidylcholine. These in v i m data support proposals that phospholipid transfer proteins in vivo can carry out a net transfer of newly formed lipid from the site of synthesis (the endoplasmic reticulum) to other membranes of the cell. What remains to be demonstrated is the actual contribution these proteins make to the intracellular movement of lipid under a variety of physiologic conditions.

D. Membrane Flow In addition to the transport of individual phospholipid molecules by specific transfer proteins, it is proposed that lipid can move from the sites of synthesis to other parts of the cell by a process of "membrane flow" (Morre et id.. 1979). By this mechanism, lipid moves from one class of membranes (such as the endoplasmic reticulum) to another (such as the Golgi, mitochondria, and plasma membranes) either by diffusion of lipid through connections between membranes or by movement of membrane vesicles from one membrane to another. Continuity between the various classes of membranes in eukaryotic cells has not been consistently demonstrated. For this reason, most investigators usually discuss the concept of membrane flow in terms of transport of discrete vesicles of membrane material from one membrane to another ("vesicle transport").

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Transport of a vesicle of endoplasmic reticulum membrane to other membranes is the proposed mechanism by which newly synthesized, membrane-bound protein moves from endoplasmic reticulum t o Golgi and possibly to other membranes (reviewed by Sabatini et al., 1982). Such a process can result in the net transport of lipid as well. Few studies have been carried out that directly address the relative importance of membrane flow versus the phospholipid transfer proteins. Recently, however, Yaffe and Kennedy (1983)investigated the movement of lipid between microsomal and mitochondria1 compartments of hepatocytes and BHK-21 cells. In their experiments, an analog of choline was used which contained a propyl group in place of one of the choline methyl groups. This analog can be incorporated by cells to form a propyl analog of phosphatidylcholine. It was found that this analog was readily transferred from microsomal compartments to mitochondria at rates nearly equal to the transfer of phosphatidylcholine in both hepatocytes and BHK21 cells. However, when the cytosol of BHK-21 cells was assayed for transfer protein activity, the propyl analog was not a substrate. These data are taken as evidence that a phospholipid transfer protein is not required for the movement of phosphatidylcholine (or its analogs) from microsomes to mitochondria. Further experiments are needed to determine the relative role of membrane flow and phospholipid transfer proteins in the intracellular movement of lipid from the sites of synthesis to other parts of the cell. The existence of transfer proteins does not exclude the need for a membrane flow mechanism, nor does transport by membrane flow exclude a role for transfer proteins. The existence of more than one mechanism responsible for intracellular lipid transport may offer a redundancy which insures that the cell will always form, maintain, and recycle membrane under a variety of physiologic conditions. It remains to be determined to what extent membrane flow and lipid transfer proteins can sort out various classes of lipid as they are transported. These mechanisms should be further studied to determine their role in the formation of different membranes with different lipid compositions.

ACKNOWLEDGMENTS

I wish to thank Christine Resele-Tiden for research assistance. This work was supported by a grant from the National Institutes of Health (HL-28619).

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REFERENCES Agranoff. €3. W. ( 1962). Hydrolysis of long-chain alkyl phosphates and phosphatidic acid by an enzyme purified from pig brain. J. Lipid R P S .3. 190-196. Agranoff. B. W.,and Hajra. A. K. (1971). The acyl dihydroxyacetone phosphate pathway for glycerolipid biosynthesis in mouse liver and Ehrlich ascites tumor cells. Proc,. Nrrrl. Accrd. Sci. U . S . A . 68. 411415. Akesson. B. ( 1969). Composition of rat liver triacylglycerols and diacylglycerols. Eitr. J . Biochctt1. 9. 463471.

Allan. D., and Michell. K. H. (1975). Enhanced synthesis ill, not’o of phosphatidylinositol in lymphocytes treated with cationic amphiphilic drugs. Biockrn. J , 148. 471478. Audubert. F.. and Vance, D. E. (1983). Pitfalls and problems in studies o n the methyliltion of phosphatidylethanolamine. J. B i d . Chum. 258, 10695-10701. Ballas. 1.. M.. and Bell, R. M. (1980). Topography of phosphatidylcholine. phosphatidylethanolamine and triacylglycerol biosynthetic enzymes in rat liver microsomes. Bioc~hitn. Bioph>’.y.Acto 602, 578-590. Ballas. L. M..and Bell, K. M. (1981). Topography ofglycerolipid synthetic enzymes. Synthesis of phosphatidylserine. phosphatidylinositol and glycerolipid intermediates occurs on the cytoplasmic surface of rat liver microsomal vesicles. Biochim. Biophys. Aclrr 665. 586-595.

Baranska. J. (1982). Biosynthesis and transport of phosphatidylserine in the cell. Adv. Lipid Res. 19. 163-184. Barenholz. Y..and Thompson. T. E. (1980). Sphingomyelins in bilayers and biological membranes. Biochim. Biophvu. A(.rci 604, 129-158. Batenburg, J. J. ( 1982). The phosphatidylcholine-lysophosphatidylcholinecycle. In ”Lung Development: Biological and Clinical Perspectives” (P. M. Farrell. ed.). Val. I. pp. 359-390. Academic Press. New York. Bell, R. M.. and Coleman. R. A. (1980). Enzymes of glycerolipid biosynthesis in eukaryotes. Annrr. Re\,. Biochein. 49, 459-487. Bell, K. M.,Ballas. L. M.,and Coleman. R. A. ( 19x1). Lipid topogenesis. J . Lipid Res. 22, 391403. Bergelson. L. D., Dyatlovitskaya, E. V.. Sorokina. I . B.. and Gorkova. N. P. ( 1974). Phospholipid composition of mitochondria and microsomes from regenerating rat liver and hepatomas of different growth rate. Biochitti. Bioppliys. Aciu 360. 361-365. Bernert, J. T., and Sprecher. H. (1977). An analysis of partial reactions in the overall chain elongation of saturated and unsaturated fatty acids by rat liver microsomes. J . Biol. Clierri. 252. 67366744. Bernert, J . T.. and Ullman, M. D. (19x1). Biosynthesis of sphingomyelin from cyvrhro-ceramides and phosphatidylcholine by a microsomal cholinephosphotransferase. Biochiin. B i o ~ p h . A(,ttr ~ ~ . 666, 99-109. Bishop. J . E.. and Hajra. A. K. (1981). Mechanism and specificity of formation of long chain alcohols by developing rat brain. J . Biol. Chrrn. 256. 95424550. Bleasdale, J. E.. and Johnston, J. M. (1982). Phosphatidic acid production and utilization. In “Lung Development: Biological and Clinical Perspectives” ( P . M . Farrell. ed.). Vol. I . pp. 259-294. Academic Press. New York. Bleasdale, J . E.. and Wallis. P. (1981). Phosphatidylinositol-inositol exchange in rabbit lung. Biociiirn. Biopli.vs. Ac,lci 664, 4 2 8 4 0 .

Bleasdale, J. E.. Davis, C. S., Hajra, A. K., and Agranoff, B. W.(1978). A rapid sensitive 87. 19-27. assay for phosphatidate phosphohydrolase. Anrrl. Bioch~~rri.

168

KENNETH J. LONGMUIR

Bleasdale, J. E., Wallis, P., MacDonald, P. C., and Johnston, J. M. (1979).Characterization of the forward and reverse reactions catalyzed by CDP-diacylglycerol:inositoltransferase in rabbit lung tissue. Biochim. Biophys. Acta 575, 135-147. Bleistein, J., Heidrich, H. G., and Debuch, H. (1980). The phospholipids of liver lysosomes from untreated rats. Hoppe-Seyler'r Z. Physiol. Chem. 361, 595-597. Bretscher, M. S. (1972a). Phosphatidyl-ethanolamine:Differential labelling of intact cells and cell ghosts of human erythrocytes by a membrane-impermeable reagent. J . Mol. B i d . 71, 523-528. Bretscher, M. S. (1972b). Asymmetrical lipid bilayer structure for biological membranes. Nature (New Biol.) 236, 11-12. Brindley, D. N., and Bowley, M. (1975). Drugs affecting the synthesis of glycerides and phospholipids in rat liver. Biochem. J . 148, 461-469. Brophy, P. J., Choy, P. C., Toone, J. R., and Vance, D. E. (1977). Choline kinase and ethanolamine kinase are separate, soluble enzymes in rat liver. ENT.J. Biochcm. 78, 491-495. Brotherus, J., and Renkonen, 0. (1977). Phospholipids of subcellular organelles isolated from cultured BHK cells. Biochirn. Biophys. Aria 486, 243-253. Brown, A. J., and Snyder, F. (1983). The mechanism of alkyldihydroxyacetone-P synthase. J. B i d . Chem. 258. 4184-4189. C h o y , P . C . , a n d V a n c e , D. E . ( 1 9 7 8 ) . Lipid r e q u i r e m e n t s f o r a c t i v a t i o n of CTP:phosphocholine cytidylyltransferase from rat liver. J . B i d . Chcm. 253,5 163-5167. Choy, P. C., Lim, P. H., and Vance, D. E. (1977). Purification and characterization of CTP:cholinephosphate cytidylyltransferase from rat liver cytosol. J . Biol. Chem. 252. 7673-7677. Colbeau, A., Nachbaur, J., and Vignais, P. M. (1971). Enzymic characterization and lipid composition of rat liver subcellular membranes. Biochim. Biciphys. Acru 249. 462-492. Coleman, R.,and Bell, R. M. (1978). Evidence that biosynthesis of phosphatidylethanolamine, phosphatidylcholine, and triacylglycerol occurs on the cytoplasmic side of microsomal vesicles. J. Cell Biol. 76, 245-253. Coleman, R. A., and Bell, R. M. (1980). Selective changes in enymzes of the sn-glycerol 3-phosphate and dihydroxyacetonephosphate pathways of triacylglycerol biosynthesis during differentiation of 3T3-LI preadipocytes. J. Bioi. Chem. 255, 7681-7687. Coleman, R. A., and Haynes, E. B. (1983). Selective changes in microsomal enzymes of triacylglycerol and phosphatidylcholine synthesis in fetal and postnatal rat liver. J. Biol. Chem. 2 5 8 , 4 5 0 4 6 . Crain, R. C., and Marinetti. G. V. (1979). Phospholipid topology of the inner mitochondria1 membrane of rat liver. Biochemistry 18, 2407-2414. Crain, R. C., and Zilversmit, D. B. (1980). Two nonspecific phospholipid exchange proteins from beef liver. 2. Use in studying the asymmetry and transbilayer movement of phosphatidylcholine, phosphatidylethanolamine,and sphingomyelin in intact rat erythrocytes. Biochemistry 19, 140-1447. de Kruijff, B., van den Besselaar, A. M. H. P., Cullis, P. R., van den Bosch, H., and van Deenen, L. L. M. (1978). Evidence for isotropic motion of phospholipids in liver microsomal membranes. A "P NMR study. Biochim. Biophys. Acta 514, 1-8. Dennis, E. A., and Kennedy, E. P. (1972). lntracellular sites of lipid synthesis and the biogenesis of mitochondria. J . Lipid Res. 13, 263-267. Eichberg. J., Gates, J., and Hauser, G. (1979). The mechanism of modification by propranolol of the metabolism of phosphatidyl-CMP (CDP-diacylglycerol) and other lipids in the rat pineal gland. Biochirn. Biophys. Acta 573. 90-106.

BIOSYNTHESIS AND DISTRIBUTION OF LIPIDS

169

Esko, J. D.. and Raetz, C. R. H. (1980).Mutants of Chinese hamster ovary cells with altered membrane phospholipid composition. Replacement of phosphatidylinositol by phosphatidylglycerol in a ~n.~o-inositol auxotroph. J. Biol. Chem. 255, 44744480. Etemadi, A. -H. (1980).Membrane asymmetry. A survey and critical appraisal of the methodology. 11. Methods for assessing the unequal distribution of lipids. Biocliim. B i o p h y s . Aciu 604, 423-475. Fitzpatrick, S. M., Sorresso, G., and Haldar, I). ( 1982).Acyl CoA:sn-glycerol-3-phosphate 0-acyltransferase in rat brain mitochondria and microsomes. J . Nerrrochem. 39. 286289. Fleming. P. J., and Hajra. A. K. (1977). I-Alkyl-sn-glycero-3-phosphate:acyl-CoA acyltransferase in rat brain microsomes. J . Biol. Chem. 252, 1663-1672. Freinkel, N., El Younsi, C.. Dawson, R. M. C. (1975).Interrelations between the phospholipids of rat pancreatic islets during glucose stimulation and their response to medium inositol and tetracaine. Eur. J . Biochrm. 59. 245-252. Fujiki, Y., Fowler, S . , Shio, H., Hubbard, A. I-.. and Lazarow. P. B. (1982). Polypeptide and phospholipid composition of the membrane of rat liver peroxisomes: Comparison with endoplasmic reticulum and mitochondrial membranes. J. Cell B i d . 93. 103-1 10. Glaumann, H., and Dallner. C. (1968).Lipid composition and turnover of rough and smooth microsomal membranes in rat liver. J . Lipid Res. 9, 720-729. Gordesky, S . E., and Marinetti, G. V. (1973).The asymmetric arrangement of phospholipids in the human erythrocyte membrane. Binclwm. Biophys. Res. Commitn. 50, 1027-1031. Hajra. A. K.(1968). Biosynthesis of phosphatidic acid from dihydroxyacetone phosphate. Biochem. Biophvs. Res. Commitn. 33. 929-93.5. Hajra, A. K. (1983).Biosynthesis of 0-alkylglycerol ether lipids. In “Ether Lipids. Biochemical and Biomedical Aspects” (H. K. Mangold and F. Paltauf. eds.). pp. 85-106. Academic Press, New York. Hajra, A. K., and Bishop, J. E. (1982).Glycerolipid biosynthesis in peroxisomes via the acyl dihydroxyacetonephosphalepathway. Ann. N.Y. Acad. Sci. 386. 170-182. Haldar, 0.. Tso. W.-W., and Pullman. M. E. (1979).The acylation of sn-glycerol 3-phosphate in mammalian organs and Ehrlich ascites tumor cells. J . Biol. Chem. 254, 4502-4509. Hallman. M.. and Epstein, B. L. (1980).Role of mvo-inositol in the synthesis of phosphatidylglycerol and phosphatidylinositol in the lung. Biochurn. Biophys. Res. Commrtn. 92, 11.51-11.59.

Harb. J. S.. Comte, J., and Gautheron, D. C. (1981).Asymmetrical orientation of phospholipids and their interactions with marker enzymes in pig heart mitochondrial inner membrane. Arch. Biochem. Biophys. 208. 30.5-31X. Hers, H. G., and Hue. L. (1983).Gluconeogenesis and related aspects of glycolysis. Annir. Rev. Biocliern. 52, 617-653. Heymans. H. S. A.. Schutgens, R. B. H.. Tan, K..van den Bosch, H., and Borst, P. (1983). Severe plasmalogen deficiency in tissues of infants without peroxisomes (Zellweger syndrome). Nuturr (London) 306, 69-70. Hirschmann. H. (1960).The nature of substrate asymmetry in stereoselective reactions. J. B i d . Cliem. 235, 2762-2767. Hokin-Neaverson. M.. Sadeghian. K.. Harris. D. W., and Merrin. J . S. (1977).Synthesis of CDP-diglyceride from phosphatidylinositol and CMP. Biorhrm. Biopliys. Res. Commrtri. 78, 364-371. Holman, R. ’r. (1977).Essential fatty acids in human nutrition. Adti. E x p . Med. A i d . 83, 5 I 5-534. Holub. B. J., and Kuksis, A. (1978).Metabolism of molecular species of diacylglycerophospholipids. Adv. Lipid Rrs. 16. 1-125.

170

KENNETH J. LONGMUIR

Hosaka, K., Schiele, U.,and Numa, S. (1977). Diacylglycerol acyltransferase from rat liver microsomes. Separation and acyl-donor specificity. Eur. J. Biochem. 76, 113-1 18. Hostetler. K . Y., and van den Bosch, H. (1972). Subcellular and submitochondrial localization of the biosynthesis of cardiolipin and related phospholipids in rat liver. Biochim. Biopliys. Acfa 260, 380-386. Hostetler, K. Y., Galesloot, J. M., Boer, P., and van den Bosch, H. (1975). Further studies on the formation of cardiolipin and phosphatidylglycerol in rat liver mitochondria. Effect of divalent cations and the fatty acid composition of CDP-diglyceride. Biochim. Biopliys. A C ~ 380, U 382-389. Hostetler, K. Y., Zenner, B. D., and Moms, H. P. (1976). Abnormal membrane phospholipid content in subcellular fractions from the Morris 7777 hepatoma. Biochim. Biophys. Aciu 441, 231-238. Hostetler, K. Y.,Zenner, B. D., and Morris, H. P. (1979). Phospholipid content of mitochondrial and microsomal membranes from Morris hepatomas of varying growth rites. Cancer Res. 39. 2978-2983. International Union of Biochemistry (1984). “Enzyme Nomenclature.” Academic Press. Orlando. loannou, P. V . , and Golding, B. T. (1979). Cardiolipins: their chemistry and biochemistry. Prog. Lipid Res. 17, 279-318. IUPAC-IUB Commission on Biochemical Nomenclature. (1978). The nomenclature of lipids. J . Lipid Res. 19, 114-128. James. A. T. (1977). The specificity of mammalian desaturases. Adv. Exp. Med. Biol. 83, 51-74. Jeffcoat, R. (1979). The biosynthesis of unsaturated fatty acids and its control in mammalian liver. Es.says Biochem. IS, 1-36. Jelsema, C. L., and Morre, D. J. (1978). Distribution of phospholipid biosynthetic enzymes among cell components of rat liver. J . Biol. Chem. 253, 7960-7971. Joutti, A.. and Renkonen, 0. (1979). The stereoconfiguration of bis(monoacylg1ycero)phosphate synthesized in vitro in lysosomes of rat liver: comparison with the natural lipid. J. Lipid Res. 20. 840-847. Joutti, A., Brotherus, J., Renkonen. 0..Laine, R., and Fischer, W. (1976).The stereochemical configuration of lysobisphosphatidic acid from rat liver, rabbit lung, and pig lung. Biochini. Bioptrys. Acia 450, 206-209. Kahlenberg, A., Walker, C., and Rohrlick, R. (lW4). Evidence for an asymmetric distribution of phospholipids in the human erythrocyte membrane. Can. J. Biochem. 52, 803-806. Kanfer, J. N. (1980). The base exchange enzymes and phospholipase D of mammalian tissue. Can. J . Bioclrem. 58, 1370-1380. Keenan, T. W., and Morre, D. J. (1970). Phospholipid class and fatty acid composition of Golgi apparatus isolated from rat liver and comparison with other cell fractions. Biochemistry 9, 19-25. Kleinig, H. (1970). Nuclear membranes from mammalian liver. 11. Lipid composition. J. Cell Biol. 46. 396-402. Krebs, J . J. R. (1982). The topology of phospholipids in artificial and biological membranes. J . Bioenerg. Biomembr. 14, 141-157. Krebs. J. J. R., Hauser, H., and Carafoli, E. (1979). Asymmetric distribution of phospholipids in the inner membrane of beef heart mitochondria. J . B i d . Cliem. 254, 5308-53 16. Kuksis, A. (1978). Fatty acid composition of glycerolipids in animal tissues. In “Handbook of Lipid Research. Fatty Acids and Glycerides” (A. Kuksis, ed.), pp. 381-442. Plenum. New York.

BIOSYNTHESIS AND DISTRIBUTION OF LIPIDS

171

Laposata. M.. Prescott. S. M.. Bross. T. E.. and Majerus, P. W. (1982). Development and characterization of a tissue culture cell line with essential fatty acid deficiency. Proc. Nir/l. Acnd. Sci. U.S.A. 79. 7654-7658. I-ongmuir. K. J . , Bleasdale. J . E., Quirk. J . G . , and Johnston, J. M. (1982). Regulation of lamellar body acidic glycerophospholipid biosynthesis in fetal rabbit lung in organ culture. Biochinr. Biophys. AcVu 712, 356-364. MacDonald. P. M., and McMurray. W. C. (1980). Piirtial purification and properties of mammalian phosphatidylglycerophosphatiise. Biodrirn. Bioplrvs. A c t a 620. 80-89. Manning. R., and Brindley. D. N. (1972). lritium isotope effects in the measuremcnt of the glycerol phosphate and dihydroxyacetontt phosphate pathways of glycerolipid biosynthesis in rat liver. Bioc.henr. J . 130, 1o03-1012. Marggraf. W.-D., Anderer, F. A.. and Kanfer, J . N . (1981). The formation of sphingomyelin from phosphatidylcholine in plasma membrane preparations from mouse tibroblasts. Bio<’hi/?1.B i i p h y . ~Ac.lu . 664, 6 1-73. Mason. R. J . (1978). Importance of the acyl dihydroxyacetone phosphate pathway in the synthesis of phosphatidylglycerol and phosphatidylcholine in alveolar type 11 cells. J . Biol. Chetri. 253. 3367-3370. Matsuzawa. Y.,Poorthuis, B. J . H. M.. and Hostetler, K. Y. (1978). Mechanism of phosphatidylinositol stimulation of lysosomal his(monoacylglycery1)phosphate synthesis. J . Biol. Chctn. 253, 6650-6653. McMurray. W. C. (1973). Phospholipids in subcellular organelles and membranes. I n “Form and Function of Phospholipids” tG. B. Ansell. J. N . Hawthorne. and R. M. C. Dawson, eds.), pp. 205-251. Elsevier, Amsterdam. McMurray. W. C.. and Dawson. K. M. C. (1969). Phospholipid exchange reactions within the liver cell. Biochem. J . 112, 91-108. McMurray. W. C.. and Jarvis. E. C. (19781. Purification and properties of phosphatidylglycerophosphate synthetase from mammalian liver mitochondria. Can. J. Biochrrn. 56. 414-419. McMurray, W. C.. and Jarvis, E. C. (1980). Partial purification of diphosphatidylglycerol synthetase from liver mitochondria1 membranes. C ‘ N ~ JI .. Biorhr/n. 58. 771-776. McMurray, W . C.. and Magee. W. L. (1972). Phospholipid metabolism. Atinrc. Rev. Biocfienr. 41. 129-160. Mogelson. S.. and Sobel. B. E. (1981). Ethanolamine plasmalogen methylation by rabbit myocardial membranes. Biockiw. Biop/ry.c. Ac/rc 666, 205-21 I . Morre, D. J.. Kartenbeck. J., and Franke, W. W. (1979). Membrane flow and interconversions ;rmong endomembranes. Bioclrirn. Biophys. Acfrr 559. 71-152. Mozri. R.. Siepi, D., Andreoli, V.. and Porcellati, G . (19x1). The synthesis of‘choline plasmalogen by the methylation pathway in rat brain. FEBS L e t / . 131. 115-1 18. Nishizuka, Y. (1984). The role of protein kinase C in cell surface signal transduction and tumour promotion. Nu/rrrP (London) 308, 693-698. Op den Kamp. J. A. F. (1979). Lipid asymmetry in membranes. Annrc. Rci!. Biochcni. 48. 47-7 I . Paltauf. I-‘. ( 1983). Biosynthesis of I-(>( 1’-alkenyl)glycerolipids(plasmalogens). I n “Ether Lipids: Biochemical and Biomedical Aspects” (H. K. Mangold and F. Paltauf, edx.). pp. 107-128. Academic Press, New York. Poorthuis. B. J. H. M.. and Hostetler, K . Y . (1978). Conversion of diphosphatidylglycerol to bis(monoacylglyceryl)phosphate by lysosomes. J . Lipid Rcs. 19. 309-3 IS. Possmayer. F.. Scherphof, G. L., Dubbelman. 7’. M. A. K.. van Golde. L. M.G., and van Deenen, L. L. M. (1969). Positional specificity of saturated and unsaturated Fatty acids in phosphatidic acid from rat liver. B i o d i i m . Biop/!?..s. A d o 176. 95-1 10.

172

KENNETH J. LONGMUIR

Raetz, C. R. H. (1978). Enzymology, genetics, and regulation of membrane phospholipid synthesis in Escherichiu coli. Microbial. Rev. 42, 614-659. Raetz, C. H. R., Dowhan, W.,and Kennedy, E. P. (1976). Partial purification and characterization of cytidine 5‘-diphosphate-diglyceridehydrolase from membranes of Escherichia coli. J . Bucteriol. 125, 8.55-863. Ray, T. K., Skipski, V. P., Barclay, M.,Essner, E., and Archibald, F. M. (1%9). Lipid composition of rat liver plasma membranes. J . B i d . Chem. 244, 5528-5536. Renooij, W., van Golde, L. M. G . , Zwaal, R. F. A., and van Deenen, L. L. M. (1976). Topological asymmetry of phospholipid metabolism in rat erythrocyte membranes. Eur. J. Biochem. 61, 53-58. Rittenhouse, H. G . , Seguin, E. B., Fisher, S. K., and Agranoff, 9 . W. (1981). Properties of a CDP-diglyceride hydrolase from guinea pig brain. J . Neurochem. 36, 991-999. Rognstad, R., Clark, D. G., and Katz, J. (1974). Pathways of glyceride glycerol synthesis. Biochem. J . 140, 249-251. Sabatini, D. D., Kreibich, G., Morimoto, T., and Adesnik, M.(1982). Mechanisms for the incorporation of proteins in membranes and organelles. J . Ce// B i d . 92, 1-22. Sanders, R. L. (1982). The composition of pulmonary surfactant. In “Lung Development: Biological and Clinical Perspectives” (P.M. Farrell. ed.), Vol. I , pp. 193-210. Academic Press, New York. Schlossman, D. M., and Bell, R. M. (1977). Microsomal sn-glycerol 3-phosphate and dihydroxyacetone phosphate acyltransferase activities from liver and other tissues. Arch. Biochem. Biophys. 182, 732-742. Schroeder, F., Perlmutter, J. F., Glaser, M., and Vagelos, P. R. (1976). Isolation and characterization of subcellular membranes with altered phospholipid composition from cultured fibroblasts. J. B i d . Chem. 251, 5015-5026. Sleight, R., and Kent, C. (1983a). Regulation of phosphatidylcholine biosynthesis in mammalian cells. 1. Effects of phospholipase C treatment on phosphatidylcholine metabolism in Chinese hamster ovary cells and LM mouse fibroblasts. J. Biol. Chem. 258, 824830. Sleight, R., and Kent, C. (1983b). Regulation of phosphatidylcholine biosynthesis in mammalian cells. 11. Effects of phospholipase C treatment on the activity and subcellular distribution of CTP:phosphocholine cytidylyltransferase in Chinese hamster ovary and LM cell lines. J. B i d . Chem. 258, 831-835. Sleight, R., and Kent, C. (1983~). Regulation of phosphatidylcholine biosynthesis in mammalian cells. 111. Effects of alterations in the phospholipid compositions of Chinese hamster ovary and LM cells on the activity and distribution of CTP:phosphocholine cytidylyltransferase. J. B i d . Chem. 258, 836-839. Sprecher, H. (1981). Biochemistry of essential fatty acids. f r o g . Lipid Res. 20, 13-22. Sribney, M.. and Kennedy, E. P. (1958).The enzymatic synthesis of sphingomyelin. J . Biol. Chem. 233, 1315-1322. Sturton, R. G., and Brindley, D. N. (1977). Factors controlling the activities of phosphatiddte phosphohydrolase and phosphatidate cytidylyltransferase. Biochem. J. 162, 25-32. Sundler, R. (1975). Ethanolaminephosphate cytidylyltransferase. Purification and characterization of the enzyme from rat liver. J. B i d . Chem. 250, 8585-8590. Sundler, R., and Akesson, 9. (1975). Regulation of phospholipid biosynthesis in isolated rat hepatocytes. J. Biol. Chem. 250, 3359-3367. Takenawa, T., and Egawa, K. (1980). Phosphatidyl inosito1:myo-inositol exchange enzyme from rat liver: partial purification and characterization. Arch. Biochem. Biophys. 202, 60 1-607.

BIOSYNTHESIS AND DISTRIBUTION OF LIPIDS

173

Taki. T.. and Kanfer, J. N. (1978). A phospholipid serine base exchange enzyme. B i ( ~ ~ / i i t i i . Bi<>phy.S.Ac’/cl 528. 309-317. Taki, T., and Kanfer. J. N. (1979). Partial purification and properties o f a rat brain phospholipase D.J . Biol. Chem. 254. 9761-976.5. Ulane, K. E. (1982). The CDPcholine pathway: choline kinase. /,I “Lung Development: Biological and Clinical Perspectives” (P. M. Farrell, ed.). Vol. 1, pp. 295-316. Academic Press, New York. Vance, D. E., and Choy, P. C. (1979). H o w i s phosphatidylcholine biosynthesis regulated? Trends Biochem. Sci. 4. 145-148. Vance, D. E., Choy, P. C., Blake-Farren. S..Lim. P. H., and Schneider. W. J. (1977). Asymmetry o f phospholipid biosynthesis. Noriire (Londoii) 270. 268-269. van den Bosch. H. (1974). Phosphoglyceride metabolism. Annrr. Rcv. Bioch~ni.43, 243277. van Golde, L. M. G . , Raben. J., Batenburg, J. J.. Fleischer. B . . Zambrano. F.. and Fleischer, S . (1974). Biosynthesis o f lipids in Golgi complex and other subcellular fractions from rat liver. Bioc/iinr. Bioppliv.~.Acttr 360, 179-192. van Heusden. G . P. H.. and van den Bosch. H. (1978). The influence o f exogenous and o f membrane-bound phosphatidate concentration on the activity o f CTP:phosphatidate cytidylyltransferase and phosphatidatc phosphohydrolase. Eur. J . Bioclirrn. M ,405412. Verkleij, A. J., Zwaal, R. F. A., Roelofsen. H.,Comfurius. P..Kastelijn. D., and van Deenen, L. L. M. (1973). The asymmetric distribution o f phospholipids in the human red cell membrane. A combined study using phospholipases and freeze-etch electron microscopy. Biochim. Biophys. A c / u 323. 178-193. Virtanen. I., Brotherus. J.. Renkonen. 0.. and Wartiovaara, J. (1977). Phospholipids of outer and inner nuclear membranes in rat liver and BHK-21 cells. B ~ ( J C / I C B WiI(.~ p l i y . ~ . R ~ sC. o t t t t ~ ~ i i76. i . 142-149. Voelker. D. K.,and Kennedy, E. P. (1982). Cellular and enLymic synthesis o f sphingomyelin. Bioc/wmi,s/ry 21, 2753-2759. Wakil. S . J.. Stoops. J. K.. and Joshi, V . C. (1983). Fatty acid synthesis and its regulation. Annii. Rev. Biocht.m. 52, 537-579. White, D. A. (1973). The phospholipid composition of mammalian tissues. 1,) “Form and Function o f Phospholipids” ( G . B. Ansell. J. N. Hawthorne, and R. M. C. Diwson. eds.), pp. 441-482. Elsevier, Amsterdam. Wirtz, K. W. A . ( 1983). Phospholipid transfer proteins. I n ”Lipid-Protein Interactions” (P. C. Jost and 0. H . Griffith, eds.), Vol. I . pp. 151-231. Wiley. New York. Wood. R.. and Snyder. F. ( 1969). Tumor lipids: Metabolic relationships derived from structural analyses o f acyl. alkyl. and alk-I-enyl moieties o f neutral glycerides and phoaphoglycerides. Arch. Biochern. Biophys. 131. 478-494. Wykle, R. L.. and Schremmer, J . M. (1979). Biosynthesis of plasmalogens by the microsomal fraction o f Fischer R-3259 sarcoma. Influence of specific 2-acyl chains on the desaturation of I-alkyl-2-acyl-.s,i-glycero-3-phosphoethan~~lamine.Biochc,nii.s/ry 18. 35 12-35 17. Yaffe. M. P . . and Kennedy. E. P. (1983). Intracellular phospholipid movement and the role o f phospholipid transfer proteins in animal cells. Biochc~mis/ry22, 149771507, Yamashita. S . . and Numa, S. (1972). Partial purification and properties o f glycerophosphate acyltransferase from rat liver. Formation o f I-acylglycerol 3-phosphate from sn-glycerol 3-phosphate and palmityl coenzyme A . Eirr. J . Binchrm. 31. 565-573. Yamashita, S . , Hosaka. K., and Numa. S . (1972). Resolution and reconstitution o f the phosphatidate-synthesizing system o f rat-liver microsomes. Proc.. N u / / . A c d . S(.i. U . S . A . 69. 3490-3493.

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Yamashita. S.. Hosaka, K., and Numa, S. (1973). Acyl-donor specificities of partially purified I-acylglycerophosphate acyltransferase, 2-acylglycerophosphate acyltransferase, and I-acylglycerophosphorylcholine acyltransferase from rat-liver microsomes. Eur. J . Biochetn. 38, 25-31. Yamashita, S., Hosaka, K . , Miki. Y . , and Numa, S. (1981). Clycerolipid acyltransferases from rat liver: I -acylglycerophosphate acyltransferase, I-acylglycerophosphorylcholine acyltransferase, and diacylglycerol acyltransferase. Methods Enzymol. 71, 528-536. Yeung, S. K . F., and Kuksis, A. (1974). Molecular species of ethanolamine phosphatides of dog and pig kidney. Can. J . Biorhern. 52, 830-837.

CURRENT TOPICS IN MEMBRANES A N D TKANSI’OKI, V O I . U M t i ?‘I

Lipid Exchange: Transmembrane Movement, Spontaneous Movement, and Protein-Mediated Transfer of Lipids and Cholesterol ELIEZAR A . D A W I D 0 WICZ

1.

INTRODUCTION

In eukaryotic cells, the endoplasmic reticulum is the principal site for the initial assembly of membrane lipids and proteins (Wilgram and Kennedy, 1963; Chesterton, 1968; Moore e t o / . , 1979). Cholesterol can also be acquired from low-density lipoprotein which enters the cell via receptormediated endocytosis (Goldstein et i i l . , 1979). Since the various membranes in a cell differ markedly in phospholipid composition (Colbeau r t a/., 1971; Renkonen et ul.. 1972; Ulsamer et id., 1971), sterol content (Ulsamer et ( I / . , 1971; Green, 1977), and proteins (Wickner, 1980; Sabatini et a / . , 1982), specific sorting and transport mechanisms must exist which ensure the correct ultimate location of these membrane components. Although several studies have begun to address the question of intracellular transport and sorting of membrane proteins (see, for example, “Membrane Recycling,” Ciba Foundation Symposium 92, Pitman, London, 1982), relatively little is known about similar processes for lipids. Lipids in biological membranes exist in a dynamic state. They can undergo lateral diffusion within the plane of the membrane (Scandella r t u l . , 1972; Sackmann et a / . , 1973) as well as possible transmembrane movement. In addition, both spontaneous and protein-mediated transfer of lipids between membranes can occur. The sorting and intracellular transport of lipids could involve these processes. Reviews concerning the inter-membrane movement of lipids (Bruckdorfer and Graham, 1976; 175

Copyright I’ 19x7 hy A c d c m i c I+\\. InL. All right, of reproduction in a n y form rewrvcd.

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Dawson, 1973; Bell, 1976, 1978, 1984), protein-stimulated lipid exchange (Wirtz, 1974; Bloj and Zilversmit, 1981; Crain, 1982; Zilversmit, 1983), and the transmembrane movement of lipids (Thompson, 1978; Etemadi, 1980; van Deenen, 1981) have been published. It is the aim of this article to include some of the more recent studies on these topics, and wherever possible to indicate possible biological implications.

II.

SPONTANEOUS EXCHANGE OF LIPIDS BETWEEN MEMBRANES

A. Cholesterol

Hagerman and Gould (1951) first reported that cholesterol could undergo spontaneous exchange between red cells and plasma in vitro with a halftime of 1 hr at 37°C. This exchange process was later shown to occur in vivo, between plasma and red cells in dogs, with a very similar half-time to that measured for the process in virro (Eckles et al., 1955). Bruckdorfer and co-workers extended these initial studies by demonstrating that exchange or net movement of cholesterol can occur between rat erythrocytes and dispersions of cholesterol and phospholipid, depending on the cholesterol content of the latter. When the dispersions are composed of equimolar proportions of phosphatidylcholine and cholesterol, exchange of cholesterol with the erythrocytes is observed (Bruckdorfer et af., 1968a). A decrease in the cholesterol content of the lipid dispersion results in depletion of cholesterol from the erythrocytes (Bruckdorfer et a f . , 1968a). The cholesterol content of these depleted membranes can be restored by incubation with an equimolar dispersion of cholesterol and phosphatidylcholine (Bruckdorfer et al., 196813). Several years later, Lange and D'Alessandro (1977) showed that the transfer of cholesterol from human erythrocytes into plasma by exchange or net movement are identical processes characterized by the same rate constant. The movement of cholesterol has been shown to occur between the following: small unilamellar vesicles (Haran and Shporer, 1977; Backer and Dawidowicz, 1979; Nakagawa et a f . , 1979; McLean and Phillips, 1981); lipid vesicles and mammalian cells in culture (Phillips et ul., 1980; Poznansky and Czekanski, 1982; Rothblat and Phillips, 1982; SIotte and Lunberg, 1983; Bellini et al., 1984);lipid vesicles and viral membranes (Moore et ul., 1978); lipid vesicles and Mycopfusmu membranes (Kahane and Razin, 1977);lipid vesicles and bovine serum high-density lipoprotein (Jonas and Maine, 1979); lipid vesicles and isolated sheep liver nuclei (Agutter and Suckling, 1981); lipid vesicles and mitochondria1 membranes (Madden et al., 1980); serum albumin-phospholipid complexes and mammalian cells

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(Bartholow and Geyer, 1981; Bartholow and Geyer, 1982), serum lipoproteins, and rat liver mitochondria (Graham and Green, 1970); serum lipoproteins and vesicular stomatitis virus (Pal et u / . , 1981); serum lipoproteins and Myccyhsmu membranes (Slutzky ct u / . , 1977); human serum high- and low-density lipoproteins (Lund-Katz et a / ., 1982);chylomicrons and high-density lipoprotein (Lippiello and Waite, 1983). Related stcroids have also been shown to undergo transfer between lipid vesicles and erythrocytes (Bruckdorfer et u / . , 1969; Green et N/., 1973). Although the physiological role of cholesterol movement between membranes is not clearly understood, there are a number of parameters which affect this process. Several of these have been documented by Bruckdorfer and Graham (1976). Studies show that the rate of cholesterol exchange is independent of metabolic energy (Murphy, 1962; Bruckdorfer and Green, 1967); is temperature dependent (Murphy, 1962); appears to be independent of ionic strength (Bruckdorfer and Graham, 1967); is independent of the surface charge on the membrane (Bruckdorfer and Graham, 1967); and is enhanced by the presence of certain organic solvents (Bruckdorfer and Green, 1967; Quarfordt and Hilderman, 1970; Bruckdorfer and Sherry, 1984). The kinetics of cholesterol exchange between donor and acceptor can be mediated by the lipid composition of the donor. Increasing the degree of unsaturation of phosphatidylcholine in small unilamellar vesicles (SUVs) composed of phosphatidylcholine and cholesterol enhances the rate of cholesterol exchange between these vesicles and either intact human erythrocytes (Bloj and Zilversmit, 1977a) or erythrocyte ghosts (Poznansky and Lange, 1978) at a given temperature. Similar results have been reported 1979; McLean for cholesterol exchange between S U V s (Nakagawa c>t d., and Phillips, 1982; Fugler et d., 1985). In addition, the rate of cholesterol exchange from S U V s containing N-palmitoylsphingomyelin was strikingly slower when compared with vesicles containing dipalmitoylphosphatidylcholine (Fugler et a / ., 1985). Enrichment of Mycwplusmu gu//isc~ptic./rrn with exogenous sphingomyelin produced an analogous effect in slowing the rate of cholesterol exchange when compared with cells that had been enriched with exogenous phosphatidylcholine (Clejan and Bittman, 1984b). An important finding by Poznansky and Czekanski (1979) is that the activation energy for cholesterol exchange between dioleoyl phosphatidylcholine/cholesterol S U V s and erythrocyte ghosts exhibits a 2-fold increase when the molar percentage of cholesterol is decreased from 30 to 20 mol 5% in both donors and acceptors. This result can be correlated with the model for cholesterol-phospholipid interactions presented by Engelman and Rothman (1972). These latter authors suggested that insertion of cholesterol molecules into a phospholipid bilayer, with no cholesterolcholesterol contact, would reach saturation at a molar ratio of 2: I leci-

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thin:cholesterol (33 mol % cholesterol). Implicit in this model, although not specifically stated, is the fact that above 33 mol % cholesterol in phospholipid cholesterol-cholesterol interactions must occur. The study by Poznansky and Czekanski (1979) is a clear demonstration that cholesterol experiences different interactions in the lipid matrix above and below 33 mol %. This conclusion is further supported by Kahane and Razin (1977), who had previously shown that dispersions of dipalmitoylphosphatidylcholine and cholesterol with a high sterol content (>35 mol %) were more effective cholesterol donors to Mycoplasma membranes than dispersions with a lower sterol content (<35 mol %). A similar ability of cholesterol to modulate its own exchange kinetics has been reported in Mycoplasma gallisepticum, where a significant decrease in the rate of cholesterol exchange between these cells and lipid vesicles is detected as the cholesterol/ phospholipid molar ratio is lowered from 0.36 to 0.25 (Clejan and Bittman, 1984a). In contrast to all of these studies, it must be pointed out that McLean and Phillips (1982) were unable to detect a change in the activation energy for cholesterol movement from egg yolk phosphatidylcholine-cholesterol vesicles with increasing cholesterol content. Cholesterol depletion is similarly influenced by the chemical composition of the acceptor. Incubation of mammalian cells in culture with serum albumin-phospholipid complexes for a given time results in an increase in cholesterol depletion with increasing chain length of the lipid (up to stearate) and a decrease in depletion with increasing degree of unsaturation (Bartholow and Geyer, 1982). Depletion of cholesterol from vesicular stomatitis virus by serum lipoproteins is affected by enriching the latter with exogenously added phospholipids (Pal et al., 1981). Addition of bovine brain sphingomyelin or dipalmitoylphosphatidylcholine results in an almost equally effective depletion of the sterol from the virion. Enrichment with egg phosphatidylcholine is slightly less effective, whereas incorporation of either bovine brain phosphatidylserine or egg phosphatidylethanolamine into the lipoproteins is relatively ineffective in cholesterol depletion. The data that have been presented are all consistent with the currently held view of the mechanism for cholesterol transfer between membranes. Concerning this process, Hagerman and Could (1951) stated in their original paper: The simplest assumption (referring to the mechanism of exchange) is that equilibria exist between cholesterol in each of the various lipoproteins concerned and cholesterol in the unbound form. This last might be present in minute concentration, possibly in true aqueous solution, and still be capable of acting as intermediate in a rapid equilibration of lipoprotein cholesterol.

It is unfortunate that these words went unheeded for almost 25 years. The suggestion by Curd (1960) that the exchange and transfer of phos-

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pholipids, triglycerides, cholesterol, and cholesterol esters would be brought about by collision between membranes appears to have dominatcd thoughts on the mechanism of these processes for several years even though this latter proposal was not based on any experimental observation but was a suggestion based solely on the view that the above mentioned lipid molecules are essentially insoluble in water. In support of the model for cholesterol exchange presented by Hagerman and Gould ( 1 9 5 Vandenheuvel ~ (1966) proposed that lipid molecules could escape from membranes as a result of the thermal motion of the water molecules. Despite this suggestion, it is interesting to follow the discussion of Quarfordt and Hilderman (1970) to obtain a sense of how strong the feelings were at the time for a collisionally dependent mechanism for cholesterol transfer. After finding that the flux of cholesterol between lipoprotein and red cells remained constant over a wide range of lipoprotein concentrations, Quarfordt and Hilderman (1970) concluded that their results were not incompatible with the “collision complex” concept proposed by Gurd (1960). Although they admitted that their data were compatible with a “probably unlikely” water-soluble intermediate, this possibility was ruled out since cholesterol exchange had not been detected when red cells and lipoproteins were separated by a semipermeable dialysis membrane. Details of these latter observations were not presented in the paper. Evidence in favor of a collisionally dependent mechanism for cholesterol exchange has been presented in several studies (Bruckdorfer and Green, 1967; Lenard and Rothman, 1976; Haran and Shporer, 1977; Poznansky 1978: Jonas and Maine, 1979; Giraud and and Lange, 1978; Moore et d., Claret, 1979; Gottlieb, 1980; Lippiello and Waite, 1983). However, it should be pointed out that in some of these studies the data that were presented in support of this conclusion were scant. In a preliminary communication, Backer and Dawidowicz (1978) presented kinetic data for the exchange of cholesterol between small unilamellar lipid vesicles (SUVs) which were not consistent with the idea that exchange was occurring via a collision complex. Evidence was also prcsented for the transfer of cholesterol between SUVs separated by a dialysis membrane, from which it was concluded that cholesterol exchange occurs via a water-soluble intermediate. Preliminary data presented by Phillips et ril. (1980) supported this point of view for the exchange of cholesterol between SUVs and between cells in culture and SUVs. I n the subsequent detailed accounts of these studies (Backer and Dawidowicz, l981b; McLean and Phillips, 1981) it was demonstrated that the rate of cholesterol exchange between SUVs remained constant over a wide range of donor:acceptor concentrations, a result which is inconsistent with a collisionally dependent transfer process. Both groups presented data in s~ipport

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of an aqueous diffusion model which indicated that cholesterol transfer between SUVs could be detected when the vesicles were separated by a dialysis membrane. These latter data, which are important direct evidence in support of a water-soluble intermediate in the exchange process, have been substantiated by other workers (Kreuter rf a / . , 1981; Bruckdorfer et a/., 1984; Clejan and Bittman, 1984a). Kinetic measurements, indicating that the rate of cholesterol transfer between membranes is independent of the acceptor concentration, were later reported to be consistent with the aqueous diffusion model for cholesterol efflux from cells (Rothblat and Phillips, 1982) and cholesterol exchange between human serum high- and low-density lipoproteins (LundKatz rt a / . , 1982) and between human erythrocytes and plasma (Lange rf a / . , 1983). Other authors (Jonas and Maine, 1979; Giraud and Claret, 1979; Gottlieb, 1980; Lippiello and Waite. 1983) have argued that their kinetic data, where a dependence of the transfer rate on acceptor concentration has been found, support a collisionally dependent exchange of cholesterol. However, Bojesen (1982) has indicated that cholesterol movement between dissimilar donors and acceptors can exhibit a dependence on the acceptor concentration yet still be consistent with an aqueous diffusion mechanism. Desorption of cholesterol appears to be the rate-limiting step in the transfer process (McLean and Phillips, 1981), yet it is not clear whether the sterol moves through the aqueous phase in a monomer or micellar form. Haberland and Reynolds (1973) reported a critical micellar concentration (CMC) for cholesterol of 25-40 nM at 25°C. Sinensky (1981) also reported the existence of cholesterol micelles with a CMC between 13 and 65 nM. Backer and Dawidowicz (1981b) detected a free cholesterol concentration of 190 nM [well below the maximum solubility of 4.7 KM reported by Haberland and Reynolds ( 197311 after incubating lipid vesicles at 37°C for I hr. These data would suggest that cholesterol transfers between membranes in a micellar form. In a careful study on the nature of dilute aqueous cholesterol suspensions, Renshaw et ul. (1983) were unable to obtain any evidence for the existence of a cholesterol micelle. These authors were only able to detect microcrystals of cholesterol above a concentration of 10 nM, which would suggest that cholesterol transfer between membranes occurs via the monomeric form. The apparent discrepancies between these aforementioned studies are not clear. More direct approaches are required in order to determine the physical state in which cholesterol transfers between membranes. The dependence of cholesterol exchange on the lipid composition of the donors, described above, is consistent with the finding that the ratedetermining step in the exchange process is desorption from the donor membrane. Furthermore, the decrease in the rate of this exchange process

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from SUVs containing N-palmitoylsphingomyelin compared with vesicles containing dipalmitoylphosphatidylcholine is consistent with the finding by Demel ef al. (1977)that cholesterol has a preferential affinity for sphingomyelin over phosphatidylcholine. With these latter data in mind, Wattenberg and Silbert (1983)tested their hypothesis that the nonuniform distribution of cholesterol among the various membranes of a mammalian cell is related to the membrane lipid composition. In particular these authors noted that in comparing endoplasmic reticulum ( E R ) , mitochondria, and plasma membrane (PM) there is an increase in both the cholestero1:phospholipid ratio and sphingomyelin content (PM > mitochondria > E R ) . Furthermore the degree of unsaturation of the phospholipid acyl chains increases from PM to mitochondria to ER. However, measurements of the equilibrium partitioning of cholesterol between egg lecithin-cholesterol vesicles and lipid vesicles prepared from the lipid extract of either PM, mitochondria, or ER showed only qualitative agreement with the . these data Wattenberg sterol content of these membranes in i . i \ ~ ~From and Silbert (1983) concluded the distribution of sterol within cells is not an equilibrium phenomenon. As indicated earlier, incubation of cells with cholesterol-free dispersions of phospholipids results in a depletion of cholesterol from the cells. This technique has been used to investigate the role of cholesterol in various cellular functions. In the development of this approach Bruckdorfer e t ul. (1969) showed that cholesterol depletion of human erythrocytes resulted in an increase in permeability of the nonelectrolyte glycerol. This is consistent with the current knowledge that decreasing the cholesterol content of red cell lipids decreases the order of the hydrocarbon chains (Demel and de Kruyff, 1976). Bruckdorfer ct r i l . (1969) also demonstrated that removal of 33% of the cholesterol from the erythrocyte (to a molar ratio for phospho1ipid:cholesterol of 2: I ) results in hemolysis of the cells. These latter data support the notion presented earlier in this chapter that the organization of cholesterol in phospholipid bilayers undergoes a change at a phospho1ipid:cholesterol molar ratio of 2 : I . Using a similar approach Deuticke and Hruska (1976)found an increase in the nonelectrolyte permeability of erythrocytes on lowering the phospho1ipid:cholesterol molar ratio to approximately 2 : I . In this latter study the authors were able to demonstrate that the effect of cholesterol depletion on glycerol permeability was reversible, an essential control in relating changes in function with changes in cholesterol content of the membrane. Hoover et crl. (1983) depleted the cholesterol content of murine lymphocytes by incubation with lipid vesicles and found that capping of surface immunoglobulin receptor was inhibited. The capping process was restored after restoring the cholesterol level by incubation with cholesterol-containing vesicles, thereby demonstrating a role of cholesterol in the capping

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process. Cholesterol depletion from liver plasma membranes in vitro using lipid vesicles is accompanied by a marked increase in 5'-nucleotidase activity (Whetton and Houslay, 1983) whereas increasing cholesterol levels above normal has no effect on activity, indicating that the enzyme is not regulated by the order of the bulk lipid in the membrane. In both of these studies (Hoover et ul., 1983; Whetton and Houslay, 1983) the authors suggested that in the native membrane the proteins under investigation exist and function in a relatively rigid microenvironment which can be modulated by cholesterol depletion. B. Phospholipids

The spontaneous exchange of phospholipids between membranes is a very slow process and is probably not of physiological significance. In fact the lack of spontaneous movement of labeled phospholipids between isolated microsomes and mitochondria lead to the proposal for the existence of transfer proteins (Wirtz and Zilversmit, 1968; McMurray and Dawson, 1969). The exchange of phospholipids that occurs between lipoproteins has been extensively covered in the review by Bell (1976), and will pot be discussed in this chapter. Phospholipid and cholesterol move spontaneously in an independent manner between membranes (Jonas and Maine, 1979; McLean and Phillips, 1981). The half-time for the transfer of cholesterol between lipid vesicles is 2.3 t 0.3 hr at 37"C, whereas the corresponding value for l-palmitoyl2-oleoylphosphatidylcholine is 48 4 5 hr (McLean and Philips, 1981). Movement of cholesterol between Mycoplusmu gullisepticum and lipid vesicles is 10-fold faster than that of phospholipid (Clejan and Bittman, 1984a). The spontaneous transfer of phospholipids is modulated by the physical state (McLean and Phillips, 1984)and cholesterol content (Clejan and Bittman, 1984a) of the donor, the rate of transfer decreasing either below the phase transition of the phospholipid or with increasing cholesterol content. Martin and MacDonald (1976) were the first to investigate the mechanism of spontaneous phospholipid transfer between lipid vesicles. These authors demonstrated that the vesicles composed of dimyristoylphosphatidylcholine (DMPC) are in equilibrium with a low concentration of monomer DMPC which is transferred from donor to acceptor in a net manner. Thus, Martin and MacDonald were the first to provide evidence for the aqueous diffusion model of lipid transfer between membranes. Comparable data were obtained by Papahadjopoulos et a/. (1976). Duckwitz-Peterlein et a / . (1977) reached a similar conclusion in their study of phospholipid transfer between vesicles prepared from Escherichiu colilipid extracts. Subsequent analyses of these kinetic data by Thilo (1977)

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and Duckwitz-Peterlein and Moraal (1978) led to the conclusion that the rates of phospholipid transfer are determined by the off rate from the donor vesicles and that this is dependent on the acyl chain length of the phospholipid, the spontaneous transfer rate of phospholipids decreasing with increasing chain length. The aqueous diffusion of phospholipids has been supported by kinetic measurements of spontaneous phospholipid exchange between lipid vesicles (McLean and Phillips, 1981; De Kuyper o r a / ., 1983; McLean and Phillips, 1984)and between Mycoplcisma gcilli.sc.pticirrn and lipid vesicles (Clejan and Bittman, 1984a).However, kinetic studies have supported a collisionally dependent transfer process (Kremer et a / . , 1977; Jonas and Maine, 1979; Lippiello and Waite, 1983). Fluorescence techniques have been widely used to investigate the mechanism of the spontaneous lipid transfer between membranes. The transfer of pyrene-containing molecules can be monitored by utilizing the property that in the donor membrane, where the concentration of the pyrene-labeled molecule is high. excimer fluorescence is observed. Upon transfer to the acceptor, a distinct monomer fluorescence is observed. Thus, the transfer of pyrene-labeled molecules is followed by the increase in monomer fluorescence (Charlton rt N / . , 1976; Sengupta et d., 1976). An alternate approach is based on fluorescence quenching. Incorporation of the fluorescent phospholipid I-acyl-2-[12-1(7-nitro-2,I ,3-benzoxadiazol4-yl)amino]dodecanoyI]phosphatidylcholine (C,,-NBD-PC) into donor lipid vesicles at concentrations greater than SO mol VJresults in a self-quenching of >98% of the fluorescence of the C,,-NBD-PC. Transfer of this tluorescent phospholipid from the donors to nonfluorescent acceptor vesicles results in an increase in fluorescence intensity (Nichols and Pagano, 1981). One caveat in all of these fluorescence measurements is that the attachment of the fluorescent group to the phospholipid may change the transfer properties of the molecule. In order to conclude that spontaneous transfer of phospholipids between membranes occurs via the aqueous phase, most workers have used the kinetic criterion that this mechanism would result in a transfer rate which is independent of the acceptor membrane concentration. However, as mentioned earlier for cholesterol transfer, a dependence of the transfer rate on acceptor concentration is not inconsistent with aqueous diffusion (Bojesen, 1982). Roseman and Thompson (1980)reported that the spontaneous transfer of I-palmitoyl-2-pyrenedecanoylphosphatidylcholine between lipid vesicles is independent of the acceptor concentration, and they concluded that this process occurs by aqueous diffusion with a half-time of 13 h r at 37°C. Correa-Freire et nl. (1982)showed that the spontaneous transfer of glucocerebroside has a half-time of approximately 32 days and reasoned that the lack of spontaneous transfer for glycolipids is consistent with

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their function as cell surface markers and receptors. The transfer of pyrenelabeled sphingomyelin between apolipoprotein-lipid complexes also proceeds via the aqueous phase (Pownall et d . , 1982). These model lipoproteins were subsequently used as donors in a systematic study of the effects of various parameters on the transfer of pyrene-labeled phosphatidylcholines (Massey et al., 1982a,b). The authors remarked that since there is no compartmentalization of lipids into an inner and outer monolayer in the apolipoprotein-lipid recombinants the analysis of the data is simplified. However, it is known that apolipoproteins can themselves undergo spontaneous movement between lipoproteins (Eisenberg, 19781, which could facilitate the movement of the phospholipid. Since the fluorescence techniques negate the necessity to physically separate donors and acceptors in order to follow the exchange process, controls must be conducted to indicate that the apoproteins are not being transferred during the experiment. Massey et af. (1982a) reported that the transfer rate of pyrene-labeled phosphatidylcholine is increased by decreasing the fatty acyl chain length or by adding neutral salts to the aqueous phase which they suggested increased the solubility of the labeled lipid. The transfer of these labeled molecules is affected by the physical state of the lipid, decreasing significantly below the lipid phase transition, and by the lipid-protein nature of the donor particles. From the transfer kinetics it was concluded that pyrene-labeled phosphatidylcholines are transferred between membranes via the aqueous phase. Transfer of fluorescent derivatives of phosphatidylglycerol, phosphatidylserine, phosphatidylcholine, phosphatidylethanolamine, and diacylglycerol occurs by the same mechanism (Massey et al., 1982b). The transfer rates of these various phospholipids appeared to be predictable, to a first approximation, based on the hydrophobic content of the molecule (Massey et al., 1982b). In a subsequent paper Massey et al. (1984) indicated that the rate of spontaneous transfer of phospholipids between plasma lipoproteins can be predicted from their hydrophobicity, as measured by retention on a reversed-phase (C,J high-performance liquid chromatography column. A conclusion reached in this study was that the predicted rate for the spontaneous transfer of naturally occurring phospholipids would be too slow to be of physiological significance. Covalent modification of one of the acyl chains of a phospholipid with the fluorescent NBD reporter group dramatically increases its rate of spontaneous transfer (Struck and Pagano, 1980). Indeed it is this property of these fluorescently labeled phospholipids that has enabled Pagano and co-workers to introduce them from lipid vesicles into the outer surface of cells (Pagano, 1983). In a detailed kinetic analysis, Nichols and Pagano (198I ) concluded that these NBD-labeled phosphatidylcholines transfer

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between lipid vesicles via the aqueous phase and that the rate-determining step for this process is the off rate from the donor membrane. Frank rt al. (1983) have correlated the spontaneous transfer of sphingomyelin with lateral organization in the donor lipid bilayer. Initially it was demonstrated that the half-time for the transfer of either pyrene or 'H-labeled sphingomyelin from a pure liquid-crystalline bilayer is about two orders of magnitude shorter than from a corresponding gel phase. The half-time for the transfer of sphingomyelin from a bilayer of I-palmitoyl-2-oleoylphosphatidylcholine(POPC) at 30°C (where POPC is in the liquid-crystalline phase) was shown to be characteristic of transfer from a gel phase. The authors suggested that this could be due to the existence of a gel phase, rich in sphingomyelin, in the liquid-crystalline matrix even at low sphingomyelin concentrations. Evidence suggesting the existence of glycosphingolipid-enricheddomains in a phospholipid bilayer has been similarly obtained from measurements of the spontaneous transfer of gangliotetraosylceramide between bilayers. which occurs via the aqueous phase (Brown et a / . , 1985). Thus, analysis of spontaneous lipid transfer appears to be able to provide new information concerning the lateral organization of lipids in membranes. C. Other Molecules

The spontaneous transfers of pyrene, free fatty acids, diglycerides, and cytochrome b, between membranes have been reported. Movement of pyrene between either lipid vesicles (Sengupta et al., 1976)or high density lipoproteins (Charlton et a/., 1976) is an extremely rapid process, occurring with a half-time of seconds or less via the aqueous phase. Pyrenedecanoic acid (Sengupta et a / . , 1976)and 943-pyreny1)nonanoic acid (Doody et al.. 1980) have been shown to transfer between membranes by aqueous diffusion with a half-time of seconds. A similar mechanism has also been reported for the rapid (l,,? of seconds) transfer of the fluorescent diglyceride rac- 1-oleyl-2-[4-(3-pyrenyl)butanoyl]glycerol(Charlton rt a / ., 1978). Cytochrome hs, a microsomal enzyme, is also capable of a spontaneous vesicle-vesicle transfer that has been shown to occur via the aqueous phase (Leto rr a / . , 1980). Gurd (1960) had ruled out the possibility that lipid transfer could occur by aqueous diffusion on the basis of the insolubility of lipids in water. I t is therefore of interest to note that the measured solubility of pyrene in water is less than 0.4 FM (Charlton et a/.. 1976) and the solubility of the fluorescent diglyceride is less than 25 nM. Nevertheless, kinetic measurements for the spontaneous transfer of these two molecules are consistent with transfer via the aqueous phase.

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In concluding this section on the spontaneous transfer of lipids between membranes, it would appear that an overwhelming amount of data supports the idea that transfer of a variety of lipid molecules between various membranes proceeds by aqueous diffusion which does not involve membranemembrane contact. 111.

PROTEIN-MEDIATED LIPID TRANSFER BETWEEN MEMBRANES

Wirtz and Zilversmit (1968) first demonstrated that the slow movement of labeled phospholipids between microsomes and mitochondria in vitro could be stimulated by the 105,000 I: supernatant from a rat liver homogenate. Similar findings were reported by McMurray and Dawson (1969). It was suggested that transfer factors, later identified as proteins, could be responsible for the movement of newly synthesized phospholipids from their site of synthesis within a cell (endoplasmic reticulum) to other membranes which are incapable of de novo lipid biosynthesis, an idea first proposed by Dawson (1966). Subcellular localization of a phosphatidylcholine transfer protein (PC-TP) in rat liver indicates that approximately 60% of this protein is present in the 105,000 g supernatant fraction of a liver homogenate, the remainder being evenly distributed among particulate fractions (Teerlink et ul., 1982), suggesting a dynamic state between membrane-bound and soluble PC-TP. Teerlink et ul. (1982) further showed that the highest levels of the PC-TP in the various tissues of the rat occur in the liver and intestinal mucosa, both of which are very active in phosphatidylcholine metabolism. Despite the identification and purification of several lipid transfer proteins from a variety of different tissues, their physiological role still remains unclear. Nevertheless, these transfer proteins have proved to be excellent probes of lipid asymmetry (Etemadi, 1980) and transmembrane movement of lipids, and they provide a means for altering membrane lipid composition in vitro (Zilversmit, 1978). In order for transfer proteins to be involved in membrane biogenesis, they must be capable of net movement of lipid between membranes. Initial studies describing purification of these proteins employed assays which measured phospholipid exchange between membranes, hence the term phospholipid exchange protein was used. More recently it has become apparent that these exchange proteins can also function in a net transfer mode, thus the term transfer protein has now found favor. To illustrate this point, the purification of a phosphatidylcholine-specific transfer protein (PC-TP) from beef liver is monitored by the exchange of labeled phosphatidylcholine between microsomes and phospholipid vesicles (Westerman et ul., 1983) If one of the membranes lacks phosphatidylcholine, the

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PC-TP catalyzes a net transfer of this phospholipid (Wilson ~t d., 1980: Wirtz e t a / . , 1980). Net transfer of phospholipids between membranes has also been demonstrated with the transfer protein from beef heart (Kagawa et d., 1973) and the nonspecific transfer proteins from bovine liver (Crain and Zilversmit, 1980a). Kasper and Helmkamp (1981) have shown that in the case of the bovine brain phospholipid exchange protein a net unidirectional flux of phosphatidylinositol into a phosphatidylcholine vesicle is accompanied by a compensatory flux of phosphatidylcholine in the opposite direction. Thus a true molecular exchange, rather than net transfer, was proposed for the action of the bovine brain exchange protein. It is difticult to reconcile this latter finding with a role for this particular protein in membrane biogenesis. Lipid transfer proteins with varying specificity for phospholipids and cholesterol have been isolated and purified from diverse sources. Transfer proteins for esterified cholesterol have been identified and purified from plasma. Properties of these latter proteins have been reviewed (Barter c’t o/., 1982) and will not be discussed here. Wirtz and his colleagues have described a large-scale purification of the phosphatidylcholine-specific transfer protein (PC-TP) from beef liver (Westerman et a/., 1983) for which 1981) and they have determined the primary sequence (Akeroyd et d., characteristics of phosphatidylcholine binding (Berkhout et a / . , 1984). Detailed purification schemes have been published for the following: nonspecific lipid transfer proteins from liver (Bloj and Zilversmit, 1983; Poorthuis and Wirtz, 1983); a phosphatidylcholine-specifictransfer protein from rat liver (Teerlink et a/., 1983);and a cerebroside transfer protein (Radin and Metz, 1983). Diversity of the sources from which transfer proteins have been isolated is indicated by the recent accounts of phospholipid transfer proteins from maize seedlings (Douady ct u / . , 1982), yeast (Daum and Paltauf, 1984), and human plasma (Albers r t a / . , 1984). Other transfer proteins have been purified. The reader is referred to the reviews on this subject, mentioned in the introduction of this chapter, for further information. The mechanism of protein-mediated lipid transfer is not clear. Transfer rates are affected by both the hydrophobic and hydrophilic nature of the donor membrane. Welti and Helmkamp ( 1984) have demonstrated that the phosphatidylcholine-specific transfer protein (PC-TP) from bovine liver will not transfer lipids in the gel state to any significant extent, and that this protein has a preference for long-chain phosphatidylcholines in the 1981) and fluid phase. The PC-TP from bovine liver (Somerhaju et d., the phosphatidylinositol transfer protein (PI-TP) from bovine brain (Somerhaju c>t d . , 1983; Yoshimura and Helmkamp, 1984) are affected by the presence of charged lipid. It has been shown that the PC-TP from bovine liver binds phosphatidylcholine in a 1: I molar ratio (Demel et d . ,

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1973; Wirtz, 1974). Similar binding of phosphatidylcholine has been reported for purified transfer proteins from bovine heart (Johnson and Zilversmit, 1975) and maize seedlings (Douady et a l . , 1982). The PI-TP from bovine brain binds both phosphatidylcholine (Helmkamp et ul., 1976) and phosphatidylinositol (Somerhaju et al., 1983). There is strong evidence that both the PC-TP from bovine liver (Demel er al., 1973) and the PI-TP from bovine brain (Helmkamp et al., 1976) function as carriers of phospholipid. The other transfer proteins, for which phospholipid binding has not been reported, could enhance the off rate of the phospholipid from the donor membrane. Since this dissociation from the membrane is the rate-determining step in lipid transfer (as discussed earlier in this chapter) enhancement of this step would facilitate the overall transfer process. Lipid transfer proteins have been used to modulate both the phospholipid and cholesterol content of membranes in vitro, to investigate the lipid dependence of certain membrane functions. This approach has the advantage that a particular membrane protein need not be purified and reconstituted. However, if the protein under investigation has a high affinity for a particular phospholipid, this lipid may not be removed by transfer proteins. Barsukov et al. (1978) used a phospholipid transfer protein to incorporate phosphatidylcholine into protoplasts of Micrococcus lysodeikricus, thereby restoring the permeability barrier of the initial protoplast membrane. Transfer proteins have been used to investigate the lipid dependence of glucose-6-phosphatase (Dyatlovitskaya er al., 1979; Crain and Zilversmit, 1981). Nonspecific lipid transfer proteins have been used to modulate the cholesterol content in synaptic membranes (North and Fleischer, 1983); to investigate the effect of cholesterol content on the activity of acyl-CoA:cholesterol acyltransferase (Poorthuis and Wirtz, 1982) and hydroxymethylglutaryl-CoA reductase (van Heudsen and Wirtz, 1984) in isolated microsomes; and to modify both the phospholipid content and composition of intact human erythrocytes (Franck et a l . , 1984). Voelker and Kennedy (1982) used the phosphatidylcholine-specifictransfer protein to demonstrate that synthesis of sphingomyelin in BHK 21 cells is localized at the plasma membrane. These authors introduced ['H]phosphatidylcholine into the plasma membrane of BHK 21 cells from lipid vesicles via the transfer protein and showed that this molecule is the source of phosphocholine in the synthesis of sphingomyelin. In an attempt to define a role for lipid transfer proteins in membrane biogenesis, Yaffe and Kennedy (1983) compared the relative rates of transfer of newly synthesized phospholipids to mitochondria in vivo in BHK-21 cells with the relative rates of transfer of these lipids from liposomes to mitochondria in vitro stimulated by a partially purified phospholipid transfer protein isolated from these cells. No correlation was observed, leading these authors to conclude that this transfer protein was

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not involved in the transport of newly synthesized lipids to the mitochondria of these cells in vivo. The physiological role for glycolipid transfer proteins (Radin and Metz. 1982) is also somewhat puzzling. The glycosylating enzymes face the lumen of the endoplasmic reticulum and Golgi, yet the glycolipid transfer proteins appear to be cytoplasmic. How is it possible that these proteins which do not penetrate across membranes ever have access to the glycolipid substrates in living cells?

IV. TRANSMEMBRANE MOVEMENT OF LIPIDS A. Phospholipids

Kornberg and McConnell (1971) were the first to report the transmembrane movement of a phospholipid in lipid vesicles using a spin-labeled phosphatidylcholine. These authors measured a half-time for this process of 6.5 hr at 30°C. Subsequently, Rothman and Dawidowicz (1975) and Johnson et al. (1975) showed that phospholipid transfer proteins could be used to investigate this process. In these studies transmembrane movement of radiolabeled phosphatidylcholine in lipid vesicles was not detectable, and from the precision of the measurements a lower limit of 1 1 days was estimated for this process at 37°C. The reason for the discrepancy between these measurements and those using the spin-labeled phospholipid is not clear. Roseman et al. (1975) were also unable to detect the transbilayer movement of phosphatidylethanolamine using a chemical labeling procedure, and from the precision of their measurements estimated a halftime for this process of at least 80 days at 22°C. Thus a spontaneous transmembrane movement of phospholipids across a pure lipid bilayer would seem to be an unlikely event. Measurements of the transmembrane movement of phospholipids in several biological membranes have been reported. It would appear that the data can be divided into three groups. In the first group, the reported half-times for transmembrane movement of phospholipids are extremely long, of the order of several days. This has only been detected in the membrane of the influenza virus (Rothman r t ul., 1976)and in a derivative of the plasma membrane of LM cells (Sandra and Pagano, 1978). In the second group, the reported half-times for transmembrane movement of phospholipids are in the range of 2-7 hr. These include vesicular stomatitis virus membranes (Shaw et ul., 1979) and intact erythrocyte membranes from humans and rats (van Meer et al., 1980; Crain and Zilversmit, 1980b). In the third group are two membranes, microsomal vesicles from rat liver (Zilversmit and Hughes, 1977; van den Besselaar et ul., 1978)and Bacillus rnegaterium (Rothman and Kennedy, 1977), where the measured half-times

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for the transmembrane movement of phospholipids are a few minutes or less. Can any sense be made from these measurements? It would appear that very slow transmembrane movement of phospholipids of days) occurs mainly in pure lipid bilayers. This process is somewhat faster (tl,?of hours) in biological membranes incapable of de n o w phospholipid biosynthesis, whereas it is extremely rapid (f,,, of minutes) in membranes capable of conducting de n o w phospholipid biosynthesis. Attempts have been made to determine possible factors that might induce transmembrane movement of phospholipids in lipid vesicles. Addition of cytochrome oxidase by reconstitution has no effect (DiCorleto and Zilversmit, 1979). Inclusion of intrinsic proteins from the human erythrocyte membrane (de Kruijff et ul., 1978; Gerritsen et a / . , 1980), cytochrome P450 (Barsukov et ul., 1982),or small amounts of detergent (Kramer c>t a l . , 1981) induce transmembrane movement of phospholipids (tl,?of hours) in lipid vesicles. Asymmetrical perturbations of the lipid bilayer with exchange proteins (de Kruijff and Wirtz, 1977; de Kruijff and van Zoelen, 1978) or digestion by phospholipase D (de Kruijff and Baken, 1978) as well as oxidation of the phospholipids (Shaw and Thompson, 1982) have resulted in transbilayer movement of the phospholipids with half-times of hours. Similar half-times have been reported for the transbilayer movement of phosphatidylcholine in lipid vesicles containing unsaturated phosphatidylethanolamine which would form “nonbilayer” structures in the membrane (Noordarn et al., 1981). However, in none of these studies has the rapid transmembrane movement of phospholipids observed in microsomes and B . meguterium been reconstituted. van den Besselaar et al. (1978) were unable to detect any transmembrane phospholipid movement in vesicles prepared from the total lipid extract of rat liver microsomes. This finding suggests the possible involvement of a protein which facilitates transmembrane movement in the intact microsomal membrane. This proposal could have some physiological significance since it has been demonstrated that the active sites of phospholipid biosynthetic enzymes are localized on the cytoplasmic surface of isolated microsomal vesicles derived from rat liver (Bell ef ul., 1981). The lipid in the membrane of the endoplasmic reticulum is organized in a bilayer, indicating that a mechanism must exist for the translocation of the newly synthesized phospholipid to the lumenal surface. Studies of phospholipid movement across lipid bilayers by various asymmetrical perturbations and the study of van den Besselaar et 01. (1978) suggest that this cannot be accomplished by lipid alone and thus might involve a protein. Bretscher (1973) proposed the existence of such proteins for which he coined the name “flippase.” Models for this hypothetical protein have been presented by Zilversmit (1978) and Langley and Kennedy (1979).

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Sleight and Pagano (1985) have recently shown that a fluorescent phosphatidylethanolamine, which does not readily traverse a pure lipid bilayer, undergoes a rapid transmembrane movement (t,,, of the order of minutes) across the plasma membrane of lung fibroblasts. In contrast these authors had previously shown that transmembrane movement of the corresponding fluorescent phosphatidylcholine across the plasma membranes of these same cells is not detectable (Sleight and Pagano, 1984). These combined data could reflect a “flippase” in the plasma membrane of the fibroblasts. An intriguing study indicates that the redistribution of spin-labeled phospholipids across the erythrocyte membrane is ATP dependent (Seigneuret and Devaux, 1984). This result is in contrast to the rapid transmembrane movement of phosphatidylethanolamine in membranes of B . megtrterirrm, which does not require metabolic energy (Langley and Kennedy, 1979).

B. Cholesterol Bruckdorfer et al. (196th) demonstrated that cholesterol could traverse a lipid bilayer and the membrane of the human erythrocyte ghost with half-times of the order of hours. Unfortunately these data were overlooked when Poznansky and Lange (1976) reported that the transbilayer movement of cholesterol in lipid vesicles was not detectable and thus placed a lower limit of 6 days at 37°C for the half-time of this process. To add to the confusion, Lenard and Rothman (1976) reported a half-time of about 13 days at 37°C for the transmembrane movement of cholesterol across the membrane of influenza virus. These extremely long half-times have not been reported in similar studies of either lipid vesicles or biological membranes. Bloj and Zilversmit ( 1977a) showed that cholesterol movement across lipid bilayers occurred with a half-time of less than 2-5 hr depending on the degree of unsaturation of the phospholipid. These data have been confirmed by Backer and Dawidowicz (1979). Since both these studies employed cholesterol exchange techniques, the transmembrane movement of cholesterol was not an artifact of cholesterol depletion as suggested by Poznansky and Lange (1978). Exchange techniques have been used to demonstrate that the transmembrane movement of cholesterol occurs readily across the membranes of the following: vesicular stomatitis virus (Patzer ef al., 1978; Sefton and Gaffney. 1979); Sindbis virus (Sefton and Gaffney, 1979); Myc.oplusn~ugallisc~ptic~m (Rottem ct d.,1981); human erythrocytes (Lange ut NI.. 1977; Kirby and Green, 1977; Bloj and Zilversmit. 1977b); and rabbit intestinal brush border (Bloj and Zilversmit. 1982). In several of these studies the complete exchange of cholesterol occurred with a single time constant, indicating that the transmembrane movement occurs more rapidly than intermembrane exchange.

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Backer and Dawidowicz (1981a) used the enzyme cholesterol oxidase to demonstrate that the transbilayer movement of cholesterol in lipid vesicles is an extremely rapid process, occurring with a half-time of 1 min or less at 37°C. Lange et al. (1981) showed that the half-time of this process in the human erythrocyte has an upper limit of 3 sec at 37”C, using a similar approach. Huang et al. (1970) measured a half-time of 42 min at room temperature [data reported by Thompson ( 1978)] for the transbilayer movement of the cholesterol analog thiocholesterol in lipid vesicles by titrating with 5,5’dithiobis(2-nitrobenzoicacid) (DTNB). Dawidowicz and Backer (198 1) later demonstrated that thiocholesterol could traverse a lipid bilayer with a half-time of 1 min or less at 20°C. Thus, although a physiological role for this process is not clear, it can be stated that cholesterol undergoes a rapid transmembrane movement across both lipid bilayers and biological membranes. Unlike cholesterol, alkyl-substituted sterols such as ergosterol, p-sitosterol, and stigmasterol do not appear to readily cross the membrane of Mycoplusma capricolurn (Clejan et al., 1981). C. Other Molecules

Doody et al. (1980) have reported that the transbilayer movement of a pyrene-labeled nonanoic acid across lipid vesicles occurs in a fraction of a second. However, Abumrad et al. (1984) have provided evidence that the transfer of long-chain fatty acids into adipocytes is protein mediated (a “flippase”?). A possible explanation is that the fatty acid investigated by Doody et a/. (1980) is not physiological and can therefore cross a bilayer more readily than longer chain fatty acids. 1,2-Diacylglycerol, undergoes a rapid transbilayer movement in erthrocytes with a half-time of 1 min or less at 37°C (Allan et al., 1978). The sulfhydryl analog of this molecule, dioleoylthioglycerol, can move across a lipid bilayer with a half-time of 15 sec or less, whereas the transbilayer movement for phosphatidylthioglycerol is in excess of 8 days (Ganong and Bell, 1984). These latter data indicated to the authors that the primary barrier to the transmembrane movement of lipids must reside in the nature of the polar head group.

ACKNOWLEDGMENTS The author wishes to acknowledge the collaboration and contributions of Jonathan Backer to several studies from the author’s laboratory quoted in this review, which were funded by grants from the National Institutes of Health and the American Heart Association.

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REFERENCES Abumrad, N. A., Park. J. H.. and Park, C. R. (1984). Permeation of long-chain fatty acids into adipocytes. Kinetics, specificity. and evidence for involvement of’ a membrane protein. J. Biol. Chem. 259, 89448-8953. Agutter, P. S., and Suckling, K. E. (1981). Effects of cholesterol on the properties of the membranes of isolated sheep liver nuclei and nuclear envelopes. Biochini. Eiophvs. AC/U 643, 182-190. Akeroyd, R., Moonen, P., Westerman. J.. Puyk, W. C., and Wirtz. K. W. A. (1981). The complete primary structure of the phosphatidylcholine-transfer protein from bovine liver. Isolation and characterization of the cyanogen bromide peptides. Eicr. 1. B i i ~ c h ~ i114, n. 385-39 1. Albers, J . J . , Tollefson, J. H., Chen, C. -H., and Steinmelz, A. (1984). Isolation and characterization of human plasma lipid transfer proteins Arterinsc./ero.si.s 4, 49-58. Allan. D., Thomas. P.. and Michell, R. H. (1978). Rapid transbilayer diffusion of I.2-diacylglycerol and its relevance to control of membrane curvature, Ntrture fLondoriJ276, 289-290. Backer, J . M.,and Dawidowicz, E. A. (1978). Mechanism of cholesterol exchange between membranes. Circihtion 58 (2), 140. Backer, J . M . , and Dawidowicz, E. A. (1979).The rapid transmembrane movement of cholesterol in small unilamellar vesicles. Biochim. Biophys. Actci 551, 260-270. Backer, J. M., and Dawidowicz, E. A . (1981a). Transmembrane movement of cholesterol in small unilamellar vesicles detected by cholesterol oxidase. J. Biol. Clirm. 256, 586588. Backer, J. M.. and Dawidowicz, E. A. (1981b). Mechanism ofcholesterol exchange between lipid vesicles. Biochemistry 20, 3805-3810. Barsukov, L. 1.. Kulikov, V. I., Simakova, I. M.. Tikhonova, G . V., Ostrovsskii. D. N.. and Bergelson, L. D. (1978). Manipulation of phospholipid composition of membranes with the aid of lipid exchange proteins. Incorporation of phosphatidylcholine into protoplusts of Micrococcus l.v.sodrikriciis. ENT.J . Biochem. 90, 33 1-336. Barsukov, L. I . , Kulikov. V. I., Bachmanova. G . I., Archakov, A. I., and Bergelson, L. D. ( 1982). Cytochrome P-450 facilitates phosphatidylcholine flip-flop in proteoliposomes. FEES Lett. 144, 337-340. Barter, P. J.. Hopkins, G. J., and Calvert. G. D. (1982).Transfers and exchanges of estenfied cholesterol between plasma lipo-proteins. Riochenr. J . 208, 1-7. Bartholow, L. C., and Geyer, R. P. (1981). Sterol release in mammalian cells. The role of ii liposomal-free albumin phospholipid complex. Biochint. Biophys. Actu 665, 4 0 4 7 . Bartholow. L. C . , and Geyer, R. P. (1982). Sterol efflux from mammalian cells induced by human serum albumin-phospholipid complexes. Dependence on phospholipid acyl chain length, degree of saturation and net charge. J. Biol. Chem. 257,3126-3130. Bell, F. P. (1976). Lipoprotein lipid exchange in biological systems. / t i “Low Density t i poproteins” (C. E. Day and R. S. Levy, eds.) pp. I 11-133. Plenum, New York. Bell, F. P. (1978). Lipid exchange and transfer between biological lipid-protein structures. Prog. Lipid Nes. 17, 207-243. Bell, F. P. (1984)The dynamic state of membrane lipids: The significance of lipid exchange and transfer reactions to biomembrdne composition, structure, function, and cellular metabolism. In “Membrane Fluidity” (M. Kates and L. A. Manson. eds.), pp. 543562. Plenum. New York. Bell, R. M.. Ballas. L. M., and Coleman, R. A. (1981). Lipid topogenesis. J . LipidRrs. 22, 39 1-403.

194

ELIEZAR A. DAWlDOWlCZ

Bellini, F.. Phillips, M. C., Pickell, C . , and Rothblat, G . H. (1984).Role of the plasma membrane in the mechanism of cholesterol eftlux from cells. B i w h i m . Biophys. Actu 777, 209-2 15. Berkhout, T. A.. Visser, A . J . W. G., and Wirtz, K. W. A. (1984).Static and time-resolved fluorescence studies of fluorescent phosphatidylcholine bound to the phosphatidyl transfer protein of rat liver. Biochemistry 23, 1505-1513. Bloj, B., and Zilversmit. D. €3. (197720. Complete exchangeability of cholesterol in phosphatidylcholine-cholesterol vesicles of different degrees of unsaturation. Biochc,mi.stry 16, 3943-3948. Bloj. B., and Zilversmit, D. B. (1977b).Transposition and distribution of cholesterol in rat erythrocytes. Proc. Soc. Exp. Biol. Med. 156, 539-543. Bloj, B., and Zilversmit, D. B. (1981). Lipid transfer proteins in the study of artificial and natural membranes. Mol. Cell. Biochem. 40, 163-172. Bloj. B.. and Zilversmit, D. B. (1982). Heterogeneity of rabbit intestine brush border plasma membrane cholesterol. J . B i d . Chem. 257, 7608-7614. Bloj. B.. and Zilversmit, D. B. (1983).Nonspecific lipid transfer protein from rat and beef liver: Purification and properties. Methods Enzymol. 98, 574-58 I. Bojesen, E. (1982).Diversity of cholesterol exchange explained by dissolution into water. Nuticre (London) 299, 216-278. Bretscher, M. S. (1973).Some general principles of membrane structure. In “The Cell Surface in Development” (A. A. Moscona. ed.). pp. 17-27. Wiley, New York. Brown. R. E., Sugar, 1. P.. and Thompson, T. E. (198.5). Spontaneous transfer of gangliotetraosylceramide between phospholipid vesicles. Biochemistry 24, 4082-409I. Bruckdorfer, K. K.. and Graham, J. M. (1976).The exchange of cholesterol and phospholipids between cell membranes and lipoproteins. I n “Biological Membranes” (D. Chapman and D. F. H. Wallach, eds.), Vol. 3, pp. 103-152. Academic Press, New York. Bruckdorfer, K. R.. and Green, C. (1967).The exchange of unesterified cholesterol between human low-density lipoproteins and rat erythrocyte “ghosts.” Biochetn. J . 104. 270277. Bruckdorfer, K. R.. and Sherry, M. K . (1984). The solubility ofcholesterol and its exchange between membranes. Biochim. Biophys. Acttr 769, 187-196. Bruckdorfer, K. R . , Edwards, P. A,, and Green, C. (1968a). Properties of aqueous dispersions of phospholipid and cholesterol. Eur. J . Biochem. 4, 506-51 1. Bruckdorfer, K. R., Graham, J. M., and Green, C. (1968b). The incorporation of steroid molecules into lecithin sols, P-lipoproteins and cellular membranes. Eitr. J . Biochcvn. 4,512-518. Bruckdorfer, K. R., Demel., R. A . , de Gier, J . , and van Deenen, L. L. M. (1969). The effect of partial replacement of membrane cholesterol by other steroids on the osmotic fragility and glycerol permeability of erythrocytes. Biochim. Biophys. Actu 183, 334345. Bruckdorfer, K. R.. Crowe, J., and Sherry, M . K. (1984).Evidence for a water-soluble intermediate in exchange of cholesterol between membranes. Biochim. Biophys. Artu 778, 4x9496. Charlton, S. C., Olson, J. S . , Hong, K-Y., Pownall, H. J., Louie, D. D., and Smith, L. C. (1976). Stopped flow kinetics of pyrene transfer between human high density lipoproteins. J . B i d . Chem. 251, 7952-7955. Charlton, S. C., Hong, K-Y., and Smith. L. C. (1978).Kinetics of ruc-l-oleyl-2-[4-(3-pyrenyl)butanoyl]glycerol transfer between high density lipoproteins. Biochernisfry 17,33043309. Chesterson, C.J. (1968).Distribution of cholesterol precursors and other lipids among rat liver intracellular membranes. J . B i d . Chcm. 243, 1147-1 151.

LIPID EXCHANGE

195

Ckjan. S . , and Bittman. R. (19x44. Kinetics of chole\terol and phospholipid exchange be-

tween Mgcwp/ersiiis gct//isepfic'rtrii and lipid vesicles. Alterations in membrane cholesterol and protein content. 1.B i d . Clrcvti. 259, 441-448. Clejaii, S., and Bittman. R. (1984b). Decreases in the rates of lipid exchange between Mycwplcrsino ,qcr//i.sepfic~ctm and unilamellar vesicles by incorporation of sphingo-myelin. J . BiO/. ChCii1. 259, 10823-10826. Clejan, S.. Bittman, R., and Rottem. S. (1981). Etfects of sterol structure and exogenoub lipids on the transbilayer distribution of sterols in the membrane of M y q h . s t n t r c w pricdroi?. Biochcvni.s/ry 20. 2200-13-04. Colbeau. A.. Nachbaur. J.. and Vignais. P. M . (1971). Enzymic characterization and lipid composition of rat subcellular mcmbranes. Bioc.hirtr. B i o p h y s . Acfer 249, 462-492. Correa-Freire, M. C.. Barenholz, Y.. and Thompson. T. E. ( 1982).Glucocerebroside transfer 21, 1244-1148. between phosphatidylcholine bilayers. Bioc~/rc/iiisrr.~ Crain. R . C. (1982). Non-specific lipid transfer proteins :is probe3 of membrane structure and function. Lipids 17. 935-943. Crain. R . C . . and Zilversmit. D. B. ( IYXOa). Net transfer of phospholipid by the nonspecific phospholipid transfer proteins from bovine liver. Bioc,/iini. B i o p h g s . Actif 620. 3748. Crain. R. C.. and Zilversmit, D. B. ( IY80b). Two nonspecific phospholipid exchange proteins from beef liver. 2. Use in studying the asymmetry and transbilayer movement of phosphatidylcholine, phosphatidylethanolamine and sphingomyelin in intact rat erythrocytes. Biodicini.s/r.v 19, 1440-1447. Crain. R. C.. and Zilversmit, D. B. (1981). Lipid dependence of glucose-6-phosphate phosphohydrolase: A study with purified phospholipid transfer protein and phosphatidylinositol-specific phospholipase C. Biochc~mis/rv20, 5310-5326. Daum. G . . and Paltauf. F. (1984). Phospholipid transfer in yeast. Isolation and partial characterization of a phospholipid transfer protein from yeast cytosol. Biochiir~.B i o p / i y s . A ( ' / ( / 794, 385-391. Dawidowicz. E. A , , and Backer. J . M . (1981). The rapid transbilayer movement of thiocholesterol in small unilamellar phospholipid vesicles. f l i o c , / i i / n ,Biop/1ys. Acfo 644,373375. Dawson, R . M. C. (1966). The metabolism of animal phospholipids and their turnover in cell membranes. In "Essays in Biochemistry" (P. N . Campbell and G. D. Greville, eds.). Vol. 1. pp. 69-1 15. Academic Press. I m d o n . Dawson. R . M . C. (1973). The exchange ofphoxpholipids between cell membranes. S U / J C . C , / / . 8ioc.hc~nr.2, 69-x9. de Kruijff. B., and Baken, P. ( 1978). Rapid Iransbilayer movement of phospholipids induced I 38-47. by an asymmetrical perturbation of the bilayer. Bioc~iiim.Biopiiy.~.A C ~ 507, de Kruijff. B.. and Wirtz. K. W. A. (1977). lnduction o f a relatively fast transbilayer movement of phosphatidylcholine in vesicles. A "C N M R study. Bicdiirti. Biophys. Ac~ci468,318326. de Kruijff. €3.. and van Zoelen. E. J . J. (1978). Effect o f t h e phase transition on the transbilayer movement of dimyristoyl phosphatidylcholine in unilamellar vesicles. Biochitn. Biopkys. ACfrj 511, IOS-I I S . de Kruijff. B.. van Zoelen. E. J . .I.,and van Deenen, L. L. M. (1978). Glycophorin facilitates the transbilayer movement o f phosphatidylcholine in vesicles. Biochim. Biophgs. Acttr 509. 537-S4?.

De Kuyper, M.. Joniau. M.. and Dangreau, H. ( 1983). Intervesicular phospholipid transfer. A free-flow electrophoresis study. Biw/ieiiii,sfry 22, 415420. Demel, R . A , . and de Kruyff. B . (1976). The function of sterols in membranes. Biochim. H i o p h . ~ .Ac'fci ~ . 457, 109-132.

196

ELIEZAR A. DAWIDOWICZ

Demel, R. A., Wirtz, K. W. A., Kamp, H. H., Geurts van Kessel, W. S. M.,and van Deenen, L. L. M. (1973). Phosphatidylcholine exchange protein from beef liver. Nature (New Biol.) 246, 102-105. Demel, R. A,, Jansen, J . W. C. M., van Dijck, P. W. M., and van Deenen, L. L. M. (1977). The preferential interaction of cholesterol with different classes of phospholipids. Biochim. Biophys. Actu 465, 1-10. Deuticke, B., and Hruska, C. (1976). Changes of non-electrolyte permeability in cholesterol loaded erythrocytes. Biochim. Biophys. Acta 433, 638-653. DiCorleto, P. E., and Zilversmit, D. B. (1979). Exchangeability and rate of flip-flop of phosphatidylcholine in large unilamellar vesicles, cholate dialysis vesicles and cytochrome oxidase vesicles. Biochim. Biophys. Acra 552, 114-1 19. Doody, M. C., Pownall, H. J., Kao, Y.J., and Smith, L. C. (1980). Mechanism and kinetics of transfer of a fluorescent fatty acid between single-walled phosphatidylcholine vesicles. Biochemistry 19, 108-1 16. Douady, D., Grosbois, M., Guerbette, F., and Kader, J. C. (1982). Purification of a basic phospholipid transfer protein from maize seedlings. Biochim. Biophys. Actri 710, 143153.

Duckwitz-Peterlein, G., and Moraal, H. (1978). Transport of lipids through water as exchange mechanism between two liposome populations. Biophys. Struct. Mech. 4, 3 15-326. Duckwitz-Peterlein. G., Ellenberger, G., and Overath, P. (1977). Phospholipid exchange between bilayer membranes. Biochim. Biophys. Acta 469: 311-325. Dyatlovitskaya, E. V., Lemenovskaya, A. F., and Bergelson, L. D. (1979). Use of proteinmediated lipid exchange in the study of membrane-bound enzymes. The lipid dependence of glucose-6-phosphatase. Eur. J. Biochem. 99, 605-612. Eckles, N. E.,Taylor, C. B., Campbell, D. J., and Gould, R. J. (1955). The origin of plasma cholesterol and the rates of equilibration of liver, plasma and erythrocyte cholesterol. J. Lab. Clin. Med. 46, 359-371. Eisenberg, S. (1978). Effect of temperature and plasma on the exchange of apolipoproteins and phospholipids between rat plasma very low and high density lipoproteins, J. Lipid Res. 19, 229-236. Engelman, D. M., and Rothman, J. E. (1972). The planar organization of lecithin-cholesterol bilayers. J. Biol. Chem. 247, 3694-3697. Etemadi, A.-H. (1980). Membrane asymmetry. A survey and critical appraisal of the methodology, 11. Methods for assessing the unequal distribution of lipids. Biochim. Biophys. Acta 604,423475.

Franck, P. F., DeRee, J. M.,Roelofsen, B., and Op den Kamp, J. A. F. (1984). Modification of the erythrocyte membrane by a non-specific lipid transfer protein. Bioc-him. Biophys. AC/CJ778, 405-41 I . Frank, A., Barenholz, Y., and Thompson, T. E. (1983). Spontaneous transfer of sphingomyelin between phospholipid bilayers. Biochemistry 22, 5647-565 I . Fugler, L., Clejan, S., and Bittman, R. (1985). Movement of cholesterol between vesicles prepared with different phospholipids and sizes. J. B i d . Chem. 260, 4098-4102. Ganong, B. R., and Bell, R. M. (1984). Transmembrane movement of phosphatidylglycerol and diacylglycerol sulfhydryl analogues. Biochemistry 23,4977-4983. Gerritsen, W. J., Henricks, P. A. J., de Kruijff, B., and van Deenen, L. L. M. (1980). The transbilayer movement of phosphatidylcholine in vesicles reconstituted with intrinsic proteins from the human erythrocyte membrane. Biochim. Biophys. Acra 600,607-619. Giraud, F., and Claret, M. (1979). A study of cholesterol transfer between erythrocytes and lipid vesicles. Possible involvement of interparticular collisions. FEBS Lett. 103, 186191.

Goldstein, J. L., Anderson, R. G. W.. and Brown, M. S. (1979). Coated pits. coated vesicles, and receptor-mediated endocytosis. Nature (London) 279, 679-685.

LIPID EXCHANGE

197

Gottlieb. M. H. (1980). Rates of cholesterol exchange between human erythrocytes and plasma lipoproteins. Biochitn. Biophys. A m i 600, 530-541. Graham. J. M., and Green, C. (1970). The properties of mitochondria enriched in ~iiirowith cholesterol. Eur. J . Biochc~ttr.12, 58-66. Green, C. (1977). Sterols in cell membranes and model membrane systems. In “International Review of Biochemistry. Biochemistry of Lipids 11’’ (T. W. Goodwin, ed.). Vol. 14, pp. 101-152. University Park Press, Baltimore, Maryland. Green. J. R., Edwards. P. A.. and Green, C. (1973). Optical rotatory dispersion studies of compounds related to cholesterol in liposomes and the membranes o f erythrocyte “ghosts.” BifJCh<’m.J . 135, 63-71. Gurd, F. R. N. (IW).Some naturally occurring lipoprotein systems. I n “Lipide Chemistry” (D. J. Hanahan. ed.), Chapt. 9. Wiley. New York. Haberland. M. E., and Reynolds, J. A. (1973). Self-association of cholesterol in aqueous solution. Proc. N d . A c d . Sci. U.S.A. 70, 2313-2316. Hagerman. J. S . . and Gould, R. G. (1951). The in vitro interchange of cholesterol between plasma and red cells. Proc. Soc. Exp. BiiJ/.Me(/. 78, 329-332. Haran, N., and Shporer, M. (1977). Proton magnetic resonance study of cholesterol transfer between egg yolk lecithin vesicles. Biochim. Biopliys. A m 465, 11-18. Helmkamp. G. M., Jr.. Wirtz. K. W. A,, and van Deenen, L. L. M. (1976). Phosphatidylinositol exchange protein. Effects of membrane structure on activity and evidence for a ping-pong mechanism. Arch. Biochetn. H i o ~ p h , ~174, . ~ . 592-602. Hoover, R. L., Dawidowicz, E. A.. Robinson. J. M.. and Karnovsky, M. J. (1983). Role of cholesterol in the capping of surface immunoglobulin receptors on murine lymphocytes. J . Cell Bi0l. 97, 73-80. Huang. C., Charlton, J. P., Shyr, C. I . , and Thompson, T. E. (1970). Studies on phosphatidylcholine vesicles with thiocholesterol and a thiocholesterol-linked spin label incorporated in the vesicle wall. Bioc.hetni.s/ry 9, 3422-3426. Johnson. L. W.. and Zilversmit, D. B. (1975). Catalytic properties of phospholipid exchange protein from bovine heart. Biochirn. Biopliy.~.Actcr 375, 165-175. Johnson. L. W.. Hughes. M. E.. and Zilversmit. D. B. (1975). Use of phospholipid exchange protein to measure inside-outside transpositions in phosphatidylcholine liposomes. Biochim. Biophys. Actci 375, 176-185. Jonas, A.. and Maine. G. T. (1979). Kinetics and mechanism of phosphatidylcholine and cholesterol exchange between single bilayer vesicles and bovine high-density lipoprotein. Bioc’het?tis/n118, 1722-1728. Kagawa. Y.,Johnson, L. W., and Racker. E. (1973). Activation of phosphorylating vesicles by net transfer of phosphatidylcholine by phospholipid transfer protein. Bioc~lrem.Bioplivs. Res. Conitnun. 50, 245-25 I . Kahane, I . , and Razin, S. (1977). Cholesterol-phosphatidylcholine dispersions as donors of cholesterol to M.vc.o~p/ostm~ membranes. Biochitn. Biophys. Acru 471, 32-38. Kasper. A. M., and Helmkamp, G. M.. Jr. (19x1). lntermembrane phospholipid fluxes catalyzed by bovine brain phospholipid exchange protein. Biochim. Bioplrys. Acw 664, 22-32. Kirby, C . J., and Green, C. (1977). Transmembrane migration (“flip-flop”) of cholesterol in erythrocyte membranes. Bioc~lwtn.J . 168, 57S.577. Kornberg. R. D.. and McConnell. H. M. (1971). Inside-outside transitions of phospholipids in vesicle membranes, Biocliemi.~/n,10, I 1 11-1 120. Krmer. R.. Hasselbach. H-J.. and Semenza, G. (1981 ). Rapid transmembrane movement of phosphatidylcholine in small unilamellar lipid vesicles formed by detergent removal. Bioc.liitn. Biophys. Acrn 643, 233-242. Kremer. J. M . H.. Kops-Werkhoven, M. M.. Pathmamanoharan. C., Gijzema. 0. L. J., and Wiersema. P. H. (1977). Phase diagrams and kinetics of phospholipid exchange for

198

ELIEZAR A. DAWIDOWICZ

vesicles of different composition and radius. Biochim. Biophys. A d a 471, 177-188. Kreuter, J., Higuchi. W. I.. Ganesan. M. G., and Weiner. N. D. (1981). Delivery of liposome-associated sterols through silastic membranes. Biochirn. Biophys. A i m 676, I 18121.

Lange, Y., and D’Alessandro, J. S. (1977). Characterization of mechanisms for transfer of cholesterol between human erythrocytes and plasma. Biochernisfry 16, 43394343. Lange, Y.,Cohen, C. M., and Poznansky, M . J. (1977). Transrnembrane movement of cholesterol in human erythrocytes. Proc. Nuri. Acad. Sci. U.S.A. 74, 1538-1542. Lange. Y., Dolde, J., and Steck, T. L. (1981). The rate of transmembrane movement of cholesterol in the human erythrocyte. J. B i d . Chem. 256, 5321-5323. Lange, Y., Molinaro, A. L., Chauncey, T. R., and Steck, T. L. (1983). On the mechanism of transfer of cholesterol between human erythrocytes and plasma. J . B i d . Chem. 258, 6920-6926.

Langley, K. E., and Kennedy, E.P. (1979). Energetics of rapid transmembrane movement and of compositional asymmetry of phosphatidylethanolamine in membranes of Bricilli4s megatrrirtm. Proc. Null. Acad. Sri. U.S.A. 76,6245-6249. Lenard, J., and Rothrmn, J. E. (1976). Transbilayer distribution and movement of cholesterol and phospholipid in the membrane of influenza virus. Pmr. Nut/. Acad. Sci. U.S.A. 73, 39 1-395.

Leto, T. L., Roseman. M. A., and Holloway, P. W. (1980). Mechanism of cytochrome 6 , exchange between phosphatidylcholine vesicles. Biochemistry 19, 191 1-1916. Lippiello, P.M., and Waite. M. (1983). Kinetics and mechanism of phosphatidylcholine and cholesterol exchange between chylomicrons and high-density lipoproteins. Biochem. J. 215, 279-286. Lund-Katz, S.. Hammerschlag, B., and Phillips, M. C. (1982). Kinetics and mechanism of free cholesterol exchange between human serum high- and low-density lipoprotein. Biochemis/r.y 21, 2964-2969. McLean, L. R., and Phillips, M. C. (1981). Mechanism of cholesterol and phosphatidylcholine exchange or transfer between unilarnellar vesicles. Biuchumistry 20, 2893-2900. McLean, L. R.,and Phillips, M. C. (1982). Cholesterol desorption from clusters of phosphatidylcholine and cholesterol in unilamellar vesicle bilayers during lipid transfer. Biochemistry 21,4053459.

McLean, L. R., and Phillips, M. C. (1984). Kinetics of phosphatidylcholine and lysophosphatidylcholine exchange between unilamellar vesicles. Biochemis/ry 23, 4624-4630. McMurray, W. C., and Dawson, R. M . C. (1969). Phospholipid exchange reactions within the liver cell. Biochcm. J . 112, 91-108. Madden, T. D., Vigo. C., Bruckdorfer, K. R., and Chapman, D. (1980).The incorporation of cholesterol into inner mitochondria1 membranes and its effect on lipid phase transition. Blochim. Biophys. Actu 599, 528-537. Martin. F. J., and MacDonald, R. C. (1976). Phospholipid exchange between bilayer membrane vesicles. Biochtvnis/ry 15, 321-327. Massey, J. B., Gotto, A. M., Jr., and Pownall, H. J . (1982a). Kinetics and mechanism of the spontaneous transfer of fluorescent phosphatidylcholines between apolipoproteinphospholipid recombinants. Biochemis/ry 21, 3630-3636. Massey, J . B.,Gotto, A . M., Jr., and Pownall, H. J. (1982b). Kinetics and mechanism of the spontaneous transfer of fluorescent phospholipids between apolipoprotein-phospholipid recombinants. Effect of the polar head group. J . B i d . Chum. 257,5444-544s. Massey, J. B., Hickson. D., She, H. S.,Sparrow, J. T., Nia, D. P., Gotto, A. M., Jr., and Pownall, H. J. (1984). Measurement and prediction of the rates of spontaneous transfer of phospholipids between plasma lipoproteins. Biochirn. Biophys. Ac/n 794, 274-280.

LIPID EXCHANGE

199

Moore, N . F., Patzer, E. J.. Shaw. J . M.. Thompson, T . E., and Wagner, R. K . (1978). Interaction of vesicular stomatitis virus with lipid vesicles: depletion o f cholestei-ol and effect on virion membrane fluidity and infectivity. J . Virol. 27. 320-329. Mone, I).J.. Kartenbeck, J.. and Franke. W. W. (1979). Membrane flow and interconversions among endomembranes. Biocliirri. Biophys. Acui 559, 7 1-15?, Murphy, J. R. ( 1962). Erythrocyte metabolism. IV. Equilibration of cholesterol-4-C" between erythrocytes and variously treated sera. J. Lab. ('h.Med. 60, 571-578. Nakagawa, Y Inoue, K., and Noiima, S. ( 1979). Transfer of cholesterol between liposonial membranes. Biocliim. Bioplrvs. ACIU 553, 307-3 19. Nichols, J. W.. and Pagano, R. E. (1981). Kinetics of soluble lipid niononier diffusion between ~ 2783-2789. vesicles. B i c x ~ l i e i n i s i20, Noordam. P. C., van Echteld, C. J. A., d e KruijtT, H.. and d e Gier. J. (1981). Rapid transbilayer movement of phosphatidylcholine in unsaturated phosphatidylethanol~imine containing model membranes. Bicwhirn. Biophys. Aciir 646,483-487. North. P., and Fleischer. S. (1983). Use of a nonspecific lipid transfer protein to modify the cholesterol content of synaptic membranes. Mailrods Dr:ymo/. 98, 5 9 9 4 1 3 . Pagano. R. E. (1983). lntracellular processing of lipids: A theory based on studies with tluorescent lipids, liposomes and cells. /ti "Liposome Letters" (A. D. Hangham. ed.), pp. 83-95. Academic Press, London. Pal. R.. Harenholz, Y.. and Wagner. R. R. (1981). Depletion and exchange of cholesterol virus by interaction with serum lipoproteins from the membrane of vesicular stomat or poly(vinylpyrro1idone) complexed with bovine serum albumin. Riocheriiisiry 20, 5%

_.

539.

Papahadjopoulos. D.. Hui, S., Vail. W. J.. and Posle, G. (1976). Studies on membrane fusion. 1 . Interactions of pure phospholipid membranes and the effect of myristic acid. lysolecithin. proteins and dimethylsulfoxide. Biodfirfr.Biopliys. Act0 448,245-764. Patzer, E. J . , Shaw. J . M.. Moore, N. F..l h o m p s o n . T. E.. and Wagner. R. K. (197x1. Transmembrane movement and distribution of cholesterol in the membrane of vesicular stomatitis virus. Bioc~/rerrri.stry17, 4192-4200. Phillips. M. C., McLean, L. R., Stoudt. 6 .W.. and Rothhlat, G . H. (19x0). Mechanism o f cholesterol efflux from cells. Arlrc~ro.sc/c~ro,~i,s 36, 409-422. Poorthuis. B. J. H. M.. and Wirtz. K. W. A. (19x2). Increased cholesterol esterification in rat liver microsomes by purified non-specific phospholipid transfer protein. B i o d i i m . B ~ C J ~ / Ac/cr I ) ~ S710, . 99-10s. Poorthuis. B. J . H. M.. and Wirtz, K . W. A . (1983). Nonspecific lipid transfer protein from rat liver. Me/hod.s Enzpmol. 98, 592-596. Pownall, H. J . . Hickson, D., Gotto. A. M., Jr.. and Massey. J. B. (1982). Kinetics ofspontaneous and plasma-stimulated sphingomyelin transfer. Biochinr. B i ~ ~ p h yAc.ro . ~ . 712, 169- 176. Poznansky, M. J.. and Czekanski. S. (1979). Cholesterol exchange 21sa function of cholesterol/ phospholipid mole ratios. Biochcvrz. J . 177. 989-99 I . Pomansky. M. J.. and Czekanski, S. (1982). Cholesterol movement between human skin fibroblasts and phosphatidylcholine vesicles. Bioc.ltirrr. Bioplrys. A c m 685, IX?-I'N). Poznansky, M. J.. and Lange. Y. (1976). Transbilayer movement of cholesterol in dipalmitoyllecithin-cholesterol vesicles. Ncirrirt, (Londori) 259, 420-42 I. Poznansky. M. J., and Lange, Y. (1978). Transhilayer movement of cholesterol in phospholipid vesicles under equilibrium and non-equilihrium conditions. Bioc4irn. Bio~plrys. Acrcl 506, 2 5 6 2 6 4 . Quarfordt. S. H.. and Hilderman. H.L. (1970). Quantitation of the in vim) free cholesterol exchange of human red cells and lipoproteins. J . Lipid Rcs. I I , 528-535.

200

ELIEZAR A. DAWlDOWlCZ

Radin, N . S., and Metz, R. J. (1982). Proteins that transfer sphingoglycolipids. Adv. Exp. B i d . Med. 152, 235-239. Radin, N . S., and Metz, R. J. (1983). Cerebroside transfer protein. Methods Enzymol. 98, 613-622. Renkonen, O., Gahmberg, C. G., Simons, K., and Kaarianen, L . (1972). The lipids of the plasma membranes and endoplasmic reticulum from cultured baby hamster kidney cells (BHK 21). Biochim. Biophys. Acta 255, 66-78. Renshaw, P. F., Janoff, A. S . , and Miller, K. W. (1983). On the nature of dilute aqueous cholesterol suspensions. J . Lipid Res. 24, 47-51, Roseman, M. A., and Thompson, T. E. (1980). Mechanism of spontaneous transfer of phospholipids between bilayers. Biochemistry 19, 439-444. Roseman, M. A., Litman, 8 . J., and Thompson, T. E. (1975). Transbilayer exchange of phosphatidylethanolamine for phosphatidylcholine and N-acetimidoylphosphatidylethanolamine in single-walled bilayer vesicles. Biochemistry 14, 4826-4830. Rothblat, G. H., and Phillips, M. C. (1982). Mechanism of cholesterol emux from cells. Effects of acceptor structure and concentration. J . Biol. Chem. 257, 47754782. Rothman, J. E., and Dawidowicz, E. A. (1975). Asymmetric exchange of vesicle phospholipids catalyzed by the phosphatidylcholine exchange protein: Measurement of inside-outside transitions. Biochemistry 14, 2809-28 16. Rothman, J. E., and Kennedy, E. P. (1977). Rapid transmembrane movement of newly synthesized phospholipids during membrane assembly. Proc. Natl. A c d . Sci. U . S . A . 74, 1821-1825. Rothman, J . E., Tsai, D. K., Dawidowicz, E. A., and Lenard, J. (1976). Transbilayer phospholipid asymmetry and its maintenance in the membrane of influenza virus. Biochemistry 15, 2361-2370. Rottem, S . , Shinar, D., and Bittman, R. (1981). Symmetrical distribution and rapid transbilayer movement of cholesterol in Mycoplasma gallisepticrrm. Biochirn. Biophys. Acta 649, 572-580. Sabatini, D. D., Kreibich, G., Morimoto, T., and Adesnik, M. (1982). Mechanisms for the incorporation of proteins in membranes and organelles. J . Cell B i d . 92, 1-22. Sackmannn, E., Trauble, H . , Galla, H.-J., and Overath, P. (1973). Lateral diffusion, protein mobility, and phase transitions in Eschericia coli membranes. A spin label study. Biochemistry 26, 5360-5369. Sandra, A , , and Pagano. R. E. (1978). Phospholipid asymmetry in LM cell plasma membrane derivatives: polar head group and acyl chain distributions. Biochemistry 17, 332338. Scandella, C. J., Devaux. P . , and McConnell, H. M. (1972). Rapid lateral diffusion of phospholipids in rabbit sarcoplasmic reticulum. Proc. Natl. Acad. Sci. U . S . A . 69, 20562060. Sefton, B. M., and Gaffney, B. J. (1979). Complete exchange of viral cholesterol. Biochemistry 18, 436442. Seigneuret, M . , and Devaux, P. F. (1984). ATP-dependent asymmetric distribution of spinlabeled phospholipids in the erythrocyte membrane: Relation to shape changes. Proc. Nut/. Acud. Sci. U . S . A . 81, 3751-3755. Sengupta, P., Sackman, E . , Kuhnle, W., and Scholz, H. P. (1976). An optical study of the exchange kinetics of membrane bound molecules. Biochim. Biophys. Actrr 436, 869878. Shaw, J. M., and Thompson, T. E. (1982). Effect of phospholipid oxidation products on transbilayer movement of phospholipids in single lamellar vesicles. Biochemistry 21, 920-927.

LIPID EXCHANGE

201

Shaw. J . M.. Moore. N . F., Patzer. E. J.. Correa-Freire, M. C., Wagner. R. R.. and Thompson, T. E. (1979). Compositional asymmetry and transmembrane movement of phosphatidylcholine in vesicular stomatitis virus. Bioc~hrtrristty 18, 538-543. Sinensky, M. (1981). A comparison of solution properties of cholesterol and 25-hydroxycholesterol. Arch. Biochem. Biophvs. 209, 321-324. Sleight, R. G . , and Pagano, R. E. ( 1984). Transport of a fluorescent phosphatidylcholine analog from the plasma membrane t o the Golgi apparatus. J . Cell Biol. 99, 742-751. Sleight. R. G.. and Pagano, R. E. (1985). Transbilayer movement of a fluorescent phosphatidylethanolamine analogue across the plasma membrane of cultured mammalian cells. J . B i d . Chivn. 260, 1146-1 154. Slotte. J . P., and Lunberg. B. (1983). Transfer of [’H]cholesterol between lipid vesicles and rat arterial smooth muscle cells in r?itro. Bioc,hirn. Biophys. Actu 750, 434439. Slutzky, G . M.. Razin, S., Kahane, I., and Eisenberg. S. (1977). Cholesterol transfer from serum lipoproteins to Mycoplastnu membranes Biochemistq~16, 5158-5 163. Somerhaju, P., Brockerhoff, H., and Wirtz, K. W. A. (1981). A new fluorimetric method to measure protein-catalyzed phospholipid transfer using I -acyl-2-parinaroylphosphatidylcholine. Biochim. Biophys. Acta 649, 521-528. Somerhaju. P., van Paridon, P., and Wirtz, K . W. A. (1983). Phosphatidylinositol transfer protein from bovine brain. Substrate specificity and membrane binding properties. Biochitrr. Biophys. Acta 731, 186-195.

Struck. D. K., and Pagano, R. E. (1980). Insertion of fluorescent phospholipids into the plasma membrane of a mammalian cell. J. B i d . C‘hem. 255, 5404-5410. Teerlink. T., van d e r Krift. T. P., Post, M.. and Wirtz, K. W. A. (1982). Tissue distribution and subcellular localization of phosphatidylcholine transfer protein in rats as determined by radioimmunoassay. Biochim. Biophvs. A(.tci 713, 61-67. Teerlink. T.. Poorthuis, B. J. H. M., and Wirtz, K. W. A. (1983). Phosphatidylcholine transfer protein from rat liver: Purification and radioimmunoassay. Methods Enzymol. 98, 586592. Thilo, L. (1977). Kinetics of phospholipid exchange between bilayer membranes. Biochim. Biophvs. Actu 469, 326334. Thompson, T. E. (1978). Transmembrane compositional asymmetry of lipids in bilayers and biomembranes. In “Molecular Specialization and Symmetry in Membrane Function” (A. K. Solomon and M. L. Karnovsky. eds.). pp. 78-98. Harvard Univ. Press, Cambridge, Massachusetts. Ulsamer, A. G., Wright. P. L., Wetzel. M. G., and Korn, E. D. (1971). Plasma and phagosome membranes of Acunthomoeha custellanii. J. Cell Biol. 51, 193-215. van Deenen. L. L. M. (1981). Topology and dynamics of phospholipids in membranes. FEBS Lett. 123, 3-15.

van den Besselaar, A. M. H. P., de Kruijff, B.. van den Bosch. H., and van Deenen. L. L. M. ( 1978). Phosphatidylcholine mobility in liver rnicrosomal membranes. Biochim. Biophys. AC‘ILI510, 242-255. Vandenheuvel, F. A. ( 1966). Lipid-protein interactions and cohesional forces in the lipoprotein systems of membranes. J . A m . Oil Chc,m. SOC. 43, 258-264. van Heusden. G. P. H., and Wirtz, K. W. A. (1984). Hydroxymethylglutaryl CoA reductase and the modulation of microsomal cholesterol content by the nonspecific lipid transfer protein. J . Lipid Res. 25, 27-32. van Meer. G., Poorthuis, B. J . H. M., Wirtz, K. W. A , , Op den Kamp. J . A . F., and van Deenen, L. L. M. ( 1980). Transbilayer distribution and mobility of phosphatidylcholine in intact erythrocyte membranes. A study with phosphatidylcholine exchange protein. Eur. J . B i o c h ~ . ~103, ~ . 283-288.

202

ELIEZAR A. DAWIDOWICZ

Voelker. D. R . , and Kennedy, E. P. (1982). Cellular and enzymatic synthesis of sphingomyelin. Eiocherrrisiry 21, 2753-2759. Wattenberg, B. W., and Silbert, D. F. (1983). Sterol partitioning among intracellular membranes. Testing a model for cellular distribution. J. Biol. Chem. 258, 2284-2289. Web, R. and Helmkamp. G. M., Jr. (1984). Acyl chain specificity of phosphatidylcholine transfer protein from bovine liver. J . Biol. Chem. 259, 6937-6941. Westerman. J., Kamp. H. H., and Wirtz, K. W. A. (1983). Phosphatidylcholine transfer protein from bovine liver. Methods Enzymol. 98, 581-586. Whetton, A. D., and Houslay, M . D. (1983). 5’-Nucleotidase is activated upon cholesterol depletion of liver plasma membranes. FEBS Lett. 157, 70-74. Wickner. W. (1980). Assembly of proteins into membranes. Science 210, X61-868. Wilgram, G. F., and Kennedy, E. P. (1%3). lntracellular distribution of some enzymes catalyzing reactions in the biosynthesis of complex lipids. J . Biol. Chem. 238, 26152619.

Wilson, D. B., Ellsworth, J. L., and Jackson. R. L. (1980). Net transfer of phosphatidylcholine from plasma low density lipo-proteins to sphingomyelin-apolipoproteinA-11 complexes by bovine liver and human plasma phospholipid exchange proteins. Biochitn. Biophvs. Acru 620, 550-561. Wirtz. K. W. A. (1974). Transfer of phospholipids between membranes. Biochim. Biophys. Acru 344, 95-1 17. Wirtz, K. W. A., and Zilversmit, D. 3.(1968). Exchange of phospholipids between liver mitochondria and microsomes in iitro. J. B i d . Chem. 243, 3596-3602. Wirtz, K. W. A., Devaux. P. F., and Bienvenue, A. (1980). Phosphatidylcholine exchange protein catalyzes the net transfer of phosphatidylcholine to model membranes. Eiochemistry 19, 3395-3399. Yaffe, M. P., and Kennedy, E. P. (1983). lntracellular phospholipid movement and the role of phospholipid transfer proteins in animal cells. Biochemistry 22, 1497-1 507. Yoshimura, T., and Helmkamp, G. M., Jr. (1984). Bovine brain phosphatidylinositol transfer protein. Effects of pH, ionic strength and lipid composition on transfer activity. Biochim. Eiopliys. Act0 793, 463-470. Zilversmit, D. B. (1978). Phospholipid-exchange proteins as membrane probes. Ann. N. Y . Acud. Sci. 308, 149-160. Zilversmit, D. B. (1983). Lipid transfer proteins: Overview and application. Methods Enzymol. 98, 565-573. Zilversmit, D. B., and Hughes, M . E. (1977). Extensive exchange of rat liver microsomal phospholipids. Eiochim. Biophys. Acru 469, 99-1 10.

CURRENT TOPICS IN MEMBRANES AND TRANSPORT. VOLUME 29

Membrane Fusion ROBERT BLUMENTHAL

1. INTRODUCTION

In eukaryotic cells there are many types of membranes which differ from one another in their structure, composition, and function. Yet the biosynthetic machinery to produce membrane constituents (lipids and proteins) is centralized. Cells must therefore have evolved an efficient system of traffic control for membrane proteins as well as membrane lipids. One important element in this traffic control is vesicular transport. Palade (1975) has outlined such a pathway for the secretory process in pancreatic exocrine cells from the site of protein synthesis to its discharge in the extracellular fluid. Vesicular membrane traffic is not only important in biosynthetic delivery of membrane constituents to their appropriate locations, but also serves a number of physiological functions. They include chemical signaling between cells by exocytosis of neurotransmitters and hormones and transport of macromolecules into and across cells by endocytosis. In all these processes, fusion between membranes is the key. Fusion must occur continuously to support the constant flow and turnover of membranes. On the other hand the process must be tightly controllcd both spatially and temporally. Fusion must occur at desired locations, and not occur at undesired locations, to allow membranes to maintain their specific structure, function, and composition. Although a host of biological processes (c.g., endocytosis, exocytosis, viral infection, parasite invasion, cell division, fertilization, polykaryon formation in bone and muscle) involve membrane fusion, 1 shall limit my discussion of biological membrane fusion mainly to two well-studied examples: exocytosis and viral membrane fusion. I focus on exocytosis, 203

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since a great deal is known in that system about triggers and controls, and on viral fusion, since the membrane components involved are well defined. Fusion is a complex phenomenon involving a whole range of biochemical and biophysical interactions. In an attempt to understand the fundamental mechanism, the system has been reduced to simpler elements. The systems best understood are, for the most part, very simple when compared to the biological systems which we wish to explain. Examples of such model systems are Ca” -induced fusion of negatively charged phospholipid vesicles and polyethylene glycol-induced fusion of membranes. These are often uncontrolled processes, with considerable membrane damage concomitant with fusion. However, they have been very useful for defining the barriers between apposing membranes and for determining essential conditions for membrane fusion.

TABLE I BARRIERS TO FUSION Barrier to overcome in order to fuse Triggering by ions or “second messengers” Movement of membranes into apposition Recognition of site at which fusion is to occur Steric constraints

Packing of the membrane

Electrostatic repulsion Dehydration repulsion

Resistance to deformation Transition from the pentalaminar structure to complete fusion of membranes Reestablishment of membrane composition after fusion

Possible mechanisms to overcome the barrier Activation of Ca2’ channels or proton pumps, increase of cellular cyclic AMP, increase of phosphatidylinositol turnover Directed movement of intracellular membranes by cytoskeletal elements, random (Brownian) motion Specific receptors (glycoproteins or glycolipids), cytosolic cross-bridging proteins (e.g. synexin) Removal of sialic acid by viral neuraminidase, lateral diffusion of glycoproteins and glycolipids, disaggregation or dissociation of cytoskeletal elements Alteration of the lipid phase or of the local cholesterol content, selective metabolism of lipids, “fluidizing” effects (temperature, ions, pH, proteins) Shielding or binding by ions. van der Waals attractions Calcium bridges, water removal by freeze-thawing or chemical dehydrants, hydrophobic attraction through “defects,” specific “junctional” contacts Osmotic forces, wedge-shaped molecules Phospholipid flip-flop, formation of intermediate structures (defects, inverted micelles)

Catabolism and synthesis of lipids and proteins, lateral diffusion, flip-flop, exchange

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In order to obtain a complete picture of membrane fusion we must consider all the factors which control fusion. Nature has set up many barriers to prevent membranes from fusing, otherwise we all would be one big syncytium. Membranes destined to fuse must overcome those barriers. This will be the theme of this article. 1 have summarized in Table I the events leading up to actual membrane fusion in terms of those barriers. The order in which those events are listed in Table 1 is not necessarily the chronological order of steps resulting in fusion. For instance, movement into apposition or recognition could precede triggering by ions or second messengers. I shall describe in detail the characteristics of these barriers, and discuss mechanisms how the barriers are overcome, based on what is known about membrane structure and interactions. Since this article is written in a conceptual vein, I shall not present a comprehensive summary of data in the literature on membrane fusion. For this 1 refer readers to a number of reviews which have come out in the past years. For instance the volume on membrane fusion edited by Poste and Nicholson ( 1978) is still an excellent compendium of facts and hypotheses about membrane fusion. A collection of more recent reviews on exocytosis and secretion can be found in Hand and Oliver (1981), and viral fusion has recently been reviewed by White et a / . (1983). A series of reviews on cell fusion can be found in a recent Ciba Foundation Symposium volume (Evered and Whelan, 1984). What I hope to achieve here is delineation of what is fact, hypothesis, or theory in membrane fusion, and use this as an appropriate basis for further study. 11.

HOW WE OBSERVE MEMBRANE FUSION

A. Definitions

Membrane fusion involves the merging of the membranes of two different organelles. In exocytosis, the plasma membrane merges with the membrane of a secretory vesicle; in viral fusion, the plasma membrane merges with the viral membrane. Merging of membranes proceeds concomitant with mixing of aqueous compartments surrounding the membranes. In exocytosis the granular material in the secretory vesicle is mixed with the extracellular space; in viral fusion the viral core material (nucleocapsid) is mixed with the cell cytoplasm.' The two fusing membranes 'Most enveloped animal viruses enter into cells by endocytosis. followed by fusion of the viral membrane with the endosome membrane at a lowered pH (see Section V ) . However, some viruses can fuse directly at the plasma membrane. A well-known example ofthis class is the Sendai virus. Since the morphology of fusion with the plasma membrane is better defined. I shall only discuss that aspect of virus-cell fusion in this section.

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are in fact bounded by three aqueous compartments, two of whose contents mix. If there is also spillover t o the third compartment, the fusion is “leaky.” Fusion of viruses with the plasma membrane seems to be accompanied by immediate and profound changes in the passive permeability of the target membrane (Pasternak and Micklem, 1974; Kohn, 1979). However those permeability changes are not a necessary concomitant of fusion, but seem to be an artifact arising from the circumstance that the membranes of the virions are damaged to start with due to aging, freezethawing, or sonication (Homma et al., 1976; Vaananen and Kaariainen, 1979; Young et d., 1983).* Exocytosis is not expected to be leaky, because products stored in the secretory granule must be protected from destruction by cytoplasmic components and, vice versa, proteolytic enzymes destined for secretion should not spill over into the cytoplasm. Moreover, in exocytotic processes such as synaptic transmission the whole package has to be released for efficient transmission. B. Biological Membranes

Fusion of biological membranes is mainly assessed by electron microscopic techniques, using thin-sectioning and freeze-fracture approaches (Orci and Perrelet, 1978; Chandler and Heuser, 1979; Ornberg and Reese, 1981; Knutton, 1979a,b,c). The cartoons in Fig. 1 are representations of stages in fusion as seen by thin sections of conventionally prepared material (i.e., aldehyde and osmium fixation followed by lead and uranyl staining). The two partners involved in fusion are distinguished by their original morphology: the secretory granule is round, the virion is round or bullet shaped, and the plasma membrane is flat. The membranes of both partners show the typical trilaminar unit structure (Robertson, 1959). The first detectable step in fusion is the merging of the two leaflets in contact (e.g.. outer leaflet of secretory granule with inner leaflet of plasma membrane), resulting in a so-called pentalaminar structure (Palade, 1975).3 The thin ‘An alternative interpretalion of leaky viral fusion is that some portion of the viral spike glycoprotein destabilizes the large1 membrane. This is consistent with the recent observation (Schlegel and Wade, 1984). that a synthetic peptide corresponding to the NH2-terminus of the spike glycoprotein of vesicular stomatis virus causes pH-dependent lysis of red blood cells. ‘In an early study by Palade and Bruns (1%8) parts of the pentalaminar sandwich seemed lo merge into two dark lines, which presumably implied one single bilayer membrane. Such a “bilayer diaphragm” has also been seen in a freeze-fracture study of exocytosis in zoospores (Pinto da Silva and Nogueira, 1977). However, according to Chandler and Heuser (1979). such ”intermediates” seem to be an artifact, resulting from exposure to high concentrations of glycerol.

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A

B

I%;. I . Schematic drawing of the morphology o f ( A ) exocytosis and (B)viral fusion. The drawing i s an interpretation of images seen in electron microscopy (see text. Section 1 1 ) . Four stages going from left to right are shown as follows: ( I)the membranes o f a secretory granule or virion are separate from the plosma memhrane: (2) triggering. movement. and recognition (see Table 1) brings the two into close apposition (prefirsion state), perhaps forming a pentalaminar complex; (3) fusion results in merging of membranes and contiguity of aqueous compartments; and ( 4 ) in the final .itage the membrane Components and aqueous components are completely mixed. The membranes of the granule and virion are stippled and the plasma membrane is cross-hatched to indicate separate membrane componcnls (lipids. proteins). After fusion they are intermixed. The granule core material (checkered) i s scen extruding into the medium during fusion. Cytuskeletal components are not shown. The nucleocapsid of the virion (drawn as two wavy lines) i s seen in the cytoplasm after fusion. The spikes on the virion represent viral spike glycoproteins. and the fuzzy "broccoli" on the plasma membrane represents cell surface carbohydrates (glycoproleins, glycolipids).

sections do not allow detection of structural changes in the membranes involved in fusion. However, use of electron-dense probes for carbohydrate residues or fenitin-labeled antibodies to specific surface components (e.g., viral membrane proteins, see Loyter and Volsky. 1982) reveal redistributions of membrane components Concomitant with fusion. For instance it has been shown that lectin binding sites are absent from the plasma membrane involved in a pentalaminar association with the membrane of a secretory granule of mast cells (Lawson et a / . , 1977). Thin sections have also revealed electron-dense bridges, which are interpreted as microfilamentous elements of the cell Webb, between two interacting membranes (Ornberg and Reese, 198 I ).

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Freeze-fracture reveals more elaborate structural organization of membranes. Since the plane of fracture passes through the middle of the phospholipid bilayer, the inside of both cytoplasmic and extracytoplasmic leaflets of the membrane are exposed (Pinta da Silva and Branton, 1970). In freeze-fracture cellular membranes appear as surfaces consisting of a smooth domain in which a variable number of small globules (7-12 nm in diameter) are interspersed. Based on a combination of morphological, biochemical, and biophysical approaches, it has been shown that the smooth background represents the lipidic component of the membrane and the 7- to 12-nm globules correspond to membrane protein^.^ The difference in size and distribution of intercalated membrane particles between the partners in fusion (plasma membrane and secretory granule membrane, or plasma membrane and viral membrane) allows us to follow the process before, during, and after fusion. In aldehyde-fixed, glycerol-impregnated mammalian cells, exocytosis is accompanied by the formation of particlefree patches at the fusion sites. Similar areas have been seen in viralplasma membrane fusion (Knutton, 1979~).Thus the process of fusion has been interpreted as involving “bare lipid patches” (Lucy, 1978). However, in protozoan exocytosis, the membranes which are about to fuse appear instead to be decorated by a specific particle arrangement, the “rosette” (Satir, 1974). Moreover, the existence of particle-free patches in aldehyde-fixed, glycerol-impregnated mammalian cells has been ascribed to an artifact, since they were not found in unfixed, unimpregnated cells freeze-fracturedby the rapid freezing method (Chandler and Heuser, 1979). Although electron microscopy using negative stain does not reveal any structural details of membranes, it has been found useful in some fusion studies. The radial projections on the surface of the membranes of enveloped animal viruses representing spike glycoproteins are easily visualized (Poste and Pasternak, 1978; Loyter and Volsky, 1982; Eidelman et ul., 1984). Those spike glycoproteins are then seen to spread over the target cell surface after fusion. Electron microscopy using negative stain is also used to monitor changes in size and morphology of vesicles after fusion (Kumar et ul., 1982; Blumenthal et al., 1983). Mixing of contents can also be observed in electron microscopy. The dense granular material in secretory vesicles is seen exuding from the granule into the medium, and the viral nucleocapsid is seen in the cyto41t has been argued that the globules might also represent “lipid particles” or inverted micelles (Verkleij and Ververgaert, 1978). A study by Hui er a / . (1983) and an analysis by Siegel (1984) indicate that the existence of such inverted micelles within a single bilayer is quite implausible. However, such structures could very well exist as interbilayer intermediates (Siegel, 1984). The possible role played by such intermediates in membrane fusion is further discussed in Section IX.

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plasm. Physiological assays of viral fusion include measurement of polykaryon formation (Toister and Loyter, 1973) and of hemolysis. The underlying assumption for the latter measurement is that hemolysis is commensurate with fusion (see Section 1l.A). Biochemical and biophysical measurements of biological fusion events are based on the secretion of materials during exocytosis. The underlying assumption is that the membranes always fuse during such a secretory event. For the most part, the amount of secreted material is measured chemically. In the neuromuscular junction release of acetylcholine during synaptic transmission can be measured electrophysiologically. From measurement of endplate potentials and miniature endplate potentials it has been shown that acetylcholine release occurs in packages with a size consistent with the contents of a presynaptic vesicle (Katz, 1966). More recently a novel technique has been developed to measure the exocytotic event. Fusion of a secretory vesicle with the plasma membrane gives rise to an increase in surface area. Since membrane capacitance is proportional to area, this increase has been monitored in terms of discrete capacitance jump with a size consistent with the area of a secretory vesicle (Neher and Marty, 1982). The permeability barrier imposed by the plasma membrane makes the secretory system inaccessible to the study of factors that control membrane fusion in exocytosis. Therefore, experimental procedures have been developed to bypass the plasma membrane permeability barrier without interfering with the ability of the remaining membrane to take part in exocytosis. I n adrenal medullary cells this has been achieved either by exposure to intense electric fields (Knight and Baker, 1982) or by treatment with saponin (Brooks and Treml, 1983). An in vitro secretory granuleplasma membrane preparation from sea urchin eggs has been developed (Vacquier, 1975) by rupturing and washing a monolayer of eggs attached to a planar surface. The secretory machinery in this system is intact, and exocytosis is observed by light microscopy. C. Artificial Lipid Membranes

Model systems provide a clear way to look at the influence of lipid state on fusion. Such model systems can be constructed in the form of liposomes or lipid vesicles. Unless I discuss a specific type of vesicle. I shall use the term liposome to include multilamellar vesicles as well as small unilamellar vesicles (SUVs) and large unilamellar vesicles (LUVs). The SUVs are usually formed by sonication to a size limit of about 20 nm in diameter. The LUV is defined arbitrarily as any single-walled vesicle larger than 100 nm in diameter. A variety of techniques have been developed to pre-

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pare and characterize liposomes. A comprehensive survey of “all you want to know about liposomes” can be found in Gregoriadis (1984). The chemical composition of liposomes can be vaned considerably from pure one-component systems to mixtures of various phospholipids differing in head group or hydrocarbon chain configuration. Thus questions relating to head group specificity, bilayer packing, phase transitions, and separations can be studied in detail. Other membrane components such as cholesterol, glycolipids, and various proteins or glycoproteins can be incorporated into the phospholipid bilayer, and their effects on fusion studied under controlled conditions. Membrane-active molecules and drugs that are known to alter fusion of natural membranes can be added externally to a well-defined system, and their influence on the mechanism of fusion assessed in detail. The results from these simple systems can then be correlated with the evidence on fusion with natural membranes. However, there are a number of objections leveled against artificial vesicles. First, in the early studies on vesicle-vesicle interaction it was not clear that fusion, rather than other forms of interaction such as lipid exchange was taking place. Second, some liposomes represent thermodynamically metastable forms following sonication or other procedures to form vesicles (Evans et a/., 1982). Third, the mechanism of lipid membrane fusion might be totally different from that of biological membrane fusion (see Section 1x1. Many studies had been carried out on the interaction of lipid vesicles with cells (Pagan0 and Weinstein, 1978; Margolis, 1984). The aim of these studies was to use lipid vesicles as vehicles to introduce biologically active substances into cells. Physiological effects of vesicle-encapsulated solutes on (nonendocytosing) cells have indicated transfer of the solute from vesicle to cell. These effects include inhibition of the light response in retinal rods by vesicle-encapsulated EDTA (Hagins and Yoshikami, 1978) and activation of histamine release from mast cells by Ca2+-containingvesicles (Theoharides and Douglas, 1978). Vesicle-encapsulated drugs such as cytosine arabinoside (Mayhew et d., 1976) and methotrexate (Koslowski et d., 1978) were found to have hharmacological effects in the cell cytoplasm in the absence of transport systems for drugs or endocytosis of liposomes. Vesicle-encapsulated RNA (Ostro et af., 1978), DNA (Mukherjee et a / . , 1978), and whole virus (Papahadjopouloset a/., 1980) were found to exhibit functional activity in the cell cytoplasm. In spite of the indications of transfer from vesicles to cell there was no clear demonstration of fusion nor quantitation of the interaction. Most of the quantitation in these studies was done by labeling the liposome with aqueous or lipid markers, and measuring the amount of cell-associated material. However, molecules from liposomes can become cell-associated by mechanisms other than

MEMBRANE FUSION

21 1

fusion, such as stable adsorption, endocytosis, and lipid exchange (Pagano and Weinstein, 1978). Weinstein ot al. (1977) developed an assay for fusion of lipid vesicles with cells in which transfer of contents can be visualized. This a based on self-quenching properties of the water-soluble dye carboxytluorescein and monitors transfer of contents of liposomes to cells. Based on measurements of the amounts of material transferred to cells the quantity of putative fusion was much less than originally estimated, and the process was saturable (Blumenthal et ( I / . , 1977). However, even using that assay the amount of fusion was overestimated. There was considerable cell-induced leakage of liposome contents (Szoka et t i / . , 1979). Moreover, fluorescence photobleaching showed that fluorescent lipid probes incorporated into lipid vesicles remained immobilized on the cell surface (Szoka r t u / . , 1980; Blumenthal et ml., 1982). This, taken together with studies on energy transfer between lipid probes incorporated into liposomes (Struck et d . , 19811, indicated that only a very small portion of the Iiposome-cell interaction was membrane fusion. Transfer of contents seems to take place by some other still undefined mechanism (Blumenthal et d . , 1982; Margolis, 1984). These considerations necessitate a cautious approach to the analysis of these (liposome) systems before identifying the process as fusion. Fusion of phospholipid vesicles with each other results in it change in size. A variety of techniques can be used to monitor size changes, including light scattering (Lansman and Haynes, 1975) and gel exclusion chromatography (Liao and Prestegard, 1979). One advantage to using light scattering techniques is that the size changes can be time resolved in the millisecond range using stopped-flow rapid mixing techniques. Such measurements were done both with natural membrane vesicles (Morris et ( I / . , 1979; Haynes et d.,1979) and with artificial vesicles (Lansman and Haynes, 1975). Since Rayleigh scattering can, in theory, be related to the size as well as the shape of the particle, this\ technique should ideally distinguish between aggregation (a dumb-bell shape) and fusion (a larger sphere). However, in practice this is not so straightforward. In fact, most methods that monitor the putative fusion event do not distinguish between real fusion (merging of bilayers concomitant with mixing of aqueous contents) and other processes. Mixing of membrane components might occur by diffusion of individual phospholipid molecules or protein molecules from one vesicle to another. The technique of differential scanning calorimetry can distinguish between molecular mixing of membranes as a result of diffusion of individual components from other processes occurring in vesicle populations (Papahadjopoulos, 1978). However, this technique fails t o distinguish between genuine fusion and a process of rupture fol-

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lowed by reannealing of the resulting membrane fragments with one another to form larger structures. During the past years a number of assays have been published which were designed to demonstrate fusion of phospholipid vesicles to each other. These fall into two classes: those which show mixing of the membrane components (labeled phospholipids or lipid-soluble probes) and those which indicate the mixing of labeled material trapped within the vesicle lumens. Both methods can be used for studying the kinetics of the fusion process. The assay for mixing of membrane components is based on fluorescence energy transfer between a donor and acceptor group, both of which are attached to a phospholipid. The concept relies on the ability of a fluorescent donor to excite an energy acceptor if both are free to diffuse in the membrane. Fung and Stryer (1978) had shown that the energy transfer efficiency depends on the surface density of the acceptor for a given donor concentration. If vesicles containing both donor and acceptor fuse with unlabeled vesicles, a change in donor quenching and sensitized emission is observed. This is consistent with lateral diffusion of the fluorescent lipids in the plane of the enlarged membrane area. It can not be excluded that the energy transfer changes are due to exchange or aggregation. However, Struck et ad. (1981) have shown that probes labeled in the head group do not spontaneously exchange between vesicles. Moreover, in studies with tubulin-vesicle complexes in the presence and absence of Ca“, it was possible to distinguish between aggregates and fused structures: aggregates exhibited no energy transfer changes, whereas fusion resulted in marked changes in energy transfer (Kumar et al., 1982). Several fluorescent assays have been described to monitor mixing of aqueous vesicle contents (reviewed by Nir et ul., 1983). One such assay for core mixing involves the fast formation of a chelation complex between Tb3+,encapsulated as a citrate complex in one population of vesicles, and dipicolinic acid in the second population (Wilschut and Papahadjopoulos, 1979). Fusion and mixing of contents results in the formation of a fluorescent Tb3’-dipicolinic acid complex. Presence of Ca’+ and EDTA in the external medium prevents the formation of the fluorescent complex outside the vesicles. This system does not work well for ‘‘leaky’’ fusion (Kendall and MacDonald, 1982; Morris et al., 1985). Ideally, both assays need to be carried out simultaneously to demonstrate fusion (Rosenberg rt al., 1983), since mixing of membrane components does not necessarily imply core mixing, and vice versa. A study by Wojcieszyn et af. (1983) on mixing of fluorescent membrane and cytoplasmic probes in polyethylene glycol (PEG)-mediated fusion of cells is of interest in this context. Fusion of liposomes with planar phospholipid bilayers is being actively pursued in a number of laboratories as a way to incorporate biologically

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functional membrane proteins (membrane channels) into planar bilayers (see review by Blumenthal and Klausner, 1982).This system is also being studied as a model for exocytosis (Zimmerberg rt al., 1980a,b; Cohen rt ul., 1980). Its morphology corresponds more to the secretory vesicleplasma membrane interaction, whereas vesicle-vesicle fusion corresponds to compound exocytosis (Pollard rt al., 1982) or to fusion between intracellular organelles. To demonstrate fusion it must be shown that vesicle contents are transferred across the planar membrane concomitant with incorporation into the planar bilayer of a membrane protein (channel) originally present in the liposome membranes. The first criterion has been satisfied in a series of experiments (Zimmerberg et al., 1980a), where transfer of fluorescently labeled material from liposomes across planar bilayers was observed. Most of the liposome-planar bilayer fusion assays demonstrate mixing of membranes by monitoring channel activity in the bilayer of a pore protein, isolated from the outer membrane of bacteria or mitochondria. This is the equivalent of the lipid mixing assay in vesiclevesicle fusion. Osmotic swelling of vesicles, attached to the planar bilayer in “pre-fusion” state, is a necessary and sufficient condition for fusion in this system (Cohen rt ul., 1982). In this assay, the same conditions led to both membrane mixing and transfer of contents. 111.

THE TRIGGERING EVENT IN MEMBRANE FUSION

In a wide range of secretory tissues an increase in cytosolic Ca” activity from a resting value of 0.1 JLMto an activated value of 1-10 p,M appears to be the trigger of the exocytotic release mechanism (Llinas and Heuser, 1977). The increase in cytosolic Ca” activity could be a result of an increase in the rate of Ca” influx from the extracellular medium across the plasma membrane or a release of Ca” from internal stores. I n nerve terminals the former appears to be more likely. There are many ways to show that Ca2’ influx is the trigger. In the classical experiments on release of neurotransmitters from presynaptic terminals, the relationship between Ca2+and exocytosis was established electrophysiologically. As the nerve impulse travels along the axon, it depolarizes the presynaptic membrane which results in the opening of highly selective voltage-dependent Ca” channels. Unlike the Na’ channels these so-called late Ca” channels are blocked by various di- and trivalent cations as well as by some pharmacological agents such as verapamil, D-600, and nifedipine (Baker. 1972). Other ways to show the importance of Ca” influx as a trigger include uptake of radioactive Ca” and intracellular injection of dyes that show a change in absorption or in fluorescence when they bind to Ca”. A highly

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effective fluorescent probe for intracellular Ca” , quin2, has been synthesized (for a review on the measurement of intracellular Ca” activity, see Blinks et nl., 1982). Exocytosis can be induced in isolated secretory cells, such as bovine adrenal medullary cells, by adding chemical agents called “secretagogues” to the bathing medium (Schneider et al., 1981; Baker and Knight, 1981). Nicotinic cholinergic agonists (such as acetylcholine), veratridine, and elevated K’ concentration in the medium are secretagogues in this system. Experiments with Ca” channel blockers indicate that the effect of acetylcholine is indirect: interaction of acetylcholine with its membrane receptor causes depolarization of the plasma membrane, which in turn triggers the opening of the voltage-sensitive Ca” channels (Schneider et a ( . , 1981;Baker and Knight, 1981).[A small portion of the Ca” influx might, however, be mediated by the acetylcholine receptor channel (Schneider et nl., 1981).]Veratridine or high K + in the medium brings about a similar effect by depolarizing the membrane. In mast cells, binding of antigen to IgE, which is itself bound to the IgE receptor on the cell surface, leads to influx of Ca” and subsequent exocytosis. Recent data indicate that cross-bridging of two IgE receptors is sufficient to open Ca” channels in basophils (Mazurek rt al., 1984). Release of Ca’+ from intracellular stores has been associated with turnover of phosphatidylinositol (PI) (Berridge, 1984;Nishizuka, 1984).A wide variety of hormones or neurotransmitters stimulate this turnover by interacting with their receptor on the cell surface. Evidence indicates that stimulation of the receptor activates a phospholipase, which rapidly mediates the degradation of phosphdtidylinositol4,5-bisphosphate(PIP-,) to produce 1,2-diacyIglycerol (DG) and dernyo-inositol 1,4,5-trisphosphate (IP,) (see Section VI1,B). The IP, is released to the cytosol, where it presumably acts as a second messenger to release Ca’+ from internal storage depots, and thereby to raise [Ca”], required for exocytosis. Reports from several laboratories indicate that IP, induces a rapid release of Ca’+ from intracellular organelles in permeabilized pancreatic acinar cells and hepatocytes (reviewed by Berridge, 1984). In permeabilized cells Knight and Baker (1982)found a K!, for exocytosis from bovine adrenal cells of about 1 pM [Ca’+],.’ Those low Ca” concentrations are insufficient to induce fusion between pure bilayers (see Section VIll for more details). The micromolar Ca’ ’ sensitivity indicates that a high-affinity Ca’+-binding protein might mediate the Ca”-induced effects at the exocytotic site. Several proteins have been proposed to play ‘Exocytosis in this system also required millimolar MgATP. However, the MgATP might be required as a source of energy, rather than as a trigger for exocytosis.

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a role of this sort. The finding that a calmodulin antagonist, tiifluoperazine, inhibits exocytosis in permeabilized adrenal cells (Knight and Baker, 1982) and in isolated sea urchin egg cortices (Whitaker and Baker, 1983) indicates that calmodulin might play this role. Other Ca’ ’ -binding proteins possibly involved in fusion, such as synexin (Creutz ri d.,1978) and an unidentified protein from brain which markedly enhances the vesicle-planar bilayer fusion (Zimmerberg et l i l . , 198Ob), will be discussed in Sections V and IX,respectively. Rink et (11. (1983)have shown that increased levels of DG resulting from receptor-mediated PI breakdown can stimulate exocytosis in platelets without raising cytoplasmic [Ca”]. DG analogs or phorbol esters added exogenously evoke similar secretion without elevating [Ca”], above the basal level of 0.1 pM. DG stimulates protein phosphorylation through activation of the Ca”-activated, phospholipid-dependent protein kinase C (Kaibuchi r t NI., 19831.‘ Knight and Bakcr (1983) have shown that a phorbol ester can shift the Ca”-activation curve for exocytosis in permeabilized chromaffin cells to lower [Ca’+],.This is consistent with the notion that a change in the affinity of the Ca”-sensitive process (i.e., protein kinase C) for Ca’ ’ controls Ca” &pendent exocytosis without altering [Ca” I,. In polarized cells, such as the pancreas acinar cell, exocytosis from thc apical surface is triggered by ligand-receptor interaction on the basolateral surface (Gardner and Jensen, 1981). The mechanism of intracellular communication of this signal is still unclear. It is possible that hormone-receptor interactions on the basolateral surface lead to depolarization of apical membrane. This depolarization could trigger release. Ca’ ‘ does not always seem to be the mediator of release. The interaction of some hormones with their receptors on the basolateral surfacc causes elevation of intracellular cyclic AMP levels. This might be the trigger for the release (Gardncr and Jensen, 1981). The Ca2+concentration in extracellular fluids is about 2 mM in mamis about 10,000, a very steep electrochemical mals. Since [Ca”],J[Ca”], gradient will tend to force Ca” into the cells. Moreover, during activation [Ca”], is raised according to mechanisms outlined above. Control of exocytosis by small changes in [Ca’+Ii necessitates maintenance of a low basal level of [Ca”],. Ca” pumps or Ca’+-Na+ exchange carriers have been established in the membranes to achieve low concentrations of Ca” (Blaustein et a / . , 1980). In secretory cells such pump or exchange systems ‘Upon activation. protein kinase C mediates the phosphorylation o f a 40-kI)a protein (Kaibuchi cr d.. 1983). Although the phosphorylation o f this protein is a prerequisite for serotonin releahe in platelets, its precise role i n exocytosis has yct to be determined.

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are localized in the plasma membrane, endoplasmic reticulum, or mitochondria (Henkart, 1980). Viral fusion events are often triggered by an increase in hydrogen ion concentration within the endosome (White et al., 1983). The virus binds to the plasma membrane and is endocytosed via the coated pit pathway. Once it arrives in the endosome, the virus membrane fuses with the endosome membrane. The fusion is triggered by the elevation of H’ activity from 0.1 t.M (pH 7) to 1-10 (pH 6-5) (White et al., 1983). Interestingly, this ion activity change is in the same range as that of the Caz+trigger in exocytosis. The H+ enter the endosome via a proton pump (Forgac et al., 1983; Stone et al., 1983). Fusion of viral membrane to the plasma membrane can be triggered directly in these viruses by lowering the pH to 5-6 outside the cell (White et al., 1983). IV. MOVEMENT INTO APPOSITION A. Directed Movement

A precondition for exocytosis is movement of the secretory vesicle to its site for exocytosis at the plasma membrane. Motions of intracellular organelles observable in cells include cytoplasmic streaming, Brownian motion, saltatory motion, and axonal transport in neurons (Allen et al., 1981). All of these except Brownian motion are characterized as intracellular motility, since they require metabolic energy. Brownian motion is the result of thermal agitation. The mechanisms of intracellular motility are very poorly understood. Cytoskeletal elements involved in intracellular motility are good candidates for having important functions in the secretory process. The actin filament interacting with other proteins has been proposed as a prime candidate for generating movements in secretion (Stossel, 1981) in analogy with its function in generating muscle motility. According to a model proposed by Pollard et al. (1982), chromaffin granules are immersed in an actin network at intracellular Ca2’ levels of less than 0.1 pM. In this model the chromaffin granule membrane binds to two or more actin filaments via a putative actin binding site on the cytoplasmic surface of the granule. This cross-linking gives rise to a large increase in viscosity in an in vitro assay (Fowler and Pollard, 1982). When the Ca” concentration rises to greater than I pLM the actin filaments dissociate from the actin binding sites on the granule membrane, thus removing the cross-links and resulting in a lower viscosity of the solution. The secretory vesicles would then be free to move to the plasma membrane. The sensitivity of this cross-linkingto low levels of Ca2+is conferred by a protein called gelsolin

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which binds to actin and dissolves cross-linked gels at a threshold Ca” concentration of 0.2 p,M (Stossel, 1981). Its maximal activity is at about 10 p M Ca”. This concentration range is consistent with the cytosolic Ca” concentration required to trigger exocytosis. The role of cytoskeletal elements has been probed using drugs such as cytochalasin B and vinblastine or colchicin, which specifically interact with microfilaments and microtubules, respectively. Schneider et al. (1981) found that these drugs do not inhibit K’-evoked release. Knight and Baker (1982) reached the same conclusion using bovine adrenal medullary cells permeabilized by exposure to intense electric fields, and Whitaker and Baker (1983) showed that neither cytoskeletal drugs nor antibody to actin had an effect on exocytosis in the sea urchin egg cortical granule preparation. Therefore, it seems unlikely that actin or tubulin participate intimately in exocytosis. For a recent review on the interaction of secretory granules with cytoskeletal elements and other cytosolic proteins see Burgoyne (1984).

8. Random Movement

In model studies with isolated chromaffin granules or with lipid vesicles the movement into apposition induced by divalent cations (or high concentrations of monovalent cations) occurs by random collisions (Lansman and Haynes, 1975; Morris el al., 1979; Nir et al., 1983). The probability of forming stable aggregates has been estimated in the framework of the Derjaguin-Landau-Verwey-Overbeek (DLVO) theory developed for flocculation of colloidal particles (Verwey and Overbeek, 1948; the DLVO theory is discussed in extenso in Section VIII). Studies using stoppedflow rapid mixing techniques have been carried out on aggregation of natural and artificial phospholipid vesicles (Lansman and Haynes, 1975; Morris r t al., 1979). It has been shown that it is a second-order process following the scheme V

+

V

--j

Vz

-j

(aggregate)

(1)

where V is a vesicle monomer, V2 a dimer, and the aggregate is any complex beyond the dimer state. Prior to reaction ( I ) there is an ion-binding step, which is very rapid. The rate constant of dimer formation kapp can be assessed by stopped-flow kinetics. If this value is determined solely by the rate at which two vesicles can diffuse up to each other (diffusion control), then the rate constant for bimolecular collision ( k , ) can be predicted from a theory developed by Smoluchowski (see Verwey and Overbeek, 1948). His treatment for spherical particles gives

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k , = 8kTNI3q

(2)

where N is the number of particles, and q the viscosity of the medium (see Lansman and Haynes, 1975). This theory predicts that the rate constant for collision will depend only on the absolute temperature (7') and viscosity of the medium and that it ill be independent of the particle size. Using this result a value of 6.8 x lo9 M - ' sec-' is calculated fork,. This is about 500 times greater than the measured rate constant for Ca"-induced dimerization of negatively charged phospholipid vesicles (Lansman and Haynes, 1975). From these data, Lansman and Haynes (1975) proposed that dimerization proceeds by a mechanism in which vesicles first collide to form an encounter complex followed by a slower conversion to a stable complex. For phospholipid vesicles, 1 in about 500 collisions results in stable dimer formatin, according to that model. Interestingly, intact chromafin granule membranes and T o r p ~ d oelectric organ synaptic vesicles form stable contacts with each other at near diffusion-controlled rates (Morris el al., 1979; Haynes et al., 1979). Modification of the membrane proteins or use of extracted membrane lipids to form artificial vesicles greatly reduces the success of collisions. This strongly suggests that the major determinants of the aggregation rates of natural membranes are proteins which protrude from the membrane surface and serve as points of contact. This contact does not seem to require full neutralization of the surface charge (see Section VIII). According to Eq. (2) k, should be independent of particle size; the lower diffusion coefficient of the larger particle is compensated by its larger surface area for contact. However, Morris et al. (1985) found that the kupp value for small unilamellar vesicles (about 250 A diameter) was 5 times greater than that for large unilamellar vesicles. Equation (2) assumes that there are no repulsive forces between aggregating particles. However, it is clear that there is a tremendous repulsive barrier between phospholipid surfaces at separation distances of about 20 A (see discussion in Section VIII). To account for this barrier the Smoluchowski theory has been modified (see Nir and Benz, 1978) to include a factor W i n the expression for the rate constant of dimerization, which is then given by

k , = 8kTNl3qW

(3)

If there is no repulsive barrier (as for instance with chromaffin granule aggregation) then W = 1. In fact, 1IW is the fraction of vesicle-vesicle collisions which lead to a stable aggregate. Nir and Benz (1978) have attempted to calculate W from first principles. However, they do not include in their calculation the hydration repulsion barrier nor possible deformation

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forces (see Section VIII). If the measured rate constant (k,,,,) is assumed to be equal to k , in Eq. ( 3 ) , then W can be obtained directly from Eq. ( 3 ) since all other parameters are known. Wong and Thompson (1982) use the value of W obtained in that way to calculate barrier height and separation distance for aggregation of DPPC vesicles. Apposition of viruses to the plasma membrane probably occurs through random collisions. However, apposition of secretory granules by random collisions is not likely to occur in physiological situations. Nevertheless, these studies do allow us to learn something about energy barriers to apposition. Moreover, according to Morris ei rrl. (19833, the fact that chromaffin granules aggregate spontaneously in rVtro but not in viw at physiological K’ concentrations indicates that those granules must be held apart in the cell by the cytoskeletal network (see Section IV,A).

V.

THE RECOGNITION EVENT

It is clear that some specificity is involved in the interaction between the plasma membrane and the secretory granule membrane. For instance in polarized cells such as the pancreatic acinar cell the secretory granule will only fuse with a specialized membrane region, the apical mcmbrane of the cell (Gardner and Jensen, 1981). Neurotransmitter release also takes place at specialized regions of presynaptic nerve endings (Llinas and Heuser, 1977). I t is not clear what determines that specit’icity. One possibility is that the cytoskeletal network localizes thc granule at the appropriate location for exocytosis. Alternatively. the presence of receptors on both membranes, granule and plasma membrane, could cause specific localized exocytosis. There could be a receptor site on the secretory vesicle which recognizes a “global” structure on the plasma membrane. I do not know of any proven cases of such receptors on either plasma membrane or secretory vesicle membrane. Moreover, it has not been demonstrated that a secretory granule will not fuse with the basolateral membrane, given a proper chance. Alternatively, a cytosolic molecule, when triggered. could mediate crossbridging of the two membranes. Indeed such molecules have been identified. A 47,000-MW protein called syncxin, which has been isolated from chromafiin cells and from liver, mediates aggregation of vesicles in a Ca’ dependent way (Creutz er d.,1978). The Ca” dependence for aggregation was estimated by turbidity changes, and the K,, value was found to be approximately 200 p M . This appears to be higher than expected for exocytosis ( K , about 1 p M Ca”, see Section I l l ) . Moreover. Morris and Hughes (1979) found that synexin is quite unselective and will lower the

220

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Ca” requirement for a variety of natural and artificial lipids. Synexin also promotes fusion of negatively charged phospholipid vesicles (Hong et al., 1982), and, if cis-unsaturated fatty acids are added, synexin will fuse chromafin granules (Creutz, 1981). Although these observations are of great interest in formulating models for protein-mediated fusion of bilayers (see Section VIII), their relevance to synexin as a specific mediator of exocytosis has been questioned (Morris et al., 1983; Burgoyne, 1984). A 33-kD protein called calelectrin, with similar CaZ+-specifcaggregation activities, has been isolated from Torpedo electric organ (Sudhof et al., 1982). We may look forward to the identification of a plethora of such Ca2’-dependent aggregation proteins. They will have to be reconstituted and characterized step by step with other cytoskeletal components of the exocytotic machinery in order for us to understand their in vivo importance. Moreover, those proteins need to be localized at the appropriate regions (where exocytosis takes place), using electron microscopic immunocytochemical techniques. Since virus infection is in many cases selective with respect to the host, the virus must interact with the cell surface in a specific way.’ The specificity could be conferred either by molecules on the viral surface or by molecules on the host cell membrane surface, or both. While little is known about receptors for viruses on the host cells, the surface glycoproteins on the viral cell membrane have been well characterized. Owing to the simplicity of the virus membrane, and the easy access to the genes, spike glycoproteins are among the best characterized membrane proteins. White et al. (1983) have reviewed spike glycoprotein composition and their sequences for a number of enveloped animal viruses. Sendai virus, the best studied member of the paramyxoviruses, has two major spike glycoproteins: HN, which is responsible for attaching the virus to cell surface sialic acid residues and for the subsequent cleavage, and F, which is presumably a fusion protein. Glycophorin A has been identified as the receptor for Sendai virus in red blood cells (Nigg et al., 1980). However, it has been shown that specific gangliosides can function as host cell receptors for Sendai virus (Markwell et al., 1981). The membrane of influenza virus, a well-characterized member of the orthomyxoviruses, also has two major spike glycoproteins which have neuraminidase, hemagglutination, and fusion activities. However, one of the spike glycoproteins (NA) is a neuramidase, and the other, the hemagglutinin (HA), contains the capacity both to bind to cell surface sialic acid residues and to catalyze fusion. The HA consists of two disulfidelinked glycopolypeptide chains, HA1 and HA2, which are derived by proteolytic cleavage from a precursor glycopolypeptide called HAO. This ’The specificity might, however, reside in the replication step.

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proteolytic cleavage renders HA fusion competent. The three-dimensional structure of the ectodomains of both HA and N A have been determined to a resolution of 3 A (Wilson et a / . , 1981, and Varghese et a / . , 1983, respectively). Rhabdoviruses consist of a bilayer membrane with a single type of glycoprotein (usually designated the G protein). The G protein from vesicular stomatitis virus (VSV) has been cloned and sequenced: it is a 70-kDa transmembrane protein with a hydrophobic run of amino acids residues close to the C-terminus that forms the intramembranous domain. Since it is the only cell surface protein, it must comprise both recognition and fusogenic functions. Direct evidence that both functions reside solely in the G protein has recently been obtained by expressing a complementary DNA clone encoding that protein in a stable cell line and showing polykaryon formation at pH 5.5 (Florkiewicz and Rose, 1984). Schlegel et al. (1982) described a saturable, high-affinity binding site for VSV on the surface of Vero monkey cells that appears to mediate viral infectivity. Analysis of a detergent extract from Vero cells, which inhibits VSV binding and infectivity, indicated that this binding site might be phosphatidylserine (Schlegel et al., 1983). VI.

STERIC CONSTRAINTS

In a previous section 1 have discussed constraints imposed by actin filaments on movement of secretory granules to the plasma membrane and how these constraints might be removed by dissociating the filaments at increased Ca” concentration in the presence of gelsolin. However, some microfilamentous material seems to remain in the fusion areas (Ornberg and Reese, 1981). Clathrin-coated pits (Steer and Klausner, 1983) provide another example how cytoskeletal elements can provide constraints on membrane fusion. Clathrin-coated pits and clathrin-coated vesicles mediate endocytosis of ligands bound to cell surface receptors (Goldstein et al., 1979) and appear to be involved in many aspects of intracellular traffic (Pearse and Bretcher, I98 I ) . These endocytosis and exchange events require membrane fusion. Morphological studies indicate that endocytosed material is rapidly sequestered into coated pits and vesicles but subsequently appears in smooth vesicles, which in turn fuse with other organelles such as the Golgi or lysosome (Pastan and Willingham, 1981). Altstiel and Branton (1983) developed an in vitro assay for fusion of coated vesicles with lysosomes. Their results indicated that the clathrin coat of coated vesicles inhibits the fusion of the vesicle membrane with that of the lysosome. Although conditions for stripping clathrin coats have been developed in the in vitro system, it is not clear what the physiological

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mechanisms for uncoating are. Recent data indicate that hydrolysis of ATP mediated by an “uncoating ATPase” is involved in the uncoating process (Braell et al., 1984). Steric constraints can also arise from integral membrane proteins, which extend out a considerable distance into the medium. This will prevent bilayers from coming together. Electrostatic repulsions due to proteins might also play a role. Theoretical calculations (Dembo and Bell, this volume) and experimental evidence suggest that there is a strong nonspecific repulsive force between most cells, arising from electrostatic repulsion and from solvent exclusion from the hydrated polymers extending from their surfaces (steric stabilization). The negative charge on surface glycoproteins is due to terminal sialic acid residues. Removal of these residues by neuraminadase dramatically increases the aggregation potential of red blood cells in dextran solutions or plasma (Buxbaum et al., 1982). The neuraminadases, which form part of the spike glycoproteins of viruses (see previous section), might serve to remove the electrostatic repulsion due to glycoproteins and glycolipids on the surface of the host cell. However, lateral movement appears to be the way to remove glycoproteins and glycolipids from the contact area between two apposing membranes (Lawson et al., 1977). Based on measurements of lateral diffusion coefficients of membrane proteins by a variety of techniques, including fluorescence recovery after photobleaching (FRAP), it has been shown that lateral diffusion of membrane proteins is about an order of magnitude slower than expected from free diffusion theory (Edidin, this volume). These considerations led to the notion that movement of membrane proteins is under cytoskeletal control. This has been borne out by experiments showing that when those controls were relaxed, for instance in blebs, the measured diffusion coefficients were close to the values expected from diffusion theory (Edidin, this volume). These observations are consistent with the idea that cytoskeletal elements can control movement of certain integral proteins into or out of an area where membranes are about to fuse. Rotational mobility of membrane proteins has also been measured as a way to monitor changes in protein-protein interactions in membranes concomitant with fusion. For example, Nigg et al. (1980) have shown that both influenza and Sendai virus markedly reduce rotational mobility of band 3 in erythrocytes. This immobilization was presumably caused by cross-linking of the virus receptor, glycophorin A, by the absorbed particle. Band 3 was then immobilized due to its association with glycophorin A. Sendai virus, but not influenza virus, fused with the plasma membrane and also caused aggregation of intramembrane particles, as shown by freeze-fracture electron microscopy. Nigg et al. (1980) hypothesized that

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MEMBRANE FUSION

during or following Sendai virus-membrane fusion. integral proteins are released from the restriction to long-range lateral diffusion normally imposed on them by the cytoskeleton. Thus control of movement of membrane glycoproteins might be an other role played by cytoskeletal elements in membrane fusion. The clearing of intramembrane particles at the sites of fusion has been attributed to a glycerination artifact (Chandler and Heuser, 1979; Ornberg and Reese, 1981). On the other hand the formation of a definite pattern of intramembrane particles at the fusion sites has been observed in protozoan cell membranes ("junctions," Satir, 1974)or in rapid frozen specimens ("exocytotic pores," Ornberg and Reese, 1981). It is. however, not inconceivable that some membrane particles (e .g., cell surface glycoproteins) need to be swept away while others (embedded membrane proteins) are required to form ':junctions" for fusion of biological membranes.

VII.

MOTION OF PHOSPHOLIPIDS

A. Studies with Natural Membranes Membrane fusion involves a catastrophic rearrangement of membrane components in its transition from two bilayers to one single one. We would therefore expect considerable rearrangements of membrane lipids during the process. Membrane lipids undergo a plethora of different motions which occur at different time scales (Smith and Oldfield, 1984). They include trans-gauche isomerization ( 107/sec),axial diffusion (IO"/sec), translational motion ( cm-'/sec). and phospholipid flip-flop. There is also a segmental motion, that is, rotational diffusion around a carboncarbon bond, whose rate constant varies from IO"/sec to I09/secdepending on its position along the fatty acyl chain in the bilayer. This motion has also been described in terms of an "order parameter." These motions have been monitored by a variety of spectroscopic techniques: Raman spectroscopy, NMR, ESR, fluorescence polarization, and lifetime analysis (Kleinfeld, this volume). The flip-flop motion is the movement of a phospholipid head group from one side of the bilayer to the other. Early studies with liposomes showed that half-times of flip-flop in unperturbed bilayers range from days to weeks; in contrast, tlip-flop in some biological membranes takes place in minutes or even seconds. It has been shown that phospholipid flip-flop is accelerated in reconstituted systems by the presence of integral membrane proteins (for a review on phospholipid flipflop, see Dawidowicz, this volume).

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In theory a decrease in motional restriction of membrane lipids should enhance fusion. In practice this is hard to demonstrate since only a small proportion of all the membrane lipids participate in the fusion reaction. Attempts have been made to change overall membrane packing (membrane “fluidity”) by altering phospholipid composition or cholesterol content. For instance, virus-induced fusion of hen erythrocytes was enhanced by enrichment of the erythrocyte membranes with cholesterol (Lucy, 1978). Horwitz et al. (1978) have carried out a thorough study on the effect of lipid alterations on fusion of cultured myogenic cells. Lowering sterol levels by inhibiting sterol synthesis inhibits fusion. This indicates a requirement for cholesterol for fusion in this system. In general cholesterol decreases fluidity of membranes, whose lipids are in the liquid-crystalline state, and these data would indicate that less fluid membranes fuse better. On the other hand, enrichment of fatty acyl chains with elaidate inhibits fusion in this system, whereas enrichment with oleate enhances fusion. These results are consistent with the data of Ahkong et al. (1973) showing that cis-unsaturated fatty acids and monoglycerides containing cisunsaturated fatty acids were effective in inducing fusion of hen erythrocytes whereas trans-saturated fatty acids were not. Similarly, Creutz (1981) has shown that fusion of aggregated chromaffin granules could be caused by addition of arachidonic acid. Other unsaturated fatty acids could mimic the effect, whereas saturated fatty acids did not enhance fusion. It has been hypothesized, based on fluorescence polarization data, that cis-unsaturated fatty acids “fluidize” membranes (Kleinfeld, this volume). This fluidization would then make those membranes more fusogenic. This is in contradiction to the cholesterol data, which could be interpreted as showing that less fluid membranes are more fusogenic. Therefore, it seems that membrane fluidity is not a very good parameter or predictor of a membrane’s fusogenic properties. The lipid phase in membranes seems to be not homogenous but rather organized in a domain structure (Kleinfeld, this volume; Hui, this volume). Although cis-unsaturated fatty acids change fluorescence polarization of a membrane probe, they do not seem to change lateral mobility (fluidity) of membrane lipids (Kleinfeld, this volume). Biological membranes are also inhomogeneous in the perpendicular direction, that is, they have different lipid compositions on either side of the membrane (Dawidowicz, this volume). Therefore, even if the overall lipid composition is modified, the membranes might still form appropriate domains for fusion. The amount of unsaturated fatty acids added to make membranes fusogenic is quite high (more than 1%). Therefore, in high concentration, they could render membranes fusogenic by creating “defects” (see Section VIII). Alternatively, fatty acids or cholesterol could affect fusion indirectly by

MEMBRANE FUSION

225

changing the lateral arrangement of proteins in the membrane or by interacting with specific proteins involved in fusion. Another aspect of the role of phospholipid mobility in fusion is that membrane fusion is often concomitant with phospholipid metabolism or turnover. In mast cell exocytosis, transmethylation seems to occur prior to fusion (Ishizaka et al., 1980). These authors believe that membranes undergo a fluidity change concomitant with lipid transmethylation. However, it also could be that lipid transmethylation has a specific effect on membrane proteins involved in fusion. Siraganian et af. (1983)have observed release of arachidonic acid concomitant with exocytosis in mast cells. Since this cis-unsaturated fatty acid can induce membrane fusion (see above), one might assume that the metabolic event generating arachidonic acid is the direct cause of fusion of the secretory granule with the plasma membrane. However, the amount of arachidonic acid release is very small compared to the amount needed for fusion in vitro. Furthermore, a transient very high local concentration of arachidonic acid in the fusion area seems unlikely in view of the rapid diffusion constant of fatty acids. Turnover of phosphatidylinositol has also been associated with exocytosis (Nishizuka. 1984). A wide variety of hormones or neurotransmitters stimulate PI turnover by interacting with their respective receptors on the cell surface. Initially, PI was regarded as the prime messenger, but more recent evidence indicates that hosphatidylinositol4,S-bisphosphate(PIP,) is degraded rapidly after stimulation of the receptor to produce 1.2-diacylglycerol (DG) and demyo-inositol I ,4,S-trisphosphate (IPJ. The 1P, is released in the cytosol, where it presumably acts as a second messenger to release Ca” from internal storage depots and thereby to raise intracellular Ca” activity required for exocytosis (see Section 111). Concomitantly, the DG initiates the activation of a specialized intracellular protein, called protein kinase C. Exocytosis caused by the synergistic action of protein kinase C activation and Ca” mobilization occurs in a wide variety of tissues (see Section 111). Here we see a clear role for lipid mobility in membrane fusion. However, the specific lipid (DG) acts as a cofactor to a protein involved in the exocytotic cascade rather than as a global mediator of the “fluidity” of the membranes involved in fusion. B. Studies with Artificial Lipid Membranes

Since liposomes can be prepared homogeneously either in the gel phase or in the fluid phase, depending on temperature, chain length, or degree of unsaturation of the lipid, the effect of fluidity on fusogenicity can readily be tested in that system. In their studies on liposome-cell interactions,

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Poste and Papahadjopoulos (1976) found that neutral phospholipids and charged vesicles composed of phospholipid mixtures such as PS/DSPC/ DPPC, which are “solid” at 37°C (i.e., below their gel to liquid crystalline transition temperature at the experimental temperature), are unable to fuse with cells. In contrast, charged lipsosomes composed of phospholipid mixtures that are “fluid” at 37°C will readily fuse with the plasma membrane of cells cultured in vitro. However, these findings were based on incomplete evidence for fusion (see Section 11). In studies on vesiclevesicle fusion, on the other hand, it was found that small unilamellar vesicles (SUVs) made of PC with saturated chains fuse spontaneously below the lipid phase transition temperature (Larrabee, 1979; Schullery et NI., 1980), albeit at a very slow rate (time for completion of the order of days). Phosphatidylcholine vesicles in the fluid phase do not fuse spontaneously. This is consistent with the large hydration repulsion brought about by the PC head group (see Section VIII). According to Evans and Parsegian (1983) the behavior of SUVs is opposite to that of large unilamellar vesicles (LUVs). In SUVs the vesicle membrane thickness is no longer neglible in relation to the vesicle radius. Therefore its mechanical properties can not be represented by simple tension and moment resultants obeying equations like the law of Laplace (Evans and Parsegian, 1983). Instead, the lateral stress distribution along the acyl chains must be considered. In the fluid phase SUVs are rigid, in contrast to LUVs which are readily deformed in the fluid phase. Small vesicles can only partially freeze, leaving large liquid-like defects because of molecular packing restrictions. Those defects can then form the sites for vesicle-vesicle fusion (see Section VIII). According to Papahadjopoulous ( 1978) the crucial event in membrane fusion is a Ca”-induced lateral phase separation which results in the formation of rigid crystalline domains of acidic phospholipids within a mixed lipid membrane. Fusion is then initiated between closely apposed membranes at the boundaries between those domains. Such boundaries represent structurally unstable points (defects), and thus offer focal points for mixing of molecules from apposed membranes. This hypothesis is based on data showing a strong correlation between Ca”-induced aggregation, increase in permeability, onset of phase changes, and fusion of negatively charged vesicles (Papahadjopoulos, 1978). In studies of the effect of the lipid phase on membrane fusion, nearly all types of behavior have been observed in one system or another. Fusion

nSilvius and Gagne (1984) observed a strong relationship between Ca’ ‘ -induced lateral phase separation and fusion in PS/PC mixtures. However. in PS/PE mixtures Ca” -induced fusion occurred without lateral phase separation.

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was found to be maximal below, at, or above the phase transition temperature (see Nir et d.,1983). Similarly, studies with liposomes of the effect of cholesterol (which tightens packing of membranes whose lipids are in the the fluid phase) on fusion have also shown no clear-cut picture. Fusion of Semliki Forest virus required cholesterol in the target liposome (White and Helenius, 1980). In contrast, fusion of influenza virus with liposomes did not show any cholesterol requirement (Maeda o r d..1981; White o r d., 1982). Similarly, cholesterol did not affect the rate of fusion of liposome fusion mediated by VSV G protein reconstituted in phospholipid vesicles (Eidelman rr ul., 1984). In the case of Sendai virus, in one study cholesterol in the target liposomes was required for fusion (Hsu er c d . , 1983), but in another study it seemed to decrease fusion (Haywood and Boyer, 1984). Both the phase state and the cholesterol content affect bilayer packing, lipid mobility, as well as vesicle shape, size, and mechanical properties. Therefore, only if all these factors are taken into account in the model systems will it be possible to come to definite conclusions regarding the role of lipid fluidity in membrane fusion. VIII.

INTERACTIONS BETWEEN BILAYER MEMBRANES

A. Forces between Bilayers A necessary condition for fusion of membranes is close contact. The likelihood of surfaces coming in close contact has been estimated in the framework of the Derjaguin-Landau-Vewey-Overbeek (DLVO) theory (Verwey and Overbeek, 1948). This theory has been developed for aggregation of colloidal particles. It considers the interplay of attractive van der Waals and repulsive electric double layer forces between charged surfaces in liquids. It is assumed that the dispersing liquid (usually water) can be treated as a structureless medium defined solely by its bulk dielectric permittivity (dielectric constant, refractive index). According to the theory, the free energy of interaction between particles G ( d )can be calculated from the electrostatic and electrodynamic contributions, as a function of distance of separation (d).This energy can be compared with the kinetic energy of translational motion of spheres, AT. The ratio p ( d ) between those particles in suspension that have and those that have not sufficient kinetic energy to come to a given distance of separation ( d ) is given by p ( d ) = exp[G(d)/kTj

(4)

Due to the combination of the opposing forces, that energy usually goes through a minimum as a function of distance of separation. The depth of

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ROBERT BLUMENTHAL

the energy minimum determines the stability of colloidal suspensions; the aggregate will be stable if GIkT >> 1. While sufficient to create ordered arrays of bilayer membranes, the energy barriers effected by van der Waals attraction are weak in any practical mechanical sense. It has become possible in recent years to test the DLVO tbeory experimentally by measuring force as a function of distance at 1 A resolution (Rand, 1981; Israelachvili, 1982). These measurements have shown a remarkable verification of the theory for mica surfaces in dilute electrolyte solution. However, there are many systems for which the DLVO theory clearly fails, the most notable example for our interests being interactions between bilayers. LeNeveu et al. (1976) measured the work of removal of water from an ordered array of phospholipid bilayers and used X-ray diffraction to observe the structural consequences of water removal. Short-range repulsive forces became evident from the fact that bilayers made from zwitterionic, electrically neutral, phospholipids repel each other in aqueous solution at separations up to 30 A. With decreasing separation the force grows exponentially with a characteristic decay length of 2.5-3.0 A. As bilayers approach contact, repulsive pressures can grow to the order of I000 atmospheres. These short-range forces have been attributed to solvation effects at surfaces, which give rise to a repulsive “hydration” force. Figure 2 is a linear plot of bilayer interaction energies in the vicinity of energy minima with and without the addition of charged lipids, The exponentially dropping hydration repulsion either is followed by a more gradual electrostatic repulsion followed by van der Waals attraction, or, for uncharged surfaces, it is balanced at shorter distance by the van der Waals attraction. Gruen et af. (1984) have developed a theoretical framework to explain the exponentially decaying hydration repulsion forces. According to their model, changes in the structure of water near an interface are associated with a nonzero value of an order parameter P, the dielectric polarization of water. It is represented as a vector with a zero value in bulk water and a maximal value in a state where water molecules are fully oriented in one direction (see Fig. 3). Although the decay of the polarizing perturbation reflects the properties of the aqueous medium, the value of P depends on properties of the membrane surface. Thus the importance of head group packing density becomes clear: zwitterionic compact surfaces, which have their charges near each other, will cause a weak polarizing perturbation. This is borne out by the data on differences in interaction energies between bilayers made of PC and PE (see Fig. 2). [For details see references in Parsegian and Rand (19831.1 The polar groups of bulky liquid-chain PC molecules pack poorly and must be surrounded with polarized water, whereas PE forms tightly packed bilayers. Figure 2 shows that the equilibrium distance for egg PE is about 10 A closer to the surface than that

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MEMBRANE FUSION

A

Separation

20

I

40

(A) 60

80

I

-.03 0

Separation 20

0

-.2

I

I

40

(A) 60 1

'

3

_ - - -_ _ _ - - - -

I

FIci. 2. Linear plots of bilayer interaction energies in the vicinity of energy minima. ( A ) Difference in depth and location of energy niinima with and without the presence of electrostatic interactions. ( B ) Difference in interaction energies of egg PC and egg PE as a function of distance. (From Parsegian and Rand. 1983. with permission.)

for egg PC, and that the energy minimum is about 10-fold as deep. Also consistent with this model is the reduced hydration repulsion of PC in the gel phase, which is tightly packed, and therefore expected to polarize less water, as compared with PC in the fluid phase (Parsegian and Rand, 1983). Since hydration poses a formidable barrier to fusion, it follows that fusion of lipid bilayers will be facilitated by treatments or agents which cause dehydration of the surface. Bilayer surfaces are dehydrated by addition of polyethylene glycol, formation of a dehydrated calcium-phos-

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Membrane

I

/ J

W\J\

FIG.3. Polarization of water near a surface. The arrows represent the dielectric polarization of water. The maximal value is fully oriented perpendicular to the surface. I t decays as a function of distance from the surface.

phatidylserine (Ca-PS) complex, or freezing and thawing (Hui et al., 1981; Strauss, 1984). As we shall see in Section VIII,D dehydration is not sufficient to induce membrane fusion; such treatments also cause destabilization of the bilayer. The dependence of vesicle-vesicle fusion on the phospholipid head group is also consistent with their relative hydration energies. Duzgunes et al. (1981a) found that inclusion of PE in PS vesicles greatly enhanced divalent cation-induced fusion whereas inclusion of PC was inhibitory. Sundler and Papahadjopoulos ( 198 1) compared divalent cation-induced fusion rates of vesicles with phospholipids containing different (negatively charged) head groups-. They found that PA was most fusogenic, followed by PS, and no fusion with PI, consistent with the bulkiness and hydration energies of the head groups (Loosley-Millman et al., 1982). Similar results where obtained with pH-dependent fusion induced by VSV G protein reconstituted in lipid vesicles (Eidelman et d.,1984). B. Membrane Electrostatics

From the discussion in the previous section it is clear that the depth of the energy minimum is considerably decreased, and the equilibrium distance is considerably increased, when the membrane surfaces are charged. [For a calculation of the electrostatic energy between two charged spheres (vesicles) see Gingell and Ginsberg (1978) or Ohki et ul. (19821.1 Therefore, it is reasonable to expect charge neutralization to be a necessary condition for aggregationlfusion. This has been borne out experimentally. The presence of fixed charges on a membrane surface gives rise to a surface potential, 9. Free cations from the medium accumulate in the interfacial region and screen the charges. The distribution of these cations can be described approximately by the classical Gouy-Chapman formulation for the diffuse double layer. A balance is struck between electrostatic attraction of the cations to the negative charges at the membrane surface

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and the entropic tendency of the cations to spread uniformly throughout the medium. At physiological ionic strength. the surface potential falls off nearly exponentially from the membrane surface with a characteristic distance of 9 A. The surface potential can be related to the surface charge density by an expression derived by Grahame (1947). Divalent cations are more effective at screening than are monovalent cations, but calculations indicate that screening is unlikely to be a major factor for charge neutralization by divalent cations, which bind more strongly to bilayers made of negatively charged lipids than to those made of zwitterionic lipids. The effects of this binding on )I can be calculated from a Stern equation, which is a combination of the Langmuir isotherm for surface adsorption of each ion, a Boltzmann expression for each ion, and the Grahame expression for surface charge density [see McLaughlin et (11. (1981) and Weinstein et nl. (1982) for more details]. Calculation of surface potentials requires data on binding constants of ions to phospholipid membranes. The binding of Ca’+ to PS membranes has been measured in a variety of ways including estimation of the effect of Ca” on monolayer and bilayer potentials, N M R relaxation, equilibrium dialysis, use ofCa’+sensitive electrodes, and 6 potential measurements. If the right corrections are made for the effect of screening, as predicted from diffuse double layer theory, and for the effect of monovalent cation binding (Na’), then the data in the literature are consistent with binding constant of 12 M ’ for the Ca-PS complex (McLaughlin et [ I / . , 1981). Using this value, McLaughlin ef a / . (1981) showed that the Stern equation is capable of accurately describing 5 potential data for PS vesicles and mixed PS/PC vesicles in 0. I M NaCl for a variety of concentrations of divalent cations. Thus, in theory, charge neutralization for mixed PS/PC vesicles can be determined for any given ionic condition. However, this does not work out experimentally. PS vesicles aggregate and fuse in about I niM Ca’ + and 100 mM Na’. According to McLaughlin r t rrl. (1981) the surface potential is about -40 mV under those conditions, and one would not expect those vesicles to aggregate and fuse. The discrepancy is due to the fact that it has been assumed that there is one affinity and a 1: I stoichiometry for the Ca-PS complex. However, under conditions in which membranes aggregate there is a dramatic discontinuity in the binding curve with a change to a 1.2 stoichiometry for the complex (Ekerdt and Papahadjopoulos, 1982). The discontinuity has been shown with pure PS. PS/PE ( I : 1 ), and PS/galactosylcerebroside (10: I ) . but not with PS/PC I: I . The higher affinity 1 : 2 complex is interpreted as the formation of a new type of cooperative binding site (possibly the anhydrous complex) between closely apposed bilayer membranes. X-Ray diffraction (Portis et (11.. 1979). N M R (Hauser et a/.. 19771, and freeze-fracture data (Papahadjopoulos r t

+

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al., 1975) also point to the existence of such a complex. The difference between PS/PC and PS/PE in their ability to form such a complex is consistent with their relative hydration energies (see Section VIILA). The equilibrium distance for PC-containing membranes is large enough to inhibit interbilayer molecular contacts, even after complete neutralization of the surface charge at higher concentrations of Ca”, which decreases JI sufficiently to allow formation of stable aggregates. Aggregation seems to be well described by the charge neutralization model, whereas for fusion a superthreshold amount of divalent cation binding is required (Duzgunes et al., 1981b). For instance, aggregation of pure PS vesicles can be induced by high concentrations (0.55 M ) of Na’ alone (Day et al., 1980). The equilibrium distance for this aggregation process is larger that for Ca2+-inducedaggregation. The process is very different from the Ca-PS complex formation between membranes, and there is no significant bilayer destabilization, which is apparently required for fusion, in this case (see Section VIIi,D). [For a more detailed review on cation binding, aggregation, and fusion, see Nir et al. (19831.1 C. Membrane Deformation

In the previous section I noted that stable contact between two surfaces will be achieved if G >> kT. With interaction energies at the energy minima of -0.01 erg/cm2 for PC and 0.1 erglcm’ for PE (Parsegian and Rand, 1983), a contact between two single apposing phospholipid head groups (area 70 A) will yield energies of 0.002 and 0.02 kT units, respectively (kT = 4 X erg). However, a contact between 100 phospholipid head groups will increase these energies 100-fold. Therefore, spherical vesicles need to be deformed to provide this area of contact. This deformation induces a tension on the membrane, and work has to be performed to deform against this countervailing tension. A contact angle 0 is produced that obeys the relationship (Parsegian and Rand, 1983). cos e

=

I

+ c/2T

(5)

where G is the interaction energy per unit area, and T the membrane tension. Rigid spheres have a very small area of contact, but flaccid vesicles can be deformed to form large contact areas before stresses build up (Evans and Parsegian, 1983). The contact area is in fact ITR’ sin’ 8, where R is the radius of the vesicle and 0 the contact angle (see Fig. 4). For a contact angle of 15” and R = 100 A the energy gained from contact (for PE) is 0.05 kT units; for a larger vesicle of 1000 A radius, this energy becomes 5 kT units. This simple calculation shows that for the same contact angle a small vesicle will form a smaller area of contact and therefore a less

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FIG.4. Two mutually deforming spherical vesicles of radius R . Tis the membrane tension and H (he contact angle.

stable contact. However, as mentioned earlier the behavior of SUVs is opposite to that of LUVs. In the fluid phase SUVs are rigid, in contrast to LUVs which are readily deformed in that phase. SUVs will more readily aggregate and fuse than large vesicles, not because of their ability to form large areas of contact but because of their susceptibility to destabilization (see Section VIILD). In considering the energetics for contact formation we must consider the energy of deformation of vesicles. According to Evans and Parsegian ( 1983) the strength of contact vis-a-vis thermal or mechanical perturbations is represented by -yA,

+

W , - AFI-R >> kT

(6)

where y is the free energy reduction per unit area of contact formation. A, the area of contact, W , the work of deformation, which includes deformation of the membrane and of interior contents, and AF,., i s the integrated long range attraction between the undeformed particles. The contact angle can be calculated by minimizing Eq. (6) with respect to 0. The deformation will give rise to stretching of the bilayer, because the round shape has the lowest surface area to volume ratio. If the volume does not change, a change in shape will lead to an increase in surface area

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and consequently the membrane will be under stress. Beyond a 3% increase in area, the bilayer will burst (Evans and Skalak, 1979). If the energy minimum is attained with a contact angle consistent with an area increase of less than 3%, a stable aggregate is formed. Such aggregates are probably formed with PS vesicles in high salt. On the other hand, if the energy minimum is beyond the 3% area increase a defect will be formed that might result in fusion or lysis. A simple calculation shows that a 3% area increase will be attained at a contact angle of about 50”. If the membrane is highly permeable to water (“leaky”), the new volume will readjust without tension on the surface and stable contacts will be formed without destabilization of the membrane. Consequently there will be no fusion. Since membrane leakiness is not well defined. the rate of water permeation must be considered relative to the rate of deformation. D. Membrane Destabilization

From the preceding discussions it is clear that charge neutralization and removal of water from the surface (dehydration) are necessary, but not sufficient, conditions for lipid bilayer fusion. Charge neutralization of certain lipid mixtures by certain mono- or divalent cations can give rise to aggregation but not fusion (see Section VII1,B). Dehydration by addition of sucrose, dextran, or glycerol does not cause membrane fusion (Boni er a / . , 1984). Dehydration by freeze-thawing can be protected against by substances such as glycerol, DMSO, ethylene glycol, and sucrose. These substances are cryprotective because they prevent formation of large ice crystals, which presumably cause the defects in membranes. The crucial event in lipid bilayer fusion seems to be membrane destabilization or the formation of defects in the lipid bilayer.” I have to be somewhat unclear about the nature of these defects, since their molecular arrangements are not well defined. However, it is reasonable to assume that in such defects the nonpolar fatty acyl chains will be transiently exposed to water. This can then give rise to hydrophobic attraction between two membranes which have such defects. Israelachvilli and Pashley (1982) have measured the attractive forces in aqueous solution between two mica surfaces that were rendered hydrophobic by adsorption of a monolayer of the cationic surfactant hexadecyltrimethylammonium. They found an exponentially decaying hydrophobic force in the same range of separation distance as the van der Waals force but with an energy of attraction about an order of magnitude higher. This hydrophobic force is not observed in ’An alternative mechanism, i.e.. formation of inverted micelle intermediates, will be discussed in Section IX.

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intact lipid bilayers. where it seems to be “neutralized” by the local structure of water molecules interacting with phospholipid head groups. According to Ohki ( 1984) the “hydrophobicity” of the membrane surface is related to the surface free energy measured in a phospholipid monolayer. The surface tension of a monolayer, consisting of negatively charged lipids, increases in the presence of divalent cations. The concentration of divalent cations required t o produce this effect corresponds to the threshold concentration required to induce vesicle fusion. Other factors, such a s the degree of vesicle curvature, temperature. and membrane expansion by osmotic swelling (see below), all exhibit a similar correlation between increased surface energy and tendency of the membranes to fuse. On the basis of these results Ohki proposed that the increased hydrophobicity of the membrane surface is responsible for membrane fusion. Hui pf rrl. (1981) have observed by freeze-fracture electron microscopy “point defects” in bilayers containing a mixture of egg PC and soybean PE, which were fused by freeze-thawing. They appeared as “rivets” between adjacent bilayers in cross-fractured areas. Although those structures are usually highly transient, they seem to have been preserved in the multilamellar samples containing highly unsaturated PC and PE. As mentioned in the previous section, bilayer defects can be formed in Ca”-induced lateral phase separation, which results in the formation of rigid crystalline domains of acidic phospholipids within a mixed lipid membrane (Papahadjopoulos, 1978). Polyethylene glycol (PEG) can induce such defects along domain boundaries by binding to the phospholipid bilayer and inducing local rigidification of the bilayer (Boni rt l l / . , 1984). In cells Honda et ( I / . (1981) found that certain components contained in a commercial grade PEG are fusogenic for cells and that recrystallized PEG is nonfusogenic. “’ However, recrystallized PEG is fusogenic for lipid vesicles (Boni ct nl., 1984). Presumably liposomes are destabilized by PEG alone, whereas cell membranes are more resistant to destabilization and need an additional perturbant. Osmotic swelling is also required for PEGinduced cell fusion (see below) but not for PEG-induced liposome fusion. As mentioned before, cis-unsaturated fatty acids at high concentrations can render membranes fusogenic by creating ”defects.” Lyso-PC, which has been shown to be a fusogen in a number of systems (Lucy, 197x1, probably also works according to that mechanism. The notion that destabilization is a crucial factor in bilayer fusion is “‘Keci-ystallized PEG did induce cell fusion in the assay of Smith r / t i / . (19x2). On the other hand. Wqjcieszyn 1’1 ( I / . (19x3) found a difference in the effects of nonfusogenic and fusogenic PEG on cellh. Presumably the polymer itself promotes dehydration and clow apposition of adjacent cell membranes. but fusion requires the destabilization provided hy the additives contained in commercial PEG.

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also consistent with the observation that small vesicles, which have a high degree of curvature, have an intrinsically greater capacity to fuse than larger vesicles (Liao and Prestegard, 1979; Wilschut et al., 1980; Nir et al., 1982; Boni et al., 1984; Ohki, 1984; Morris et al., 1984). The strain imposed on the strongly curved bilayer of an SUV (Mason and Huang, 1978) makes it highly susceptible to destabilization. I have noted in a preceding section that SUVs made of PC with saturated chains will spontaneously fuse below T, (the liquid phase transition temperature). SUVs can only partially freeze because of molecular packing restrictions, and consequently “defects” in the boundaries between solid and liquid domains will be created. Presence of bilayer defects are indicated by leakage of internal contents of liposomes during fusion. Morris et al. (1985) found that SUVs made of PS/PE rapidly lose their contents during Ca*’-induced fusion, consistent with their susceptibility to destabilization, whereas large vesicles with the same lipid composition are nonleaky during the first few cycles of fusion. However, the extent of leakage through the defects is dependent on a number of factors, such as how transient the defects are, how large they are, and how closely apposed the membranes are. Therefore, leakage might or might not be observed in a fusion event mediated by defects. Formation of gross defects in membranes by high-intensity electric field pulses has proved to be a very effective means to induce membrane fusion (Zimmermann and Vienken, 1982). Membrane contact between at least two cells is achieved by “dielectrophoresis” generated by an alternating nonuniform electric field, while reversible electric breakdown in the membrane contact zone is achieved by application of a field pulse of high intensity. This technique has promising biotechnological applications. In the previous section I noted that membrane deformation induced by apposition could lead to destabilization. However, membrane stretching induced by osmotic swelling is an even more powerful driving force for membrane fusion. The intracellular or intravesicular volume expansion gives rise to the membrane tension (stretch). Defects will be formed that might result in fusion when the membrane area increases to about 3% (see Section IX,C). In their studies of liposome-planar bilayer fusion, Akabas et al. (1984) have clearly shown that osmotic swelling of liposomes, which are tightly bound to the planar bilayer in a “prefusion” state, is an essential condition for fusion of vesicular and planar membranes. Osmotic swelling is also a necessary condition for PEG-induced fusion of cells. The PEG induces close apposition of the cell membranes (“prefusion” state), but fusion requires osmotic swelling brought about by removal of PEG or by dilution (Knutton, 1979a; Wojcieszyn et al., 1983).

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Experiments showing that Ca”- or ATP-evoked catecholamine release from isolated chromaffin granules can be inhibited by increasing the osmotic strength of the medium led to the suggestion that osmotic swelling might be an important step in granule-plasma membrane fusion during exocytosis (Edwards et u / . , 1974; Pollard r t d . , 1976). The fact that hyperosmotic solutions inhibit exocytosis in intact secretory cells (see references in Pollard et NI.. 1982, and in Akabas et NI., 1984) and in the cortical granule preparation (Zimmerberg, 1984) lends support to this idea. Moreover, in mast cells secretory vesicle swelling concomitant with ex1984). Although osocytosis has been visualized directly (Curran (’[ d., motic swelling seems to be required for fusion in these systems, it is still unclear what the mechanisms for generating the osmotic force are (Baker and Knight, 1984). E. Bilayer Fusion Mediated by Soluble Peptides or Proteins

1 have summarized in Table I1 a number of polymers, peptides, or proteins that have been reported to induce vesicle-vesicle fusion. This list is most probably incomplete, and 1 apologize if I have left out anyone’s favorite fusion protein. These soluble proteins are often regarded as models for natural fusion proteins. The list reveals two basic requirements for the ability of the protein to mediate fusion: cross-linking and destabilization of bilayers. In almost all cases shown in Table 11, there is also leakage of liposome contents, although not necessarily with the same time course or at the same protein concentration as required for fusion. With S U V s . protein-induced destabilization alone is in some instances sufficient to induce bilayer fusion. For example, uncharged PC vesicles are not likely to be cross-linked by agents such as alamethicin, melittin, or bovine serum albumin. As I noted before, S U V s in the solid phase are inherently unstable, and they will fuse by themselves due to interactions between hydrophobic edges (see Section VII1,D). Similarly, the proteins or peptides in Table I1 may create such edges in fluid phase SUVs, and thereby induce fusion. The interaction of (basic) proteins or peptides with acidic liposomes results in fusion by cross-linking as well as by destabilization. This has been illustrated by Eytan and Almary (1983), who compared the eficiency of various cations to induce fusion in PC/PE/CL vesicles. The efficiency was in the following order: Ca” < La” < polylysine < polymyxin B = melittin. Both polymyxin B and melittin are amphipathic molecules with a hydrophobic moiety and a hydrophilic one bearing 5 positive charges. Since it has been shown that at higher concentrations these molecules will lyse cells and disrupt membranes, it is reasonable to assume

TABLE 11 SOLUBLE PEP~IDES AND PROTEINS THATINDUCE BILAYER FUSION (Po1y)peptide Alamethicin Melittin Bovine serum albumin Albumin fragment Concanavalin A Tubulin Clathrin Myelin basic protein Polylysine, cytochrome c Polylysine, cytochrome c Polyhistidine Polymyxin B Melittin Polyamines Synexin

Liposome type"

suv suv suv

suv suv

suv suv

suv SUV SUV

suv suv

suv

LUV LUV

Liposome compositionh PC PC PC PC

Pc PC PC PCIP a PE PCIPEIPPCIPEIPPCIPEIP PCIPEIPPCIPEIPPE/P~

Reference Lau and Chan (1973) Morgan et a/.(1983) Schenkman er a/.(1981) Garcia et a/.(1984) van der Bosch and McConnell (1975) Kumar et a / . (1982) Blumenthal e? a / . (1983) Lampe and Nelsestuen (1982) Stollery and Vail (1977) Gad et a/.(1982) Wang and Huang (1984) Gad and Eytan (1983) Eytan and Almary (1983) Schuber et a/.(1983) Hong er a/.(1982)

SUV, Small unilamellar vesicle, LUV, large unilamellar vesicle. PC. Phosphatidylcholine; P - , acidic phospholipids (PA, PS, or CL): PA. phosphatidic acid; PS, phosphatidylserine; CL. cardiolipin; PE. phosphatidylethanolaine. "

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that the hydrophobic tail will insert into the bilayer and destabilize it while the charged moiety will serve as the cross-linker. Some proteins or peptides listed in Table I 1 will not destabilize the bilayer by themselves. Therefore, additional factors arc required. such as the use of solid phase S U V s (myelin basic protein and concanavalin A), passage through the phase transition (melittin-PC and concanavalin A), incubation at elevated temperature (alamcthicin). addition of lyso-PC (myelin basic protein), high amounts of PE in the liposome (polylysine, polyhistidine, polymyxin B, melittin, polyamines, synexin), lowering the pH (polyhistine, clathrin, bovine serum albumin), and addition of niillimolar divalent ions (tubulin). The majority of proteins shown in Table I1 use S U V s , which are easier to destabilize, as targets for fusion. Exceptions are synexin (Hong (/I., 1982) and polyamines (Schuber P / ( i / . , 1983). which mediate fusion of large vesicles. These molecules presumably enhance aggregation, while fusion is induced by interaction of divalent cations with the lipid bilayer, causing phase separation and destabilization, according to Papahadjopoulos ( 1978). However, in liposomes containing high mole fractions of PE, spermine and spermidine induced fusion in the absence of divalent cations. In this case the lipid head group provided the destabilization necessary for fusion. Fusion mediated by these soluble proteins helps sharpen our concepts concerning mechanisms of membrane fusion mediated by natural fusion proteins. For example. based on the three-dimensional structure of the ectodomain of the HA glycoprotein of influenza virus (see Section V ) , and on physicochemical studies of pH-dependent conformational changes of that domain (Skehel (/I., 1987), the following picture has emerged for pH-dependent fusion mediated by influenza virus (see White c/ (/I., 1983): The protein is a trimeric rod-shaped molecule 135 A in length, consisting of a stem and three globular. highly folded domains (stalks) at the top. The globular domains are composed of the HA1 chain, and, a s they contain the binding sites for sialic acid. they provide the cross-linking feature. The stem contains the hydrophobic HA2 N-terminal sequences implicated in fusion (fusion peptide). As the pH is lowered, the stalks open up and expose the fusion peptide which then interacts with the target membrane, destabilizes it, and thereby induces fusion. Although this seems to be an attractive scheme, one of its main problems is that the stalk reaches about 100 A out of the viral membrane, and consequently the fusion peptide has to move up all that way to bind to and destabilize the target membrane. Perhaps structural and physicochemical studies on viral fusion proteins reconstituted into lipid bilayers (see Section IX,D) might yield further insights into their mode of action.

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IX. MEMBRANE FUSION: FACT, HYPOTHESIS, OR THEORY? A. Introduction

The field of membrane fusion has been plagued by a great deal of confusion and controversy as to both what is the correct theory and what are the facts. Therefore it might be worthwhile to examine this problem according to the “Scientific Method.” The facts consist of what we observe, and the theories consist of what we suppose or invent. The suppositions are made sometimes to fill the gaps in our factual knowledge, sometimes to increase the understanding of facts, and sometimes for other reasons. For instance, the theory of evolution is not an account of a process which anyone has observed from beginning to end. It is a supposition, albeit not a complete supposition for it incorporates some observed facts. It has been invented to bring some sense, some intelligibility, into what otherwise would have been brute senseless facts. The “Scientific Method” is a logical sequence of steps followed in any scientific research activity of observing, forming hypotheses, experimenting, testing hypothesis, and controlling or predicting subsequent action on the basis of test results. In doing so we construct models which we can manipulate and which allow us to interpret and test hypotheses against reality. Through tests, analyses, and modifications we construct an acceptable model. The final structure and logic of the model are the basis for predictions that allow solution of the problem. In the literature on the philosophy of science there is some diversity in views as to what constitutes a “hypothesis” or “model” or “structure” on the one hand, and a “theory” on the other hand. I consider an “hypothesis” to be a result of observation and inference, followed by some deductive testing of the conclusions. A “theory,” on the other hand, is the conclusion of a thorough analysis of observations and inferences, examination of the deductive tests, invention, and analysis and testing of multiple hypotheses. Finally, it is a conclusion that has gained acceptance by the scientific community and withstood the test of time. B. Artificial Lipid Membranes

What makes lipid bilayers come close together or what holds them apart can at this point be fitted into the framework of a theory. The foundations of the theory are based on the chemistry and physics of surfaces and on the theory of colloid stability. Both areas have been studied intensively during the past century. Part of the problem can be described according to DLVO, and this has been supplemented in the past 17 years by con-

24 1

MEMBRANE FUSION

sideration of the short-range repulsive forces. The nature of these forces has been established by the highly accurate measurements and theoretical insights of the Bethesda-St. Catherine group (Parsegian, Rand, and company) and the Canberra school (Israelachvilli, Pashley, Marcjela, Grucn, and company). However. what happens after the membranes come together is still unclear. Figure 5 shows two possible intermediate structures in fusion of lipid bilayers. The inverted micelle has been proposed by a number of authors as a possible intermediate (Neher, 1974; Lau and Chan, 197.5; Pinto da Silva and Nogueira, 1977; Cullis and Hope, 1978; Rand, 1981). This inverted micelle is related to the H I , phase of phospholipids and is detected as a lipidic particle by freeze-cleavage (Verkleij and Ververgaert, 1978) and by "P-NMR (Cullis and de Kruijff, 1978). The inverted micelle is likely to exist as an intrvbilayer rather than as an inrvtrbilayer intermediate (see footnote 4). Cullis and Hope (1978) observed that certain agents (fusogens), which induce fusion between erythrocyte membranes, also induce the Hi, phase in a portion of the isolated erythrocyte membrane. Certain cone-shaped phospholipids such as PE and CL-Ca' ' favor the formation of the Hi, phase. Verklcij r ? 611. (1979) showed that C a L t induced fusion of vesicles composed of egg PC/CL ( I :I ) was associated with formation of lipidic particles. On the other hand, with the same lipid

Pentalarninar

It Defect A H u i et al., 1981)

Fused

FIG 5

Two model5 for intermediate\ in membrane fuqion. See Section 1X.B for diwuswm.

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ROBERT BLUMENTHAL

mixture Bearer et ul. (1982) observed no formation of lipidic particles during fusion events arrested by quick freezing in the absence of glycerol as a cryoprotectant. Siegel (1984) pointed out that it would be very hard to detect intermediate structures in vesicle-vesicle fusion even by quick freezing, since they have very short half-lives. Moreover, the lowering of the temperature required for the freeze-cleavage might perturb the intermediate structures, which are temperature sensitive. The other possible intermediate, shown in Fig. 5, is a “defect.” Formation of such “defects” can be induced by a variety of mechanisms, discussed in Section VII1,D. The nature of the defect might remain quite elusive since it is transient and its possible irregular structure is not amenable to techniques, such as X-ray diffraction, designed to resolve structures. They have been observed by Hui et 01. (1981) in mixtures of egg PC and soybean PE that which were fused by freeze-thawing. In the resulting multilayers, numerous lipidic particles were observed by freezefracture electron microscopy. They appeared as “rivets” between adjacent bilayers in cross-fractured areas. Since both the defect (Hui et al., 1981) and the inverted micelle (Cullis and de Kruijff. 1978) display an isotropic 3’P-NMRsignal superimposed on the broad bilayer signal, the NMR technique will not distinguish between the two structures. Moreover, the size of the defect should be about equal to that of the inverted micelle intermediate, which is about 12.5 nm in diameter (Siegel, 1984). It is therefore hard to decide on morphological grounds whether the “rivets” observed by Hui et al. (1981) are defects rather than inverted micelle intermediates. Since defects as well as inverted micelle intermediates have both concave and convex surfaces, similar cone-shaped lipids will favor their formation. Since the intermediate might be either a defect or an inverted micelle depending on the experimental conditions (lipids, temperature, the way fusion is induced), it is not necessary to postulate either one as part of a general mechanism of membrane fusion. Perhaps more insight will be gained into the nature of the intermediates or types of intermediates required through development of sophisticated biophysical techniques (NMR, fluorescence) to monitor local perturbations, which are a very small fraction of the overall global structure. In addition, theoretical insight into the nature, energetics, and probability of such intermediates might be gained from molecular dynamics modeling of the bilayer structure. C. Natural Membranes

Fusion of natural membranes is still in the realm of hypothesis. Many of the factors, such as charge neutralization and dehydration, that are

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required to make lipid bilayers come together and fuse do not pertain to natural membranes. The areas of contact between membranes are usually small in viral fusion and exocytosis. Moreover we are still unclear about the facts concerning biological membrane fusion, specifically whether it occurs in “bare lipid patches,” or via proteinaceous structures (see the discussion in Section 11). The studies of the early stages of exocytosis captured by rapid-freezing of Litnrrlirs amebocytes (Ornberg and Reese, 1981) make an argument for the latter proposition. Within seconds of stimulation, the plasmalemma buckles inward to form broad appositions with secretory granules lying near the cell surface. The distribution of intramembrane particles is not different from that over the rest of the plasmalemma. Numerous punctate pentalaminar contacts form between granular and plasma membranes. and an exocytotic opening begins at a minute pore at one of these contacts. The pore then progressively widens as the granule contents expand and diffuse into the space around the amebocyte. The capacitance patch clamp studies of mast cell degrdnuhtion support the notion of an exocytotic pore. Fernandez and Neher (1984) captured single exocytotic events using this technique and observed rapid increases and decreases (”tlickering”) of the capacitance steps, consistent with the opening and closing of an “exocytotic” pore. The hypothesis of a pore or junction structure in the fusion event is attractive since in general proteins rather than lipids provide regulation of biological events. However, in viral fusion it has been shown in a number of studies that presence of protein in the target menibrane was not required (White and Helenius, 1980: Maeda ct ( I / . , 1981; White ct u / . , 1982; Hsu et ( i / . , 1983; Haywood and Boyer, 1984). Although the viral membrane obviously contains the regulatory protein(s) required for t’usion. the possibility of forming a junctional complex between proteins in hotli membranes becomes less likely. Therefore we must leave open the qucstions of the reality of a junctional complex or of the universality of biological fusion mechanisms. D. Toward Reconstitution of Membrane Fusion A possible critique (as 1 noted in Section 11) of the use of liposomes to model membrane fusion is that artificial bilayer fusion might be completely different from natural membrane fusion. So why use model systems? A valuable lesson may be learned from the study of ionic channels. Extensive studies on model channels (such as gramicidin and EIM) in bilayers had been carried out long before single channel behavior had been observed in natural membranes. Many of the techniques and the concepts of the mechanism of ion permeation through channels (reviewed by Blumenthul

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ROBERT BLUMENTHAL

and Klausner, 1982), and the nature of opening and closing of voltagedependent channels (Ehrenstein et al., 1974), had been developed. Thus the foundations had been laid to study the problem of ionic channels in biological systems. Similarly, a conceptual framework for approaching the problem of membrane fusion is being developed with these model systems. Since proteins are one way or another involved in natural membrane fusion, a happy medium between artificial and natural fusion is proteinmediated fusion. Early attempts to model protein-mediated involved the effects of (fusion) irrelevant proteins (see Section VII1,E). However, the most promising avenue is reconstitution of proteins that have been destined by nature to fuse. Reconstitution has proven to be a very effective way to study the mode of action of transport proteins and of membrane receptors in a well-defined environment (Klausner ef al., 1984). A reconstitution of a fusion protein in a planar bilayer has been reported by Zimmerberg et ul. ( 1980b). A water-insoluble membrane-associated protein with a high affinity for Ca” was isolated from calf brain synaptosomes and inserted into a planar bilayer. Fusion of liposomes with this reconstituted bilayer could then be induced at 10 p M Ca”. The Ca”-binding protein has not been characterized structurally, and its role in promoting fusion in vivo is not known. On the other hand, viral spike proteins have been very well characterized both structurally and functionally (see Section V). X-Ray crystallography and studies of pH-dependent conformational changes have yielded some insights into their mode of action (see Section VII1,E). They have been reconstituted into lipid vesicles by detergent dialysis. Reconstituted Sendai virus envelopes have been used extensively to introduce macromolecules into animal cells and to implant membrane proteins (receptors, transport proteins) into plasma membranes of living cells (reviewed by Loyter and Volsky, 1982). Reconstitution requires solubilization of a membrane protein in an appropriate detergent in the presence of lipid followed by removal of detergent by dialysis or other means. Removal of detergent results in formation of closed vesicles. The nature of the detergent, the molar ratio of protein, lipid, and detergent, and the mode of detergent removal are all very important factors in successful reconstitution (Klausner et ul., 1984). Use of detergents with a low critical micelle concentration (cmc) such as Triton X-100 have not been as effective in reconstitution studies as detergents with a high cmc, such as deoxycholate and octylglucoside. Reconstitution studies with Sendai virus have predominantly been carried out with Triton X-100, since other detergents such as deoxycholate fail to solubilize the relevant spike glycoproteins. Loyter and co-workers (see Loyter and Volsky, 1982) have very successfully injected macromolecules into cells and implanted membrane proteins into plasma membranes, using Sendai envelopes reconstituted from Triton X-100. However,

245

MEMBRANE FUSION

those systems suffer from similar problems as encountered with liposomecell interaction in that it is not clear what components of cell-associated Sendai envelopes are due to fusion, endocytosis, stable adsorption, or exchange (see Section 111). On the other hand, the efficiency of the transfer process is much higher than that for pure liposomes." In order to study mechanisms reconstituted systems that are better defined physicochemically need to be developed. One such system has recently been developed by Eidelman et NI. (1984). Purified G protein of VSV was reconstituted in egg PC vesicles by detergent dialysis of octylglucoside. Conditions for obtaining a homogeneous population of fusion-competent reconstituted vesicles, as shown by electron microscopy and fluorescence energy transfer, include a high protein to lipid ratio and a slow removal of the detergent. The fusion activity is nonleaky and is dependent on pH, temperature, and presence of negatively charged lipid such as PA or PS in the target membrane. The system is still imperfect, since its pH dependence does not match the biological pHdependence and fusion with biological membranes is not as efficient as with liposomes as targets (0.Eidelman and R . Blumenthal. unpublished observations). However, with the improvement of reconstitution techniques and with appropriate target membranes, the possibility of functional reconstitution that matches the biological situation is very close.

ACKNO W L E D 6 M ENTS

I thank Drs. Sen-Wek Hui and Jacob Israelachvili for helpful dibcussions. and Drs. Ofer Eidelman, Stephen Moms, Richard Omberg. Adrian Parsegian, Harvey Pollard, David Siegel. Anne Walter. and Joshua Zimmerberg for their careful consideration of the mnnuscript and useful comments. The reference list was compiled with the aid of MEDLINE and completed in December. 1984.

REFERENCES Ahkong. Q. F.. Fisher, D.. Tampion, W., and Lucy. J . A. (1973). The fusionoferythrocytes by fatty acids. esters, retinol and a-tocopherol. Bio~hern.J . 136, 147-155. Akabas. M. H.. Cohen. F. S . . and Finkelstein. A. (1984). Separation of the osmotically driven fusion event from vesicle-planar membrane attachment in a model system for exocytosis. .I. Cell Biol. 98, 1063-1071. Allen. R. D., Travis. J. L., Hayden. J . H., Allen, N. S . . Breuer, A. C.. and Lewis, L. J. ( I98 I ). Cytoplasmic transport: Moving ultrastructural elements common to many cell types revealed by video-enhanced microxopy . Cold Spring Harbor S y l n p . Qrccrnt. B i d . 46, XS-87.

"Recently Vainstein el d.(1984) have reconstituted highly fusogenic Sendai virus envelopes by removal of Triton X-100by direct addition of SM-2 Biobeads. These virosomes resemble the intact Sendai virus in their composition. sire. and ability to induce fusion.

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Altstiel, L.. and Branton, D. (1983). Fusion of coated vesicles with lysosomes: Measurement with a fluorescence assay. Cell 32, 921-929. B ~~o.J ~ / I > ~ . s . Baker, P. F. (1972). Transport and metabolism of calcium ions in nerve. P ~ c J Mol. B i d . 24, 177-223. Baker, P. F., and Knight, D. E. (1981). Calcium control of exocytosis and endocytosis in bovine adrenal medulary cells. Philos. Trons. R . Soc. London Ser. E . 296, 83-103. Baker, P. F.. and Knight, D. E. (1984). Chemiosomotic hypotheses of exocytosis: A critique. B ~ ( J . s Rep. c ~ . 4, 285-298. Bearer. E. L.. Duzgunes, N., Friend, D. S., and Papahadjopoulos, D. (1982). Fusion of phospholipid vesicles arrested by quick-freezing. The question of lipidic particles as intermediates in membrane fusion. BkJChh. Biophys. Acfu 693, 93-98. Berridge, M. J . (1984). lnositol trisphosphate and diacylglycerol as second messengers. Biodrein. J . 220, 345-360. Blaustein. M. P.. Ratzlaff, R. W., and Schweitzer, E. S. (1980). Control of intracellular calcium in presynaptic nerve terminals. Fed. Proc.. F e d . Am. Soc. Exp. B i o / . 395, 2790-2795. Blinks, J . R., Wier. W. G., Hess. P., and Prendergast, F. G . (1982). Measurement of Ca2' concentration in living cells. Prog. Eiophys. Mol. B i d . 40, 1-1 14. Blumenthal. R.. and Klausner, R. D. (1982). The interaction of membrane proteins with black lipid membranes. C d Sir/:/: R e v , 8, 43-82. Blumenthal, R.. Weinstein, J. N.. Sharrow. S. 0.. and Henkart, P. (1977). Liposome-lymphocyte interactions: Saturable sites for transfer and intracellular release of liposome contents. Proc. Natl. Accrd. Sci. U . S . A . 74, 5603-5607. Blumenthal. R., Ralston, E., Dragsten, P., Leserman, L. D., and Weinstein, J . N . (1982). Lipid vesicle-cell interactions: Analysis of a model for transfer of contents from adsorbed vesicles to cells. Mernhr. Biochem. 4, 283-303. Blumenthal. R., Henkart, M., and Steer, C. J . (1983). Clathrin-induced pH dependent fusion of phosphatidylcholine vesicles. J . Bio/. Clrcm. 258, 340-3415. Boni. L. D., Hah, J. S.. Hui, S. W. Mukhejee. P., Ho, J. T.. and Jung, C. Y. (1984). Aggregation and fusion of unilamellar vesicles by polyethylene glycol. Biockim. Biophvs. Acttr 775, 409-418.

Braell, W. A., Schlossman, D. M.,Schmid. S. L., and Rothman, J. E. (1984). Dissociation of clathrin coats coupled lo the hydrolysis of ATP: Role of an uncoating ATPase. J. Cell Biol. 99, 734-141. Brooks, J. C., and Treml, S. ( 1983). Catecholamine secretion by chemically skinned cultured chromaffin cells. J . Ne~rnclicw.40, 468-473. Burgoyne. R . D. ( 1984). Mechanisms of secretion from adrenal chromaffin cells. BiocAi,,I. B i ( ~ p / i y .A~ . m 779, 201-216. Buxhaum. K.. Evans. E., and Brooks, D. (1982). Quantitation of surface afinities of red blood cells in dextran solutions and plasma. B ~ ~ J c h J n i . S /21, / y 3235. Chandler, D. E.. and Heuser. J. E. (1979). Membrane fusion during secretion. Cortical granule exocylosis in sea urchin eggs as studied by quick freezing and freeze-fracture. J . C d / Bin/. 83, 91-108. Cohen, F. S . . Zimnierberg, J.. and Finkelstein. A. (1980). Fusion of phospholipid vesicles with planar phospholipid bilayer membranes. 11. Incorporation of a vesicular membrane marker into the planar membrane. J . Gen. Physiol. 75, 252-269. Cohen. F. S . , Akabas. M. M., and Finkelstein, A. (1982). Osmotic swelling of phospholipid vesicles causes them to fuse with a planar phospholipid bilayer membrane. S&YIIY> 217, 458-460. Creutz, C. E. (1981). Cis-unsaturated fatty acids induce the fusion of chromafin granules aggregated by synexin. J. Cell Biol. 91, 247-2.56.

MEMBRANE FUSION

247

Creutz. C. E.. Paroles. C. J . . and Pollard, H. B. 1 197X). Identification and purification o f an adrenal medullary protein (synexin) that causes calcium-dependent aggregation of isolated chromaffin gritnules. J . Biol. C l r c ~ 253, . 2X5X-2866. Cullis, P. R . . and de Kruijff. B. (1979). Lipid polymorphism and functional roles o f lipid\ in biological membranes. Bioc4itti. B i t ~ p l r ~ Culli\. P. R.. and Hope, M. J . (197X). Effects c enihrane structure o f erythrocyte ghosts and the mechanism o1niemhr;ine fusion. Ntrrrrr~(Lorrhtr)271, 672674. Curran. M. J . . Brodwick. M. S.. and Edwards. C. (19x4). Direct visualization ofexocytosis in mast cells. Biophys. J . 45, 170:~ Day. E. P., Kwok, A . Y . W.. Hark, S . K.. Ho. J . T.. Vail, W. J . . Bentz, J . , and Nil-, S. ( 1980). Sodium induced reversible aggregation o f sonicated phosphatidylserine ve5icles. P ~ w .Ntrrl. Actrtl. S ( , i . U . S . A . 77, 4026. Ihzgunes. N.. Wilschut. J . , Praley. R.. and Papahadjopoulos. I). (1981a). Studies on the mechanism o f membrane fusion. Role o f heaclgroup composition in calcium- and magnesium-induced fusion o f mixed phospholipid vesicles. Bicdiiru. Biop/iy.r. Acvtr 642,

182-195. DuLgunes, N . . Nir. S.. Wilschut. J . , Bentz. J.. Newton. C.. Portis, A., and Papahadjopou10s. D. ( 1981b). Calcium- and magnesium-induced fusion o f mixed phosphatidylserine/phosphatidylcholine vesicles: Effect on ion binding. J . M r r t h . B i d . 59, I 15-

125. Edwards. W.. Phillips. J . H . . and Morris. S. 1 . (1974). Structural changes in chrornaffin granules induced by divalent cations. Bioc.lritii. B i o p h y s . Acrtr 356, 164-173. Ehrenstein. G.. Blumenthal, R . . Latorre. R . . m d Lecar. H . (1974). The kinetics o f thc opening and closing o f individual E I M channels in ;I lipid bilayer. J . C k r 7 . Phy.siol. 63, 707-72 I. Eidelman. 0.. Schlegel. R..Tralka, T. S.. and Blumenthal. R . (19x4). p H Dependent fusion induced hy vesicular stomatitis virus glycoprotein reconstituted into phospholipid ve\~ c l e s .J . Biol. Clrertr. 259, 46224618. Ekerdt. K.. and Papahadjopoulos. D.( 1982). Intermemhrme contact affects calcium binding to phospholipid vesicles. Pro(,. Ntrrl. Ac t i d . .S(,i. U . S . A . 79, 2273-2277. Evan\. E. A.. and Parsegian. V . A. (19x3). Energetics o f membrane deformation and adhesion in cell and vesicle aggregation. Arrn. N . Y . A ( , d . .S(,i. 416, 13-33. Evans. E. A , . and Skalak. R. ( 1979). Mechanics and thermodynamics o f biomembranes. C’KC Crir. Rcji.. Biorr7g. 3 and 4, IXI-41X. Evans. E.. Gershfeld, N.. Ginsberg. L.. and Parsegian, V . A . (1982). Caveats against the use o f thermodynamically unstable vesicles as models for biological systems. Biophvs. J . 37, 164a. Evered. I>., and Whelan. J . (1984). Cell fusion. (‘ihir Foirrrd. Sytrip. 103, 1-291. Eytan. G . D., and Almary, T. (19x3). Melittin induced fusion o f acidic liposomes. FEHS.

lxrr. 156. 29-32, Fernandez. J . M . . Nehrr. E . . and Gomperts. B . D. (19x4). Capacitance measurement\ reveal stepwise fusion events i n degranulating mazt cells. Ntrrrrrr (Loridorr) 312, 453455. FlorkiewicL. R. Z.. and Rose, J . K . (19x4). A c e l l line expres\ing vesicular stomatiti\ ~ 71 1-73. virus glycoprotein fuses as low p H . S c . i o r c ~ c225. Forgac, M.. Cantley. L.. Wiedenman, 8.. Altstiel. L., and Uranton. D. ( 1983). Clathnn-coated vesicles contain an ATP-dependent proton pump. I’rrw. Ntrrl. Actid. St,i. U . S . A . 80, 1300- I 303. Fowler. V . M.. and Pollard. H . B . (1982). Irr ~ i r r oreconstitution o f chroniaffin granule membranes. J . Ccll. Rioc/ic~rtr.18. 295-3 I I .

248

ROBERT BLUMENTHAL

Fung, B. K.. and Stryer, L. (1978). Surface density determination in membranes by fluorescence energy transfer. Biochemistry 17, 5241-5248. Gad, A. E.. and Eytan, G. D. (1983). Chlorophylls as probes for membrane fusion: Polymyxin B-induced fusion of liposomes. Biochim. Biophys. Acla 727, 170-176. Gad, A. E., Silver, B. L., and Eytan, G. D. (1982). Polycation-induced fusion of negatively charged vesicles. Biochim. Biophys. Acla 690, 124-132. Garcia, L. A., Araujo, P. S., and Chaimovich, H. (1984). Fusion of small unilmellar vesicles induced by a serum albumin fragment of molecular weight 9000. Biochirn. Biophys. Acfr1 16, 231-234. Gardner, J. D., and Jensen, R. T. (1981). Regulation of pancreatic exocrine secretion in vitro: the action of secretagogues. Philos. T r m s . R . Soc. London 296, 17-26. Gingell. D., and Ginsberg, L. (1978). Problems in the physical interpretation of membrane interaction and fusion. Cell. Siirj: Rev. 5, 791-833. Goldstein, J . L., Anderson, R. G., and Brown, M . S. (1979). Coated pits, coated vesicles, and receptor-mediated endocytosis. Nutiire (London) 279, 679-685. Grahame, D. C. (1947). The electrical double layer and the theory of electrocapillarity. Chem. Rev. 9, 447-463. Gregoriadis, G . (ed.) (1984). “Liposome Technology.” Vols. I . 2, and 3. CRC Press, Boca Raton, Florida. Gruen, D. W. R.. Marcelja, S.\ and Parsegian, V. A. (1984). Water structure near the membrane surface. In “Cell Surface Dynamics” (A. S. Perelson, C. DeLisi, and F. W. Wiegel, eds.). pp. 59-92. Dekker, New York. Hagins, W. A., and Yoshikama, S . (1978). lntracellular transmission of visual excitation in vertebrate retinal photoreceptors: Electrical effects of chelating agents introduced into rods by vesicle fusion. In “Vertebrate Photoreceptors” (P. Fatt and H. B. Barlow, eds.), p. 97. Academic Press, New York. Hand, A. R., and Oliver, C. (eds.) (1981). ”Basic Mechanisms of Cellular Secretion” (Methods in Cell Biology, Vol. 23). Academic Press, New York. Hauser, M., Finer, E. G., and Darke. A. (1977).Crystalline anhydrous Ca-phosphatidylserine bilayers. Biochem. Biophys. Res. Commun. 76, 267-274. Haynes. D. H., Lansman, J., Cahill, A. L., and Morris, S. J. (1979). Kinetics ofcation-induced aggregation of Torpedo electric organ synaptic vesicles. Biochim. Biopltys. Actci 557, 340-353. Haywood, A. M., and Boyer, B. P. (1984). Effect of lipid composition upon fusion of liposomes with Sendai virus membranes. Biochemistry 23, 4161-4166. Henkart, M. (1980). Identification and function of intracellular calcium stores in neurons. Fed. Proc., Fed. Am. Soc. Exp. Biol. 39, 2776-2777. Homma, M.. Shimizu, Y. K.,and Ishida. N. (1976). On the study of Sendai virus hemolysis. I. Complete Sendai virus lacking in hemolytic activity. Viro/ogy 71, 41-47. Honda, K.. Maeda, Y., Sasakawa, S.. Ohno, H., and Tsuchida, E. (1981). The components contained in polyethylene glycol of commercial grade (PEG-6,000) as cell fusogen. Biochem. Biophys. Res. Commun. 101, 165-171. Hong, K., Duzgunes, N., Ekerdt. R., and Papahadjopoulous, D. (1982). Synexin facilitates fusion of specific phospholipid membranes at divalent cation concentrations found intracellularly. Proc. N d . Acrid. Sci. U . S . A . 79, 46424644. Horwitz, A. F., Wright. A., Ludwig, P., and Cornell. R. (1978). Interrelated lipid alterations and their influence on the proliferation and fusion of cultured myogenic cells. J. Cell Biol. 77, 334-357. Hsu, M. C.. Scheid, A., and Choppin, P. W. (1983). Fusion of Sendai virus with liposomes: Dependence on the viral fusion protein (F) and the lipid composition of liposomes. ViroIOgv 145, 361-369.

MEMBRANE FUSION

249

Hui. S. W.. Stewart, T . P., Boni, L. T.. and Yeagle, P. L. (1981).Membrane fusion through point defects in bilayers. Science 212, 921-923. Hui. S . W., Stewart, T. P.. and Boni. L. T. (1983). The nature of lipidic particles and their roles in polymorphic transitions. Chetn. P l i y . Lipids 33, 113-126. Ishizaka, T.. Hirata, F., Ishizaka, K.. and Axelrod. J . (1980). Stimulation of phospholipid methylation, Ca” influx and histamine release by bridging of IgE receptors on rat mast cells. Proc. Nrrtl. Aciid. Sci. U.S.A. 77, 1903-1906. Israelachvili, J. N. (1982). Forces between surfaces in liquids. Adv. Coll. ln/erf Sci. 16, 31-47. Israelachvilli, J.. and Pashley. R. (1982). The hydrophobic interaction is long range decaying exponentially with distance. Nurure (London) 300, 341-342. Kaibuchi, K., Takai. Y., Sawamura, M., Hoshijima, H.. Fujikura, T., and Nishizuka, Y. (1983). Synergistic functions of protein phosphorylation and calcium mobilization in platelet activation. J . Biol. Cliern. 258, 6701-6704. Katz, B. (1966). “Nerve, Muscle and Synapse.” McGrdw-Hill. New York. Kendall. D. A,. and MacDonald. R . C. (1982). A fluorescence assay to monitor vesicle J . Biol. Clietn. 257, 3892-3895. Klausner, R. D., van Renswoude, J.. Blurnenthal, R.. and Rivnay, B. (1984). Reconstitution of membrane receptors In “Receptor Biochemistry and Methodology: Molecular Characterization of Membrane Receptors” ( J . C. Venter and L. C. Harrison, eds.). Vol. 3. pp. 209-239. Liss, New York. Knight. D. E.. and Baker, P. F. (1982). Calcium-dependence of catecholamine release from bovine adrenal medullary cells after exposure to intense electric fields. J . Mi.mhr. Biol. 68, 107-140. Knight. D. E.. and Baker. P. F. (1983). The phorbol ester TPA increases the affinity of exocytosis for calcium in “leaky” adrenal medullary cells. FEBS L e u . 22, 98-100. Knutton, S. (1979a). Studies of membrane fusion. 111. Fusion of erythrocytes with polyethylene glycol. J . Cell Sci. 36, 61-72. Knutton. S. (1979b). Studies of membrane fusion. IV. Fusion of HeLa cells with Sendai virus. J . Cell Sci. 36, 85-96. Knutton, S . ( 1 9 7 9 ~ ) Studies . of membrane fusion. V. Fusion of erythrocytes with non-haemolytic Sendai virus. J . Cdl Sci. 36, 85-96. Kohn, A. (1979). Early interactions of viruses with cellular membranes. Adv. Virus Res. 24, 223-276. Koslowski, M.. Rosen. F., Milholland, R., and Papahadjopoulos, D. (1978). Effect of lipid vesicle (liposome) encapsulation of methotrexate on its chemotherapeutic efficacy in solid rodent tumors. Cancer Reps. 38, 2848-2853. Kumar. N . . Blumenthal, R., Henkart, M . . Weinstein. J . N.. and Klausner. R. D. (1982). Aggregation and Ca’+-induced fusion of phosphatidylcholine vesicle-tubulin complexes. J . B i d . Chc.tt/. 257, 15137-15144. Larnpe. P. D., and Nelsestuen, G. L. (1982). Myelin basic protein-enhanced fusion of membranes. Biochirn. Biopliys. Ac/cr 693, 320-325. Lansman, J . . and Haynes. D. H . (197.5).Kinetics of a Ca”-triggered membrane aggregation ~ I 335-347. reaction of phospholipid membranes. BiocViitn. Biophys. A ~ , I394, Larrabee. A. L. ( 1979). Time-dependent changes in the size distribution of distearoylphosphatidylcholine vesicles. Biorhmis/ry 18, 332 I . L N I .A. L.. and Chan, S. I . (1975). Alaniethicin mediated fusion of lecithin vesicles. Proc. Ntrrl. A c i d . Sci. U . S . A . 72, 2170-2174. Lawson. D.. Raff, M. C., Comperts. B.. Fewtrell, C . . and Gilula, N . B. (1977). Molecular events during membrane fusion: A study of exocytosis in rat peritoneal mast cells. J . Cc.11 Biol. 72, 242-259.

250

ROBERT BLUMENTHAL

LeNeveu. D. M.. Rand, P. M., and Parsegian. V. A. (1976). Measurement offorces between lecithin bilayers. Ntifirre (London) 259, 601-603. Liao, M.-J., and Prestegard, J. H. (1979). Fusion of phosphatidic acid phosphatidylcholine mixed lipid vesicles. Biochirn. Biopliys. Acfci 550, 157-173. Llinas. R. R., and Heuser, J . E., (eds.) (1977). Depolarization-release coupling systems in neurons. Neiirosci. Res. Proguu~nBirll. 15. Loosley-Millman. M.. Rand. R. P., and Parsegian. V. A. (1982). Effects of monovalent ion binding and screening on measured electrostatic forces between charged phospholipid bilayers. Biophys. J . 40, 22 1-232. Loyter. A,. and Volsky, D. J . (1982). Reconstituted Sendai virus envelopes as carriers for the introduction of biological Material into animal cells. Cell Sirrf. Rev. 8, 215-266. Lucy, J. A. (1978). Mechanism of chemically induced fusion. Cell Srwf Rev. 5 , 267-304. McLaughlin. S., Mulrine, N . , Gresalfi. T., Viao, G . ,and McLaughlin, A. (1981). Adsorption of divalent cations to bilayer membranes containing phosphatidylserine. J . Gen. PIiysiol. 77, 445-473. Maeda, T., Kawasaki. K.. and Ohnishi. S.-I. (1981). Interaction of influenza virus hemagglutinin with target membrane lipids is a key step in virus-induced hemolysis and fusion at pH 5.2. Proc. N m l . A c ~ d S. c i . U.S.A. 78, 4133-4137. Markwell, M . A . K.,Svennerhold. L..andPaulson,J.C. (1981). Specificgangliosidesfunction as host-cell receptors for Sendai virus. Puoc. Narl. A c d . Sci. U.S.A. 78, 5406-5410. Margolis. L . B. (1984). Cell interaction with model membranes. Probing, modification and simulation of cell surface functions. Biochim. Biophvs. Acfci 779, 161-189. Mason, J . T.. and Huang, C. (1978). Hydrodynamic analysis of egg phosphatidylcholine vesicles. Ann. N . Y . Accrd. Sci. 308, 2 9 4 9 . Mayhew, E.. Papahadjopoulos. D., Rustum. Y . M., and Dave. C. (1976). Inhibitionof tumor cell growth in vi/ro and in viw by I-o-arabinofuranosylcytosineentrapped within phospholipid vesicles. Cancer. Res. 36, 4406-441 I . Mazurek, N . , Schindler, H., Schiirholz, T.. and Pecht, I . (1984). The cromolyn binding protein constitutes the Ca” channel of basophils opening upon immunological stimulus. Proc. N o / / . Actid. Sci. U . S . A . 81, 6841-6845. Morgan. C. G . . Williamson, H.. Fuller, S . , and Hudson, B. (1983). Melittin induces fusion of unilamellar phospholipid vesicles. Biochirn. Biophys. Actri 732, 668-674. Morris, S . J . . and Hughes, J . M. X . (1979). Synexin protein is non-selective in its ability to increase Ca’ +-dependentaggregation of biological and artificial membranes. Biocllern. Biopliys. Rc,.s. Comnrirn. 91, 345-350. Morris, S. J . , Chiu. V. C. K . , and Haynes, D. H. (1979). Divalent cation-induced aggregation of chromaffin granule membranes. Memhr. Biocliem. 2, 163-201. Morris, S . J . . Costello, M. J.. Robertson. J . D..Sudhof. T. C., Odenwald. W. F.. and Haynes. D. H. (1983). The chromaffin granule as a model for membrane fusion implications for exocylosis. J . Aicfon. Nerv. S y s f . 7, 19-33. Morris, S . J.. Gibson, C. C . . Smith. P. D., Greif, P. C., Stirk, C. W., Bradley, D., Haynes, D. M., and Blumenthal, R. (1985). Rapid kinetics of Ca”-induced fusion of phosphatidylserine-phosphatidylethanol vesicles. J . B i d . Clrew~.260, 4122-4127. Mukherjee, B., Orloff, S . . De Butler, J.. Triche, T., Lalley, P., and Schulman, J . D. (1978). Entrapment of metaphase chromosomes into phospholipid vesicles (lipo-chromosomes): Carrier potential in gene transfer. Proc.. Ncirl. Actid. S c i . U.S.A. 75, 1361-1365. Neher, E. (1974). Asymmetric membranes resulting from the fusion of two black lipid bilayers. B i o ~ h i / ? rBiophys. . AC/O373, 327-336. Neher. E., and Marly, A. (1982). Discrete changes of cell membrane capacitance observed under conditions of enhanced secretion in bovine adrenal chromaffin cells. Proc.. Nrrrl. A c u ~ S. C ~U.S.A. . 79, 67124716.

MEMBRANE FUSION

25 1

Nigg. E. A , , Cherry. R. J.. and Bachi. 7’. (1980). Influence of influenza and Sendai virus on the rotational mobility of hand 3 proteins in human erythrocyte membranes. Viroloqv

107. 552-556. Nir, S.. and HentL, J . (1978). On the forces between phospholipid bilayers. J . Coil. / t i / c r : / ; Sci. 65, 399-4 14. Nir. S.. Wilschut. J.. and Hentz. J. (1982).Thc rate o f fusion of phospholipid vesicles and the role of hilayer curvature. B i ( ~ h i / i iBiopliy.s. . Ac,rir 688. 27s-278. Hentz. J.. Wilschut. J.. and Duzgunes. N . (1983). Aggregation and fusion o f phoslipid vesicles. Pro,y. Sur Nihhizuka, Y . (19x4). Turnover

holipids and signal trancduction. Sc.icnt.c

225, 1365-1370. Ohki. S. ( 1984). Effects of divalent cation\. tcniperatui-e, osmotic pressure gradient. and vesicle curvature on phosphatidylserine vesicle fusion. J . Mcr77br..Biol. 77, 265-275. Ohki. S..Duzgunes. N., and I,eonards. K . (1982). Phospholipid vesicle aggregation: Effect of monovalent and divalent ions. Bioc~Iictni.\/ry21, 2 127-2 133. Orci, I>.. and Perrelet. A . (1978). Ultrastructurul aspects of exocytotic membrane fusion. Ccll S/fr:/: ROI’. 5 , 629-656. Ornherg. K. L.. and Keese. 7’. S . (1981). Beginning ofexocytosis captured by rapid-freeAng of /.;iirrr/rt,\ amebocytes. J . C d l Riol. 90, 40-54. Ostro. M., Giacomoni. D.. Lavelle. I>.. Piixton. W., and Dray. S.(1978). Evidence for the translation of rabbit globin niKNA after liposome mediated insertion into a human ccll line. Ntrrurc (Loritloti) 274, 92 1-923. I’agano. K. E . . and Weinstein. J. N. (1978). lntefiictions ofliposonies with mammalian cells. .4unrf. Rcl.. Biopl1y.c. Biocr7g. 7, 435-463. Palade, G. (1975). Intracellular aspects o f the process o f protein synthesis. S(,it,ric t’ 189,

347-358. Palade. Ci. E.. and Bruns. R. R. ( 1908). Structural modulations of plasmalemmal vesicles. J . CP// Biol. 37, 633-649. Papahad.jopoulos. D. ( 1978). Calcium-induced phase changes and fusion in natural and model membranes. Cell Sw/: Rci.. 5, 765-7YO. Papahadjopoulos. D.. Vail. W. J.. Jacohson. K . . and I’oste. G . (1975). Cochleatc lipid cylinders: Formation hy fusion of unilaniellar lipid vesicles. Bioc,lrirn. Biopliys. At.rri 394,

483-49 I. Papahadjopoulos, D., Wilson. T., and Taber. K. (19x0). Liposonies a s vehicles for cellular incorporation of biologically active maci-oiiiolcculr~./ r r Virro 16, 49-54. Parsegian. V . A,. and Rand. K. 1’. ( 19x3). Membrane interaction and deformation. Sur-fxc forces in biological systems. Arrti. N . Y . Ac t r d . S(.i. 416, 1-12. Pastan. 1. H., and Willingham. M. C. (19x1 ). Jour-ney to the center of the cell: Role of the 214, 504509. rcceptosome. Scierrc~c~ Pastcrnak. C. A , . and Micklem. K . J . (1974). The biochemistry of virus-induced cell fusion. Changes i n membrane integrity. Bioc~/ic,rii.J . 140, 405-41 I . Pearse. H . M . F . . and Bretcher. M . S. (19XI). Membr;ine recycling by coated vesicles. Arirrrt, Rcv. Rifdfc’rr7. 50. 85-101, Pinto da Silva. P.. and Hranton, D.(1970). Membrane splitting in freere etching. Covalently hound ferritin as a membrane marker. J . C’c// / j i o / . 45, 598-605. Pinto da Silva. P.. and Nogueira. M . L. (1977). Membfiine fusion during secretion. A hyzoospores pot hesis based on electron microscope observation of /’/iytop/rrhortr ptr/r~ii~ortr during encystment. J . Cell Biol. 73. Ihl-1x1. Pollard. H. H . . Zinder. 0 . .Hoffman. 0. G . . and Nikode-icvic. 0 . (1976). Regulation of the transmembrane potential of isolated chromafiin granules by ATP. ATP analogs and external pH, J . B i d . Clic~rii.251, 4544-4550.

252

ROBERT BLUMENTHAL

Pollard. H. B., Creutz, C. E., Fowler, V . , Scott, J . , and Pazoles, C. J . (1982). Calcium-dependent regulation of chromaftin granule movement, membrane contact, and fusion during exocytosis. Cold Spring Harbor Symp. Quant. Biol. 46, 819-834. Portis, A,, Newton, C., Pangborn, W., and Papahadjopoulos, D. (1979). Studies on the mechanism of membrane fusion: Evidence for an intermembrane Ca2+ phospholipid complex synergism with MgLt and inhibition by spectrin. Biochemistry 18, 780-790. Poste, G . , and Nicholson, G . C. (eds.) (1978). Membrane fusion Cell Surf. R e v . 5. Poste, G . , and Papahadjopoulos, D. (1976). Lipid vesicles as carriers for introducing materials into cultured cells. Influence of vesicle lipid composition on mechanisms of vesicle incorporation into cells. Proc. N u l l . Acad. Sci. U . S . A . 73, 1603-1607. Poste. G.. and Pasternak, C. A. (1978). Virus induced cell fusion. Cell Surf. R e v . 5 , 305367. Rand, R. (1981). Interacting phospholipid bilayers: Measured forces and induced structural changes. Annu. R e v . Biophys. Bioeng. 10, 277-3 14. Rink, T. J., Sanchez, A., and Hallan, T. J . (1983). Diacylglycerol and phorbol ester stimulate secretion without raising cytoplasmic free calcium in human platelets. Nature (London) 305, 317-319. Robertson, J. D. (1959). The ultrastructure of cell membranes and their derivatives. Biocltem. Soc. S y m p . 16, 3-43. Rosenberg, J., Duzgunes, N., and Kayalar, C. (1983). Comparison of two liposome fusion assays monitoring the intermixing of aqueous contents and of membrane components. Biochim. Biophys. Acta 735, 173-180. Satir. B. (1974). Ultrastructural aspects of membrane fusion. J. Supramol. Strucrr. 2, 529537. Schenkman, S., Araujo, P. S. , Dijkman, R., Quina, F. H., and Chaimovich, H. (1981). Effects of temperature and lipid composition on the serum albumin-induced aggregation and fusion of small unilamellar vesicles. Biochim. Biophys. Acia 649, 633-641. Schlegel, R.. and Wade, M. (1984). A synthetic peptide corresponding to the NH2 terminus of vesicular stomatiti's virus glycoprotein is a pH-independent hemolysin. J. B i d . Chem. 259, 4691-4694. Schlegel. R., Willingham, M. C.. and Pastan, I. H. (1982). Saturable binding. Saturable binding sites for vesicular stomatitis virus on the surface of Vero cells. J . Virol. 43, 871-875. Schlegel, R., Tralka, T. S., Willingham, M. C., and Pastan, I. (1983). Inhibition of VSV binding and infectivity by phosphatidylserine: Is phosphatidylserine a VSV-binding site'? Cell 32, 639-646. Schneider, A. A., Cline, H. T., Rosenheck, K., and Sonenberg, M. (1981). Stimulus secretion coupling in isolated adrenal chromaffin cells: Calcium channel activation and possible role of cytoskeletal elements. J. Neurochern. 37, 567-575. Schuber, F., Hong, K., Duzgunes, N.. and Papahadjopoulos. D. (1983). Polyamines as modulators of membrane fusion: Aggregation and fusion of liposomes. Biochemistry 22, 6 134-6 140. Schullery, S . E., Schmidt, C. F., Felsner. P., Tillack, T. W., and Thompqon. T. E. (1980). Fusion of dipalmitoylphosphatidylcholinevesicles. Biochemistry 19, 3919-3923. Siegel, D. P. (1984). Inverted micellar structures in bilayer membranes. Formation and half-lives. Biopkys. J. 45, 399-420. Silvius, J. R., and Gagne, J. (1984). Calcium effects on lipid lateral distribution and fusion in mixtures of synthetic phosphatidylserines, phosphatidylcholines and phosphatidylethanolamines. Biopphys. J . 45, 169a. Siraganian, R. P.. Urata, C., and McGivney, A. (1983). Arachidonic acid release during IgE

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253

and Ca”4onophore activation of rat basophilic leukemia cells. Monogr. Allergv 18, 120-123. Skehel, J., Bayley, P., Brown, E., Martin, S. . Waterfield. M., White, J.. Wilson. I . , and Wiley. D. (1982). Changes in conformation of influenza virus hemagglutinin at the pH optimum of virus-mediated membrane fusion. Pro<,.N d . Acod. Sci. U.S.A. 79, 968972. Smith, R . L.. and Oldfield, E. (1984). Dynamic structure of membranes bydeuferiom NMR. Scirnce 225, 280-288. Smith, C. L., Ahkong. Q. F., Fisher. D., and Lucy. J. A . (1982). Is purified poly(ethy1ene glycol) able to induce cell fusion’! Biocliim. B i ~ ~ p h yAcra . ~ . 682, 109-1 14. Steer, C. J., and Klausner, R. D. (1983). Clathrin-coated pits and coated vesicles: Functional and structural studies. Hepatolog.v 3, 437454. Stollery, J . G., and Vail, W. J. (1977). Interactions of divalent cations or basic proteins with phosphatidylethanolamine vesicles. BiocAirii. Biopliys. Acra 97, 372-390. Stone. D. K., Zie, X . S . . and Racker. E. (1983). An ATP-driven proton pump in clathrin-coated vesicles. J . Biol. C h m i . 258, 4059-4062. Stossel. T. P. (1981). Actin filaments and secretion: The macrophage model. Part 111. Translocation of secretory granules. / t i “Methods in Cell Biology,” (A. R. Hand and C . Oliver, eds.). Vol. 2 3 . pp. 215-230. Academic Press, New York. Strauss. G. (1984). Freezing and thawing of liposome suspensions. In “Liposome Technology” ( G . Gregoriades, ed.), Vol. I . pp. 197-219. CRC Press. Boca Raton, Florida. Struck. D. K.. Hoekstra. D., and Pagano, R . E. (1981). Use of resonance energy transfer to monitor membrane fusion. Biochernisrp 20, 4093-4099. Sudhof. Th. Chr., Walker, J. H., and Obroki, J. (1982).Calelectrin self-aggregates and promotes membrane aggregation in the presence of calcium. EMBO J . 1, 1167-1 170. Sundler. R., and Papahadjopoulos, D. (19811. Control of membrane fusion by phospholipid headgroups. I , Phosphatidate/phosphatidylinositolspecificity. Biochirn. Biopliys. Acra 649, 743-750. Szoka, F. C . . Jr.. Jacobson, K.. and Papahadjopoulos, D. (1979). The use of aqueous space markers to determine the mechanism of interaction between phospholipid vesicles and cells. Biochir?i. Biupkys. Actci 551, 3 5 - 3 0 ) , Szoka. F.. Jacobson. K . . Perzko, Z., and Papahadjopoulos. D. (1980). Fluorescence studies ~ . 600, on the mechanism of liposome-cell interactions iri ijirro. Bioc-him. B i ~ p l i V Acrri 1-18. Theoharides. T. C.. and Douglas. W. W. (1978). Secretion in mast cells induced by calcium entrapped within phospholipid vesicles. Scicnce 201, 1143-1 145. Toister. Z., and Loyter. A. (1973). The mechanism of cell fusion. 11. Formation of chicken erythrocyte polykaryons. J . B i d . Chern. 246, 422-432. Vaananen, P.. and Kaariainen. L. (1979). Haeniolysis by two alphaviruses: Semliki Forest virus and Sindbis virus. J . Gen. Virol. 43, 593-601. Vacquier. V. D. (1975). The isolation of intact cortical granules from sea urchin eggs. Calcium ions trigger granule discharge. Drv. Biol. 43, 62-74. Vainstein. A., Hersckovitz. M., Israel, S . , Rabin. S . . and Loyter, A. (1984). A new method for reconstitution of highly fusogeneic Sendai virus envelopes. Biockirn. Biophys. Acrtr 773, 181-188. Van der Bosch, J., and McConnell. H. M. (197.5).Fusion of dipalmitoylphosphatidylcholine vesicle membranes induced by concanavalin A. Pro(,. Nml. A ~ i dSci. . U.S.A. 72. 44094413. Varghese, J. N., Laver. W. G., and Colman. P. M. (1983). Structure of the influenza virus glycoprotein antigen neuraminidase at 2.9 A resolution. Niiritre (London) 303, 35-40,

254

ROBERT BLUMENTHAL

Verkleij. A. J., and Ververgaert, P. H. J . Th. (1978). Freeze-fracture morphology of biological membrane. Biochim. Biophys. Acta 515, 303-327. Verkleij. A. J . , Mombers, C., Gerritsen. W. J.. Leunissen-Bijvelt, L.. and Cullis, P. R. (1979). Fusion of phospholipid vesicles in association with the appearance of lipidic particles as visualized by freeze fracturing. Biochim. Biophys. Acta 555, 358-361. Verwey. E. I. W., and Overbeek, T. Th. G. (1948). “Theory of Stability of Lyophobic Colloids.’’ Elsevier, Amsterdam. Wang, C.-Y., and Huang, L. (1984). Polyhistidine mediates an acid-dependent fusion of negalively charged liposomes. Biochemistry 23, 4409-4416. Weinstein. J. N., Yoshikami, S . . Henkart, P., Blumenthal, R., and Hagins, W. A. (1977). Liposome-cell interaction: Transfer and intracellular release of a trapped fluorescent marker. Scicvce 175, 489-492. Weinstein, J . N., Blumenthal, R., Kempf, C., van Renswoude, J., and Klausner. R. D. (1982). Charge clusters and the orientation of membrane proteins. J . Membr. B i d . 66, 203-212. Whitaker, M. J . , and Baker, P. F. (1983). Calcium dependent exocytosis in an in vitro secretory granule plasma membrane preparation from sea urchin eggs and the effects of some inhibitors of cytoskeletal function. Proc. R . Soc. London ( B i d . ) 218, 397-413. White, J., and Helenius, A. (1980). pH-Dependent fusion between the Semliki Forest virus membrane and liposomes. Proc. Nail. Acud. Sci. U . S . A . 77, 3273-3277. White, J . , Kartenbeck, J., and Helenius, A. (1982). Membrane fusion activity of influenza virus. E M B O J . 1, 217-222. White, J . , Kielian, M.. and Helenius, A. (1983). Membrane fusion proteins of enveloped animal viruses. Q.Rev. Biophys. 16, 151-195. Wilschut, J . , and Papahadjopoulos, D. (1979). Ca’+ -induced fusion of aqueous contents. Nature (London) 281, 690-692. Wilschut, J., Duzgunes, N . , Fraley, R . , and Papahadjopoulos, D. (1980). Studies on the mechanism of membrane fusion: Kinetics of calcium ion induced fusion of phosphatidylserine vesicles followed by a new assay for mixing of aqueous vesicle contents. Biochemistry 19, 601 1-6021. Wilson, I . A.. Skehel, J . J . , and Wiley, D. C. (1981). Structure of the haemagglutinin membrane glycoprotein of influenza virus at 3 A resolution. Nature (London) 289, 366-373. Wqicieszyn, J. W., Schlegel, R. A., Lumley-Sapanski, K., and Jacobson, K . A. (1983). Studies on the mechanism of polyethylene glycol-mediated cell fusion using fluorescent membrane and cytoplasmic probes. J . Cell B i d . 96, 151-159. Wong. M., and Thompson, T. E. (1982). Aggregation of dipalmitoylphosphatidylcholine vesicles. Biocliemistr?, 21, 4133-4139. Young, J . D.-E.. Young, G . P. H., Cohn, Z. A , , and Lenard, J. (1983). lnteraction of enveloped viruses with planar bilayer membranes: Observations on Sendai, influenza, vesicular stomatitis, and Semliki Forest viruses. Virology 128, 186-194. Zimmerberg, J. (1984). Light-scattering microscopic measurement of the kinetics of exocytosis in isolated planar cortices from eggs of sea urchin Stron~l.vocenfrofusprtrprtrat~stRetardation of rate with hyperosmotic solutions. Biophys. J . 45, 88a. Zimmerberg, J., Cohen, F. C., and Finkelstein, A. (1980a). Fusion of phospholipid vesicles with planar bilayer membranes, 1. Discharge of vesicular contents across the planar membrane. J . Gen. Physiol. 75, 241-250. Zimmerberg, J., Cohen, F. C., and Finkelstein, A. (1980b). Micromolar Ca2’ stimulates fusion of lipid vesicles with planar bilayers containing a calcium-binding protein. Science 219, 906-908. Zimmermann, U., and Vieken, J. (1982). Electric field-induced cell-to-cell fusion. J . Mernbr. B i d . 67, 165-182.

C'UKRENT TOPICS IN MEMBRANES A N D TRANSPORT, V O L U M E ?Y

The Control of Membrane Traffic on the Endocytic Pathway IRA M E L L M A N , CHRISTINE HOWE, A N D ARI HELENIUS Depiirtnienr q f Cell Biology Yule University School of Medicine New H a r m , Connecticur 06510

1.

INTRODUCTION

One of the fundamental principles of eukaryotic cell biology holds that the transport of membrane between intracellular organelles occurs via vesicular carriers (Palade, 1975). While the vesicular traffic can be extremely rapid and involve extraordinary amounts of membrane, it is highly specific and regulated. During endocytosis, for example, vesicles involved in the uptake of extracellular macromolecules can result in the internalization of an area of membrane equivalent to 200% of a cell's total surface area every hour (Steinman ef d.,1976). However, in spite of their great number, these vesicles interact only with a restricted subset of organelles, typically resulting in the delivery of their internalized contents to lysosomes. In the biosynthetic pathway, vesicles form at the endoplasmic reticulum carrying newly synthesized membrane and secretory proteins to the Golgi complex from which other vesicles are then selectively targeted to the plasma membrane, lysosomes, storage granules, or other destinations. The mechanisms that control the intracellular traffic of membrane vesicles and their contents are only now beginning to be understood. While no firm answers are as yet at hand, some basic principles have already emerged, mostly from the study of the endocytic pathway. In this article, we shall summarize our current understanding of these principles as they apply to both endocytosis and exocytosis. with special emphasis on recent concepts pertaining to the function of endocytic organelles. N o attempt will be made to provide a comprehensive view of the field; for such information, the reader is referred to any of a number of more complete reviews (Steinman ef al., 1983; Kelly, 1985; Mellman, 1984; Farquhar, 255 Copyright Q 1987 hy Academic Preha. Inc All rights of reproduction in any Term reserved

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1985; Simons and Fuller, 1985; Pastan and Willingham, 1985; Mellman, 1986; Mellman et al., 1986). II. ENDOCYTOSIS AND MEMBRANE RECYCLING

A. Types of Endocytosis

In mammalian cells, two types of endocytosis are generally distinguished: phagocytosis and pinocytosis (Steinman et al., 1983). Phagocytosis, first described by Elie Metchnikoff in the late 1800s, is associated with the uptake of a large particle (>0.5 pm in diameter), such as a bacterium or another cell. While characteristic of “professional” phagocytic leukocytes (neutrophils, macrophages), this form d endocytosis occurs in most cells. Pinocytosis, on the other hand, involves the formation of much smaller vesicles (0.1-0.2 pm) resulting in the internalization of extracellular fluid and dissolved solutes (“fluid phase pinocytosis”) as well as any macromolecules bound to the plasma membrane at the site of vesicle formation (“adsorptive endocytosis”). The internalization of macromolecules bound to specific cell surface receptors is referred to as receptormediated endocytosis. In many cells, all pinocytosis is initiated by the invagination and subsequent vesiculation of clathrin-coated pits at the plasma membrane (Marsh and Helenius, 1980; Steinman et a l . , 1983). Apart from the sizes of the vesicles involved, the actual differences between pinocytosis and phagocytosis are not clear. While the formation of large ( > I km) phagocytic vesicles can be blocked by microfilamentdisrupting agents such as cytochalasin D, pinocytosis and the phagocytosis of smaller particles are generally unaffected (Steinman et al., 1983). Both types of endocytosis typically result in the delivery of internalized contents to hydrolase-rich lysosomes for digestion, and both are inhibited at low temperature (4°C). In addition, at least some types of phagocytosis may be associated with the formation of clathrin coats on the cytoplasmic surface of the nascent phagocytic vacuole (Aggeler and Werb, 1982; Aggeler et al., 1985; Montesano et al.. 1983). One important difference between the two processes, however, involves their regulation. Phagocytosis is a highly regulated event in mammalian cells: a phagocytic vacuole forms and is custom-fit in direct response to a particle binding to the appropriate receptors on the cell surface. In contrast, pinocytosis is generally thought to be a constitutive process, whereby pinocytic vesicles form continuously without the need for external signals. Pinocytosis can thus be likened to a continuously operating escalator, whereas phagocytosis behaves more in the manner of an elevator called into service only when needed.

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B. Membrane Recycling 1. QUANTITATIVE CONSIDERATIONS

As mentioned above, endocytosis can involve the internalization of enormous quantities of membrane. During one round of phagocytosis, a cell such as a macrophage can internalize as phagocytic vesicles an amount of plasma membrane equivalent to 50% of its initial surface area (Werb and Cohn, 1972; Steinman et ul., 1983; Petty et ul., 1981). During pinocytosis, typical mammalian cells in culture will continuously internalize 50-200% of their plasma membrane surface area every hour (Steinman et al., 1976). Nevertheless, a cell’s size and surface area are maintained at relatively constant values. Since the amounts of membrane involved are far in excess of a cell’s biosynthetic capacity, and since most cell surface glycoproteins are relatively long Lived ( t , , , 10-20 hr), it has long been presumed that internalized membrane components must be returned intact to the cell surface, i.e.. recycled (Steinman et ul., 1976; Silverstein et NI., 1977). Indeed, during the last several years a considerable body of direct and indirect evidence has been obtained for the existence of extensive and rapid membrane recycling during endocytosis (for review, see Steinman et ul., 1983; also, see below).

2. SORTING OF ENDOCYTIC VESICLEMEMBRANEA N D CONTENTS Since membrane recycling must occur via vesicular transport, a major conceptual problem is apparent: How can the continuous bidirectional vesicular traffic occur simultaneously with the continuous accumulation and/or degradation of internalized macromolecules in lysosomes? In fact, the intracellular accumulation of fluid-dissolved solutes is not quite unidirectional. Up to one-third may be lost from the cells back to the extracellular medium in the first few minutes following internalization. an expected consequence of vesicular recycling (Besterman et ul., 198 I ; Adams et ul., 1982; Swanson et u l . , 1985). Nevertheless, it is clear that there is a net accumulation of endocytosed solutes, indicating that at some stage after uptake there must be a sorting of endocytic vesicle membrane from contents: the membrane container returns to the cell surface while internalized solutes remain inside the cell. To a first approximation, this sorting event may simply reflect the geometry of endocytic and recycling vesicles: internalization in vesicles of low surface to volume ratio, recycling of membrane (and proportionately less fluid) in vesicles of relatively high surface to volume ratio. While the recycling vesicles have yet to be definitively identified, initial morphological evidence suggests that they may consist of small diameter (50 nm) vesicles

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or tubules (van Deurs and Nilausen, 1982; Geuze et uf., 1983; Beguinot et uf., 19841, structures having a much higher surface to volume ratio than the large, spherical coated vesicles (usually 0.2 Fm in diameter) which serve as the primary endocytic compartment. The available evidence strongly suggests that sorting of membrane from contents occurs soon after endocytosis, i.e., prior to the delivery of internalized tracers to lysosomes. As mentioned above, loss of endocytosed solutes due to membrane recycling is significant only during the first few minutes after uptake. Following delivery to lysosomes, there is relatively little reflux out to the extracellular medium (Besterman et af., 1981; Steinman et al., 1983). Similarly, in spite of extensive endocytosis and recycling, lysosomal enzymes-endogenous markers of lysosomal contents-are not lost at appreciable rates (Gonzalez-Noriega et al., 1980; Steinman et al., 1983). Recent work on receptor-mediated endocytosis has defined the prelysosomal site of sorting and recycling as a class of uncoated vesicles and tubules referred to as endosomes (Helenius et af., 1983), organelles whose properties will be considered in detail below. 111.

RECEPTOR-MEDIATED ENDOCYTOSIS

A. Receptor Recycling

The central importance of the endocytic pathway has been established by numerous findings over the last several years demonstrating that a variety of extracellular macromolecules bind to specific plasma membrane receptors and are internalized by endocytosis. Approximately 50 such examples have now been described, including receptor-mediated uptake systems for cell nutrients [e.g., cholesterol via low density lipoprotein (LDL), iron via transferrin, cobalamin via transcobalamin 111, polypeptide hormones [e.g., insulin, epidermal growth factor (EGF)], antibodies, lysosomal enzymes, modified plasma glycoproteins, bacterial toxins, and RNA and DNA viruses (e.g., influenza virus, poliovirus) (Steinman ef al., 1983). The existence of cell surface receptors greatly enhances the efficiency of endocytosis of specific macromolecules both by dramatically increasing their concentration at the plasma membrane and by targeting their uptake to selected cell types. The efficiency y is enhanced still further by the fact that receptors can be reutilized during ligand uptake. In many cases, the number of ligand molecules taken up per hour greatly exceeds the number of available receptors, even in the absence of new synthesis (i.e., cycloheximide-treated cells). Kinetic data show that an individual receptor can mediate the uptake of up to 10-20 ligand molecules every hour and

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therefore hundreds during the course of its lifetime (Steinman et al., 1983). Accordingly, it is clear that after internalization, receptors must recycle back to the plasma membrane. 6. Pathway of Receptor-Mediated Endocytosis

A general consensus has been reached concerning the basic features of the intracellular pathway taken by receptors and their ligands (for review, see Helenius et al.. 1983; Brown et al., 1983; Hopkins and Trowbridge, 1983). As diagramed in Fig. I , ligands first bind to their receptors, which results in the accumulation of the receptor-ligand complex at clathrincoated pits. Most investigators believe that the coated pits then pinch off to form coated vesicles (see Pastan and Willingham. 1983, for a different view), which rapidly lose their coats and fuse with endosomes, a heterogeneous population of vesicles and tubules in the peripheral and perinuclear cytoplasm. Due to their slightly acidic internal pH, endosomes

4

recycling vesicles pinocytic vesicles

endosomes

FIG. I ,

The basic pathway of receptormediated cndocytosis.

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facilitate the dissociation of many receptor-ligand complexes, thus allowing the return of free receptors to the cell surface and the transfer of the discharged ligand, now free in the endosome’s lumen, to lysosomes for degradation. This scheme (Fig, 1) clearly illustrates the role of endosomes as the central sorting station in the endocytic pathway, providing an intracellular site for receptor-ligand dissociation and recycling without requiring transit through the proteolytically perilous lysosomal environment. It also implies that acidic intraendosomal pH is crucial to maintaining the orderly traffic of membrane, receptors, and ligands in to and out from the cytosol. Having summarized the basic aspects of receptor-mediated endocytosis, in the next section we shall consider what is known (or suspected) concerning the mechanisms that control each step of the pathway. IV. MECHANISMS AND FUNCTIONS OF ENDOCYTIC ORGANELLES A. Coated Pits and Coated Vesicles 1. STRUCTURE AND ASSEMBLY

The composition and structure of the polyhedral lattice comprising the cytoplasmic coat of coated pits and coated vesicles has been well studied (for review, see Pearse and Bretscher, 1981; Harrison and Kirchhausen, 1983). The coat is composed primarily of the 180-kDa protein clathrin and its two associated light chains of 30-36 kDa. Clathrin normally exists as a trimer which takes the shape of a triskelion (Ungewickell and Branton, 1981; Kirschhausen and Harrison, 1981).Three light chains are associated with the clathrin heavy chains at the vertex of each triskelion. Several other nonclathrin proteins are associated with the coat including one or more 100-kDa species, tubulin, and a SO-kDa phosphoprotein (Pfeffer et al., 1983). The most important known property of clathrin coats is their capacity for self-assembly. Given the appropriate conditions of pH and ionic strength, clathrin triskelions will spontaneously form baskets, with or without an enclosed membrane vesicle. Conceivably, the favorable free energy change associated with the self-assembly process provides the driving force for membrane vesiculation, initiating the formation of an endocytic vesicle. Only clathrin and clathrin light chains are needed for assembly; thus, the role of the other associated coat proteins is unclear, although there is some suggestion that one or more 100-kDa species is required for attachment of the lattice to the membrane (Unanue ef al., 1981).

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In intact cells, the factors which regulate when and precisely where clathrin [or the 100-kDa protein(s)] attaches to the plasma membrane are unknown (see also Section IV,B). There is definite specificity to the attachment process, since not all organelles can serve as substrates for coated pit formation. Similarly, iri vitro, certain membranes (e.g., the erythrocyte membrane) are known not to permit clathrin binding and assembly (Unanue e t a / . , 1981). Large plaques of assembled clathrin in hexagonal arrays are often found at sites of focal adhesion to substrates or to other cells (Aggeler and Werb, 1982);whether adhesion signals clathrin assembly or simply prevents the subsequent vesiculation event is unclear. The only known inhibitor of coated pit formation (and the subsequent formation of coated vesicles) is depletion of cytoplasmic K’ concentrations (Larkin et al., 1983; Moya et a / . , 1985).The depolymerized clathrin presumably enters the preexisting pool of soluble clathrin (Goud et a / . , 1985) from which it can be recruited when K’ is returned to normal levels. Why K’ concentrations should exert this effect, however, is unknown.

2.

FORMATION AND UNCOAI’ING OF COATED VESICLES

The progressive transformation of a coated pit to a spherical coated vesicle is associated with the insertion of pentameric faces of triskelions into the planar hexagonal lattice (Heuser and Evans, 1980). While the addition of pentons serves to increase the curvature of the structure, pentamer formation may represent a cause or an effect. Whatever triggers the coated pit to coated vesicle conversion, it is clear that coated vesicles begin to lose their coats only moments after they form. Uncoating is presumably important not only to allow the coated vesicle to fuse with its intracellular target(s), but also to permit the reutilization of clathrin. Uncoating is unlikely to be spontaneous since the assembly process appears to be energetically favorable. Thus, it is of particular interest that the cytoplasms of most cells contain an ATP-dependent uncoating activity (Schmid et a / . , 1984; Braell rt al., 1984; Rothman and Schmid, 1986).The “uncoating ATPase” is a 70-kDa protein which binds both ATP and assembled clathrin heavy and light chains (Schmid et al., 1984). ATP hydrolysis is coupled to depolymerization of the clathrin basket.

B. Localization of Receptors at Coated Pits Since coated pits represent regions of the plasma membrane with a high probability of internalization, another manifestation of the efficiency of receptor-mediated endocytosis is the ability of receptors and receptorligand complexes to accumulate at coated pits. Some receptors, such as

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the LDL receptor, appear to be localized at coated pits even in the absence of bound ligand; others only after ligand attachment (Steinman et al., 1983). Thus, the signal for accumulation at coated pits cannot be ligand binding per se. Similarly, ligand binding cannot be the unique signal for the de n o w formation of a coated pit beneath the receptor. In this regard it is important to point out that coated pits are relatively catholic: a single coated pit can accumulate and mediate the internalization of several distinct receptors (Anderson et al., 1982). One possible mechanism for the localization of receptors at coated pits postulates a specific and conserved recognition event between the cytoplasmic domain of a receptor and some coat component (Pearse and Bretscher, 1981). While attractive, the nature of this recognition system is not apparent when one considers the structures of a variety of receptors. As illustrated in Fig. 2, “coated pit receptors” differ greatly in their overall

N

I I

COOH I

I

I 5

COOH

N

27aa

COOH

COOH

LDL

_-__*--

IU~W~LUI-

transferrin

EGF

_____ --recepror

_____I__

rscspror

COOH

N

N

C ASGP

Fc

...._ & ..

recepror

recepror

*..

insulin

VSV G

rscepror

prorein

-.

FIG.2.- Comparison of plasma membrane receptors that localize at coated pits. Each of the schematically diagrammed receptors mediate the uptake of their respective ligands via coated pits and coated vesicles. Only in the case of the LDL receptor is it clear that the receptor localizes preferentially at coated regions of the plasma membrane in the absence of bound ligand. N and COOH refer to amino- and carboxyl-termini, respectively. The sizes of the cytoplasmic domains (where known) are indicated in number of amino acid residues (aa). The overall molecular weights for each receptor are as follows: LDL receptor (160K). transferrin receptor (2 x YOK, disulfide-linked), EGF receptor (170K), asialoglycoprotein receptor (ASGP) (-54K). Fc receptor (55-60K). insulin receptor (2 x 90K. 2 x 125K. disulfide-linked tetramer), VSV G protein (5OK). For references, see Brown et rrl. (1983, review), Yamamoto et ul. (1984). Ullrich et ul. (1984, 1985). Chiacchia and Drickamer (19841, Green ef ril. (1985), Lewis e t a / . (1986), and Roth e t a / . (1986).

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sizes, in the sizes of their cytoplasmic tails, and in their orientations in the bilayer. Some receptors are phosphorylated or fatty acylated, some have multiple subunits joined by interchain disulfide bridges. A comparison of the receptors whose amino acid sequences are known reveals no unique primary structure homology (Brown et ul., 1983). Even several nonreceptor membrane proteins, such as vesicular stomatitis virus (VSV) G protein and leucine aminopeptidase, enter coated pits either constitutively or fol1986; Louvard, 1980; lowing the addition of specific antibody (Roth r t d.. H. Reggio and D. Louvard, personal communication). 1 . GENETIC EVIDENCE

Nevertheless, genetic evidence supports the importance of a membrane protein’s transmembrane and/or cytoplasmic domains in coated pit localization. The best example is provided by the human LDL receptor, a 160-kDa transmembrane glycoprotein with a single 50-amino acid cytoplasmic domain (Yamamoto et d . , 1984). Of the many mutations of the receptor known to disrupt receptor activity. one class blocks the efficient uptake of LDL by coding for a receptor protein that is unable to accumulate at coated pits (Goldstein et a / . , 1979). cDNAs corresponding to several mutant alleles have been cloned and sequenced, and in each case the defect was found to yield a receptor with an altered or truncated endodomain (Lehrman P I a l . , 1985; Russell et al., 1985). One mutation resulted from a single amino acid change (Tyr to Cys) at the eighteenth residue from the membrane’s cytoplasmic surface. In a second example, the transmembrane and cytoplasmic domains of VSV G protein were spliced onto the extracellular domain of the influenza virus spike glycoprotein, hemagglutinin ( H A ) , thus converting a protein which does not normally enter 1986). coated pits (the HA) to a chimeric protein which does (Roth et d., 2 . COMPOSITION OF COATED PITS

Given the great diversity in the structure of membrane proteins which enter coated pits, it is a priori unlikely that any recognition between a receptor and a coat component will turn out to be specific, in the sense of a ligand-receptor interaction. Instead, some aspect of a receptor’s oligomeric structure, which may be altered in response to ligand binding, rnay be a determining factor. For example, if receptors are simply aggregated by intermolecular interactions or by ligand binding, they rnay be relatively slow to diffuse through a coated region. Conceivably, the dense layer of coat proteins closely apposed to the cytoplasmic face of the membrane may physically impede the movement of a receptor’s endodomain. The slower rate of diffusion would increase the net residence time of a receptor at a coated pit, increasing the probability of internalization. Im-

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portantly, such a mechanism does not necessarily require the existence of specific interactions between receptors and coat components. Irrespective of the mechanism of receptor localization at coated pits, any interaction may not involve clathrin directly. Even a 50-amino acid long cytoplasmic tail is only 10-20 A long, and the clathrin basket may be as far as 150 A away from the membrane. Other coat proteins, such as the 100-kDa protein which is probably more closely apposed to the membrane than clathrin, may be more likely candidates. Clearly, coated pits generate distinct differentiated microdomains of the cell surface. At the very least, they differ biochemically from adjacent segments of the plasma membrane with respect to an increased concentration of receptors. Little quantitative or biochemical data concerning the concentration of other membrane proteins in coated pits have yet been obtained. However, one might expect that such nonreceptor proteins may be at least partially excluded from steric considerations alone: the accretion of receptor-ligand complexes may simply not leave much room. Exclusion is not likely to be 100% effective, i.e., coated pits probably are not perfect “sorters” which select out only receptors. All plasma membrane proteins exhibit finite turnover rates, and therefore are likely to be internalized and delivered to lysosomes (Steinman et al., 1983). In addition, mutant LDL receptors which fail to localize selectively at coated pits still mediate LDL uptake at a much slower rate (see Table 2 in Brown and Goldstein, 1976) that is comparable to the estimated rate at which plasma membrane surface area is internalized (see above): approximately one cell surface equivalent of bound LDL per hour. This would be the expected result if the mutant receptors were randomly distributed on the cell surface. Since coated pits occupy only I-2% of the surface area, and given the resolution of current morphological techniques, the detection of a randomly distributed components in coated pits would be difficult at best. C. Endosomes 1. DEFINITION OF ENDOSOMES

Following the uncoating of coated vesicles, the next station on the endocytic pathway is the endosome (for review, see Helenius et d.,1983). Although these structures were first described as intermediates on the pathway to lysosomes over 20 years ago (Straus, 1964), the extent of their importance in regulating membrane traffic during endocytosis has only recently been appreciated. Morphologically, endosomes are indistinct. They comprise a heterogeneous population of narrow tubules and vesicular elements distributed throughout the peripheral and perinuclear cytoplasm. Moreover, endosomes are not yet known to have any specific marker

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enzyme or other unique biochemical feature. The identification and definition of endosomes, therefore, is strictly functional: any uncoated vesicle which labels with endocytic tracers prior to their appearance in lysosomes. Endosomes in the cell periphery often seem to be labeled with internalized macromolecules prior to those in the perinuclear or lysosomal region (e.g., Wall er ul., 1980). Time-lapse photomicrographs also suggest that the peripheral endosomes actually migrate in toward the perinuclear region, perhaps along microtubule tracks, and perhaps fuse directly with lysosomes (e.g., Helenius et ul., 1983; Hirsch et ul., 1968; Herman and Albertini, 1984). The known functions of endosomes include the following (Helenius et ul., 1983): ( 1 ) dissociation of receptor-ligand complexes; (2) targeting of receptors back to the cell surface (or some other organelle); (3) targeting of dissociated ligands to lysosomes; (4) discharge of iron from internalized transferrin; ( 5 ) site of penetration of enveloped animal viruses (influenza virus, VSV, Semliki Forest virus) into the cytosol; and (6) site of penetration of many bacterial toxins, such as diptheria toxin, into the cytosol.

2. STRUCTURE OF ENDOSOMES By analyzing computer-generated three-dimensional reconstructions, we have found that “typical” endosomes consist of a large central vesicle (0.4-1 .O p,m in diameter) from which 2-10 narrow (
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vesicles and would thus be derived entirely from plasma membrane components. Alternatively, endosomes may contain unique constitutents which are not present (or are present in greatly reduced amounts) in the plasma membrane or other organelles. Since until recently endosomes have been difficult to isolate in purified form, little information exists concerning the biochemical composition of the endosomal membrane. Early studies which relied on the use of internalized lactoperoxidase to radioiodinate the major luminally disposed glycoproteins of the endosomal membrane suggested the qualitative presence of several major plasma membrane glycoproteins, but also indicated a number of potentially signifcant differences (Mellman et al., 1980; Mellman, 1982; Steinman et al., 1983). Also unknown is whether individual endosomes are stable, long-lived organelles or are transient structures which form, mature, and ultimately transform into lysosomes-either by direct fusion with preexisting lysosomes or by repeated interaction with Golgi-derived vesicles (“primary lysosomes”) containing newly synthesized lysosomal enzymes (Helenius e t a / . , 1983). While the time-lapse photomicroscopy referred to above would appear to support the concept of the transient endosome, one cannot yet exclude the possibility that long-lived endosornes transfer membrane and contents among themselves and finally to lysosomes via specialized transport or “shuttle” vesicles. Whichever view is correct, it is clear that the overall population we have referred to as “endosomes” must actually consist of several functionally and biochemically distinct subcompartments. At the very least, endosomes involved in the recycling of receptors can be distinguished from those which mediate transport to lysosomes. For example, endosomes containing transferrin, the iron-transport protein which recycles along with its receptor after internalization, are largely distinct from endosomes carrying lysosome-bound ligands such as EGF or az-macroglobulin (Yamashiro et al., 1984). The questions of endosome ontogeny and heterogeneity will remain until detailed information is available concerning the biochemical composition of the endosomal membrane.

4. ISOLATION

OF

ENDOSOMES

Progress on the structure, function, and biogenesis of endosomes has thus far been limited by the lack of convenient and effective isolation procedures. Separation of low density endosomes from much higher density lysosomes is readily accomplished by centrifugation of crude cell homogenates in Percoll density gradients (van Renswoude et d . ,1982; Merion and Sly, 1983; Galloway et ul., 1983; Marsh et al., 1983b). Thus,

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endocytic tracers which sediment with the low density fractions after brief periods of uptake (10-15 min) are progressively transferred to the high density lysosomal fractions beginning after 15-30 min of chase. Percoll gradients do not, however, resolve endosomes from most other smooth membranes (Golgi, endoplasmic reticulum, plasma membrane) which also sediment in the low density fractions (Merion and Sly, 1983). One novel approach to isolation has relied on selective modification of endosome density (Courtoy et a / . , 1984). In this approach, cells are first permitted to briefly internalize a ligand coupled to horseradish peroxidase (HRP). Cell homogenates are then exposed to peroxide and diaminobenzidine, and the intraendosomal HRP catalyzes the deposition of the diaminobenzidine reaction product; the net result is a sufficient increase in the endosomes' density to cause them to shift their sedimentation position in sucrose gradients. However, neither this nor more conventional approaches (relying on density gradient centrifugation and column chro1983) have thus far generated endosomal matography: Dickson et d., fractions of sufficient purity or yield to allow detailed biochemical and functional analysis. Recent work from our laboratory has demonstrated that free flow electrophoresis provides an effective and noninvasive method for the purification of endosomes from cultured cells and rat liver (Marsh er a / . , 1987). Crude cell homogenates or microsomal fractions are injected into a laminar curtain of buffer flowing perpendicular to an electric field. A combined endosome-lysosome fraction shifts toward the anode, well separated from a major peak containing most cell proteins and virtually all of the marker enzymes specific for other organelles. The shifted fraction is then resolved into highly purified endosomes and lysosomes by Percoll gradient centrifugation. Not only is the procedure rapid ( < 4 hr) and based only on the intrinsic properties of the organelles of interest, but 0.1-1 mg of endosomal protein can be generated per experiment. Most importantly, the isolated endosomes and lysosomes are still functionally intact with respect to ATP-dependent acidification (see below). Preliminary biochemical analysis has revealed that the endosomal membrane contains a distinct subset of intrinsic and peripheral membrane proteins relative to the membrane proteins of other organelles. Although endosomes may have components in common with some of the major iodinatable proteins of the plasma membrane, many endosomal proteins appear to be novel. They also appear to be largely different from those found in the lysosomal membrane. While these results must still be considered tentative, they support the concept that endosomes exist as independent organelles with their own unique composition.

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5 . ACIDIFICATION OF ENDOSOMES

The only known functional activity of endosomes which has been characterized at the biochemical level is acidification (for a detailed review, see Mellman et al., 1986). Although lysosomes have long been known to maintain a low internal pH (pH 4.7-4.8) (Ohkuma and Poole, 1978), the first evidence that endosomes were also acidic was provided by whole cell experiments which used internalized ligands coupled to the pH-sensitive dye fluorescein (van Renswoude et al., 1982; Tycko and Maxfield, 1982). Similar results were obtained using the low pH-triggered fusogenic activity of Semliki Forest virus (SFV) as an intracellular pH probe (Marsh et al., 1983a,b). While the internal pH of acidic endosomes seems to vary over a wide range (pH 5-6), there is evidence suggesting that the first endosomes encountered by endocytosed tracers are generally less acidic than “older” endosomes (Merion et al., 1983; Murphy et al., 1984). For example, SFV, like most enveloped animal viruses, penetrates into the cytosol by fusing its envelope with the endosomal membrane of the host cell. This fusion activity is mediated by pH-sensitive conformational changes in one or more viral spike glycoproteins (for review, see White et al., 1986). While wild-type SFV fuses with membranes at pH 6.1, a mutant virus has been isolated which has a lower fusion threshold, pH 5.2. When the kinetics of penetration of the two viruses are compared, the mutant is found to enter the cytosol much more slowly than wild type (although the relative rates of endocytosis are similar and the fusion event can still be localized to prelysosomal vacuoles). Thus, it takes longer for incoming virus particles to reach endosomes of pH 5.2 than it does to reach endosomes of p H 6. I (Kielian et al., 1986). The only known functions of acidification on the endocytic pathway are ( I ) the facilitation of receptor-ligand dissociation in endosomes and (2) the provision of an optimal environment for the activity of acid hydrolases in lysosomes. Acidic intravesicular p H is not an absolute requirement for the delivery of internalized macromolecules from endosomes to lysosomes (Ukkonen et al., 1986). However, treatment of cells with agents such as ammonium chloride, chloroquine, or monensin, all of which elevate the pH of endosomes and lysosomes, can cause gross alterations in the distribution and fate of internalized membrane (see Mellman et al., 1986, for review). a. The Endosomal Proton Pump. Experiments using isolated organelles have demonstrated that endosomes, like lysosomes, contain an ATP-driven proton pump (Galloway et al., 1983; Merion et af., 1983). The ATPase is similar with respect t o its ion requirements and inhibitor specificity t o proton pumps found in other organelles of the vacuolar system, namely, secretion granules, Golgi membranes, lysosomes, and coated vesicles

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(Mellman et a / . , 1986). These "vacuolar ATPases" can be distinguished from other classes of ion transport ATPases by their insensitivity to vanadate (a potent inhibitor of the Na+,K'-ATPase and related enzymes) and to agents such as oligomycin, azide, and efrapeptin (inhibitors of the mitochondrial-type F,F,, ATPases). Only the relatively nonspecific alklylating agents N-ethylmaleimide and NBD-CI- have been found to be effective inhibitors of the endosomal proton pump and other vacuolar ATPases. Initial studies of endosome acidification made use of crude fractions prepared by Percoll gradient centrifugation (Galloway et ul., 1983) (see above). Since these fractions were heavily contaminated by other organelles, it was necessary to selectively label the endosomes with fluoresceinconjugated endocytic tracers prior to isolation. More detailed study of the bioenergetics of the acidification process has now been possible with the availability of highly purified endosomes prepared by free flow electrophoresis (R. Fuchs and I. Mellman, manuscript in preparation). The basic mechanism of rat liver endosome acidification is summarized in Fig. 3. Endosome acidification is electrogenic: the proton pump transports protons from the cytosol into the endosome lumen, lowering the internal pH but also creating an internal-positive membrane potential. Dissipation of this electrical gradient is needed to allow continued proton transport; thus, the endosome permits the passive influx of cytoplasmic anions (e.g. CI-) and/or the efflux of internal alkali cations (Na' or K') to retain charge balance. In the absence of external CI the rate and extent of ATPdependent acidification is decreased. One additional permeability characteristic of the endosomal membrane is its significant conductance to protons. Following the removal of ATP, the generated pH gradient dissipates at a rate just slower than the rate at which it was formed. Thus, there is presumably a continuous bidirectional

AiP

FK;. 3. Mechanism of endosome acidificalion.

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flux of protons across the endosomal membrane, indicating that pH regulation probably occurs through regulation of the rates of proton influx and efflux as opposed to the absolute number of protons translocated. This may be accomplished by controlling the number or activity of proton pumps or by altering the activity of the anion, cation, or proton conductances. While we have not yet documented distinct ion permeabilities in different endosome subpopulations, it is already clear that endosomes differ from lysosomes by possessing a higher intrinsic permeability to protons and K’ and a lower permeability to CI- (R. Fuchs and I. Mellman, manuscript in preparation). An important unknown concerns the origin and biogenesis of the endosome proton ATPase. The observation that bovine brain coated vesicle fractions contain a vacuolar-type proton pump (for review, see Mellman et af., 1986) suggests the possibility that the endosome ATPase may be derived from the plasma membrane by endocytosis. The pump could then conceivably be transferred to and collect in lysosomes. However, there is no evidence to support this view. It is not at all clear that endocytic coated vesicles contain an ATPase. Coated vesicles isolated from brain or liver represent a mixed population consisting of both endocytic and Golgi-derived coated vesicles. Therefore, one cannot know whether the acidification activity observed in vitro was due to one or to both populations. Electron microscopic evidence, using pH-sensitive cytochemical stains, has yet to demonstrate that peripheral (i.e., endocytic) coated vesicles are acidic; in contrast, coated vesicles in the Golgi region can often be shown to accumulate these dyes (Anderson ef al., 1984; Anderson and Pathak, 1985; Schwartz et al., 1982). If the endosomal ATPase is not derived from the cell surface, it is conceivable that the pump represents one of the “unique” components of the endosomal membrane referred to above. Progress in identifying the endosomal ATPase has been limited by the difficulty in preparing sufficient quantities of endosomes to facilitate purification and also by the difficulty in isolating the ATPase even from more readily accessible source membranes (e.g., coated vesicles, secretion granules) (Mellman et a / . , 1986). As the ATPases of these other organelles become better characterized, the chances of identifying the endosomal proton pump using the highly purified free flow fractions will improve. h. Genetics of Endosorne Acidification. As mentioned above, many viruses and bacterial toxins require acidic endosomal pH to gain access to and subsequently kill cells. Thus, mutant cell lines selected for resistance to viruses and toxins might exhibit a defect in endosome acidification. Several such cell lines have been generated, and several have been shown to express a decreased capacity for ATP-dependent acidification in vitro

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(Robbins ef a / . , 1983, 1984; Merion el ( I / . . 1983; Klausner et a / . , 1984; Draper rt a / . , 1984). Among the best characterized mutants is the series of Chinese hamster ovary (CHO) cell lines isolated by Robbins and colleagues. These cells were found to be relatively resistant to both toxins and viruses and to exhibit pleiotropic defects in receptor-mediated endocytosis (e.g., inability to discharge iron from transfemn; ineficient uptake of lysosomal enzymes via the mannose 6-phosphate receptor) (Robbins ei d., 1983; Klausner et a / ., 1984). In vitro, isolated endosomes were defective in ATP-dependent acidification although lysosomes were found to be normal (Robbins et ( I / . , 1984). The mutant acidification phenotype was observed in endosomes labeled with fluorescein coupled to dextran (as a marker of fluid endocytosis) as well as to transferrin (a marker of receptor-mediated endocytosis) (R. Fuchs and I. Mellman. manuscript submitted). Genetic complementation analysis revealed that the mutations fell into two complementation groups, designated end-l and end-2, suggesting that at least two genes are involved in endosome acidification (Robbins ef d.,1984). Mutants of both classes were phenotypically similar, and temperaturesensitive as well as nonlethal alleles could be isolated for each complementation group (R. Fuchs and Mellman, manuscript submitted). One particularly interesting aspect of the end-1 and end-2 mutations is that in addition to affecting endosome acidification. they cause aberrant Golgi function (Robbins et a / . , 1984). Cells of both classes were unable to complete terminal glycosylation (sialic acid or galactose addition) of Sindbis virus spike glycoproteins E l and E2 or of a variety of endogenous secretory proteins (e.g., fibronectin). Since endosomal and Golgi membranes contain similar ATPases and acidification mechanisms (Galloway et d.,1983; Glickman et a / . . 19831, these findings raise the possibility that the endocytic and biosynthetic pathways share a common gene product related to ion transport which is required for the normal function of each. Confirmation of this possibility, however, must await identification of the biochemical defects in the mutant cells. 6. MOLECULARSORTING

IN

ENDOSOMES

By triggering the dissociation of ligand-receptor complexes, acidic pH in endosomes is clearly responsible for mediating the critical initial event in endosomal sorting. The mechanisms underlying the subsequent events, such as the selective segregation of internalized receptors into recycling vesicles and the targeting of dissociated ligands to lysosomes, are less well understood. it is possible that the geometry of endosomes accounts for all of the observed specificity: recycling in vesicles derived from the

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formation of high surface to volume tubules, transport to lysosomes in the low surface to volume vacuole region (see above). However, unless the fraction of endosomal membrane present as tubules has been greatly underestimated and/or the tubules have a relatively rapid rate of turnover (both are distinct possibilities), the efficiency of receptor recycling may also require a mechanism which selectively localizes receptors in the nascent recycling compartment. If a typical receptor with a tllz of 10 hr can mediate five rounds of internalization every hour, it would have only a I% chance of degradation at each round of endocytosis. Although the probability that an internalized receptor will be recycled may be as high as 99%, only 70-75% of the endosomal membrane is present as putative recycling tubules at any one time (Marsh et al., 1986). This percentage may be too low to account for the efficiency of recycling of receptors randomly distributed in the endosomal membrane. Whether the “solution” to receptor recycling is strictly Euclidean or depends on some biochemical mechanism which restricts the distribution of receptors in the endosomal membrane represents a major unknown. a . Receptor Transport to Lysosomes. Another physiologically important aspect of endosomal sorting is illustrated by the fact that a number of receptors fail to recycle, or recycle only inefficiently, during ligand uptake. Among this class are receptors for various polypeptide hormones (EGF, insulin) (Kasuga et al., 1981; Stoscheck and Carpenter, 1984; Beguinot et al., 1984) and those for IgG-containing antibody-antigen complexes (Mellman et a / . , 1983; Mellman and Plutner, 1984; Ukkonen et al., 1986). In several of these cases, it has been shown that ligand binding results in the delivery of receptor and ligand to lysosomes where both are degraded. Consequently, there is a selective loss of these receptors from the plasma membrane and the cell becomes insensitive to subsequent additions of ligand, a situation classically referred to as “down regulation.” Control of receptor expression in this way provides a cell with a rapid mechanism to regulate either its physiological response to the environment or its ligand-elicited effector functions. Receptors that do not recycle appear to be internalized via the same basic pathway as do recycling receptors. Indeed, EGF has been shown to enter fibroblasts in the same coated vesicles as particles of LDL, a receptor which efficiently recycles during ligand uptake (Carpentier et af., 1982). Ligands bound to down-regulating receptors are also delivered to typical endosomes prior to appearance in lysosomes (Haigler et al., 1979; Merion and Sly, 1983; Beguinot et al., 1984; Ukkonen et al., 1986). Thus, endosomes must be capable of distinguishing among receptors in order to target some back to the plasma membrane and others to lysosomes. Although it seems likely, additional information will be required to confirm that this sorting event can occur in a single endosomal vacuole.

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h. Possible Role qf Oligomcric Strricture. The molecular mechanisms responsible for directing some receptors to lysosomes are still obscure. An examination of the structural features of this class of receptors (cf. the EGF, insulin, and Fc receptors in Fig. 2 ) reveals no obvious distinguishing characteristic such as amino acid sequence, disposition in the membrane, phosphorylation, etc. Our work on the mouse macrophage Fc receptor, however, has begun to lend some insight into the process. The receptor is a 55-kDa transmembrane glycoprotein, consisting of four asparagine-linked oligosaccharides and a large (8- to 12-kDa) cytoplasmic domain (Green el al., 1985). We have recently cloned and sequenced a full-length cDNA for the receptor, and its deduced amino acid sequence reveals that it bears significant homology to immunoglobulins and also appears to exist as a single monomeric chain (Lewis et d . , 1986). When the receptor is bound to a monovalent ligand, a Fab fragment of a high affinity antireceptor monoclonal antibody (directed to the receptor's ligand binding site), we have found that the receptor is internalized, delivered to endosomes, and then rapidly recycled back to the plasma membrane (Mellman et al., 1984). In contrast, when a polyvalent ligand is usedeither a multivalent 1gG-antigen complex or a preparation of Fab fragment aggregated by adsorption to colloidal gold-the receptor is delivered to lysosomes and degraded (Mellman and Plutner, 1984; Ukkonen et ul., 1986). Since neither ligand is discharged from the receptor by acidic endosomal pH, these findings suggest that ligand valency is an important determinant of whether internalized Fc receptors recycle or are transported from endosomes to lysosomes. Conceivably, adjacent receptors are crosslinked by multivalent ligands, a situation which either prevents their recycling or signals their transport to lysosomes (Mellman, 1984). It is already clear that ligand. and perhaps the oligomeric state of a receptor, can influence its fate after endocytosis. Several receptors, many of which normally recycle, can be artificially directed to lysosomes when allowed to bind nondissociating multivalent ligands such as antireceptor antibody or colloidal gold-conjugated monovalent ligand. These include receptors for LDL (Anderson et id,, 1982), lysosomal enzymes (lee.,mannose 6-phosphate) (von Figura et ul., 1984), transferrin (Neutra e f a / . , 1985; Hopkins and Trowbridge, 1983; Ekblom et d.,1983; Weismann e t al., 1986), EGF (Schreiber et al., 1983), and ricin (van Deurs r t al., 1986). Whether alterations in the state of oligomeric assembly of a receptor are of general importance in physiological examples of down regulation is not clear. Certainly, it seems likely in the case of the Fc receptor, whose physiological "down-regulating" ligand is multimeric. However, in the case of the EGF receptor, for example, where the ligand triggering lysosomal transport presumably binds as a monomer, one would have to postulate that binding alters the conformation of the receptor in a way

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which increases its propensity for intermolecular interaction. In some cases, ligand-induced receptor phosphorylation may be related to these events. c . Role of Multivesicular Bodies. How might endosomes select out specific receptors for transport to lysosomes? Endosomal pH itself does not appear to be involved since agents which elevate intravesicular pH (e.g., monensin, ammonium chloride, chloroquine) do not block the normal intracellular transport of Fc receptor-bound ligands (Ukkonen et al., 1986). Similar results have been obtained for chorionic gonadotropin (Ascoli, 1984), another ligand which does not dissociate from its receptor at acidic pH. Thus, the ability of acidotropic agents to block lysosomal transport of pH-dissociable ligands to lysosomes (Merion and Sly, 1983; Harford et al., 1983) may reflect an inhibition of ligand discharge as opposed to a direct effect on the transport pathway itself (Mellman et al., 1986). One interesting possibility, however, has been suggested by electron microscopy. Ligands bound to two receptors destined for lysosomes, the EGF and Fc receptors, have been found to be tightly associated with the membrane of the small vesicular inclusions in endosomes (Fig. 4) (Haigler et al., 1979; Ukkonen et al., 1986). Presumably, these structures arise from small invaginations of the endosome’s limiting membrane (Hirsch et ul., 1968). Segregation of a receptor on these vesicular inclusions would effectively prevent recycling since in most cells internal vesicles would be transported to lysosomes along with other endosomal contents. Oligomerization of a down-regulating receptor may favor its inclusion in intraendosornal vesicles. d . Kinetic Considerations. Whatever its molecular mechanism, endosomal sorting of receptors may not reflect a dramatic all-or-none signal. Slight alterations in any one of the kinetic rate constants that govern endocytosis, recycling, and transport to lysosomes may exert large effects on a receptor’s intracellular distribution and fate. As shown in Fig. 5, a slight increase in the rate of internalization ( K , )without a corresponding increase in the rate of recycling (K,)would result in a net decrease in the number of surface receptors and an increase in receptor number in endosomes. In some cases, such as the asialoglycoprotein receptor of hepatocytes, ligand binding is believed to cause at least a slight increase in the forward rate constant for internalization (Schwartz ei al., 1982). A similar situation would arise if K , decreased while Ki remained unchanged. In either event, the increased concentration of receptors in endosomes might be expected to result in an increased probability that a receptor would be transferred to lysosomes and degraded. A third possibility is the rate of receptor transport to lysosomes (K,) may be directly affected by ligand binding.

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FIG.4. Association o f Fc receptor-bound ligiind with internal vesicles in macrophage endosomes. 5774 cells (a mouse macrophage cell line) were exposed to colloidal gold-labeled IgG-containing antibody-antigen complexes for 10 min at 37°C: under these conditions. virtually all internalized ligand i s found in endosonies (Ukkonen 1'1 ( I / . , 19%). Gold pailicles were observed closely associated (150-200 A) with multivesicular elements within the endosomes. Little gold was associated with the limiting membrane of the endosomes. Since antihody-antigen complexes do not dissociate from Fc receptors at low pH. i t i s likely that the gold also marks the localization of the receptors (Ukkonen Y I d.,1986). Bar, 0 . 2 pni.

That the alterations leading to down regulation may be comparatively subtle has been illustrated by a recent study of the transferrin receptor (Weissman rr id.. 1986). Uptake of a bivalent antireceptor antibody was found to increase the rate of receptor degradation ?-fold (corresponding to a decrease in half-life from 8 to 4 hr). At the same time, transferrin uptake (determined as cellular accumulation of transferrin-bound iron) was only slightly diminished. Kinetic analysis revealed that the observed increase in receptor turnover could be explained by a small (< 10%) decrease

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plasma membrane

1.y Y

Y ’ J

Lr

0

endosome

0

lyeoeome

FIG.5 . Kinetic parameters governing intracellular membrane traffic on the endocytic pathway. This diagram simply illustrates some of the more important rate constants which must be considered when evaluating possible mechanisms of receptor down regulation (see text for details). The kinetic parameters are defined as follows (in receptorshin): K,.rate constant for internalization; K,.,rate constant for receptor recycling; K,, rate constant for transport of internalized receptors from endosomes to lysosomes; K,. rate of synthesis of new receptors. The diagram is adapted from the kinetic model for receptor-mediated endocytosis of Jenny Lindermann and Douglas Lauffenberger (University of Pennsylvania).

in the probability of recycling of internalized receptors. Conceivably, this alteration in transferrin receptor transport could be triggered by receptor cross-linking by the bivalent antireceptor IgG used. In contrast to these results, however, other physiological systems of receptor down regulation are often characterized by a rapid accumulation of receptors in lysosomes (e.g., Fc receptor, EGF receptor; see Ukkonen et ul., 1986; Beguinot et a / . , 1984) and thus may display far more dramatic alterations in one or more kinetic rate constants. Preliminary kinetic analysis of Fc receptormediated endocytosis, for example, suggests that receptors bound to multivalent versus monovalent ligands display a decrease in recycling probability from approximately 98 to 25% (J. Lindermann, D. Lauffenberger, V. Lewis, and I. Mellman, unpublished results). Considerably more quantitative data will be needed before any generalizations can be made regarding the mechanisms of receptor down regulation. 7. POLARIZED EPITHELIAL CELLS

In addition to directing transport of certain receptors to lysosomes, endosomes clearly may play a role in maintaining the polarized distribution

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of membrane proteins in epithelial cells (for review, see Simons and Fuller, 1985). The transcellular transport of IgG or IgA across intestinal and hepatic epithelia (respectively) is well known to involve an endosomal intermediate (Abrahamson and Rodewald, 1981; Geuze ef al., 1983; Hoppe et nl., 1985). In addition, ectopic implantation of VSV G protein in the apical plasma membrane of cultured MDCK cells results in G protein’s rapid internalization and transit through endosomes en route to the basolateral surface (Matlin ef al., 1984; Pesonen and Simons. 1984; Pesonen ef al., 1984).

D. Lysosomes Lysosomes are the main digestive compartment in the cell, being rich in hydrolytic enzymes (proteases, glycosidases, lipases) capable of digesting almost all biological macromolecules to their component parts (Dingle et d.,1984). These enzymes typically have acidic pH optima (pH 4 . 5 - 5 3 , a property which reflects the low internal lysosomal p H , estimated to be between pH 4.6 and 4.8 (Ohkuma and Poole, 1978). The lysosomal membrane contains an ATP-driven proton pump, similar to that found in endosomes, which is responsible for acidification (Ohkuma et ai., 1982; Harikumar and Reeves, 1983; Mellman el al.. 1986). The membrane also displays discrete conductances for anions (e.g., CI-) and cations, both of which may be important in regulating electrogenic proton transport (see above). Lysosomes generally represent the final destination of extracellular rnacromolecules internalized during endocytosis. As discussed earlier, little membrane or content appears to escape from cells once delivered to the lysosomal compartment. Lysosomes are clearly not obligate intermediates in the pathway of receptor recycling. Several receptors, such as the transferrin receptor, Fc receptor, and mannose 6-phosphate receptor, have been shown directly (by Percoll gradient centrifugation) not to enter lysosomes during ligand uptake (van Renswoude ef a/., 1982; Mellman ef ul., 1984; Sahagian and Neufeld, 1983). Furthermore, incubation of cells at temperatures below 2PC, conditions which prevent the transfer of internalized material from endosomes to lysosomes (Dunn et ul., 19801, does not disrupt receptor recycling (see Mellman, 1984). Finally, if the lysosomal transport of a receptor which usually recycles is induced (e.g., Fc receptor, mannose 6-phosphate receptor), rapid receptor degradation usually results (Mellman and Plutner, 1984; von Figura ef al., 1984) (see above). The isolation of the lysosomal compartment from the constitutive recycling pathway is further emphasized by the fact that the lysosomal membrane is biochemically distinct from either the endosomal or plasma membranes. This has been indicated by SDS-gel analysis of organelles

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purified by free flow electrophoresis (Marsh et al., 1987) (see above) and also by the production of highly specific antilysosome monoclonal and polyclonal antibodies (Reggio et al., 1984; Lewis et al., 1985; Chen et al., 1985). These antiorganelle reagents have indicated that the lysosomal membrane is characterized by a unique set of glycoproteins. These lysosomal glycoproteins (Igp’s) are intrinsic membrane proteins which range in molecular weight from 20 to I50 kDa. The most extensively studied proteins, lgp120, lgpl10, and lgp55, are heavily N-glycosylated (containing up to 18 asparagine-linked oligosaccharide chains) and extremely acidic (pl < 4) due entirely to their high sialic acid content. Their polypeptide portion can actually be quite small. lgp120, for example, has a polypeptide molecular mass of only 38 kDa; thus, approximately one out of every seven amino acids is involved in generating a canonical tripeptide N-glycosylation site. Although intimately associated with the membrane, Igp’s appear to have short, if any, cytoplasmic domains. Finally, unlike most lysosomal enzymes, Igp’s do not contain mannose 6-phosphate and therefore must be targeted to lysosomes in a manner independent of the mannose 6-phosphate receptor (Lewis et al., 1985; Granger er a/., 1985; B. Granger, C. Howe, V. Lewis, N. Walworth, S. Green, A. Helenius, and I. Mellman, unpublished results). OF LYSOSOMES THEBIOGENESIS

The formation of lysosomes has long been of interest and represents a central problem in understanding membrane traffic on both the endocytic and biosynthetic pathways. The most extensively studied aspect of lysosomal biogenesis concerns the synthesis and targeting of lysosomal hydrolases (for review, see Creek and Sly, 1984). While it is clear that newly synthesized enzymes, tagged with the mannose 6-phosphate recognition marker, exit from the Golgi apparatus bound to one of two mannose 6phosphate receptors, the precise pathway by which these enzymes find their way to lysosomes is unknown. One possibility is that the Golgi-derived vesicles, “primary lysosomes,” fuse with endosomes where the low internal pH favors the dissociation of the enzyme-receptor complex resulting in the transport of enzymes (along with other endosomal contents) to lysosomes and the recycling of receptors either back to the Golgi or to the cell surface. This view is supported by observations that the mannose 6-phosphate receptor is rarely detected in lysosomes (Sahagian and Neufeld, 1983; V. Lewis and I. Mellman, unpublished results). It is also in agreement with the concept that endosomes are gradually converted into lysosomes as they migrate from the cell periphery to the perinuclear region (see above) (Helenius et al., 1983). However, the possibility that primary lysosomes continuously recycle between the Golgi and “preformed” sec-

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ondary lysosomes (i.e., high density structures already rich in hydrolases and containing endocytosed markers) cannot yet be eliminated. An analogous problem exists for the targeting of membrane components, such as the Igp family, to lysosomes. Preliminary immunocytochemical studies on ultrathin frozen sections has indicated the presence of tubular structures, rich in Igp’s, which may mediate transport to lysosomes from the Golgi ( H . Geuze, J. Slot, S. Green, A. Helenius, and 1. Mellman, unpublished results). Even assuming a transport function for these structures, however, it is not yet clear where in the pathway they are inserted (i.e., maturing endosomes or preformed secondary lysosomes). The discovery of the Igp family has created one additional consideration: How is this class of membrane glycoproteins preferentially targeted to and accumulated in the lysosomal membrane when all other membrane glycoproteins in mammalian cells fastidiously avoid transport to lysosomes? While Igp targeting clearly does not depend on the mannose 6phosphate receptor (see above), it is nonetheless attractive to postulate the existence of some other carbohydrate or protein recognition marker which signals transport of these proteins to their unique destination. However, other possibilities must be considered. Given the apparent role played by oligomeric assembly in triggering the transport of plasma membrane receptors to lysosomes, it is conceivable that Igp’s assume an analogous tertiary or quaternary structure which favors lysosomal targeting. The absence of any obvious primary structural feature would also be reminiscent of the high degree of structural diversity among receptors which localize at plasma membrane coated pits (see above and Fig. 2). It is also possible that, to some extent, the accumulation of Igp’s in lysosomes is an indirect consequence of their resistance to degradation by lysosomal hydrolases. By this view, targeting would not need to be highly selective if Igp’s, in contrast to all other membrane proteins, are able to survive the lysosomal environment. Transport to and accumulation of Igp’s in lysosomes would thus simply reflect the continuous transfer of endocytosed material from endosomes to lysosomes.

V. THE EXOCYTIC PATHWAY

Although far beyond the scope of this chapter, many of the principles which are just now being derived for membrane traffic on the endocytic pathway will no doubt also be applicable to understanding the exocytic pathway. For a consideration of the problems of exocytic membrane traffic and protein secretion, the reader is referred to reviews by Kelly (1985), Farquhar (1983, and Simons and Fuller (1985).

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In direct analogy to endocytosis, membrane vesicles and their secretory protein contents are formed and transferred through successive biosynthetic compartments en route to their final destinations. While the plasma membrane is the most common destination, two major routes are generally distinguished. The so-called regulated pathway involves the targeting to, concentration, and storage of characteristic secretory proteins in specialized secretion granules. These granules fuse with the plasma membrane only in response to the appropriate stimulus. The constitutive pathway, on the other hand, is characterized by the continuous, nonconcentrative exocytosis of newly synthesized proteins. It is thought that new plasma membrane components reach the cell surface via the constitutive pathway, as opposed to the regulated pathway. What factors govern the transport of individual membrane and secretory proteins on the constitutive versus the regulated pathways? How are constitutively secreted proteins sorted from those to be stored in secretion granules? Little information is available on either of these points. One view postulates the involvement of distinct intracellular receptors which mediate the transport of newly synthesized proteins from endoplasmic reticulum to Golgi, and then from Golgi to storage granules or directly to the cell surface (Kelly, 1985). If such receptors exist, however, they would have to be extraordinarily nonselective. For example, a mouse pituitary tumor cell line (AtT-201, which normally stores the polypeptide hormone ACTH in granules for regulated secretion, will also store proteins such insulin and trypsinogen synthesized from transfected cDNAs (Moore et al., 1983; Burgess ef al., 1985). Although these proteins are subject to granule storage and regulated secretion in their cells of origin (endocrine and exocrine pancreas, respectively), they are never expressed in pituitary cells. There is specificity in the system, however, since proteins which are not normally subjected to regulated secretion, such as albumin, are not targeted to the AtT-20 granules. In addition, a role for acidic intravesicular pH in targeting has been implied by the finding that chloroquine diverts ACTH from the regulated to the constitutive secretory pathway (Moore et a [ . , 1983). Similarly broad specificity would apply to any putative receptor on the constitutive exocytic pathway. Cells not only synthesize and secrete a wide variety of their own polypeptides but also are capable of expressing many “exogenous” proteins such as viral spike glycoproteins and the products of transfected cDNAs. Therefore, it seems unlikely that specific intracellular receptors can exist for each individual protein. While there is insufficient information to reach any firm conclusions concerning the role of receptors in transport on the exocytic pathway, it is important to stress that the role of quaternary structure, or state of oligomeric assembly, should be considered as a possible controlling factor.

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As discussed above for the case of the Fc receptor on the endocytic pathway, interaction with either monovalent or multivalent ligands exerts profoundly different effects on the fate of internalized receptors. It seems likely that ligand valency would in turn be expected to influence the degree of receptor aggregation. Conceivably, the molecular mechanisms underlying the intracellular membrane transport react differentially according to the oligomeric state of the proteins being transported. Accordingly, secretory proteins which are targeted to storage granules on the regulated pathway may signal their transport by a tendency to selfassociate as they enter the trans-most Golgi cisternae. Morphological evidence for such accretion is often observed for proteins destined for storage granules (see Kelly, 1985). All secretory proteins which do not self-associate in this way might be exocytosed via the constitutive-or nonselective-pathway. Direct evidence for the role of quaternary structure even in constitutive exocytic transport can be seen from studies demonstrating that immunoglobulin heavy chain is not transported out from the endoplasmic reticulum unless complexed with immunoglobulin light chain (Bole et al., 1986). Analogous results have been obtained for membrane proteins such as the class I histocompatibility antigen heavy chain, which does not reach the cell surface unless noncovalently complexed with p,-microglobulin (Severinsson and Peterson, 1984). These few examples certainly do not prove a unique role for quaternary structure in controlling the intracellular traffic of membrane and secretory proteins. They do, however, emphasize that structural features other than those which may be obvious from amino acid sequence data are likely to be important in the exocytic pathway just as they appear to be important at crucial stages of the endocytic pathway.

ACKNOWLEDGMENTS Research for this article was supported by grants from the National Institutes of Health (GM-29765, GM-33093. and the Yale Liver Center) and a Fellowship from the American Cancer Society. REFERENCES Abrahamson, D. R.. and Rodewald. R. (1981). Evidence for the sorting ofendocytic vesicle contents during the receptor-mediated transport of I& across the newborn rat intestine. J . Cell Biol. 91, 270-280. Adarns. C. J . , Maurey, K. M., and Stome. B. (1982). Exocytosis of pinocytic contents by Chinese hamster ovary cells. J . Cell Biol. 93. 632437. Aggeler, J . , and Werb, Z. (1982). Initial events during phagocytosis by macrophages viewed from outside and inside the cell: Membrane-particle interactions and clathrin. J . Ce// B i d . 94. 613-623.

282

IRA MELLMAN ET AL.

Aggeler, J . , Takemura, R., Nichols, B. A., and Werb, Z.A. (1985).Macrophage membranes and clathrin. I n “Mononuclear Phagocytes: Characteristics, Physiology, and Function” (R. van Furth, ed.), pp. 87-98. Martinus Nijhoff, Dordrecht. Anderson, R. G . W., and Pathak. R. K. (1985).Vesicles and cisternae in the trans Golgi apparatus of human fibroblasts are acidic compartments. Cell 40,635-643. Anderson, R. G . W., Brown, M. S. , Beisiegel, U., and Goldstein, J . L. (1982).Surface distribution and recycling of the low density lipoprotein receptor as visualized with antireceptor antibodies. J . Cell Biol. 93, 523-531. Anderson, R. G . W., Falck, J . R., Goldstein, J. L., and Brown, M. S. (1984).Visualization of acidic organelles in intact cells by electron microscopy. Proc. Narl. Arad. Sci. U . S . A . 81, 48384842. Ascoli, M. (1984).Lysosomal accumulation of the hormone-receptor complex during receptor-mediated endocytosis of human choriogonadotropin. J . Cell Biol. 99, 1242-1250. Beguinot, L., Lyall, R. M., Willingham, M . C., and Pastan, I. (1984).Down regulation of the EGF receptor in KB cells is due to receptor internalization and subsequent degradation in lysosomes. Proc. Natl. Acad. Sci. U.S.A. 81, 2384-2388. Besterman, J . M., Airhart, J . A,, Woodworth, R. C., and Low, R. B. (1981).Exocytosis of pinocytosed fluid in cultured cells: Kinetic evidence for rapid turnover and compartmentation. J . Cell Biol. 91, 716-727. Bole, D. G . , Hendershot, L. M., and Kearney, J. F. (1986).Posttranslational association of immunoglobulin heavy chain binding protein with nascent heavy chains in nonsecreting and secreting hybridomas. J . Cell Biol. 102, 1558-1566. Braell, W. A., Schlossmam, D. M., Schmid, S. L., and Rothman, J. E. (1984).The dissociation of clathrin coats coupled to the hydrolysis of ATP: Role of an uncoating ATPase. J . Cell B i d . 99, 734-741. Brown, M . S . , and Goldstein, J. L. (1976).Analysis of a mutant strain of human fibroblasts with a defect in the internalization of receptor-bound low density lipoprotein. Cell 9. 663-674. Brown. M. S . , Anderson, R. G . W., and Goldstein, J . L. (1983).Recycling receptors: The round trip itinerary of migrant membrane proteins. Cell 32,663-667. Burgess, T. L., Craik, C. S . , and Kelly, R. B. (1985).The exocrine protein trypsinogen is targeted into the secretory granules of an endocrine cell line: Studies by gene transfer. J . Cell B i d . 101. 639-645. Carpentier, J.-L., Gorden, P., Anderson, R. G. W., Goldstein, J. L., Brown, M. S . , Cohen, S ., and Orci, L. (1982).Co-localization of ‘251-epidermalgrowth factor and ferritin lowdensity lipoprotein in coated pits: A quantitative electron microscopic study in normal and mutant human fibroblasts. J . Cell B i d . 95, 73-77. Chen, J . W., Murphy, T. L., Willingham, M. C., Pastan, I., and August, J. T. (1985).Identification of two lysosomal membrane glycoproteins. J . Cell Biol. 101,85-95. Chiacchia, K. B., and Drickamer, K. (1984).Direct evidence for the transmembrane orientation of the hepatic glycoprotein receptors. J . B i d . Chem. 259, 15440-15446. Courloy, P. J . , Quintart, J . . and Baudhuin, P. (1984).Shift of equilibrium density induced by 3,3’-diaminobenzidine cytochemistry: A new procedure for the analysis and purification of peroxidase-containing organelles. J . Cell B i d . 98, 870-876. Creek, K. E., and Sly, W. S. (1984).The role of the phophomannosyl receptor in the transport of acid hydrolases to lysosomes. I n “Lysosomes in Biology and Pathology” (J. T. Dingle, R. T. Dean, and W. S. Sly, eds.), Vol. 7, pp. 63-82. Elsevier, Amsterdam. Dickson. R. B., Beguinol, L., Hanover, J . A., Richert, N. D., Willingham, M . C., and Pastan, I. (1983).Isolation and characterization of a highly enriched preparation of

THE CONTROL OF MEMBRANE TRAFFIC

283

receptosomes (endosomes) from a human cell line. Proc. Ntrrl. Actrd. S c i . U . S . A . 80. 5335-5339. Dingle. J . T.. Dean, R . T.. and Sly, W. S. (eds.) (1984). "Lysosomes in Biology and Pathology." Vol. 7, Elsevier, Amsterdam. Draper. R. K., O'Keefe, D. 0.. Stookey, M.. and Graves. J . (1984). Identification of a coldsensitive step in the mechanism of modeccin action. J . B i d . C h m . 259, 4083-4088. Dunn. W. A , . Hubbard. A. L., and Aronson. N . N . (1980). Low temperature selectively inhibits fusion between pinocytic vesicles and lysosomes during heterophagy of '"1' rlofetuin by the perfused liver. J . B i ~ l Chem. . 255, 5971-5978. Ekblom. P., Thesleff, I . . Lehto, V.-P.. and Virtanen. 1. (1983). Distribution of the transferrin receptor in normal human fibroblasts and tibrosarcoma cells. l n i . J . Ctrticcr 31, I I I117. Farquhar, M. G. (1985). Progress in unraveling pathways of Golgi traffic. Annrr. Rev. Ccll Bi<J/. I , 447-488. Galloway, C. J., Dean, G . E., Marsh, M., Rudnick. G . . and Mellman. I. (1983). Acidification of macrophage and fibroblast endocytic vesicles in iirro. Pro(.. N o r / . Accrd. Sci. U.S.A. 80. 3334-3338. Geuze. H. J.. Slot, J . W., and Strous, G. J . A. M. (1983). Intracellular site of asialoglycoprotein receptor-Egand uncoupling: Double-label immunoelectron microscopy during receptor-mediated endocytosis. Cell 32, 277-287. Geuze, H. J . , Slot. J . W., Strous. G . J . A. M.. Peppard. J . , von Figura. K., Hasilik. A,, and Schwartz, A. L. (1984). lntracellular receptor sorting during endocytosis: Comparative immunoelectron microscopy of multiple receptors in rat liver. C P / /37. 195204. Glickman, J . , Croen, K., Kelly, S. . and Al-Awqati, 0. (1983). Golgi membranes contain an electrogenic proton pump parallel to a chloride conductance. J . Cpll Biol. 97. 13031308. Goldstein. J . L.. Anderson. R. G.. and Brown. M. S. (1979). Coated pits. coated vesicles, and receptor-mediated endocytosis. Ntitrirc (Loudon) 279, 679-685. Gonzalez-Noriega, A , . Grubb. J. H., Talkad. V.. and Sly, W. S . (1980). Chloroquine inhibits lysosomal enzyme pinocytosis and enhances lysosomal enzyme secretion by impairing receptor recycling. J . Cell Biol. 85. 839-852. Goud. B.. Huet. C., and Louvard, D. (1985). Assembled and unassembled pools ofclathrin: A quantitative study using an enzyme immunoassay. J . Cell Biol. 100. 521-527. Granger, B. L., Plutner, H., Green, S . A , . Mellman, I . . and Helenius, A. (1985). Defining the lysosome membrane: Characterization of specific glycoproteins. J . Cc4l Biol. 101. I 99a. Green, S . A,, Plutner, H., and Mellman. I . ( l Y 8 5 ) . Biosynthesis and intracellular transport of the mouse macrophage Fc receptor. J . Biol. C h n . 260. 9867-9874. Haigler. H. T., McKanna, J. A,, and Cohen. S . (1979). Direct visualization of the binding and internalization of a ferritin conjugate of epidermal growth factor in human carcinoma cells A-43 I . J . Crll Biol. 81, 382-395. Harford. J . , Bridges. K., Ashwell, G . . and Klausner, R . D. (1983). lntracellular dissociation of receptor-bound asialoglycoproteins in cultured hepatocytes: A pH-mediated nonlysosomal event. J . B i d . Chcm. 258. 3191-3197. Harikumar. P.. and Reeves. J . P. (1983). The lysosomal proton pump is electrogenic. J . Biol. C k r n . 258, 10403-10410. Harrison. S. C . , and Kirchhausen, 1'. (1983). Clathrin. cages. and coated vesicles. Cell 33. 650-652.

284

IRA MELLMAN ET AL.

Helenius, A., Mellman, I., Wall, D., and Hubbard, A. (1983). Endosomes. Trends Biochem. Sci. 8, 245-250. Herman, B., and Albertini. D. F. (1984). A time-lapse video image intensification analysis of cytoplasmic organelle movements during endosome translocation. J . Cell Biol. 98, 565-576. Heuser, J., and Evans, L. (1980). Three dimensional visualization of coated vesicle formation in fibroblasts. J . Cell Biol. 84, 560-583. Hirsch, J . G., Fedorko, M. E., and Cohn, Z. A. (1968). Vesicle fusion and formation at the surface of pinocytic vesicles in macrophages. J . Cell Biol. 38, 629-632. Hopkins, C. R., and Trowbridge, I. S. (1983). Internalization and procession of transferrin and the transferrin receptor in human carcinoma A431 cells. J . Cell Biol. 97, 508-521. Hoppe, C. A., Connolly, T. P., and Hubbard, A. L. (1985). Transcellular transport of polymeric IgA in the rat hepatocyte: Biochemical and morphological characterization of the transport pathway. J . Cell Biol. 101, 21 13-2123. Kasuga, M., Kahn, J. A., Hedo, E., Obbergen, V., and Yamada, K. M. (1981). Insulininduced receptor loss in cultured human lymphocytes is due to accelerated receptor degradation. Proc. Natl. Acad. Sci. U . S . A . 78, 6917-6921. Kelly, R. B. (1985). Pathways of protein secretion in eukaryotes. Science 230, 25-32. Kielian, M. C., Marsh, M., and Helenius, A. (1986). Kinetics of endosome acidification detected by mutant and wild type Semliki Forest virus. EMBO J . , in press. Kirchhausen, T., and Harrison, S. C. (1981). Protein organization in clathrin trimers. Cell 23, 755-761. Klausner, R. D., van Renswoude, J., Kempf, C., Rao, K., Bateman, J. L., and Robbins, A. R. (1984). Failure to release iron from transferrin in a Chinese hamster ovary cell mutant pleiotropically defective in endocytosis. J. Cell Biol. 98, 1098-1 101. Larkin, J. M., Brown, M. S., Goldstein, J. L.. and Anderson, R. G. W. (1983). Depletion of intracellular potassium arrests coated pit formation and receptor-mediated endocytosis in fibroblasts. Cell 33, 273-285. Lehrman, M. A., Schneider, W. J., Sudhof, T. C., Brown, M. S., Goldstein, J . L., and Russell, D. W. (1985). Mutation in LDL receptor: Ala-Ala recombination deletes exons encoding transmembrane and cytoplasmic domains. Science 227, 140-146. Lewis, V . , Green, S., Marsh, M., Helenius, A., and Mellman, I. (1985). Glycoproteins of the lysosomal membrane. J . Cell Biol. 100, 1839-1847. Lewis, V., Koch, T., Plutner, H., and Mellman, I. (1986). Characterization of a cDNA clone for a mouse macrophage-lymphocyte Fc receptor. Nature (London) 324, 372-375. Louvard, D. (1980). Apical membrane aminopeptidase appears at site of cell-cell contact in cultured kidney epithelial cells. Proc. Natl. Acad. Sci. U.S.A. 77, 4132-4136. Marsh, M., and Helenius, A. (1980). Adsorptive endocytosis of Semliki Forest viru’s. J. Mol. Biol. 142, 439454. Marsh, M., Bolzau, E., White, J., and Helenius, A. (1983a). Interactions of Semliki Forest virus spike glycoprotein rosettes and vesicles with cultured cells. J . Cell Biol. 96, 455461. Marsh, M., Bolzau, E., and Helenius, A. (1983b). Penetration of Semliki Forest virus from acidic prelysosomal vacuoles. Cell 32, 93 1-940. Marsh, M., Griftiths, G., Dean, G. E., Mellman, I., and Helenius. A. (1986). Three-dimensional structure of endosomes in BHK-21 cells. Proc. Natl. Acad. Sri. U . S . A .83, 2899-2903. Marsh, M., Schmid, S. , Kern, H., Harms, E., Male, P., Mellman, I., and Helenius, A. (1987). Rapid analytical and preparative isolation of functional endosomes by free flow electrophoresis. J . Cell Biol. (in press).

THE CONTROL OF MEMBRANE TRAFFIC

285

Matlin. K.. Bainton, D. F.. Pesonen. M., Louvard, D., Genty, N.. and Simons, K. (1983). Transepithelial transport of a viral membrane glycoprotein implanted into the apical plasma membrane of Madin-Darby canine kidney cells. I. Morphological evidence. J. Cell Biol. 97. 627-637. Mellman. I. (1982). Endocytosis, membrane recycling, and Fc receptor function. hi “Membrane Recycling” (D. Evered. ed.), Ciba Foundation Symposium No. 92. pp. 35-58. Pitman Press, London. Mellman, I. (I 984). Membrane recycling during endocyfosis. I n “Molecular and Cellular Aspects of Lysosomes.” Lysosomes in Biology and Pathology, Vol. 7, (J. T. Dingle. R. Dean, and W.Sly, eds.), pp. 201-229. Elsevier, Amsterdam. Mellman, 1. (1986). Mechanisms of molecular sorting in endosomes. I n “New Insights into Cell and Membrane Transport Processes” (G. Poste and S. T. Crooke. eds.), pp. 377401. Plenum, New York. Mellman. I., and Plutner, H. (1984). Internalization and degradation of macrophage Fc receptors bound to polyvalent immune complexes. J . Cell B i d . 98, 1170-1176. Mellman, I., Steinman. R. M.,Unkeless. J. C., and Cohn, Z. A. (1980). Selective iodination and polypeptide composition of pinocytic vesicles. J . Cdl B i d . 86, 712-722. Mellman, I., Plutner, H., Steinman, R. M., Unkeless, J . C., and Cohn, Z. A. (1983). lnternalization and degradation of macrophage Fc receptors during receptor-mediated phagocytosis. J . Cell B i d . 96, 887-895. Mellman, I., Plutner, H., and Ukkonen, P. (1984). Internalization and rapid recycling of macrophage Fc receptors tagged with monovalent antireceptor antibody: Possible role . 1163-1169. of a prelysosomal compartment. J . Cell E i ~ l 98, Mellman, I., Fuchs, R., and Helenius, A. (1986). Acidification of endocytic and exocytic pathways. Annu. Rev. Bicichem. 55,663-700. Merion, M., and Sly, W. S. (1983). The role of intermediate vesicles in the adsorptive endocytosis and transport of ligand to lysosomes by human fibroblasts. J. Cell Biol. 96, 644-650 * Merion, M., Schlesinger. P., Brooks, R. M., Moehring, J. M.. Moehring, R. J . , and S l y , W. S. (1983). Defective acidification of endosomes in Chinese hamster ovary cell mutants ”cross-resistant” t o toxins and viruses. Pro<..N a f l . A m d . Sci. U . S . A . 80, 5315-531 9. Montesano, R., Mossaz, A., Vassalli, P., and Orci, I.. ( 1983). Specialization of the macrophage plasma membrane at sites of interaction with opsonized erythrocytes. J. Cell B i d . 96, 1227-1233. Moore, H.-P., Gumbiner, B., and Kelly, R. B. (1983). Chloroquine diverts ACTH from a regulated to a constitutive secretory pathway in AtT-20 cells. Nufure (London) 302, 434-436. Moya, M.,Dautry-Varsat, A., Goud, B., Louvard, D., and Boquet, P. (1985). lnhibition of coated pit formation in HepG2 cell blocks the cytotoxicity of diptheria toxin but not that of rich toxin. J. Cell Biol. 101, 548-559. Murphy, R. F., Powers, S., and Cantor, C. R. (1984). Endosome pH measured in single cells by dual fluorescence flow cytometry: Rapid acidification of insulin to pH 6 . J. Cell B i d . 98, 1757-1762. Neutra, M. R., Ciechanover, A., Owen, L. S., and Lodish, H. F. (1985). lntracellular transport of transfenin and asialoorosomucoid colloidal gold conjugates lo lysosomes after receptor mediated endocytosis. J. Hisrochem. Cyfochem.33, 1134-1 144. Ohkuma, S.. and Poole. B. (1978). Fluorescence probe measurement of the intralysosomal pH in living cells and the perturbation of pH by various agents. Proc. N o t / . Acad. Sci. U.S.A. 75, 3327-3331.

286

IRA MELLMAN ET AL.

Ohkuma, S., Moriyami, Y., and Takano, T. (1982). Identification and characterization of a proton pump in lysosomes by fluorescein isothiocyanate dextran fluorescence. Proc. Null. Acud. Sci. U . S . A . 79, 2758-2762. Palade, G. (1975). Intracellular aspects of the process of protein synthesis. Scirnce 189, 347-358. Pastan, I., and Willingham, M. C. (1983). Receptor-mediated endocytosis: Coated pits. receptosomes, and the Golgi. Trends Biochem. Sci. 8, 250-254. Pastan, I . , and Willingham, M. C. (1985). The pathway of endocytosis. In "Endocytosis" (I. Pastan and M. C. Willingham, eds.), pp. 1-44. Plenum, New York. Pearse, B. M. F., and Bretscher, M. S. (1981). Membrane recycling by coated vesicles. Annu. Rev. Biochern. 50, 85-101. Pesonen, M., and Simons, K. (1984). Transcytosis of the G protein of vesicular stomatitis virus after implantation into the apical plasma membrane of Madin-Darby canine kidney cells. 1. Involvement of endosomes and lysosomes. J . Cell B i d . 99, 796-802. Pesonen, M., Bravo, R.. and Simons, K. (1984). Transcytosis of the G protein of vesicular stomatitis virus after implantation into the apical membrane of Madin-Darby canine kidney cells. 11. Involvement of the Golgi complex. J. Cell B i d . 99, 803-809. Petty, H. R., Hafeman, D. G., and McConnell, H. M. (1981). Disappearance of macrophage surface folds after antibody-dependent phagocytosis. J. Cell Biol. 89, 223-229. Pfeffer, S. R., Drubin, D. G., and Kelly, R. B. (1983). Identification of three coated vesicle components as alpha- and beta-tubulin linked to a phosphorylated 50.000 dalton polypeptide. J . Cell Biol. 97, 4 0 4 7 . Reggio, H., Bainton, D., Harms, E., Coudrier, E., and Louvard, D. (1984). Antibodies against lysosomal membranes reveal a 100,000-mol-wt protein that cross-reacts with purified H',K'-ATPase from gastric mucosa. J . Cell B i d . 99, 1511-1526. Robbins, A. R., Peng, S. S., and Marshall, J. L. (1983). Mutant Chinese hamsterovary cells pleiotropically defective in receptor-mediated endocytosis. J . Cell Biol. 96, 1064-1071. Robbins, A. R., Bateman, J., Krag, S., Oliver, C., Galloway, C. J., and Mellman, 1. (1984). A single mutation in Chinese hamster ovary cells impairs both Golgi and endosomal functions. J. Cell Biol. 99, 1296-1308. Roth, M. G., Doyle, C., Sambrook, J., and Gething, M.-J. (1986). Heterologous transmembrane and cytoplasmic domains direct functional chimeric influenza virus hemagglutinins into the endocytic pathway. J. Cell B i d . 102, 1271-1283. Rothman, J . E., and Schmid, S. L. (1986). Enzymatic recycling of clathrin from coated vesicles. Cell 46, 5-9. Russell, D. W., Schneider, W. J., Yamamoto, T., Luskey, K. L., Brown, M.S., and Goldstein, J . L. (1984). Domain map of the LDL receptor: Sequence homology with the epidermal growth factor precursor. Cell 37, 577-585. Sahagian, G. G., and Neufeld, E. F. (1983). Biosynthesis and turnover of the mannose 6phosphate receptor in Chinese hamster ovary cells. J . Biol. Chem. 258, 7121-7128. Schmid. S., Braell, W. A., Schlossmdn, D. M., and Rothman, J. E. (1984). A role for clathrin light chains in the recognition of clathrin cages by "uncoating ATPase." Nutrtre (London) 311, 228-231. Schreiber, A. B., Liberman, T. A., Lax, I., Yarden, Y., and Schlessinger, J. (1983). Biological role of epidermal growth factor receptor clustering: Investigation with monoclonal antireceptor antibodies. J. B i d . Chem. 258,846-854. Schwartz, A. L., Fridovich, S. E., and Lodish, H. F. (1982). Kinetics of internalization and recycling of the asialoglycoprotein receptor in a hepatoma cell line. J . Biol. Cliem. 257, 423M237.

THE CONTROL OF MEMBRANE TRAFFIC

287

Severinsson. I... and Peterson, P. A. (1984). P,-Microglobulin induces intracellular transport of human class 1 transplantation antigen heavy chains in Xcrioptrs k m i v oocytes. J . C ~ l Bit)/. l 99. 226-23 1. Silverstein, S. C., Steinman. R . M.. C'ohn. Z. A . (1977). Endocytosis. Annir. Rcv. B i o c ~ h m . 46, 669-722. Simons. K.. and Fuller. S. D. (1985). Cell surface polarity in epithelia. Aanu. Re\.. C d R i d . I . 295-340. Steinman. R. M.. Brodie, S.E.. and Cohn. Z. A . (1076). Membrme flow during pinocytosis. J . C ~ lBl i ~ l68. . 665-687. Steinman. R. M., Mellman. I.. Mullcr. W. A . . and Cohn, Z.A . (1983). Endocytosis and the recycling of plasma membrane. J . CrlI Bird. 96. 1-27. Stoscheck. C. M.,and Carpenter, G . (1984). Down regulation of epidermal growth factor receptors: Direct demonstration of receptor degradation in human fibroblasts. J . Cell Riol. 98, 1048-1053. Straus, W. ( 1964). Cytochemical observations on the relationship between lysosomes and phagosomes in kidney and liver by combined staining for acid phosphatase and intravenously injected horseradish peroxidase. J. Cell Riol. 20, 497-507. Swanson. J. A., Yirinec, B. D., and Silverstein. S . C. (198.5). Phorbol esters and horseladish peroxidase stimulate pinocytosis and redirect flow of pinocytosed fluid in macrophages. J. Cell B i d . 100, 851-859. Tycko, B . , and Maxfield, F. R. (1982). Rapid acidification of endocytic vesicles containing cr,-macroglobulin. Cell 28. 6 4 3 4 5 I. Ukkonen. P.. Lewis, V., Marsh. M..Helenius. A . . and Mellman, I. (1986). Transport of macrophage Fc receptors and Fc receptor-bound ligands to lysosomes. J . Exp. M c d . 163, 952-971. Ullrich, A.. Coussens. L.. Hayflick, J . S.. Dull, T. J., Gray. A.. Tam, A. W.. Lee. J.. Yarden, Y.. Livemann, T. A.. Schlessinger. J.. Downward. J., Mayes, E. L. V..Whittle. N.. Waterfield. M. D., and Seeburg. P. ti. (19x4). Human epidermal growth factor receptor cDNA sequence and aberrant expression of the amplified gene in A431 epidermoid carcinoma cells. Natrrrc, ( L o r i c l o n ) 309, 418-425. Ullrich, A.. Bell, J. R., Chen, E. Y., Herrera, K.. Petrurclli. L. M.. e t d.(1985). Human insulin receptor and its relationship to the tyrosine kinase family of oncogenes. Ncrtrrr-e (Lorrdon) 313, 75676 I . Unanue. E.. Ungewickell, E.. and I h n t o n . D. (1981). The binding of clathrin triskelions to membranes from coated vesicles. Ccll 26. 439-446. Ungewickell, E . . and Branton. D. ( 19x11, Assembly units ufclathrin coats. N ~ t r r r r(Loridorr) 289, 420-427. van Deurs, R.. and Nilausen. K. ( 1982). Pinocytosis in mouse L-fibroblasts: Ultrastructural evidence for a direct shuttle between the plasma membrane and the lysosomal compartment. J . Cell B i d . 94. 279-286. van Deurs. B.. Tdnnessen. T. I . . Petersen, 0. W . , Sandvig. K., and Olsnes. S . (1986). Routing of internalized r i c h and r i c h conjugales to the Golgi complex. J . Cell B i d . 102. 37-47. van Kenswoude, J . , Bridges, K.. Harford, J . . and Klausner, R. D. (1982).Receptor-mediated endocytosis of transferrin and the uptake of Fe in K562 cells: Identification of ;I nonlysosomal acidic compartment. Procs.Ntitl. A w d . St,;. U . S . A . 79. 6186-6100. von Figura. K., Gieselmann. V., and Hasilik, A. (1984). Antibody t o mannose 6-phosphate specific receptor induces receptor deficiency in human fibroblasts. EMBO J . 3, 12x11282.

288

IRA MELLMAN ET AL.

Wall, D. L.. Wilson. G., and Hubbard, A. L. (1980). The galactose-specific recognition system of mammalian liver: The route of ligand internalization in rat hepatocytes. Cell 21, 19-93. Weissman. A. M., Klausner, R. D., Rao, K.,and Harford, J. B. (1986). Exposure of K562 cells to anti-receptor monoclonal antibody OKT9 results in rapid redistribution and enhanced degradation of the transfemn receptor. J . Cell B i d . 102, 951-958. Werb, Z., and Cohn, Z. A. (1972). Plasma membrane synthesis in macrophages following the phagocytosis of polystyrene latex. J . B i d . Chem. 247, 2439-2446. White, J . , Doms, R. W.,Gething, M.-J., Kielian, M., and Helenius, A. (1986). Viral membrane fusion proteins. Microbiol. Rev. 50, 54-59. Willingham, M. C., and Pastan, I. (1980). The receptosome: an intermediate organelle of receptor-mediated endocytosis in cultured fibroblasts. Cell 21, 67-77. Yamamoto, T.,Davis, C. G., Brown, M. S., Schneider, W. J., Casey, M. L., Goldstein, J. L., and Russell, D. W. (1984). The human LDL receptor: A cysteine-rich protein with multiple Ala sequences in its mRNA. Cell 39, 27-38. Yamashiro, D. J . , Tycko, B., Fluss, S . R., and Maxfield, F. R. (1984). Segregation of transferrin to a mildly acidic (pH 6.5) para-Golgi compartment in the recycling pathway. Cell 37, 789-800.

Index

A Acetylcholine receptor lateral diffusion coefficients, 109-1 10, I I4 low resolution study, 13 Actin filaments, see Cytoskeleton Alamethicin, high resolution study, 14 Alkyldihydroxyacetone-phosphatesynthase. in phospholipid biosynthesis ether bond formation and, 146 peroxisomal location. 147 Amino acid sequences bacteriorhodopsin, 60 glucagon, 60 glycophorin, 60 ATPase Ca” -, in sarcoplasmic reticulum irregular distribution, 33-34, 5 I lateral diffusion, I14 proton endosomal acidification and, 268-270 defects in mutant phenotypes, 27027 I origin and biogenesis, 270 lysosomal, acidification and, 277

B Bacteriorhodopsin a-helical configuration circular dichroism. 13 electron diffraction and microscopy, 13, 14 amino acid sequence, 60 lateral diffusion, I14

molecular structure 49, 61 in purple membrane, Hulohacreriitm. 58-59 rotational diffusion, 98-99 X-ray diffraction, 2 Bis(monoacylg1ycero)phosphate biosynthesis, 155-156. 164 lysosomal location, 136-137

C Calelectrin, from Torpedo electropax, exocytosis and. 220 Cardiolipin. mitochondria1 biosynthesis. 155 content in membranes, 136 fatty acid composition. 139 CDPdiac ylgl ycerol conversion to cardiolipin, IS5 phosphatidylglycerol. 1 5 4 1 5 5 phosphatidylinositol. 153-154 formation from phosphatidic acid and CTP. 152-153 Cell-substrate adhesion, thermodynamic model, 71-89 chemical potentials, 74-76 configuration variables, three phases of changes, 83-89 free energy minimizing. 78-82 physical parameters. 82-83 receptor-site bond formation. 72-74 repulsive potential, 67-78 Cholesterol complexes with filipin, 42 concentration in membranes. 42-43

289

INDEX

content in membranes, 137 depletion from cells, functional changes, 181-182 effect on lipid bilayers, 39-40 liposome fusion and, 227 spontaneous intermembrane exchange desorption as rate-limiting step, 180 interactions between erythrocytes and plasma, 176 lipid vesicles and cell membrane, 176-177 kinetics acceptor chemical composition and, 177- 178, 180-1 8 1 activation energy, 177-178 mechanism aqueous diffusion, 179-180 collisionally dependent, 179-180 transmembrane movement in lipid vessels, 192 in various biomembranes, 191 Chromaffin granules, movement to site of exocytosis directed, Ca"-sensitive, 216-217 random. aggregation in v i m . 218-219 Clathrin pit coating, 260-261 vesicle coating, 261 CTP, conversion to CDPcholine, 148-149 CDPdiacylglycerol, 152-153 CDPethanolamine, 148-149 Cytochalasin B, lateral diffusion in membranes and, 1 I 1 Cytochrome c oxidase low resolution study, 13 reconstituted into liposomes, lipid immobilization. 17-18 Cytochrome P-450, rotational diffusion in liposomes, l00-101 in microsomes, 99-100 in reconstituted vesicles, 100 Cytoskeleton chromaffin granule movement into apposition and, 216-217 exocytosis and, 219, 221 lateral diffusion control, I 11-1 14 in erythrocyte membrane, 112-1 13 mechanism of, 112, I14

D Diac ylgl ycerol conversion to ethanolamine plasmalogen, 149-150 phosphatidylcholine, 149 phosphatidylethanolamine, 149 triacylglycerol, 150 exocytosis triggering by, 215 formation from phosphatidic acid, 147. 148 transbilayer movement in erythrocytes, I92 Diacylglycerol acyltransferase, microsomal, 150 Dihydroacetone phosphate, acylation to phosphatidic acid. 145-146 Dihydroacetone-phosphate acyltransferase, microsomal and peroxisomal, 145 Dipalmitoylphosphatidylcholine, phase transition in bilayers, 9

E Electron microscopy biomembranes, diversity and similarity, 30-3 I freeze-fracture bilayer defects, 45-46 IMP distribution in biomembranes, 33-35, 40-42 membrane fusion, 208-209 membrane proteins, 58 synaptic vesicle exocytosis, frog, 63, 64 new technique for membrane study, 6265 nonbilayer lipid structures, 45, 47 Electron spin resonance (ESR) lipid bilayer polarity, 8 probes for diffusion assay, 92 protein-lipid interface properties, 17-19 saturation-transfer, rotational diffusion study, 94-95, 98, 100-101 Endocytosis membrane recycling, 257-258 vesicle sorting of from content, 257258 phagocytosis, 256-257

INDEX

pinocytosis, 256-257 receptor-mediated pathway of, 259-260 receptor recycling. 2%-259 Endosomes acidification, 268-27 I ATP-driven proton pump and. 268-270 genetics, 270-271 mechanism, 269 biogenesis, 265-266 definition, 264-265 epithelial cell polarization and, 276-277 isolation, 266-267 free flow electrophoresis, 267 receptor sorting. 271-277 kinetic parameters. 274-276 recycling. 271-272 lransport to lysosomes. 272 oligomeric structure role, 273-274 vesicular inclusion function. 274275 structure, 26s Erythrocyte membrane band 3 lateral diffusion, cytoskeletal control. 112-1 13 rotational diffusion. 95-97 trypsin digestion effect, 96-97 Sendai virus fusion and. 222 cholesterol depletion, effect on cellular functions. 181 spontaneous exchange with plasma, I76 electron microscopy. 30-31 freeze-fracture, 33-34 ferritin-labeled glycoprotein, 50 IMP distribution, 33-34 identification, 53 immobilization by cytoskeletal network, 56 radial distribution, 54-55 lipids lateral ditrusion, I13 nonbilayer-forming, 49 (table) proteinllipid ratio. 34-35 transniembrane movement of cholesterol, 191

291 diac ylglycerol~192 phospholipids. 189-191 X-ray diffraction, 32-33 ESR. S ~ Electron P spin resonance Ethanolaniine plasmalogen, biosynthcsis, 149-150 Exocytosis definition, 205-206 liposome fusion as model of, 212-213 pathways for membrane proteins, 280-281 quulernary structure role, 280-281 for mretory proteins, 279-28 I phospholipid metabolism and, 225 secretion during. 209 specific localization calelectrin and, 220 synexin and. 219-220 stages of. 206-207 triggering by Ca" influx. 213-216 adrenal medullary cells, 214-215 phosphatidylinositol turnover and. 214 presynaptic membranes, 213-214 diacylglycerol, 215 ligand-receptor interaction, 21.5

F Patty acids n-(!knthroyloxy), fluorescence decay anisotiopy. 7-8 cis-unsaturated, membrane fusion induction, 224-225, 23.5 desaturation, 141-144 microsomal enzymes, 142 n-doxylstearic spin-labeled. ESR assay. 8 e tonga tion, I4 I - 144 microsomal enzymes, 142 nomenclature, 130-1 3 I in phospholipids, 138-140 composition in subcellular membranes, 139-140 specificity in phospholipid synthesis. 156 transfer into adipocytes. 192 Fc receptor. in niacrophage endosomcs association with internal vesicles, 274- 275 transport to lysosomes, 273

292

INDEX

Filipin, complexes with cholesterol, 42 distribution in membranes, 4 2 4 3 Fluorescence probes for diffusion assay, 92 rotational diffusion study, 93-94 Fluorescence photo-bleaching and recovery (FPR), lateral diffusion study artifactually low values, 110-111 basic technique, 106-107 coefficients for acetylcholine receptor, 109-1 10, L14 glycoproteins, 109-109 lipids, 109 variations, 107-108

I IMP, see Intramembraneous particles Influenza virus, fusion glycoprotein effects, 220-221 mechanism of, 239 Intramembraneous particles (IMP) with one major protein, distribution, 3335, 50-51 with several major proteins distribution electric field and, 57 lectin receptor cross-linking and, 56 statistical analysis, 52, 54-55 identification methods, 51-53 Iron, colloidal, in IMP identification, CHO cells, 54

G

Glucagon as amphiphilic peptide. 58, 60 complex with lipids, 58-59 GIycerofipids nomenclature, 131-133 stereochemistry , 132 sn-Glycerol 3-phosphate acylation to phosphatidic acid, 144-145, 146 Glycerophosphate acyltransferase, microsornal and mitochondrial, 144-145 Glycophorin amino acid sequence, 58, 60 in reconstituted vesicles, 58 Glycophorin A, in erythrocytes as receptor for Sendai virus, 220, 222 GI ycoproteins lateral diffusion coefficients, 108-104 in lysosomal membranes, 278, 279 spike, viral, role in fusion, 220-221, 239, 244 Gold, colloidal, in IMP identification, 5152

H Halobacteriitm halobium, proteins in purple membrane, 58-59 Hydrogen ion, in endosomes, virus fusion triggering, 216

L Lateral diffusion, in membranes control by cytoskeleton, I 1 1-1 14 mechanism of, 112, 114 membrane protein concentration. 111

erythrocyte, 112-1 13 free-living cells, I I5 uncoupler effects, 116 function coupling reactions and, 117-1 18 in mitochondria1 electron transport, 118-1 19 dimensionality and, 117 methods frequencies of molecular collisions, 103 large-scale label distribution, 103-108 FPR, 106-108 in fused cells, 104 relaxation after electrophoresis in siru, 104-106 native and synthetic membranes, 114 restriction in cells in tissues, 115-1 16 junctional complex and, 115-1 16 Lectin receptors, IMP redistribution and, 56-57 Lincoleic acid, desaturation and elongation, 143-144

293

INDEX

Lipid bilayers conditions for formation, 5-6 disruption points. 45-46 fusion charge neutralization, ionic conditions and, 230-232 defect formation, 234-237 Ca' '-induced lateral phase separation, 235 content leakage and. 236 high- intensity electric field pulses and, 236 hydrophobicity and, 235 osmotic swelling and, 236-237 dehydration, 229-230 interaction energies, 227-229 soluble protein-mediated, 237-239, 244 spherical vesicle deformation, 232-234 heterogenous structures, 35-43 cholesterol effect, 3 9 4 0 cholesterol-enriched areas. 42-43 I M P distribution, 40-42 in bacteria, 40-41 ripple formation, electron microscopy diffraction contrast method, 37-39 freeze-fracture, 38-39 matching cross sections for head groups and acyl chains. 4 3 4 order parameter profile, 7-9 phase transitions, 9. 3 6 3 7 protein insertion into, 63 Lipids crystal structure N M R , 2-3 X-ray diffraction, 2-4 effects on protein conformation, 16-17 hexagonal tube phase, 4-6 lateral diffusion coefficients, 109 micelles, 4-6 mixtures binary model membranes. 10-1 I biological membranes, 11-1 2 nonbilayer structures, 45-50 electron microscopy. 45. 47 in energy transducent membranes, 47, 49-50 protein role in formation of. 4 7 4 8 in trypsin-digested Mycoplmnrtr i ~ r p s i cu)n membrane, 4 7 4

protein effects acyl chain packing and, 19-20 binding to protein surface, 18-19 immobilized fraction and, 17-18 rotational diffusion, 93 Lipid vesicles. S C P cilso Liposomes deformation before fusion. 232-234 fusion. 209-213 with each other content mixing, 212 as exocytosis model, 212-213 methods, 212 size changes and. 21 I interaction with cells, 210-21 I content transfer, 21 I phospholipid spontaneous transfer, I82 random movement into apposition. 217218 small unilamellar. cholesterol exchange, 179- I80 transbilayer movement of cholesterol. 192 phospholipids, 189 Lipophilin. in myelin, irregular distribution, 33-34, 5 I Liposomes, see olso Lipid vesicles cytochrome P-450. rotational diffusion, 100-101

fusogenicity. effects of cholesterol. 227 fluidity, 225-226 lateral stress distribution, 226 polyethylene glycol, 235 osmotic swelling and, 235, 236 soluble proteins, 237-239, 244 Lysosomes acidification, 277 ATP-driven proton pump. 277 biogenesis. 278-279 bis(monoacylg1ycero)phosphate biosynthesis. 155- I 56. I64 location, 136- I37 fusion with coaled vesicles in i i r r . 0 , 22 I hydrolases, origin, 278-279 isolation from recycling pathway. 277 membrane composition, 277-278 glycoproteins, 278, 279

294

INDEX

M Melittin, high-resolution study, 14 Membrane fusion, see ctlso Exocytosis, Viral fusion artificial membranes, 209-213, see ulso Lipid vesicles models of, 240-242 barriers to, 204-205 biomembranes freeze-fracture microscopy, 208209 hypothesis, 242-243 stages of, 206-207 definition, 205-206 interactions between bilayers, see Lipid bilayers phospholipid motions artificial lipid membranes. 225-227 biomembranes, 223-225 steric constraints, membrane proteins and, 222-223 Membranes, see ~ l s ospecific membranes biological, similarity and diversity, 30-35 diffusion of components lateral, 102-1 19; see also Lateral diffusion probes for, 92 rotational methods, 93-95 of proteins, 95-102 flow, phospholipid transport and, 165I66 Mitochondria1 membranes lateral diffusion function, 118-1 19 lipids biosynthesis enzymes. 164 composition, 134, 137-138 nonbilayer-forming, 49 (table) Mycoplosmu cripsiciim membrane, lipids structure, 47-48 Myelin nonbilayer-forming lipids, 49 (table) structure electron microscopy, 30-31 freeze-fracture, 33-34 lipophilin distribution. 33-34. 51 proteinflipid ratio, 34-35 X-ray diffraction, 32-33

N Nuclear magnetic resonance ( N M R ) lipid bilayer properties, 7 lipid structure, 2-3 probes for diffusion assay, 92 protein-lipid interactions, 18, 20 5'-Nucleotidase, cholesterol depletion effect. 182

0 Oleic acid, desaturation and elongation, 142- 143 Osmotic swelling, condition for fusion of cell membranes, 236 chromaffin granules with plasma membranes, 237 liposomes with planar bilayers, 236

P Palmitoleic acid, desaturation and elongation, 142-143 Phagocytosis, 256-257 Phosphatidate phosphohydrolase, diacylglycerol formation, 147-148 Phosphatidic acid conversion to CDPdiacylglycerol, 152153 formation by acetylation of d ihydrox yacet one, 145- I 46 sn-glycerol 3-phosphate. 144-145, 146 hydrolysis to diacylglycerol, 147-148 Phosphatidylc holine bios y nthesis CDPcholine formation, 148-149 choline phosphorylation, 148 diacylglycerol combination with CDPcholine, 149 phosphatidylethanolamine methylation, 150 content in membranes. 135 fatty acid composition, 138 in model membranes, 10-11 phase transition in bilayers, 9 pyrene-labeled, transfer between membranes, 184 X-ray crystallography, 3-4

INDEX

295

Phosphatidylcholine-specifictransfer protein loca I i zat ion, I86 phospholipid transfer mediation. 187I88 purification and identification, 1x6-187 Phosphatidy lethanolamine bios y nthesis CDPethanolamine formation. 148-149 diacylglycerol combination with CDPethanolamine, 149 ethanolaniine phosphorylation, 148 phosphatidylserine decarboxylation, 15 1- 152 content in membranes, 135-136 fatty acid composition, 138 methylation to phospha1idylchr)line. I50 X-ray crystallography. 3-4 Phosphatid ylgl ycerol hiosynthesis. 154-155 content in membranes. 136 Phosphatid ylinositol biosynthesis. microsomal. 153-154 content in membranes, 136 fatty acid composition, 138-139 turnover, exocytosis and. 214, 22s Phosphatidylserine biosynthesis. 150-1.5 I content in membranes, 136 decarboxylation to phosphatidylethanolamine. 151-152 fatty acid composition. 138-139 Phosphatidylserine decarboxylase. mitochondrial, 163 Phospholipids asymmetry in membranes, 140-141 hiosynthesis CDPdiacylglycerol pathway, 152-156 deacylation-reacylation cycles, 1561.57 diacylgl ycerol pathway, 147- I50 enzymes, 159-161 (table) tOpology. 162-164 ether bond formation, 146-147 Patty acid specificity, 156 nitrogen-containing, interconversion.

ISO-l52 pathways. 158 (table)

phosphatidic acid formation by acylation of dihydroxyacetone phosphate, 145I46 srr-glvcerol 3-phosphate. 144-145. I46 content in membranes, 133-138 acidic. 136- I37 cholesterol. 137 comparative data. 134 (table). 137-138 nitlogen-containing. 135-136 fatty acid composition, 138-140 motions in biomembranes, 223 membrane fusion and, 224-225 transfer between membranes biosynthesis and, 163-164 iiiembrane flow and. 165-166 protein-mediated, 164-1 6s. 186-189 mechanism, 187-188 physiological role, 188-189 spontaneous aqueous diffusion mechanism. 1x2185

pyrene-labeled. fluorescence assay, 183- I85 transmembrane movement factors affecting rate of. 190 measurements, 189- I90 protein involvement. 190-191 Phospholipid/water systems. types of molecular packing, 44-45 Photoreceptor disk membrane, retinal electron microscopy, 3 l L 3 I freeze-fracture, 33-34 nonbilayer-forming lipids. 49 (table) proteinilipid ratio, 34-35 rhodopsin distribution. 33-34. 5 I X-ray diffraction, 32-33 Pinocytosis, 256-257 Pits. clathrin-coated conversion t o coated vesicles. 261 nonreceptor proteins. 264 receptors for LDL, mutations, human, 263, 264 -1igand intei-actions. 263-264 IocaliLation. 261-263 structure and assembly. 260-261 Plasma membrane cholesterol depletion, 5'-nucleotidase activation, 182

INDEX

internalization and recycling, 257 sphingomyelin biosynthesis, 152, 164 vesicles sorting of from content, 257-258 Polyethylene glycol liposome fusion induction, 235 osmotic swelling and, 235, 236 Proteinflipid ratio, biomembrane comparison, 34-35 Proteins membrane effects on lipids acyl chain order and, 19-20 binding to protein-lipid surface, 1819 immobilization, 17-18 exocytosis constitutive pathway, 28028 1 insertion into lipid bilayers, 63 lipid composition effects on conformation, 16-17 rotational diffusion measurements, 95-102 optical methods, 93-94 saturation-transfer ESR, 94-95, 98, 100- 101

R Rhabdoviruses, fusion, G protein and, 221 Rhodopsin, bacterial, see Bacteriorhodopsin Rhodopsin. retinal lateral diffusion, 114 in photoreceptor disk membrane, 33-34, 51 rotational diffusion, 97-98 S

Sarcoplasmic reticulum Ca” -ATPase irregular distribution. 33-34, 51 lateral diffusion, 114 electron microscopy, 30-31 freeze-fracture, 33-34 nonbilayer-forming lipids, 49 (table) proteinllipid ration, 34-35 X-ray diffraction, 32-33 Secretory vesicles, movement into apposition directed, cytoskeleton role, 216-217, 219, 221 random, mathematical models, 2 17-219 Sendai virus, fusion glycophorin A as receptor in erythrocytes, 220, 222 membrane protein immobilization by, 222-223 spike glycoproteins and, 220, 244 Sphingomyelin biosynthesis in plasma membranes, 152, I64 spontaneous transfer between membranes, 184, 185 Synaptic vesicles, exocytosis sequence, 63, 64 Synexin Ca2+-dependent exocytosis and, 2 19-220 large vesicle fusion and, 239

structure, 12-16 a-helical, 13-15, 59 conformation, 57-62 hydrophobicity and hydrophobic moment, 15-16 tryptophan residue location, 15 membrane fusion and, 222-223 phospholipid transfer, 164-165, 186-189 classes of, 165 effects on membrane composition, 188 localization. 186 mechanism of action. 165, 187-188 membrane biogenesis and, 188-189 purification and identification, 186-187 phospholipid transmembrane movement and, 190-191 secretory, exocytosis regulated pathway, 279-28 I soluble, bilayer fusion mediation, 237- 239 in liposomes, 244-245 T Pyrene phospholipid labeling, tranfer study, Triacylglycerol, diacylglycerol conversion 183-184 to, 150 spontaneous transfer between memTryptophan, in membrane protein, locabranes, 185 tion, 15

INDEX

297 V

X

Viral fusion. see crlso speci$c virrtsrs random movement into apposition. 219 spike glycoproteins and. 220-22 I 244 stages of, 206-207 steric constraints, 222-223 triggering by H ' increase, 216 ~

X-ray diffraction bacteriorhodopsin, 2 electron density profiles for membranes, 3 1-33 lipid bilayers, 6 lipids, 2 4

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Contents of Recent Volumes Volume 19 Structure. Mechanism, and Function of the NalK Pump

PART I. THERMODYNAMIC ASPECTS OF MEMBRANE TRANSPORT What is a Coupled Vectorial Process'? WIILIAM P. JHNCKS Thc Membrane Equilibrium with Chemical Reactions FKIEI)KICH A. SALIIZR PART 11. STKUCTURAI, ANALYSIS OF Na. K-ATPase Structural Aspects of Na.K-ATPase ROBERT L. POST Detergent Solubilization of Na.K-ATPase MlKAEL ESMANN Methods for the Cleavage of the Large Subunit of Na.K-ATPase and the Resolution of the Peptides Produced HI:NKYRODRIWJEZ.RICHAKD A N D JACK KYTE HAKKINS, Selective Purification of Na.K-ATPase and Ca" .Mg"-ATPase from Eel Electroplax I.. M. AMENDI:.s. P. CHOCK.A N D R . W. AIJERS High-Performance Gel Chromatography of Horse Kidney Na-K-ATPase NAKAO. MAKOTO NAKAO,TOSHIKO T O M D K O OHNO, YOSHIHIRO

FUKUSHIMA, YUKICHIHARA,AND HASAKO ARAI

Native Membranes trom Dog Kidney Oute r Medulla, Enriched in Na,K-ATPase. and Vesicular in Nature BLISSFORBUSH 111

Ultrastructure of Na,K-ATPase in Plasma Membranes Vesicles AHVID B. EI.ISAHETH SKKIVEK. MALINSRACII. A N D PETEK LETH JC9RGI:NSEN Electron Microscope Analysis of Two-Dimensional Crystals of Membrane-Bound Na,K-AlPnse B . MAUNSBACH. EI.ISARI,TH AKVII) SKKIVLK. HANSHEHrKr. A N [ ) Pl-:I'l:K LlTH JQKCiliNSEN

Organiration of the Transmembrane Segment\ of Na,K-ATPase. Labeling of Lipid Embedded and Surface Domains of the uSubunit ;ind Its Tryplic Fragments with I "'IIlodonaphthylazide. I"P]ATP. and Photolabeled Ouahain PBII:KL m i J~RGENSEN. STEVEN J. I). KAIWSH, A N D CARLOS GITI.EW Structural Studies on Lamb Kidney Na.KATPase J . H. COI.I.INS, BLISSFORHIJSH 111, L. K. LANE.E. LING.ARNOI.I) SCHWAK'IY. A N D A. (RMvES) Zo'r Two Slightly Different a-Subunit Components of Kidney Na,K-ATPase Induced by Heat Ti-eatment 1'. OHTA,M. KAWAMUKA. T. A W A . H. ISHIKUKA, A N D K. N AGANO

Radiation Inactivation Analysis of Na.KA'T'I':QLP PAUL-O~-I.OI.ENC;HI. J. c l . l V l i kl.l.ORY. AN11 ROGERA. KIXIN Stoichiometrical Binding of Ligands to Less than 11% Kilodaltons of Na.KArPase H. H. MATSIII.Y. HAY.ASHI. A N U M. TAGLIC'HI HOMAREDA. The Active Site Structure of Na.K-

299

300 ATPase: Location of a Specific Fluorescein Isothiocyanate-Reactive Site CYNTHIA T. CARILLI,ROBERTA. FARLEY, AND LEWISC. CANTLEY Subunit Distribution of Sulfhydryl Groups and Disulfide Bonds in Renal Na,KATPase M. KAWAMURA, T. OHTA,AND K. NAGANO Lipid Regions of Na,K-ATPase Examined with Fluorescent Lipid Probes KIMBERLY A. MUCZYNSKI, WARDE. HARRIS,AND WILLIAM L. STAHL Role of Cholesterol and Other Neutral Lipids in Na,K-ATPase J. J . H . H . M . D E P o N T , W . H . M . PETERS, AND s. L. BONTING PART 111. LIGAND INTERACTIONS: CARDIAC GLYCOSIDES AND IONS Cardiotonic Steroid Binding to Na,KATPase 111 BLISSFORBUSH Binding of Monovalent Cations to the Na,K-ATPase M. YAMAGUCHI, J. SAKAMOTO, AND Y. TONOMURA Half-of-the-Sites Reactivity of Na,KATPase Examined by the Accessibility of Vanadate and ATP into Enzyme-Ouabain Complexes 07To HANSEN Binding of Rb' and ADP to a PotassiumLike Form of Na,K-ATPase JORGEN JENSENAND PAUL OTTOLENGHI Side-Dependent Ion Effects on the Rate of Ouabain Binding to Reconstituted Human Red Cell Ghosts H. H. BODEMANN, T. J. CALLAHAN, H.REICHMANN, A N D J. F . HOFFMAN Intracellular Sodium Enhancement of Ouabain Binding to Na,K-ATPase and the Development of Glycoside Actions TAIAKERA,KYOSUKETEMMA, AND SATOSHI YAMAMOTO Lithium-Catalyzed Ouabain Binding to Canine Kidney Na,K-ATPase GEORGER. HENDERSON

CONTENTS OF RECENT VOLUMES

Ouabain Binding and Na,K-ATPase in Resealed Human Red Cell Ghosts AND P. K. LAUF D. G. SHOEMAKER Stereoelectronic Interaction between Cardiotonic Steroids and Na,K-ATPase: Molecular Mechanism of Digitalis Action F. DITTRICH. P. BERLIN, K. KOPKE. AND K. R. H. REPKE Use of Prophet and MMS-X Computer Graphics in the Study of the Cardiac Steroid Receptor Site of Na,K-ATPase DWIGHTS. FULLERTON, DOUGLAS C. ROHRER,KHALIL AHMED,ARTHURH. L. FROM, EITARO KITATSUI1. AND TAMBOUE DEFFO Photoaffinity Labeling of the Ouabain Binding Site of Na,K-ATPase E. CLIFFORD c. HALLAND ARNOLD RUOHO New Ouabain Derivatives to Covalently Label the Digitalis Binding Site BERNARD Rossi, MAURICE GOELDNER. GILLESPONZIO, CHRISTIAN HIRTH, AND MICHELLAZDUNSKI Ouabain Sensitivity: Diversity and Disparities JOHN S. WILLISAND J. CLIVEELLORY PART IV. LIGAND INTERACTIONS: NUCLEOTIDES, VANADATE, AND PHOSPHORY LATlON Ligand Interactions with the Substrate Site of Na,K-ATPase: Nucleotides, Vanadate, and Phosphorylation JENSG . N0RBY Conformational Changes of Na,K-ATPase Necessary for Transport LEWISC. CANTLEY, CYNTHIA T. CARILLI, RODERIC L. SMITH. A N D DAVIDPERLMAN On the Mechanism behind the Ability of Na,K-ATPase to Discriminate between Na' and K' JENSCHR. SKOU Characteristics of the Electric Eel Na,KATPase Phosphoprotein ATSUNOBU YODA AND SHlZUKO YODA Sulfhydryl Groups of Na,K-ATPase: Effects of N-Ethylmaleimide on Phosphory-

30 1

CONTENTS OF RECENT VOLUMES

lation from ATP in the Presence of Na * + Mg' MIKAEL. ESMANN A N D IRENAKi.oims Alternative Pathways of Phosphorylation of Na,K-ATPase Regulated by Na ' Ions on Both Sides of the Plasma Membrane HORSTWALTER Structurally Different Nucleotide Binding Sites in Na,K-ATPase HERMANNKOEPSELLA N D DORISOLLlCi Study of Na,K-ATPase with ATP Analogs WILHELMSCHONER, HAKTMLIT PAUI.S, ENCINH. SEKPERSU. GEROLD REMPETEKS.ROSEMARIE PATZRI.TWENCZLER. AND MARIONHASSELHERG Affinity Labeling Studies of the ATP Binding Site of Canine Kidney Na,K-ATPase JAMESB. COOPER.CARLJOHNSON, A N V CHARLES G . WIN'rEK "P['*O]NMR Kinetic Analysis of InOExchange Reaction between P, and H,O Catalyzed by Na,K-ATPase A. STEPHENDAHMS AND Jotuf; E. MIAKA PART V. CONFORMATIONAL CHANGES, STRUCTURE/FUNCTION, A N D ACTIVE SITE PROBES Principal Conformations of the a-Subunit and Ion Translocation PETERL. J ~ K C E N S E N Magnesium-Induced Conformational Changes in Na,K-ATPase S. L. BONTING,H. G . P. SWARTS, W. H. M. PETERS,F. M. A. H . ANI) J . J. SCHUURMANS STEKHOVEN. H. H. M. DE PONT Ruhidium Movements in Vesicles Reconstituted with Na,K-ATPase, Measured in the Absence of ATP and P,, in the Presence of Either Ligand, and in the Presence of Both Ligands: Role of the "Occluded State" in Allowing for the Control of the Direction of Ion Movements s. J . D. KARLISHAND W.D. SI'EIN Eosin: A Fluorescent Probe of ATP Binding to Na-K-ATPase J. C. SKOU AND MIKAELESMANN Interaction of Divalent Cations with Fluorescein-Labeled Na.K-ATPase

MARCIASTEINHERC. JAMESG. KAPAKOS,A N D PAKIMAI. c . StiN Cation Activation of Na.K-ATPase after Treatment with Thimerosal MANISHAD. MONE AND JACK H. KAPLAN Alteration of Conformational Equilibria in Na.K-ATPase by Glutaraldehyde Treatment E. ELHANANY. R. D A V I DM . CHIPMAN, BEKGER.A N D A. LEV Conformational Transition between ADPSensitive Phosphoenzyme and PotassiumSensitive Phosphoenzyrne TANICUCHI. K U N I A KSLMUKI. I KAZUYA A N D SHOICHI I l D A

Relation between Red Cell Membrane Na,K-ATPase and Band 3 ERICT. FOSSEI.A N D A. K. SOI.OMON PART VI. REACTION MECHANISM A N D KINETIC ANALYSIS Kinetic Analyses and the Reaction Mechanism of the Na.K-ATPase JOSEPHD. ROI3lNSON Evidence for Parallel Pathways of Phosphoenzyme Formation in the Mechanism of ATP Hydrolysis by Electrophorus Na,K-ATPase JWFKEYP. FRORHIKH, A N NS. HOHBS,A N V R. WAYNE AI.Ht:RS Evaluation of the Reaction Mechanism of the Sodium Pump by Steady-State Kinetics JOHN R. SACHS Kinetic Evidence in Favor of a Consecutive Model of the Sodium Pump D. A. EISNER AND D. E. RICHAKUS Kinetic Models of Na-Dependent Phosphorylation of Na,K-ATPase from Rat Brain STANLEY J. DONALDM. FOSTER, RUSSELL,AND KHALII.AHMEI) Reinvestigation of the Sequence of Sensitivity of Phosphoenzyme of Na.K-ATPase to ADP and K ' during the Presteady State of the Phosphorylation by ATP Y . FUKUSHIMA A N D M. NAKAO Interaction of Na' , K ' , and ATP with Na. K-ATPase P. J . GAKKAHAN. R. ROSSI, A N D A. F. RF.Ci.4

302 Sodium Ion Discharge from Pig Kidney Na,K-ATPase YUKlCHI HARAA N D MAKOTONAKAO ADP Sensitivity of the Native and Oligomycin-Treated Na,K-ATPase ANNS. HOHHS,R. WAYNEALHERS, A N D JEFFREYP. FROEHLICH Three (at Least) Consecutive Phosphointermediates of Na-ATPase I. KLODOS. J. G . NORBY. A N D N. 0 . CHRISTIANSEN Aspects of the Presteady State Hydrolysis of ATP by Na,K-ATPase A. C . LOWEA N D L. A. REEVE Identity of the Na Activation Sites in ATPase with the K Activation Sites in p-Nitrophen ylphosphatase L. A. PARODl, J . F. PINCUS. L. JOSEPHSON. D. J. SORCE, A N D S. R. SIMON On the Existence of Two Distinct Hydrolysis Cycles for Na,K-ATPase with Only One Active Substrate Site ICOR W. PLESNER Kinetic Analysis of the Effects of Na' and K' on Na,K-ATPase A N D ICOR W. LISELOITE PI.ESNER PLESNEK Divalent Cations and Conformational States of Na,K-ATPase JOSEPHD. ROBINSON

PART VII. ION TRANSLOCATION AND REACTION MECHANISM Na,K-ATPase: Reaction Mechanisms and Ion TrdnSlOcdting Steps PAULDE WEER Existence and Role of Occluded-Ion Forms of Na,K-ATPase I . M. GLYNN A N D D. E. RICHARDS Na and K Fluxes Mediated by ATP-Free and ATP-Activated Na.K-ATPase in Liposomes BEATRICE M. ANNER Sidedness of Cations and ATP Interactions with the Sodium Pump L. BEAUGB AND R. DIPOLO Sidedness of Sodium Interactions with the

CONTENTS OF RECENT VOLUMES

Sodium Pump in the Absence of K ' RHODABLOSTEIN Magnesium Dependence of Sodium PumpMediated Sodium Transport in Intact Human Red Cells P. w.FLATMAN AND v. L. LEW K'hdependent Active Transport of Na' by Na,K-ATPase MICHAEL FORGAC AND GII.RERT CHIN ADP-ATP Exchange in Internally Dialyzed Squid Giant Axons PAULDE WEER.GERDAE. BREITWIESER, BRIANG. KENNEDY AND H . GILBERT SMITH Sodium Pump-Catalyzed ATP-ADP Exchange in Red Blood Cells: The Effects of lntracellular and Extracellular Na and K lons JACKH. KAPLAN Ouabain-Sensitive ATP-ADP Exchange and Na-ATPase of Resealed Red Cell Ghosts J. D. CAVlERES Effect of Internal Adenine Nucleotides on Sodium Pump-Catalyzed Na-Na and Na-K Exchanges BRIANG . KENNEDY, CORMLUNN, A N D JOSEPHF. HOFFMAN N d K Pump in Inside-Out Vesicles Utilizing ATP Synthesized at the Membrane E. ROBERTW. MERCEK,BEVERLEY FARQUHARSON, AND PHILIP B. DUNHAM Anion-Coupled Na Efflux Mediated by the NalK Pump in Human Red Blood Cells S. DISSINC AND J. F. HOFFMAN Effect of Trypsin Digestion on the Kinetic Behavior of the Na/K Pump in Intact Erythrocytes DONNAL. KROPP Sodium Movement and ATP Hydrolysis in Basolateral Plasma Membrane Vesicles from Proximal Tubular Cells of Rat Kidney F. PKOVERBIO, T. PROVERBIO. A N D R. MAKIN

Stoichiometry of the Electrogenic Na Pump in Barnacle Muscle: Simultaneous Measurement of Na Efflux and Membrane Current M. T. NELSONAND W. J. LEDERER

CONTENTS OF RECENT VOLUMES

PART VI11. HIOSYNTHESIS. MULI‘I PLE FORMS. AND IMMUNOLOGY Regulation of Na,K-ATPase by Its Biosynthesis and Turnover NOKMANJ. KARINA N D JOHNS. COOK Biosynthesis of Na.K-ATPase in MDCK Cells J. SHERMAN. T . MOKIMOIO,ANI) D. D. SABAIINI

Possible Functional Differences between the Two Na.K-ATPases of the Brain KATHI.EEN J. SWEADNEK Antigenic Properties of the a , p. and y Subunits of Na.K-ATPase WII.LIAM BALL..JH., JOHNH. COLLINS.L. K. LANE.AND ARNOLD SCHWARTT Antibodies to Na,K-ATPase: Characteriiation and Use in Cell-Free Synthesis Studies HIATT. A I K I AMCDONOUGH, ANDREW AND ISIIX>RE EDELMAN Inimunoreactivity of the a- and a( t )-Subunits of Na,K-ATPase in Different Organs and Species GHKAHI) D. SCH~LLENBEKG. I K I ~ NV.E PIX’H. A N D WII.l.IAM L. STAtil. Role of Na + and Ca” Fluxes in Terminal Differentiation of Murine Erythroleukemia Cells 1. G. MACARA. R. 0. SMITH. AND LEWISC. CANTI.I:.Y Na/K Pumps and Passive K ‘ Transport in Large and Small Reticulocytes of Anemic Low- and High-Potassium Sheep P. K. L A U FA N D G . VALET Enhancement of Biosynthesis of Na,KATPase in the Toad Urinary Bladder by Aldosterone But Not T, c. BRON. K. G I ~ E K I NM. ( ; .GIKAKDE.~, J.-P. K K A E H E N R ~AH N DL B. . C. ROWER Na.K-ATPase Activity in Rat Nephron Segments: Effect of Low-Potassium Diet and Thyroid Deficiency LAI.c. GAKG AN13 c. CRAIG TlSHEK Axonal Transport of Na,K-ATPase in Optic Nerve of Hamster SUSAN C. SPECHI

303 PART IX. Na.K-ATPase AND POSITIVE INOTKOPY; ENDOGENOUS GLYCOs I DES Positive lnotropic Action of Digitali\ and Endogenous Factors: Na.K-ATPase and Positive lnotropy; ”Endogenous Glycosides” SCHWARTL AKNOI.~) Endogenous Glycoside-Like Substances JK. GAKNER T. HAUPERT, Monovalent Cation Transport and Mechanisms of Digitalis-Induced Inotropy THOMAS w.SMITH A N D WILLIAM H. BAHKY Effects of Sodium Pump Inhibition on Contraction in Sheep Cardiac Purkinje Fibers W. J . LEDEKEK, AND R. D. A. EISNEH. D,VALIGHAN-JONES Quantitative Evaluation of [>H]Ouahain Binding to Contracting Heart Muscle, Positive Inotropy, Na,K-ATPase Inhibition. and ““Kh+ Uptake in Several Species EHIAND ERDMANN, 1,INI)SAY BROWN, KARI.WEKI)AN, A N D WOL.I‘GAN(I KKAWIETZ Contractile Force Effects of Low Concentrations of Ouabain in Isolated Guinea Pig. Rabbit, Cat. and Rat Atria and Ventricles GUNTER GHUPP. INGRID L. G R U P P , J . GHYSEL-BUKTON, T . GOI)T:HAINI). A. DE POVEK.A N D ARNOLDSCHWARTZ Difference of Digitalis Binding to Na,KATPase end Sarcolemma Membranes 1. KOROHANE.D. L. NANIN,A N D G. T. OKITA Pharmacological and Biochemical Studies on the Digitalis Receptor: A Two-Site Hypothesis for Positive Inotropic Action ARNOLD SC’HWARTZ. INGKII) I,. ~ R l l l l l ~ , ROBEKTJ. ADAMS,TREVOR POWELI.. GUNTER GKUPP.AND E. T. WAI.LICX Hypothesis for the Mechanism of Stimulation of the Na/K Pump by Cardiac Glycosides-Role of Endogenous Digitalis-Like Factor T. GOI~FKAIND. G. CASTANEDAHEKNANI)K%.J. GHYSE1:BURTON. A N D A . 111: Povr;K

CONTENTS OF RECENT VOLUMES

M. DELUISE,P. USHER, AND J. FLIER lmmunochemical Approaches to the Isolation of an Endogenous Digoxin-Like Factor Decreased Na,K-ATPase Activity in Erythrocyte Membranes and Intact ErythKENNETH A. GRUBER, JANICEM. rocytes from Obese Man WHITAKER, AND VARDAMAN M. DAVIDM. MOT, IWAR KLIMES,AND BUCKALEW, JR. RANDILL. CLARK Demonstration of a Humoral Na/K Pump Functionally Abnormal Na/K Pump in Inhibitor in Experimental Low-Renin HyErythrocytes from a Morbidly Obese Subpertension ject MOTILAL PAMNANI, STEPHEN HUOT, J. FLIER, P. USHER,AND M. DELUISE DAVIDCLOUGH, JAMESBUGGY,AND Specific Insulin Binding to Purified Na,KFRANCIS 3. HADDY ATPase Associated with Rapid Activation Absence of Ouabain-Like Activity of the of the Enzyme Na,K-ATPase Inhibitor in Guinea Pig JULIEE. M. MCGEOCH Brain Extract Mechanism for Cholinergic Stimulation of GEORGER. KRACKE Sodium Pump in Rat Submandibular Gland Brain Na,K-ATPase: Regulation by NorDAVIDJ. STEWART A N D AMARK. epinephrine and an Endogenous Inhibitor SEN ALANC. SWANN Evidence for an Aldosterone-Mediated, Inhibitory and Stimulatory Effects of Vanadate on Sodium Pump of Cultured Heart Na-Dependent Activation of Na,K-ATPase in the Cortical Collecting Tubule Cells from Different Species KEVINJ . PETTY, JUHAP. KOKKO,AND KARLWERDAN, GERHARD BAURIEDEL, DIANAMARVER AND ERLAND WOLFGANG KRAWIETZ, Vanadate and Somatostatin Having DiverERDMANN gent Effects on Pancreatic Islet Na,KEndogenous Inhibitor of Na.K-ATPase: “Endodigin” ATPase K. R. WHITMER, D. EPPS, AND KENJIIKEJlRl A N D SEYMOUR R. ARNOLD SCHWARTZ LEVIN Phosphorylation of a Kidney Preparation of Na,K-ATPase by the Catalytic Subunit of CAMP-Dependent Protein Kinase SVENMARDH PART X. PHYSIOLOGY AND PATHOModulation of Na,K-ATPase Activity in PHYSIOLOGY O F T H E N d K PUMP Rat Brain by Adenosine 3’,5’-Monophosphate RUSSELLB. LINGHAM AND AMARK. Disorders in Molecular Assemblies for Na SEN Transport in Essential Hypertension Stimulation and Inhibition by Plasma of Mirzv L. CANESSA.NORMAC. Ouabain-Sensitive Sodium Efflux in HuADRAGNA, ISABELBIZE, HAROLD SOLOMON, A N D DANIELc. TOSTESON man Red Blood Cells A. R. CHIPPERFIELD The Na-K Cotransport System in EssenInhibition of the Na Pump by Cytoplasmic tial Hypertension Calcium in Intact Red Cells R. P. GARAY,C. NAZARET, AND P. A. M. BROWNAND V. L. LEW HANNAERT Loss of Na,K-ATPase Activity during Cat- Involvement of Calmodulin in the Inhibition of Na,K-ATPase by Ouabain aract Formation in Lens LIONEL G. LELIEVRE, M. T. PIASCIK. PARIMAL c. SEN AND DOUGLAS R. J. D. P O ~ E RE.. T. WALLICK.AND PFEIFFER ARNOLD SCHWARTZ Na/K Pump: Effect of Obesity and NutriIndex tional State (

CONTENTS OF RECENT VOLUMES

lmmunolabeling of Frozen Thin Sections and Its Application to the Study of the Molecular Approaches to Epithelial Biogenesis of Epithelial Cell Plasma Transport Membranes I V A N EMANLIILOV IVANOV, H ~ o e PART 1. FREQUENCY DOMAlN PLBSKEN, 1)AVID D. SABATINI. A N I ) ANALYSIS O F ION TRANSPORT MICHARL J . RINDLEK Development of Antibodies to Apical Fluctuation Analysis of Apical Sodium Membrane Constituents Associated with Transport the Action of Vasopressin T . HOSHIKO J A M I ~ SB. WADE,VICI'OKIA GUCKIAN. Impedance Analysis of Nrctirrrts GallbladAN11 IN(IEB0RG KOIPPEN der Epithelium Using Extra- and IntracelMolecular Modification of Renal Brush lular Microelectrodes Border Maltase with Age: Monoclonal AnJ . J . L I M ,G. KOTIRA. L. KAMPMANN. tibody-Specific Forms of the Enzyme AND E. FROMrEK A N D Uzr REIss BERTRAMSACKTOR Membrane Area Changes Associated with Proton Secretion in Turtle Urinary Bladder PART 111. BIOCHEMICAL CHARACStudied Using Impedance Analysis TechTERIZATION OF TRANSPORT niques PROTEINS CHRIS CL.AUSEN A N D TROY E. DlXON Mechanisms of Ion Transport by the MamSodium-o-Glucose Cotransport System: malian Colon Revealed by Frequency DoBiochemical Analysis of Active Sites main Analysis Techniques R. KINNE, M. E. M. DACHUZ. AN11 N. K. W1Li.s J . T. LIN Analysis of Ion Transport Using FrequenProbing Molecular Characteristics of Ion cy Domain Measurements Transport Proteins P. SIMONA. LEWISA N D WILLIAM D.FANESTIL, RALPHJ . DAHRELL A I AS KESSLER,A N D C H U N SIK PARK Use of Potassium Depolarization to Study Aldosterone-Induced Proteins in Renal Apical 'I'ransport Properties in Epithelia Epithelia LAWRENCE G. PALMER MALCOLMCOX A N D MICHAEL. GliHEB PART 11. USE OF ANTIBODIES TO EP- Development of an Isolation Procedure for Brush Border Membrane of an Electrically ITHELIAL MEMBRANE PROTEINS Tight Epithelium: Rabbit Distal Colon B. P. MICHAEL C . GUSTINA N I ) DAVIL) Biosynthesis of Na',K'-ATPase in Am. G()OI)MAN phibian Epithelial Cells Itidex B. C. RossrEn Use of Antibodies in the Study of Volume 21 N a ' ,K '-ATPase Biosynthesis and Structure Ion Channels: Molecular and ALKIAM. MCDONOLKH Encounters with Monoclonal Antibodies to Physiological Aspects Na ' , K -ATPase Ionic Selectivity of Channels el the End MICHAEL. KASHGAKIAN, DANIEL. BIEMESDEKt'ER, AND BLISSFORBUSH111 Plate PlirrrR H. BAKKY A N D PETERW. G?zc;l,. Monoclonal Antibodies as Probes of EpiGating of Channels in Nerve and Muscle: thelial Cell Polarity A S t o c h a h c Approach GEORGE K. O.lAKl.4N A N D DORISA. RICHAKI) HOKN HEKZLINGER Volume 20

,I

'

CONTENTS OF RECENT VOLUMES

306 The Potassium Channel of Sarcoplasmic Reticulum CHRISTOPHER MILLER,JOANE. BELL, AND ANAMARIAGARCIA Measuring the Properties of Single Chdnnels in Cell Membranes H.-A. KOLB Kinetics of Movement in Narrow Channels DAVIDG. LEVITT Structure and Selectivity of Porin Channels R. BENZ Channels in the Junctions between Cells WERNER

Chloride in the Squid Giant Axon JOHNM. RUSSELL Axonal Calcium and Magnesium Homeostasis P. F. BAKERAND R. DIPOLO Regulation of Axonal pH WALTERF. BoKoN Hormone-Sensitive Cyclic Nucleotide Metabolism in Giant Axons of Loligo P. F. BAKER A N D A. CARKUTHhRS

PART 111. EXCITABILITY

R. LOEWENSTEIN

Channels across Epithelial Cell Layers SIMON A. LEWIS,JOHNW. HANRAHAN, AND w. V A N DRIESSCHE Water Movement through Membrane Channels ALANFINKELSTEIN Channels with Multiple Conformational States: Interrelations with Carriers and Pumps P. LAUGEK Ion Movements in Gramicidin Channels S. B . HLADKY A N D D. A. HAYDON Index

Volume 22 The Squid Axon

Hodgkin-Huxley: Thirty Years After H. MEWS Sequential Models of Sodium Channel Gating AND DONALDR. CLAYM. AKMSTRONG MATTESON Multi-Ion Nature of Potassium Channels in Squid Axons TEDBEGENISICH A N D CATHERINE SMITH Noise Analysis and Single-Channel Recordings FKANCO CONTI Membrane Surface Charge DANIELL. GILBERTAND GERALD

EHRENSTEIN

Optical Signals: Changes in Membrane Structure, Recording of Membrane PotenPART I. STRUCTURE tial, and Measurement of Calcium LAWHENCB B. COHEN,DAVID LANDOWNE. LESLIEM. LOEW, A N D Squid Axon Ultraslructure BRIANM. SALZBERG A N D RAIMUNDO GLORIAM. VILLEGAS Effects of Anesthetics on the Squid Giant VIILEGAS Axon The Structure of Axoplasm D. A. HAYDON,J . R. ELLIOTT, A N D B. RAYMOND J . LASEK M. HENDKY Pharmacology of Nerve Membrane Sodium PART 11. REGULATION OF T H E AXOChannels PLASMIC ENVIRONMENT TOSHIO NARAHASHI Biochemistry and Metabolism of the Squid Giant Axon HAROLDGAINER,PAULE. GALLANT, ROBERTGOULD,AND HARSH C. PANT Transport of Sugars and Amino Acids P. F. BAKERA N D A. CARRUTHERS Sodium Pump in Squid Axons Luis BEAUGB

PART IV. INTERACTION BETWEEN GIANT AXON AND NEIGHBORING CELLS The Squid Giant Synapse RODOLFOR. LLINAS

307

CONTENTS OF RECENT VOLUMES

Axon-Schwann Cell Relationship J o H w VII.I.OC~AS It1ttr x

The Proton- ATPase of Escli~vklriiic.oli A. E. SENKIK The Kdp System: A Bacterial K ' TI-ansport ATP'ise WOl.l'CiANCi b~I'STE1N

Volume 23 Genes and Membranes: Transport Proteins and Receptors

PART I. RECEPTORS A N D RECOGNITION PROTEINS Sensory Transduction in Bacteria MIIVIN I. SIMON. AI.EXANDRA KRIKOS.NORIHIRO MuT()fi. ANI) ALAN Bovo Mutational Analysis of the Structure and Function of the Influenza Virus H eniaggl ut i nin MARY-JANE GSTHING. CAROLYN I)OYt.e. MICHAEL ROTH,ANI) JOE

S A M U KOOK PART 11. C H A N N E L S Ca' ' Channels of Pmcrtncchm: A Multidisciplinary Study CHINGKUNG A N D YOSHIROSAIMI Studies of Skukcv Mutations Affecting a K ' Channel in Drosophilu L I L YYEH J A N , SANDRA BAKB~L, L t x i E TIMPE, CHERYL LAFFER, LAWRENCE S A I . K ~ FPATKICK F, ~ ' F A R R E AI ,N~D .YLIH , N U N GJAN Sodium Channels in Neural Cells: Molecular Properties and Analysis of Mutants WILLIAM A . CATTEKALL. TOHRCI GONOI. A N D MARIACOSTA PART 111. TRANSPORT S Y S T E M S The Histidine Transport System of Strlnionc4a ~ ~ p l ~ i m r r r i ~ i m GIOVANNA FERRO-LUZ'I AMES A Study of Mutants of the Lactose Transport System of E.sciic~ric,iiiucoli T. HASI'INGSWILSON. DONNA SEI'OYOLING. S Y l . V l E BEIIU, RESHAM. PLKZKATH. AND B E N N OMUI.I.I:R-HII.I.

Molecular Cloning and Characterization of ; I Mouse Ouabain Resistance Gene: A Genetic Approach to the Analysis of the N a ' .K ' -ATPase Rl)Bl:KT L b : V t i N S O N ltll~l~.r

Volume 24 Membrane Protein Biosynthesis and Turnover

Application of the Signal Hypothesis t o the Incorporation of Integral Membrane Proteins TOMA. KAIJ(.)POK'I'A N D M A K I I N W I ED M A N N Structure and Function of the Signal Peptide GtlY D. DtJl;FAUII, SUSANK. L.l~liNIiAKI)I'. PAUL. E. MARCH,A N D MASAYORI INOUYE The Use of Genetic 'Techniques to Analyze Protein Export in E.w/ierichiu coli VYIASA. BANKAIIW,J. PA-I'RIC.K K Y A N ,BETH A. RASMUSSEN, AND Ptiii.it1 J . BASSFORII.J K . Structural and Thermodynamic Aspects of the Transfer of Proteins into and across Membranes GLINNAR VON H E I J N ~ : Mechanisms and Functional Kole of Glycoaylation in Membrane Protein Synthesis SHARON S . KKAC Protein Sorting in the Secretory Pathway ENKIQUF, R I ~ I ) K I ~ ~ I ~ ~ - BDAVID ~uI.AN. E. M I S I ~ Kh .) K A VEGA DE SAI.AS. PEDKOJ. I. SALAS,AND ENZO BAND Transport of Proteins into Mitochondria GRAEMI'A. KI;II) Assembly of the Sarcoplasmic Reticulum during Muscle Development D A V I I ) H. MACI.ENNAN, ~l.l7.ABl~H ZUUKZYCKA-GAAKN. A N D ANNILISI: 0. JOR(iI
308 Receptors as Models for the Mechanisms of Membrane Protein Turnover and Dynamics H. STEVEN WILEY The Role of Endocytosis and Lysosomes in Cell Physiology YVES-JACQUESSCHNEIDER, JEAN-NOEL OCTAVE, AND ANDRBTROUET Regulation of Glucose Transporter and Hormone Receptor Cycling by Insulin in the Rat Adipose Cell IAN A. SIMPSON A N D S A M U E L w. CUSHMAN Index Volume 25 Regulationof Calcium Transport across Muscle Membranes

Overall Regulation of Calcium Transport in Muscle ADILE. SHAMOO PART I. REGULATION OF CALCIUM TRANSPORT IN PLASMA MEMBRANES Sarcolemmal Enzymes Mediating P- Adrenergic Effects on the Heart R. JONES LARRY Properties of Myocardial Calcium Slow Channels and Mechanisms of Action of Calcium Antagonistic Drugs NICK SPERELAKIS, GORDONM. WAHLER,AND GHASSAN BKAILY The Sarcolemmal Sodium-Calcium Exchange System JOHN P. REEVES PART II. REGULATlON OF CALCIUM TRANSPORT IN SUBCELLULAR ORGANELLES

CONTENTS OF RECENT VOLUMES

Role of Calmodulin in the Regulation of Muscle Contraction ANNIEMOLLA,SIDNEY KATZ,AND JACQUESG. DEMAILLE Calcium Release from Sarcoplasmic Reticulum MAKOTOENDO The Regulation of Mitochondria1 Calcium Transport in Heart MARTIN CROMPTON PART Ill. CELLULAR ION REGULATION AND DISEASE Cellular Ion Regulation and Disease: A Hypothesis BENJAMIN F. TRUMP A N D IRENE K.

BEREZESKY Index Volume 26 Na+-H+ Exchange, lntracellular pH, and Cell Function

PART 1. GENERAL ASPECTS OF INTRACELLULAR pH REGULATION AND Na'-H ' EXCHANGE Intracellular pH Regulation by Leech and Other Invertebrate Neurons R. c. THOMAS A N D w. R. SCHLUE Approaches for Studying Intracellular pH Regulation in Mammalian Renal Cells WALTERF. BORON Aspects of pH, Regulation in Frog Skeletal Muscle ROBERTw. PUTNAM AND ALBERT

Roos Molecular Properties and Physiological Roles of the Renal Na+-H' Exchanger AND PETER PETER s. ARONSON IGARASHI

PART 11. Na'-H' EXCHANGE AND CELL VOLUME REGULATION Regulation of Calcium Transport in Cardiac Sarcoplasmic Reticulum ADlL E. SHAMOO, ~ N D US. AMBUDKAR,Volume-Sensitive Alkali Metal-H Transport in Ampliiumu Red Blood Cells MARCS. JACOBSON, AND JEAN PETERM. CALA BIDLACK

309

CONTENTS OF RECENT VOLUMES

Na-Proton Exchange in Dog Red Blood Cells JOHN C. PARKER Activation of the Na '-H' Antiport by Changes in Cell Volume and by Phorbol Esters: Possible Role of Protein Kinase S. GRINSTEIN, S. COHEN,J. D. G o ~ r z , A. ROTHSTEIN. A. MELLORS,A N D E. W. GELPAND PART 111. Na'-H' EXCHANGE AND CONTROL O F CELL GROWTH The Generation of Ionic Signals by Growth Factors W. H. MOOLENAAR, L. H. K. DEFIZH. P. T. V A N DER SAAG. A N D s. w. DE LAAT Control of Mitogenic Activation of Na '-H ' Exchange D. CASSEL.P. ROTHENBERG, B. WHi.rELEY. D. MANCUSO. P. SCHI.ESSINGER. L. R ~ u s s E. . J. CRAGOE,AND L. GLASEK Mechanisms of Growth Factor Stimulation of Na '-H ' Exchange in Cultured Fibroblasts MITCHEL L. VILLEKEAL, LESLIEL. MIX-MULI)OON, LUCIAM. VICENTINI. GORDON A. JAMIESON. JK.. AND NANCYE. OWEN B Lymphocyte Differentiation: Role of Phosphoinositides. C Kinase, and Na '-H * Exchange PHILIP M. ROSOFFAND LLWIS C. CANTLEY Na '-H * Exchange and Growth Control in Fibroblasts: A Genetic Approach JACQUES POUYSS6C;UR. ARLEITE FRANCHI. MICHIAKI KOHNO,GILLES AND SONIA PARIS L'ALLEMAIN, PART IV. ROLE O F Na'-H' EXCHANGE IN HORMONAL AND ADAPTIVE RESPONSES Hormonal Regulation of Renal Na'-H' Exchange Activity SACKTOK AND JAMES L. BERTRAM KINSELLA Adaptation of Na'-Hi Exchange in the

Proximal Tubule: Studies in Microvillus Membrane Vesicles A N D RAYMOND C. JUI.IAN L. SEIFTER HAKRIS The Role of Intracellular pH in Insulin Action and in Diabetes Mellitus RICHAKD D. MOORE The Proton as an Integrating Effector in Metabolic Activation WII.I.IAM B . BUSA 1nde.r Volume 27 The Role of Membranes ln Cell Growth and Dlfferentiatlon

PART I. DESCRIPTION O F ION TRANSPORT SYSTEMS I N ACTIVATED CELLS Mitogens and Ion Fluxes L U I SREUSS.DANCASSEL,PAUL ROTHENBERG, BRIANWHITELEY, A N D L U I SG L A S F ~ K DAVID MANCUSO, Na '-H and Na'-Ca2 + Exchange in Activated Cells MrrcHEL L. VII.LEREA1. Chloride-Dependent Cation Cotransport and Cellular Differentiation: A Comparative Approach PETER K. LAUF +

PART II. TRIGGERS FOR INCREASED TRANSPORT DURING ACTIVATION External Triggers of Ion Transport Systems: Fertilization. Growth. and Hormone Systems JOANBELL, LORETTANIELSEN,SARAH SAKIBAN-SOHRABY. A N D DALEBENOS Early Stimulation of Nit'-H ' Antiport, Na'-K' Pump Activity, and Ca" Fluxes in Fibroblast Mitogenesis ENKIQUE ROZENCURT A N D STANLEY A . MENDOZA Volume-Sensitive Ion Fluxes in Amphirtmn Red Blood Cells: General Principles Governing Na-H and K-H Exchange Transport and CI-HCO, Exchange Coupling PETERM. CALA

31 0 PART 111. CONSEQUENCES O F T H E ALTERATIONS IN ION TRANSPORT OBSERVED DURING ACTIVATION IntracelIular Ionic Changes and Cell Activation: Regulation of DNA, RNA, and Protein Synthesis KATHIGEERING Energy Metabolism of Cellular Activation, Growth, and Transformation LAZARO J. MANDEL Index

Volume 28 Potassium Transport: Physiology and Pathophysiology PART I. C E L L MECHANlSM O F POTASSIUM TRANSPORT Role of Potassium in Epithelial Transport Illustrated by Experiments on Frog Skin Epithelium H. H. USSING Na , K ' , and Rb ' Movements in a Single Turnover of the Na/K Pump I11 BLISSFORBUSH Properties of Epithelial Potassium Channels DAVID c . DAWSON Role of Potassium in Cotransport Systems ROLFKINNEA N D ERICHHEINZ Functional Roles of lntracellular Potassium in Red Blood Cells JOSEPHF. HOFFMAN +

CONTENTS OF RECENT VOLUMES

GERHARD H. GIEBISCH Adrenal Steroid Regulation of Potassium Transport ROGERG. O'NEIL Potassium and Acid-Base Balance RICHARD L. TANNEN Renal Potassium Adaptation: Cellular Mechanisms and Morphology BRUCEA. STANTON Quantitative Analysis of Steady-State Potassium Regulation DAVIDB. YOUNG PART 111. RENAL AND EXTRARENAL CONTROL OF POTASSIUM: DlURETICS AND PATHOPHYSIOLOGY Regulation of Extrarenal Potassium Homeostasis by Insulin and Catecholamines RALPHA. DEFRONZO Diuretics and Potassium CHRISTOPHER S. Wrr,cox Pathogenesis and Pathophysiological Role of Hypoaldosteronism in Syndromes of Renal Hyperkalemia MORRIS SCHAMRELAN A N D ANTHONY SEEASTIA N H ypokalemia JORDAN J. COHEN Metabolism and Potassium JAMESP. KNOCHEL PART IV. POTASSIUM TRANSPORT IN MUSCLE AND COLON

Effects of Potassium Deficiency on Na,K HomeostasisandNa+,K*-ATPase inMuscle TOKEEN CLAUSEN AND KELD PART 11. RENAL AND EXTRARENAL KJELDSEN CONTROL OF POTASSIUM: Relationship between Cell Potassium and PHYSIOLOGY Hydrogen Ion in Muscle SHELDON ADLER Electrophysiology of Active Potassium Overview: Renal Potassium Transport Transport across the Mammalian Colon along the Nephron N . K. WILLS,C. CLAUSEN, A N D W.c. FREDS. WRIGHT CLAWS Potassium Recycling AND ROLAND MULLEK- Potassium Adaptation in Mammalian Colon REXL. JAMISON HENRY J. BINDEU. JOHNP. HAYSLETC, SUirR A N D MICHAEL KASHGARIAN Cell Models of Potassium Transport in the Index Renal Tubule