The spontaneous incorporation of proteins into preformed bilayers

Biochimica et Biophysica Acta 906 (1987) 33-68

33

Elsevier BBA 85302

The spontaneous incorporation of proteins into preformed bilayers Mahendra K. Jain

a

and David Zakim b

a Department of Chemistry, University of Delaware, Newark, DE and b Division of Digestive Diseases, Department of Medicine, Cornell University Medical College, New York, N Y (U.S.A.) (Received 13 May 1986)

Contents I.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

33

II.

Interactions of proteins with bilayers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Adsorbed proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Anchored proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Embedded proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

34 37 37 43

III. The phenomenon of spontaneous incorporation of embedded proteins into preformed bilayers . . . . . . . . . . . . . . . . . . . A. The mechanism of incorporation of proteins into preformed bilayers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Defects vs. general disorder . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. The state of proteins and bilayers in reconstituted systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

53 53 55 57

IV. Biologic implications of spontaneous incorporation of proteins into membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

60

V.

Epilog . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

61

Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

62

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

62

I. Introduction

Reconstitution of proteins into bilayers is required for establishing several aspects of the funcAbbreviations: D P G , diphosphatidylglycerol; lysoPC, lysophosphatidylcholine (or other lysophosphatide); PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PS, phosphatidylserine; ULV, unilamellar lipid vesicle. Correspondence: D. Zakim, Division of Digestive Diseases, Department of Medicine, Cornell University Medical College, New York, NY 10021, U.S.A.

tions of membrane proteins and lipids and for elaborating and articulating models of naturally occurring membranes. This is so because of the complexity of the composition of lipids and proteins in natural membranes and the lack of methods for determining directly the nearest neighbors of a given molecule of phospholipid or protein within a complex. Spectroscopic and biochemical methods, as, for example, immunochemical, enzymatic and binding assays, can be used to study natural biomembranes, but these techniques can provide considerably more information when applied to studies of reconstituted systems [1-6].

34 Moreover, high-resolution structural information about membrane proteins, e.g., X-ray crystallography [7], depends on placing the proteins in a two-dimensional lamellar matrix such that all molecules of the protein have identical orientations. Thus, incorporation of soluble or solubilized membrane-proteins into bilayers is necessary for isolation, identification, elucidation of mechanisms of action, and for elaboration of the significant functional interrelationships between membrane components. An especially appealing goal of reconstitution studies is the spontaneous insertion of purified membrane-bound proteins into preformed bilayers of well-defined composition and dispersity. Not only is this attractive esthetically, but studies of the spontaneous insertion of proteins into preformed bilayers could provide insights into the underlying kinetic and thermodynamic constraints on the assembly of membranes, as, for example, the mechanism of post-translational insertion of proteins into cellular membranes [8-10]. An understanding of the interactions between plasma proteins and liposomes also could lead ultimately to realizing the potential of liposomes for use as drug-delivery systems [11]. In spite of considerable progress in the methodology for reconstitution of functional lipid-protein complexes, a successful reconstitution still appears to be more art than science [2,4,12]. Most of the commonly used procedures for reconstituting membrane proteins involve removal of detergents from detergent-solubilized mixtures of lipids and protein, i.e., the bilayer is reconstituted around a 'solubilized' protein. Such systems have been useful in the study of membrane function, but they leave several questions unanswered. Some of these are intrinsic to the method of reconstitution. Others arise from technical difficulties posed by residual detergents, uncontrolled lipid compositions, and formation of particles of uncontrolled size and dispersity. Our goal in this review is to consider the conditions that have been used for incorporating membrane proteins into preformed bilayers of phospholipid. It is not difficult to achieve these conditions; yet, the extent to which this goal can be achieved has not received the attention it seems to deserve. In addition, data from experiments in which membrane proteins insert spontaneously into preformed bilayers indi-

cate that complex properties of a bilayer are involved in creating conditions that promote spontaneous insertion. Obviously, these must be defined if we are to understand how membranes are assembled. Therefore, a broad objective of this review is to elaborate the underlying constraints on spontaneous incorporation of proteins into bilayers, and to suggest that under appropriate conditions it may be possible to modulate these constraints in order to achieve spontaneous reconstitution of most membrane proteins into preformed bilayers of almost any composition.

II. Interactions of proteins with bilayers Examples of interactions between bilayers and proteins that have been studied are listed in Table I. The list is not exhaustive, but is meant to provide key references and to illustrate that the range of proteins interacting with bilayers is large in terms of function, source, and properties. The stoichiometry, function, and properties of reconstituted lipid-protein complexes also are quite varied, ranging from formation of stoichiometric lipid-protein particles, solubilization with phase change, and incorporation with retention of the bilayer organization. Membrane-bound proteins are classified as extrinsic or intrinsic to the membrane. Although this classification is useful, it does not reflect the fact that extrinsic proteins may be associated with membranes in more than one way. We prefer to consider that there are two types of extrinsic membrane-bound proteins: proteins that are adsorbed to the membrane and proteins that are 'anchored' to it. This classification is useful for the purposes of the ideas we review because we are concerned in a broad sense with the mechanisms leading to associations between membranes and proteins and because 'anchored' proteins may be 'mistaken' for intrinsic proteins. For example, that the binding of a protein to a membrane alters some bulk phase property associated with the apolar region of a membrane or that binding changes the function of the protein is no assurance that the protein has inserted into the apolar regions of a membrane. In addition, there is only limited direct information in most in-

35 TABLE I (continued)

TABLE I PROPERTIES BILAYERS

OF

PROTEINS

RECONSTITUTED

IN

Adenyl cyclase: Detergents needed for reconstitution; regulation of activity by cholesterol content [13-15].

Adrenocorticotropic hormone: Probably forms an amphiphilic helix at lipid/water interface [16]. Albumin: Fragment (307-385) promotes fusion of PC vesicles at low pH [17]. Aminopeptidase N: Reconstitution into vesicles is promoted by lysophospholipids [18]. Anion channel: Band III of erythrocyte membrane in proteoliposomes exhibits many of the properties of the channel [19]. Apolipoprotein A: Solubilization of DPPC [20], DMPC [21,22] and mixed lipid vesicles [23,24]. Kinetics of solubilization of vesicles [25]. Competitive removal by amphipathic peptides [26]. Kinetics of leakage and disruption [27]. Related peptides solubilize vesicles and form disk-shaped particles [28,29]. Apolipoprotein C: Solubilization of DMPC vesicles [30,31]. Asialoglycoprotein receptor: In DPPC reconstituted vesicles it does not cause leakage of carboxyfluoresceine although Tm is perturbed; CD and fluorescence changes indicate a conformational change [32]. ATPase, Ca2+-: Reconstituted in DMPC by cholate dialysis method [33]. Disrupts lipid domains in bilayers [34]. Effect of residual detergent on activity [35]. ATPase, (Na ++ K +): Reconstituted into BLM by fusion of liposomes (Ref. 36; see also Ref. 36a). Prefers anionic lipids [37,38]. ATPase, (Na++ Mg2)., from Acholeplasma laidlawii: Increases Tm of reconstituted vesicles [39]. Bacteriorhodopsin: Effect of lipid structure on the properties of reconstituted proteoliposomes [40]. Reconstituted by cholate dialysis method [41]. For results of spontaneous reconstitution method see text. General review on the properties of the protein see Ref. 42. Orientation of the protein backbone studied by NMR [43]. Thermotropic properties and evidence against boundary lipid [44] and on organization of lipids [45]. Deuterium-labelled amino-acid substitution used to define the aqueous interface of the lipid-protein complex [46]. Bungarotoxin: Fatty acid coupled molecule binds tightly to the bilayer and retains its activity [47]. Calcitonin: Solubilizes DMPG and D M P C + DMPG containing vesicles to form disk-shaped particles [48]. Clathrin: Does not penetrate the bilayer but appears to promote disorder of acylchains [49,50]; promotes fusion and leakage [51]. Ionic interactions may be involved in binding. Coat protein from fd phage: Transmembrane sequence data [52]. Coat protein, M13: Tendency to self-aggregate; 70 DMPC form boundary lipids; disorders the gel phase [53]. Ureacholate methods for reconstitution; motion of the protein is influenced by the gel phase [54].

Coat protein, M: The protein from stomatitis virus promotes phase separation of PC + PG bilayers [55].

Coat protein, pro-: Spontaneous transmembrane insertion [9]. Colicins: increase in helix content in bilayers (Review: Ref. 56).

Cytochrome bs: Prefers anionic lipids, but it can bind to zwitterionic lipids [57,58]. Promotes transbilayer movement of phospholipids in the early stages of reconstitution, presumably when the helix is not fully inserted [59]. Spontaneous insertion studies described in text. Cytochrome c: A spherical protein with eight net positive charges at pH 8 lowers Tm and enhances cation permeability [60], induces phase separations [61]. Interacts with anionic but not with zwitterionic lipids. Promotes bilayer to hexagonal II phase change in DPG, but not in PS and PG bilayers [62]. Cytochrome c, apo-: Transmembrane incorporation occurs [63]. Random coil structure, binds to anionic fipids like DMPG, and appears to assume greater helix content on binding to lipid [64]. Cytochrome oxidase: In DMPC vesicles 55 lipid molecules/ protein are immobilized [65], and 77 molecules in DMPC or DPG [66]; prefers DPG in mixed lipid systems [67]. Disorders acyl chains [68,69] and leakage [70] is due to residual detergent. See text for spontaneous reconstitution studies. Cytochrome P-450: In DMPC bilayer about 350 molecules do not participate in gel to fluid transition [57]. Preferentially binds to PA [71]. Duramycin: Apparently prefers PE and monogalactosyldiacylglycerol in bilayer [72], and inserts into microsomal membranes [73]. Dyanorphin: Fragment 1-13 undergoes a conformational change to helix on binding to PC bilayer [74]. Enkephalins: Bind to gangliosides and anionic lipids [75,76]. Factor Va: Reconstituted in DOPC+DOPA [77]; effect of salts and pH suggest ionic binding [78]. Fumarate reductase: Transbilayer sequence data for the protein from E. coli [55]. Glucagon: Solubilizes a variety of lipids to form disk-shaped particles [79,80]. Increases Tm and lowers enthalpy; heat of mixing is apparently due to a redistribution of gel and fluid phases [81]. Solubilization occurs at 10 mol% but not at 20 mol% cholesterol; particles are cholesterol-free [82]. Glucose transport protein: Reconstitution by reverse phase evaporation; activity depends upon the phase properties [83]. Hexagonal phase in the lipid is not necessary for reconstitution or activity [84]. Glycophorin: Transmembrane sequence of 23 residues is a hydrophobic helix. Reconstituted from premixed components in DMPC and mixed lipids shows a preference for fluid phase; lateral diffusion data show two populations [86]; DSC study [49,87]; shows some preference for PS [88]; reconstituted in BLM by freezing and thawing [84]. Stabilizes bilayer as large unilamellar vesicle [90,91] reconstituted systems are leaky [92] and exhibit a higher rate of

36 TABLE I (continued)

TABLE I (continued)

flip-flop [93]. In PCs it perturbs packing of the acyl chains and lowers enthalpy [94]. In PE retains bilayers at 1:25, and hexagonal II phase at > 1 : 50 [90]. Binding of wheatgerm agglutinin promotes hexagonal phase [91]. Glycoprotein of vesicular stomatitis virus: Reconstituted in POPC vesicles; shows an ordering effect [95]. Gramicidin A: A linear peptide which forms an asymmetric dimer across the bilayer with N-terminus in the middle [96,97] and increases gauche content [98] for PCs; effect of cholesterol [99]. Prefers fluid phase by a factor of three [100]. Promotes bilayer to hexagonal II phase change in unsaturated PE and PC [101,102], and forms a bilayer phase with LPC [103]. Halorhodopsin: Light-driven chloride pump that is uniformly distributed in Halobacterium membrane; the membrane fragments fuse with BLM [104]. Lactate dehydrogenase: The enzyme from E. coil exhibits 1.7fold increase in helix content on binding to lipids [105]. Lectins: Bind to globoside and induce fusion in the presence of calcium [106]. Review of binding to and agglutination of vesicles [107]. Lipophilin, see Myelin proteolipid. Mastoparan: Assumes helix conformation on binding to bilayer

Polyhistidine: Promotes fusion of PS+ PE vesicles [139,140]. Polylysine: Interacts with many anionic bilayers [60,141], and

[108]. Mellitin: Review [109]. Promotes fusion [110,111]. Lowers Tm in tetrameric form [112]. Disorder and rotational mobility of anion channel is reduced, presumably due to induced phase separation [113]. Reconstitution and aggregation in DSPC vesicles [114]. Myelin basic protein: Interacts with sphingomyelin [115] and gangliosides [116]. Requirement for anionic lipids or degradation products [117,118]; see text for details. A tryptic fragment capable of inducing allergic encephalitis interacts with anionic lipids [119]. NMR study interaction with anionic groups on bilayers [120]. Myelin proteolipid." Degradation product necessary for incorporation into DMPC vesicles [121,122]. Binds to PS vesicles [123]. Raman study [124,125]. ESR study of the motionally restricted lipids suggests preference for PA > DPG > PS > PG = PC > PE > steroids [126]. Parathyroid hormone, fragment 1-34: Solubilizes DMPC vesicles in the gel phase, and the bilayers above Tm remain intact [127]. Pentagastrin and related peptides: Increase ordering below Tm [128]; enhance leakage; effect seems to depend upon the primary sequence [129]. Peptides: Synthetic peptides of Lys2.Gly'(Leu),-Lys2"Ala amide (n =16, 20, 24) form transmembrane hydrophobic helix in DPPC bilayer [130]. The proportion of the protein in the fluid and the gel phases changes by varying the lipid-to-protein ratio [131,132]. Phospholipase A2: The enzyme from pig pancreas exhibits preference for anionic interface. It does not bind to the ether analog of PCs [133-137]. See text for details. Phospholipid exchange protein: Interaction with bilayers and the effect of the phase properties [138].

in DPG vesicles it protects from bilayer to hexagonal transition induced by calcium [62,142]. Polymyxin: Binds to bilayers of anionic lipids [143], and induces interdigitation [144]. See text for details. Proacrosin: Self-activated in the presence of anionic lipid vesicles [145]. Rhodopsin: Transbilayer sequence [42]; boundary lipid [146]. Signal peptide: From lambda receptor protein of E. coli interacts with lipid/water interfaces presumably as a hydrophobic helix [147]. Spectrin-actin: Attached to band III and glycophorin via coupling proteins like ankyrin and band 4.2. Spectrin is a basic protein that binds to DMPG [148] and PS, and causes fluid to gel transition. Different domains in the molecule are apparently involved in different interactions. Synexin: Facilitates fusion at low calcium concentration [150].

Toxins Review on translocation of toxins across cell membrane [151]; on channels formed by a-toxin, streptolysin, and C5b-9 complement complex [152]; predominant B-sheet structure in many toxins that form channels by aggregation [153]. Cardiotoxin IV from snake venom [154] promotes bilayer to hexagonal phase transition [155]. Cerebratulus lacteus toxin A-III: Lytic, forms channels [156]. Cholera toxin: Undergoes a conformational change on binding of the A chain to bilayer [157,158]. Binds to gangliosides [159,160]. Diphtheria toxin: The B-chain forms channels [161] and promotes translocation of chain A across the bilayer [162]. Binding to lipid induces conformational change to helix [162] and fusion [163]. Ionic interactions involved [164,165]. Endotoxin from Bacillus thuringiensis: Disorders bilayer of zwitterionic but not of anionic lipids [166]. Sea anemone toxin: Shows slight leakage in PC vesicles [167]; slight preference for SM [168]. Streptolysin O: Preference for cholesterol containing membranes [169]. Streptolysin S: Binds to most lipid but vesicles of the unsaturated lipids are more leaky; binding is promoted by the presence of cholesterol in saturated lipids [170]. Tetanus toxin: Binds to ganglioside containing vesicles [171,172]. Tubulin: Incorporated into vesicles at Tm with an increase in helix content. Causes leakage of carboxyfluoresceine [173].

s t a n c e s o n h o w m u c h o f a m e m b r a n e p r o t e i n is within a bilayer. Interactions between membranes and adsorbed proteins are dominated by ionic interactions. There

37 are, however, no specific binding interactions between the protein and the bilayer, and, the interactions are sufficiently weak that neither polar groups on the membrane nor protein become desolvated during adsorption. The underlying rate constants for formation of the complexes will be large, and the life-times of the complexes are expected to be quite brief. Proteins adsorbed to a bilayer will exchange readily between populations of vesicles, on a time-scale of less than a few seconds. For example, adsorbed proteins will not co-elute with vesicles when chromatographed on Sepharose 2B [133]. There are two mechanisms for anchoring of proteins to membranes. The first is specific binding of the protein to ligands that are components of the membranes, as carbohydrates and charged groups. The second is covalent attachment of a protein to a non-polar component of the membrane [47,174-177]. Interactions in the former mechanism involve local desolvation of the 'binding sites' on the proteins. The dissociation constant of an anchored protein is expected to be relatively small because it is the product of dissociation constants for several sequential steps. Spontaneous intervesicle exchange of 'anchored' proteins is expected to be slow. Exchange should be promoted, nevertheless, by salts or specific ligands that compete with the bilayer for binding sites on the protein. Perturbations within the hydrophobic region of the bilayer can affect anchoring of proteins to the interface and vice versa. Intrinsic proteins are embedded within the hydrophobic region of a bilayer. Insertion of these proteins, can take several forms. The protein may penetrate only partially into the bilayer. The protein may form a single looped structure in which C- and N-terminal portions are on the same side of the bilayer; or it may span the bilayer, i.e., Cand N-terminal ends are in water but on opposite sides of the bilayer; or the protein may span the membrane bilayer several times. The dissociation constant of an embedded or transmembrane protein is expected to be extremely small. Therefore, the rate of intervesicle exchange of an embedded protein will be immeasurably small (discounting fusion between vesicles), and will not be influenced by perturbations within the polar and apolar regions of the bilayer.

II.A. Adsorbed proteins Soluble proteins like polylysine, albumin, hemoglobin, lysozyme and ribonuclease change the surface pressure of monolayers of phospholipids (cf. Table I and references therein). Occasionally, they effect a modest change in the phase transition properties of bilayers [178]. Binding of these proteins to anionic lipids like PS, PG, DPG or phosphatidic acid increases Tm of the lipid phase. Polylysine prevents Ca2÷-induced phase changes, e.g., from bilayer to HII phase in bilayers of DPG [62,142]. The effects of soluble globular proteins, like albumin, on the leakage of ions across bilayers depend upon the presence of additives [179] or impurities present in commercial preparations of the proteins [11]. In addition, a fragment derived from proteolytic digestion of albumin apparently interacts with vesicles of PC to promote fusion between them [17]. These are important observations, because effects on the bilayer that have been attributed to proteins may truly reflect the presence of impurities. Thus the phase properties of bilayers of anionic lipids are perturbed strongly by water-soluble cations [180] that can produce isothermal phase changes and leakage of polar solutes trapped within vesicles. Unfortunately, dissolution of anionic lipids by cationic proteins, i.e., disruption of vesicles, has not been investigated carefully.

11.B. Anchored proteins The membrane per se presents no barrier to the association of adsorbed proteins, except in the sense that it does not contain appropriate charged groups for interacting with a given protein. Examples to be discussed show, however, that the structure of a membrane may present a barrier to association of 'anchored' proteins. That is, association does not occur even for membranes that appear to contain the information necessary for the event of association. Phospholipase A 2

This is an example of a water-soluble protein that is 'anchored' to a lipid/water interface, primarily by ionic interactions. However, this proteinbilayer system also displays many features of hy-

38 drophobic interactions. The catalytic activity of phospholipase A z, an sn-2-acylhydrolase, is activated as much as several thousand-fold when the enzyme binds to a lipid/water interface [18]. Studies with phospholipase A 2 from pig pancreas show that binding of this enzyme to the substrate interface [133] leads to catalysis 'along' the interface, i.e., the bound enzyme remains at the substrate interface (it 'scoots') for several thousand catalytic cycles [134]. Phospholipase A 2 does not bind to bilayers of 2,3-diacyl-sn-PC or 1,2-dialkyl-sn-PC, [133] or to interdigitated bilayers of alkylphosphocholine [182]. However, when lipid soluble additives such as alkanols, or both products of phospholipase A2-catalyzed hydrolysis (fatty acid and lysoPC) are co-dispersed with substrate vesicles (ternary codispersions), the binding of the enzyme is promoted, as observed by enhanced tryptophan fluorescence. Binding of phospholipase A 2 to vesicles also occurs when lysophospholipids alone are added to preformed vesicles, but not when the lysophospholipids are co-dispersed with phospholipids prior to formation of vesicles. In addition to facilitating binding, the latency phase in the progress curve for hydrolysis disappears in the presence of the above additives [183]. Thus it appears that phospholipase A 2 will not bind to some types of vesicles comprised of substrates, except in the presence of additives. There is no evidence that these additives are co-factors for the enzyme, yet in some way they facilitate binding of phospholipase A 2 to molecules of substrate arranged as a bilayer. Of importance in this regard is that the additives promoting binding of phospholipase A 2 to vesicles of PC are fusogens [184]. A variety of lipid soluble additives diminish the binding of phospholipase A 2 to ternary codispersions [184]. Additives of this type also inhibit fusion of vesicles [185]. The relationship between the propensity of vesicles to fuse with each other and to bind phospholipase A2 suggests, therefore, that organizational defects in the bilayer may be important for binding and catalytic action of the enzyme. Binding of .the phospholipase A 2 to dispersions of phospholipids can be described quantitatively by a binding curve arising from the interfacial binding equilibrium of the type, E + nL ~ EL, in

which n lipid molecules provide a site for binding of each molecule of the complex [133,136]. The values of n (20 to 50) and the dissociation constant ( K d < 0.2 /~M to about 10 t~M) depend on the nature of the lipid/water interface, the structure of the substrate, and the presence of ions. The presence of lipid-soluble additives is not required for binding of phospholipase A 2 to vesicles of anionic phospholipids. Moreover, the apparent dissociation constant for the enzyme in anionic vesicles is at least 50-times smaller vs. that for binding to ternary codispersions of PC. In fact, binding of phospholipase A 2 to vesicles of some anionic phospholipids is so strong that enzyme activity can be measured when only one molecule of enzyme is bound per vesicle. Also, bound enzyme does not dissociate from the vesicle to which it is bound, even after all the substrate in the outer monolayer of the vesicle has been hydrolyzed completely. An excess of substrate vesicles are not hydrolyzed under these conditions. Nevertheless, intervesicle exchange of the bound enzyme is induced readily by several anions: SO42-> C I - > C N S - [135]. In addition to stressing the idea that the interfacial binding of phospholipase A 2 is dominated by ionic interactions, the variability in dissociation caused by different anions suggests that the binding of enzyme to an interface has the characteristics of specific binding to a site. However, as discussed below, other types of interaction also may be important for binding. Ca 2+ is required for catalytic activity of phospholipase A 2 as well as for the interfacial binding of the enzyme. It is difficult to reach a firm conclusion about the exact function of Ca 2+ in the binding of phospholipase A 2 to the bilayers. There could be an ordered kinetic mechanism - binding of Ca 2+ to the enzyme must precede binding of the enzyme to the substrate in the bilayer. On the other hand, Ca 2+ could facilitate binding because it induces isothermal lateral phase separation in bilayers of anionic phospholipids. Two sets of observations may be significant in resolving this dilemma. The bilayer in ternary codispersions of PC is phase-separated, but the enzyme does not bind to it in the absence of Ca 2+ [133]. Such studies suggest collectively that binding of phospholipase A 2 to the interface requires that the enzyme actually bind to a molecule of substrate in

39 the interface and that this step depends on prior binding of Ca 2+ to the enzyme. The available evidence is compatible with the idea that phospholipase A 2 does not penetrate into apolar regions of the bilayer. For example, the increase in fluorescence associated with binding of phospholipase A 2 to bilayers is not associated with a change in fluorescence lifetime, which indicates a loss of static quenching of the single tryptophan in the protein [135,186]. The fluorescence of the bound enzyme is quenched, however, by water-soluble quenchers like succinimide, as well as by brominated lipids, which presumably are localized in the bilayer. These results can be rationalized by the hypotheses that the enzyme does not penetrate the bilayer when bound at the bilayer/water interface, but that the environment of the tryptophan is altered secondary to binding of a substrate molecule at the active site of the enzyme. Binding of substrate is not sufficient, however, to alter the environment of the tryptophan. Binding of monomeric lipids to enzyme in aqueous solution does not lead to the change in fluorescence observed secondary to binding of enzyme at the bilayer/water interface. The acyl chains of the molecule of substrate bound at the active site of phospholipase A 2 anchor the E. S complex to the apolar region of the bilayer [137]. Binding of enzyme to a monomer of substrate within the interface would be promoted, therefore, by factors that weaken association between a single molecule of phospholipid and other lipids, as for example, substrates with shorter vs. longer chain lengths or molecules of substrate located in the defect regions of the interface. Thus, factors that lead to a loss of defect structures, such as amphipaths, would inhibit the binding and catalysis by phospholipase A 2 [184]. In short, ionic binding of phospholipase A 2 to the interface followed by specific binding to a molecule of phospholipid in the bilayer leads to the overall high affinity binding of phospholipase A 2 to the anionic interface. Defects are induced in the packing of phospholipids in bilayers by the presence of impurities or phase separations [137]. These defects can weaken the associations between molecules of phospholipids within the region of the defect, which facilitates binding of substrate to the active

site of phospholipase A2 and thereby anchors the enzyme to the interface. A close contact between the enzyme and the bilayer interface also could be facilitated by defects that promote desolvation of the microinterface in the anchoring region, i.e., the array of phospholipids in direct contact with the protein. The bound enzyme could 'scoot' along the substrate interface, in the absence of cycles of solvation-desolvation as the enzyme moves from product to substrate in between catalytic events [137]. It also can be emphasized that vesicles containing bound phosphotipase A 2 retain the bilayer organization, (based on X-ray, NMR, and electron microscopic evidence) and the content of their aqueous compartment (Jain and Maliwal, unpublished data). In fact, leakage of calcein from phospholipase A2-treated vesicles is as slow as in untreated vesicles. These observations exclude the HII phase's being needed for binding of phospholipase A 2 or its being induced by binding of phospholipase A 2 to vesicles (Jain and Maliwal, unpublished).

fl-Hydroxybutyrate dehydrogenase This enzyme is a component of the inner mitochondrial membrane, with the active site on the matrix side. fl-Hydroxybutyrate dehydrogenase has been purified to homogeneity in a lipid-free state [187-190]. Although detergents are used during purification by most laboratories, the enzyme has been purified also in the absence of detergents [191]. The properties of fl-hydroxybutyrate dehydrogenase, in experiments examining reconstituted enzyme, are the same for enzyme purified in the presence or absence of detergents. Residual detergent in some preparations hence would seem to have no importance for explaining reconstitution. Review of the literature on fl-hydroxybutyrate dehydrogenase suggests that, like phospholipase A 2, fl-hydroxybutyrate dehydrogenase is anchored to bilayers by specific interactions with glycerophosphocholine [3,192-209]. As with phospholipase A2, it appears that the structure of the membrane can present a barrier to establishing specific interactions with fl-hydroxybutyrate dehydrogenase. Thus, reconstitution of fl-hydroxybutyrate dehydrogenase activity in bilayers is best when there is a mixture of phospolipids.

40 Lipid-free fl-hydroxybutyrate dehydrogenase has no catalytic activity. It is activated only by PC and mixtures of phospholipids containing PC [192-199]. In fact, activation is more efficient for mixtures of PC and other lipids versus pure PC [193-197]. Extensive structure/activity studies show that whereas the polar region of the PC is crucial for reconstituting function, the structural features of the apolar region are not [193,195,197, 200]. Reconstitution of the catalytic function of pure, delipidated fl-hydroxybutyrate dehydrogenase does not even depend on inserting the enzyme into a bilayer. Activity appears to be restored by monomeric PC's, e.g., by addition of enzyme to short chain PC at concentrations below the critical micelle concentration [197,200,201]. An analysis of the activity of fl-hydroxybutyrate dehydrogenase as a function of the concentration of monomers of phospholipid has been interpreted as showing that apo fl-hydroxybutyrate dehydrogenase (Mr= 31000) has two identical binding sites for PC and that activity requires only that both sites be occupied [197]. More recent data suggest that this might not be so [202], but the experiments in [197] and [202] were not carried out on identical systems. It has not been possible unfortunately to confirm the data for the stoichiometry of binding of short chain PC's by direct binding studies. One, therefore, cannot exclude formation of microaggregates (E. PCn) containing more than 2 molecules of PC per molecule fl-hydroxybutyrate dehydrogenase. Increasing the length of the acyl chains appears to increase the avidity of binding of PC's to enzyme, as determined from titrations of activity as a function of the concentration of phospholipid. The length of the acyl chain of short chain PC did not influence activity at Vm, however. Nor were there differences in activities at Vm for fl-hydroxybutyrate dehydrogenase activated by monomeric short-chain PC's or by bilayers of long chain PC [197,199-203]. /3-Hydroxybutyrate dehydrogenase binds to bilayers of PE plus DPG, but has no activity under this condition. Nevertheless, /~-hydroxybutyrate dehydrogenase does not dissociate readily from complexes with PE plus DPG. Thus, addition of vesicles of PC to a mixture of/3-hydroxybutyrate dehydrogenase and vesicles of PE plus DPG does

not reconstitute activity [191]. The complexes of fl-hydroxybutyrate dehydrogenase-PE plus DPG do dissociate, however, in the presence of 0.4 M LiBr, which leads to close to maximal activation of fl-hydroxybutyrate dehydrogenase when vesicles of PC are added. There are several possible ways in which to explain these data. One is 'hetero'-fusion of vesicles [134]. Another is intervesicle exchange of fl-hydroxybutyrate dehydrogenase promoted by LiBr [135]. An implication of this idea, which is compatible with data for fl-hydroxybutyrate dehydrogenase in several other systems, is that the enzyme can be anchored to bilayers both via interaction with anionic phospholipids and by specific binding of PC at two sites per molecule fl-hydroxybutyrate dehydrogenase. Thus, binding and catalytic activation specifically due to interactions with 2 mol PC/mol fl-hydroxybutyrate dehydrogenase are likely to be separate events, fl-Hydroxybutyrate dehydrogenase in the presence of high salt has greater avidity for bilayers of PC vs. anionic phospholipids. By contrast, direct comparisons of the amount of fl-hydroxybutyrate dehydrogenase bound per tool phospholipid (in the absence of salt) show that fl-hydroxybutyrate dehydrogenase binds more avidly to vesicles of PE plus DPG (non-activating) vs. vesicles of PC. It appears that a great deal remains to be elaborated about the exact energetic basis for interactions between fl-hydroxybutyrate dehydrogenase and phospholipids. In addition to the relationship between the structure of lipids used to reconstitute fl-hydroxybutyrate dehydrogenase and the activities of reconstituted preparations, studies with PC containing photoactivated groups suggest that fl-hydroxybutyrate dehydrogenase interacts primarily with the polar region of PC's in membranes. El Kebbaj et al. [204] synthesized PC's containing myristate at the 1-position. The 2-position contained a nitrene, which was attached either to the 12-position of dodecanoic acid (an analog of DMPC) or directly to the glycerol backbone (an analog of lysomyristoylPC). The extent of covalent labelling of fl-hydroxybutyrate dehydrogenase was examined for enzyme added to vesicles prepared from mixtures of the photoactivatable phospholipids (about 40 mol% of total lipids) and mitochondrial phospholipids. Vesicles containing

41 the nitrene-labelled analogs of DMPC and lysomyristoylPC reconstituted activity as well as vesicles containing 40 mol% DMPC. Labelling of fl-hydroxybutyrate dehydrogenase in these systems was 3-times greater in vesicles containing the lipid with the nitrene group near the polar region (direct attachment to the 2-position of glycerol) vs. the lipid with the nitrene at C-12 of the fatty acid attached at position 2 of glycerol. As compared with this result for fl-hydroxybutyrate dehydrogenase, cytochrome c was labelled to a slightly greater extent by the latter as compared with the former photoactivatable compound. Although reconstitution of the catalytic activity of fl-hydroxybutyrate dehydrogenase depends on binding 2 mol PC/mol fl-hydroxybutyrate dehydrogenase, the best reconstitution of activity occurs in bilayers of phospholipids extracted from mitochondria [194,195,202,205,206]. It is not clear why this is so because, as mentioned already, the acyl chains of PC do not seem to influence Vm. The explanation why mixing PC with other phospholipids enhances the catalytic activity versus PC alone may be related, as for binding of phospholipase A 2 to bilayers, to differences in energies for interactions between PC's in pure bilayers versus mixed bilayers. That is, energies for interactions between a given molecule of PC and other phospholipids, which are likely to be lower in a mixed bilayer versus a pure bilayer, may determine the equilibrium position of the reaction forming the catalytically active complex of the type E. PC 2. An especially interesting set of observations in this regard is that fl-hydroxybutyrate dehydrogenase is released from mitochondrial membrane when PC is hydrolyzed to lysoPC [207,208]; and although LPC alone can activate fl-hydroxybutyrate dehydrogenase [195], lysoPC interferes with maximal reactivation of pure fl-hydroxybutyrate dehydrogenase added to bilayers [191,209]. If lysophosphatides exchange readily between bilayers, their presence could weaken the anchoring of fl-hydroxybutyrate dehydrogenase to the membrane. For example, the lysoPC could displace PC from the two lipid-binding sites/ molecule enzymes, which would weaken the attachment of protein to membrane in addition to altering catalytic function. The inhibition by lysoPC of reactivation of fl-hydroxybutyrate dehy-

drogenase by lipid bilayers does not appear to be a nonspecific effect of detergents. A variety of neutral or negatively charged detergents do not inhibit reactivation. Interestingly, however cetyltrimethylammonium bromide (CTAB ÷) and cetylamine (C 16- N + H 3) do inhibit [191,209].

Polymyxin Not all the proteins that are 'anchored' to bilayers via ionic interactions can be removed with salt. Cations do not promote desorption of spectrin [148,210], myelin basic protein [118] and polymyxin [211] from anionic bilayers. These results may mean that a varying degree of electrostatic and hydrophobic interactions account for anchoring in some instances. Polymyxin and circulins are cyclic decapeptides containing a high proportion of cationic diaminobutyric acids. Binding of polymyxin to bilayers is believed to be via anionic lipids, and depends clearly on ionic strength and pH [143]. On the other hand, these antibiotics induce leakage in bilayers. This ultimately leads to lysis, which suggests that interactions between the hydrophobic tail of polymyxin and apolar regions of the membrane also are important for binding. This idea is supported by studies of colistin, which is identical with polymyxin except for lacking the C6 hydrophobic tail. Colistin does not have significant affinity for anionic vesicles (Jain, unpublished data). In addition, the order parameter for 16-doxylstearate in membranes containing polymyxin becomes similar to that for 5-doxylstearate. This observation suggests that the 'fluidity gradient', found normally in the gel phase, is abolished by polymyxin [141]. Together with X-ray diffraction data these results have been interpreted as suggesting that polymyxin causes interdigitation of the acyl chains of the two halves of a bilayer [2121. Polymyxin may be an example of a protein that interacts with membranes in two ways - electrostatically and hydrophobically. More important, the latter interaction may not be possible in the absence of the former. That is, the electrostatic interaction between bilayer and protein could lower the energy barrier for insertion of part of the protein into the apolar region of the membrane, by effects on the bilayer, the protein, or both. This could be one way for solving the prob-

42

lem of inserting a protein into a bilayer, as suggested first by Wickner [8,147]. An especially interesting example of this kind of mechanism for the spontaneous insertion of proteins into preformed bilayers is emerging from recent studies of apocytochrome c (p. 51).

Myelin basic protein Myelin basic protein constitutes about 30% of the total myelin proteins and is rich in basic amino-acid residues [213]. Interactions between myelin basic protein and phospholipid dispersions have been studied extensively. The phase state of bilayers is altered significantly in the presence of the protein. The order parameter, Tm, enthalpy, and cooperativity of the gel-to-liquid crystalline transition generally decrease. In most cases multiple phase transitions are observed, and the amplitudes and temperature dependencies of these depend on the history of the sample. Such hysteresis indicates that multiple phases coexist in the test sample. Partial interdigitation of the acyl chains in the two halves of the bilayer is induced by myelin basic protein. Myelin basic protein interacts weakly with bilayers of PC even when dispersed in the presence of the protein and taurocholate. However, the protein affects the phase properties of sphingomyelin [115], suggesting that it binds spontaneously to bilayers of this lipid. Incorporation of myelin basic protein into acidic phospholipids is rapid and in the order phosphatidic acid = PG > PS > cerebroside sulfate > PE (Table I). The exposure of the antigenic determinants depends on the type of lipid in a similar way. The nature and extent of binding apparently change with temperature, pH and concentration of salt [178]. The perturbing effect of myelin basic protein on dispersions of several lipids increases with increasing pH in the range of pH 5 to 9. Results of several different kinds of experiment [141,178] have been interpreted as demonstrating that ionic interactions and H-bonds between myelin basic protein and the polar regions of bilayers are important for incorporating myelin basic protein into bilayers. Ionic interactions of myelin basic protein with anionic lipids very likely facilitate further specific interactions that lead to extremely tight binding presumably through a network of H-bonds. Such

an anchoring of the protein to the lipid/water interface could promote an appropriate conformation for the antigenic behavior of myelin basic protein. The change in the phase properties of the host bilayer could be a manifestation of the reorganization of the acyl chains necessary to accomodate the functional groups of the protein that desolvate the microinterface between the bilayer and myelin basic protein.

Binding of proteins to bilayers and to specific head groups in bilayers The incorporation of proteins into membranes reflects the partitioning of solutes between two immiscible phases. In its simplest form the binding equilibrium for a solute (S) between a membrane (M) and a site comprised of n lipid molecules (L) is expressed as (1) or (2) M + S ~ MS

(1)

S + nL ~ SL~

(2)

The relationship can be quantitated by a dissociation constant [3] and the numerical value of n for the interaction between the solute and a binding site [133,214,215]. Ko

[Sleree[Mlfree [SI[M] [SI[PLI [complex] [Ms] [S(PL)n]

(3)

A variety of toxins act on lipid bilayers and perturb their organization enough to interfere with physiological functions. These include mellitin, toxin A-III from Cerbratulus lacteus, alfa-toxin from staphylococcus, diphtheria toxin, and Stoichactis helianthus toxin. Many of these toxins are embedded within the hydrophobic regions of a bilayer. But, in many instances interactions between toxins and polar groups promote binding and subsequent reorganization that ultimately leads to lysis. Glycolipids have been implicated as specific receptors for immunoglobulins [216,217], lectins [218], cations [219,220], hormones, interferons [221], growth factors [222], viruses [223,224], and for tetanus toxin [225], cholera toxin [226], botulin toxin [227], and sea-wasp toxin [228,229]. Specific binding of proteins to other lipids has been implicated [230]. However, in most cases, the precise mechanism of action or biological significance

43 of the interaction of proteins with glycolipids has not been established. Nevertheless, it is believed that binding of these proteins to glycolipid receptors somehow regulates the distribution and accessibility of membrane components so as to elicit coupling of the receptor to effector functions. The binding of lectin to phospholipid vesicles containing carbohydrate receptor has been reviewed recently [107]. Binding appears to be insensitive to the phase state of bilayers. On the other hand, the distribution of receptors is sensitive to the phase of the bilayer. Interaction of a protein with its ligand in a bilayer often leads to secondary manifestations such as agglutination induced by lectins, antigens, or protein by calcium-induced fusion. The concentration dependence of such a process exhibits a threshold effect, i.e., the response is observed only above a certain critical concentration of the inductor. The origin of these effects probably is the statistics of the density of receptors on vesicles and the concentration of vesicles [231], which manifest ultimately in ligand-induced aggregation, vesicle and cells. 11. C. Embedded proteins

Embedded membrane proteins do not dissociate from membranes to any extent that has been measured. Removal of integral proteins from membranes depends on destroying the bilayer structure and extracting the proteins into an excess of a suitable detergent, which usually is found by testing empirically an extensive array of different detergents. This is different from the situation of adsorbed and anchored membrane proteins. Many of these latter proteins can exist either as membrane-bound or soluble forms, and function can be regulated by the balance between the two forms. Apparently, hydrophobic effects dominate the association of the integral membrane proteins with bilayers. Review of the literature on such systems indicates that nearly all the published work centers on interactions between integral membrane proteins and lipids subsequent to formation of the lipid-protein complexes. Many such studies were designed to determine whether a protein has a specific avidity for one or another type of phospholipid, the minimum number of phospholipid

molecules required to reconstitute function, or the effects of different lipids on the functiofi of an integral membrane protein. None of this work provides insight into how a protein of interest gets into a membrane. The general experience is that there is an insurmountable energy barrier to the spontaneous insertion of 'solubilized' integral membrane proteins into preformed bilayers. This has meant, in terms of the design of experimental studies of reconstituted systems, that integral membrane proteins are reconstituted into membranes by forming the membrane in the presence of the protein, phospholipids and detergent. This experimental experience has led in turn to the idea that insertion of integral membrane proteins into membranes in cells is a complex, facilitated event. According to such arguments, no intrinsic property of the lipid bilayer is considered important for insertion of integral membrane proteins into membranes [8,10,232]. The material reviewed in this section consists of data for the spontaneous insertion of several 'solubilized' integral membrane proteins into preformed bilayers of phospholipid. These data have not received adequate attention. Thus, it appears that the spontaneous insertion of 'solubilized' integral membrane proteins into preformed bilayers can be facilitated by apparently simple modifications of a bilayer. Microsomal cytochrome b s

This protein inserts spontaneously into preformed unilamellar bilayers of PC [233-235] as reflected by enhanced fluorescence of tryptophan in the hydrophobic 'tail' region of the protein, which becomes buried in the apolar region of the bilayer [236]. The phase state of the bilayer and size of vesicles influence the spontaneous rate of incorporation of cytochrome b5 into phospholipids. Incorporation proceeds considerably faster for fluid versus gel phases. And when mixed with large and small vesicles of identical phospholipid composition, the protein associates preferentially with the small vesicles [237,238]. The energetic basis for this last result is not completely clear; but it appears to have general importance whenever such systems have been studied. Thus, spontaneous association of large integral membrane proteins with preformed ULV's appears to

44 depend on the radius of curvature of the ULV [238-240]. Cytochrome b 5 reconstituted spontaneously into preformed ULV's of pure single phospholipids is competent in electron transfer [233], but the protein transfers rapidly between different populations of phospholipid vesicles. Transfer is via the aqueous phase, i.e., it is not due to fusion between vesicles [241-243]. By contrast, cytochrome b5 is attached firmly to microsomal membranes. It cannot be extracted from microsomes by mixing with phospholipid vesicles [244]. This means that the protein does not insert spontaneously into preformed vesicles with the same o r i e n t a t i o n / o r g a n i z a t i o n displayed by cytochrome bs in biological membranes. Apparently, however, it can be made to do so by forming cytochrome bs-lipid vesicles via detergent dialysis. For, as compared with spontaneously incorporated cytochrome b5, the protein is attached irreversibly to bilayers when incorporated by detergent dialysis [244]. Moreover, whereas the C-terminal end of the protein is hydrolyzed by carboxypeptidase Y after simple mixing of the protein with vesicles of pure phospholipid, it is not hydrolyzed in vesicles of phospholipid-cytochrome b 5 formed by detergent-dialysis [244-247]. Structural data on the specific cytochrome bs-lipid systems are compatible with the conclusion that the hydrophobic tail of the spontaneously incorporated protein forms a loop with the N- and C-terminal ends in the aqueous phase on the same side of the membrane. By contrast, the C-terminal region of the protein appears to extend into the bilayer as a result of incorporation during detergent dialysis [244,248]. These data indicate that simple mixing of cytochrome b s and ULV's does not form a complex similar to the naturally occurring complex in microsomes, which suggests in turn that there is a relatively high energy barrier to spontaneous transmembrane insertion. Surprisingly, however, this barrier is removed easily and the tightly-bound orientation for cytochrome b5 can be achieved in several relatively simple ways. For example, addition of lysophosphatide to mixtures of cytochrome b 5 and phospholipid vesicles leads to irreversible association of cytochrome b 5 with the lipid bilayer: and small amounts of deoxycholate

convert loosely-bound cytochrome b s to the tightly-bound form, when added to mixtures of proteins and vesicles of egg PC [244,246,247]. Cytochrome b 5 inserts spontaneously in the tightlybound configuration into ULV's of DMPC containing 5 mol% DMPE or into ULV's of egg PC containing the microsomal enzyme stearoyl CoA desaturase [244,249]. Moreover, cytochrome b 5 is incorporated in this orientation on addition to microsomal membranes. The major question posed by these observations on spontaneous irreversible insertion is what properties of the cytochrome b5 a n d / o r of the ULV's are required for insertion that leads to the transmembrane configuration of bound cytochrome b 5. Lysophosphatides used in the method of reconstitution described by Christiansen and Carlsen [246,247] could alter either the cytochrome b 5, the vesicles, or both so as to cause insertion under conditions that lead to a non-dissociable complex of protein and phospholipid. It is highly unlikely, however, that DMPE or stearoyl CoA desaturase, as components of U LV's, could alter the conformation of cytochrome b 5 prior to its insertion into the membranes. Thus, it is reasonable to propose that the insertion of cytochrome b 5 into preformed ULV, in a configuration that makes binding tight (no dissociation), can be facilitated by changing the properties of the ULV's without changing the properties of the cytochrome b 5. An important corollary to this conclusion, for which further evidence is developed later, is that fully folded, hydrated membrane proteins may contain information that allows them to alter their conformation and thereby to insert rapidly into ULV's modified by additives.

UDP glucuronosyltransferase This enzyme, when delipidated, has a small residual activity [250-253]. It is activated to a variable extent by addition of suitable phospholipids [251,252]. The delipidated, pure enzyme that has been dialyzed extensively to remove residual detergent activates spontaneously when added to lipid vesicles [250]. The association between the enzyme and UEV's in this setting is not stable, as the complexes dissociate during density gradient centrifugation [240]. The protein recovered in such gradients behaves catalytically as if it were free of

45 lipids, which has been validated by chemical analysis. However, some preparations of delipidated enzyme exhibited stable association with preformed ULV's as shown by density gradient centrifugation. The variation between preparations in the ability to form stable association with ULV's was traced to a variable amount of residual detergent in the delipidated preparations. Preparations that did not show stable association could be made to do so by addition of small amounts of deoxycholate or other detergents, in amounts below the critical micelle concentrations [240]. These amounts of detergent were insufficient to destroy the bilayer. Of importance was that detergent had to be added to the protein, not to the lipid vesicles, prior to mixing of delipidated UDPglucuronosyltransferase and phospholipids. Prior addition of cholate of ULV's, prepared by sonication, did not facilitate reconstitution when detergent-free enzyme was added. This result suggested that the stable association of the enzyme with the vesicles depended on detergent-dependent properties of the protein, but this has proved not to be so. Deoxycholate appears to facilitate the stable insertion of UDPglucuronosyltransferase into preformed ULV's by modifying the properties of ULV's. The clue to the effects of cholate on reconstitution was the observation that optically clear preparations of enzyme that inserted stably produced immediate turbidity when added to ULV's prepared by sonication. Detergent-depleted enzyme had no such effect. Moreover, the effect of cholate on reconstitution was limited to temperatures above the phase transition for the ULV's used. The turbidity caused by mixing some preparations of pure enzyme with ULV's was attributed to detergent-induced fusion of ULV's in the liquid crystal phase [4,254-256]. Purified UDPglucuronosyltransferase forms large aggregates. Addition of small amounts of detergent could have facilitated insertion of enzyme into bilayer by comminuting the aggregates of proteins. On the other hand, the effect of detergent on insertion depended on the phase state of the ULV's. Insertion occurred only under the condition that deoxycholate was a fusogen for ULV's of DMPC, which is for lipids in a fluid-like phase. Moreover, enzyme did not insert into vesicles that had been allowed to fuse in the

presence of deoxycholate [240]. Insertion appeared to depend, therefore, on the presence of small ULV's, which fuse readily. It was proposed [240], that deoxycholate-induced fusion between the ULV's somehow facilitated the spontaneous insertion of UDPglucuronosyltransferase into preformed ULV's. To test the hypothesis that fusion between ULV's was important for the stable insertion of UDPglucuronosyltransferase into preformed bilayers, reconstitution of this enzyme was examined in a system in which fusion was induced by incorporating 10 mol% myristate into ULV's of DMPC [257-259]. Myristate is a known fusogen for ULV's of DMPC at temperatures below the gel-to-fluid phase transition [257-259]. UDPglucuronosyltransferase and ULV's of DMPC formed stable complexes when the ULV's contained myristate and the system was mixed below the phase transition for DMPC [240]. Reconstitution in the above system was a two-step process. The first step was a rapid and complete incorporation of protein into a small fraction of the total ULV's. Thus, within a few seconds of mixing enzyme and ULV's, all the enzyme was recovered from density gradients as lipid-protein complexes. Only a small fraction of the total amount of lipid was recovered in lipid-protein complexes, however. If mixtures of protein and ULV's were allowed to stand at 18°C for minutes to about 2 h, there was a continuous increase in the ratio of lipid/protein recovered from gradients as lipidprotein complexes. Therefore, subsequent to the initial rapid insertion of protein into ULV's, there was a slower incorporation of excess phospholipid into the proteoliposomes formed initially. Eventually (in about 2 to 4 h), most of the lipid was found in association with protein. The two steps of the reconstitution process could be separated because fusion of proteoliposomes with excess ULV's comprised of DMPC and myristate was interrupted by inducing the gel to fluid phase transition by raising the temperature of the system. The relationship between fusion of ULV's with each other and insertion of UDPglucuronosyltransferase into ULV's of DMPC that are fusogenic is considered further in the section on bacteriorhodopsin. The spontaneous insertion of UDPglucuronosyltransferase into preformed vesicles appeared to

46 be unidirectional. All the UDPglucuronosyltransferase binding sites were on the outside of the vesicles. This conclusion was based on measurements of the n u m b e r of U D P g l u c u r o n o syltransferase-binding sites per mole enzyme in reconstituted systems. The orientation of the enzyme in liver microsomes also seems to be with the UDPglucuronosyltransferase-binding site facing the outside of the microsomal vesicles [260-262]. Hence, the pure enzyme inserts spontaneously into ULV's with the same orientation it has in intact microsomes. As mentioned already, delipidated purified UDPglucuronosyltransferase has an aggregated size of several million daltons. The available data indicate that the aggregates of UDPglucuronosyltransferase inserted directly into ULV's. That is, the aggregates did not seem to dissociate as a prerequisite for insertion of individual molecules of holoenzyme. The evidence on this point was that the number of aggregates was far smaller than the molecules of enzyme and the number of ULV's in a typical reconstitution experiment. Simple calculations based on these numbers and the ratio of lipid to enzyme in complexes isolated by density gradient centrifugation indicated that the aggregates themselves must have inserted into ULV's [240]. This conclusion also accounts for observations made in other laboratories that only a small portion of the ULV's present often appear to be involved in the reconstitution of lipid-protein complexes under conditions leading to spontaneous insertion of membrane-bound proteins other than UDPglucuronosyltransferase into preformed ULV's [239]. As a result, only a portion of the ULV's interact directly with lipid-free aggregated protein. UDPglucuronosyltransferase had nearly the same specific activity in proteoliposomes that were stable or in complexes that subsequently could be dissociated by high g-force. There are no structural data on the organization of the lipids and proteins in either of these complexes. On the other hand, methods that produced a stable enzymephospholipid vesicle reconstituted a form of the enzyme with the aliosteric properties displayed by enzyme in intact, untreated microsomes. Moreover, the allosteric properties of UDPglucuronosyltransferase in such vesicles were sensitive to the

phase state of the phospholipids. By contrast, enzyme reconstituted under conditions that produced an unstable or dissociable lipid-protein complex did not display the allosteric properties observed for enzyme in microsomes, nor were any of the properties sensitive to the phase state of the phospholipids in the vesicles (Zakim and Scotto, unpublished data).

Bacteriorhodopsin This is not an especially large membrane protein ( M r-- 28000), but the purified protein aggregates in the form of an extensive two-dimensional crystalline array containing residual phytolipids [263]. Following reconstitution into vesicles in a fluid phase, bacteriorhodopsin is present as trimers [264,265]. The most popular method for reconstituting bacteriorhodopsin into a lipid-protein complex is by detergent dialysis [266-269]. Co-sonication of protein and phospholipid also has been used [270]. Bacteriorhodopsin is functional in vesicles reconstituted by either method. It has been observed recently, however, that bacteriorhodopsin also will reconstitute spontaneously into preformed vesicles of DMPC, provided the vesicles contain impurities [240,271]. Bacteriorhodopsin will insert, for example, into ULV's o_f DMPC that contain myristate [240] or cholesterol [271]. Both types of impurity are fusogens for ULV's of DMPC in a gel phase. Bacteriorhodopsin in vesicles reconstituted by spontaneous insertion into preformed ULV's is active as an H + pump and has the same orientation as bacteriorhodopsin reconstituted by cholate-dialysis or sonication. H + pumping is from the outside to the inside of the vesicles [240,266269,271], which is opposite to the orientation of the H ~ pump in the intact bacteria. Insertion of bacteriorhodopsin into preformed ULV's has been obtained at concentrations of myristate or cholesterol that are below the threshold for inducing fusion between the vesicles. As little as 0.1 mol% myristate (the smallest amount tested) is sufficient, for example, to allow spontaneous insertion of the protein into ULV's. Myristate and cholesterol, or contaminants in ULV's of DMPC, also will promote the spontaneous insertion of bacteriorhodopsin into these vesicles at temperatures at which these lipid con-

47 taminants are fusogens. Myristate, for example, is maximally fusogenic in ULV's of DMPC at 18°C. It does not induce fusion of these vesicles at 5°C. Nevertheless, bacteriorhodopsin will insert spontaneously into ULV's of DMPC plus myristate at 5°C as well as at 18°C [243]. Similar results are obtained with cholesterol as the fusogenic impurity in vesicles of DMPC [271]. Cholesterol is maximally fusogenic at 21°C in ULV's of DMPC, and is not fusogenic at 5°C. But the presence of cholesterol in ULV's of DMPC at 5 °C facilitates spontaneous insertion of bacteriorhodopsin into the ULV's. The protein will not insert into ULV's of pure DMPC at any temperature [240,271]. The rate of formation of phospholipid-protein complexes between ULV's and bacteriorhodopsin (or UDPglucuronosyltransferase) is more rapid than is fusion between ULV's. Since the concentration of aggregates of protein in reconstitution experiments is far lower than that for ULV's, these kinetic observations are not compatible with formation of ternary complexes (two fusing ULV's plus protein) during insertion of the proteins into bilayers. Our interpretation of the above results is that contamination of DMPC with myristate (or cholesterol) induces fusion between ULV's and dispersed aggregates of purified membrane proteins under conditions that do not necessarily facilitate fusion between ULV's. An important aspect of these studies on spontaneous reconstitution was that the rates of insertion of bacteriorhodopsin and UDPglucuronosyltransferase into preformed ULV's was on the order of mixing time (e.g., a few seconds or faster). Decreasing the temperature of the system from 18 °C to 5°C did not slow the process so that it could be measured without using optical methods. In other words, when conditions allow for spontaneous insertion of membrane proteins into preformed bilayers, the rate of the insertion event is extremely rapid. By contrast, the rate of spontaneous insertion of an aggregated membrane protein like bacteriorhodopsin into pure bilayers of a single, pure species of phospholipid is immeasurably slow. Hence the presence of impurities in lipid bilayers has a remarkably large rate-enhancing effect on this process. The data for myristate or cholesterol-facilitated insertion of aggregated membrane proteins into

ULV's provide no direct evidence for how the process of reconstitution proceeds. On the other hand, they show that the observed rates for the overall process can be modulated in surprisingly simple ways. That is, bilayers containing a single, pure phospholipid can be 'activated' by incorporating impurities. Although it does not seem that the aggregates of integral membrane proteins need to be modified in order to effect spontaneous reconstitution into activated bilayers, sonication of the crystalline arrays of pure bacteriorhodopsin does affect reconstitution [270]. For example, crystalline arrays of pure bacteriorhodopsin did not insert into vesicles of DMPC plus myristate or cholesterol that had been allowed to enlarge via fusion prior to addition of the protein. Sonicated bacteriorhodopsin did insert, however, into such large ULV's, and the extent of insertion depended directly on the duration of sonication. The effect of sonication in this context was not a kinetic one, i.e., the effect of an increased concentration of particles of aggregated protein colliding with ULV's. Thus, under the conditions described, no matter how long the duration of mixing, no unsonicated bacteriorhodopsin formed proteoliposomes. Therefore, the state of aggregation of bacteriorhodopsin per se had a significant effect on spontaneous reconstitution. The idea that the size of aggregates can influence spontaneous reconstitution is supported as well by observations on interactions between ULV's and the apo-proteins of serum lipoproteins (see below). Cytochrome c oxidase and other mitochondrial proteins

Cytochrome c oxidase is a multicomponent complex of membrane-spanning proteins [272-275]. Pure complex can be reconstituted into bilayers by cholate dialysis, which yields proteoliposomes with two different orientations of the cytochrome c oxidase [276]. The enzyme in these preparations is about 50% inside-out. Racker and co-workers have described a variety of vesicles that are suitable for demonstrating the spontaneous incorporation of cytochrome c oxidase into preformed bilayers [239,276-280]. A technically important advantage of spontaneous incorporation of cytochrome c oxidase, as compared with forming proteoliposomes by detergent dialysis, was

48 that enzyme inserted unidirectionally into preformed vesicles [277]. Spontaneous incorporation of cytochrome c oxidase into preformed vesicles depended on the properties of the vesicles. Racker and co-workers showed that the protein inserted spontaneously into several complex mixtures of phospholipids: ULV's made by sonicating mixtures of PE, PC and PS, or another phospholipid, lipids extracted from bovine heart, soy bean lipids or mitochondrial lipids [276-280]. The complex inserted into preformed vesicles of neutral lipids (egg PC) in the presence of small amounts of deoxycholate. Addition of stearylamine also facilitated insertion of cytochrome c oxidase into vesicles containing mixtures of egg PC and CL. Although the insertion events examined by Racker and his colleagues occurred in the liquid crystal phase, the conditions for spontaneous reconstitution of cytochrome c oxidase were like those for cytochrome bs, UDPglucuronosyltransferase and bacteriorhodopsin in that spontaneous insertion of cytochrome c oxidase into preformed vesicles depended on introducing 'impurities' into the bilayers and establishing conditions for spontaneous fusion between vesicles [8,239,240,271,276-281]. It appears, therefore, that the spontaneous insertion of a variety of membrane-proteins into preformed bilayers can be achieved for bilayers either in gel or liquid crystal phases, and that in each instance the crucial property of the bilayer is the presence of impurities. Vesicles prepared from bovine heart mitochondrial phospholipids or crude soy bean phospholipids contain substantial amounts of lysophosphatides [277], and hence are similar to vesicles made from mixtures of lysophosphatides plus single species of diacylphosphatides. Small amounts of cholate, approximately equivalent to the amounts of lysophosphatide on a mol% basis, also facilitated spontaneous reconstitution of cytochrome c oxidase in the systems in which lysophosphatides were effective [277]. Moreover, mixtures of PE, PC and acidic phospholipids facilitated the spontaneous insertion into preformed ULV's of the oligomycin-sensitive ATPase from bovine heart mitochondria and QH 2 cytochrome ¢ reductase [239]. As for cytochrome c oxidase, the ATPase and QH 2 cytochrome c reductase inserted uni-

directionally in systems reconstituted spontaneously, but inserted with bidirectional orientations when reconstituted by cholate dialysis. Cytochrome c oxidase inserts spontaneously into preformed ULV's of DMPC if the vesicles contain small amounts of myristate or cholesterol as impurities [243,274]. Reconstitution into ULV's of DMPC plus myristate or cholesterol occurred only for DMPC in a gel phase. Reconstitution in these experiments was assessed by separating proteoliposomes from lipid-free protein on density gradients and measuring the amount of added protein associated with lipid vesicles. Activity was not measured. However, measurements of protein indicated that nearly all the cytochrome c oxidase became incorporated into proteoliposomes. Thus, the exact conditions needed for spontaneous reconstitution of microsomal UDPglucuronosyltransferase or bacteriorhodopsin from Halobacterium halobrium apply as well to a complex mitochondrial protein. This reinforces the emerging idea that spontaneous reconstitution of purified membrane proteins is a general phenomenon. Apoproteins from serum lipoproteins The protein components of serum lipoproteins are not membrane proteins, nor do they interact normally with phospholipids to form bilayers. These proteins are hydrophobic, however; and they do interact spontaneously with preformed phospholipid bilayers [282-288]. Under the typical experimental conditions employed for studying interactions between the apoproteins of serum lipoproteins and phospholipid bilayers, the latter are solubilized, forming disc-like lipid-protein aggregates of 105 t o 10 7 Da [289]. This 'solubilization' leads to a decrease in the turbidity of large, multilamellar phospholipid vesicles, which can serve as the means for measuring interactions between vesicles and proteins [288]. The interactions between apoproteins and phospholipid bilayers have several interesting features in common with the interactions between large integral membrane proteins and preformed bilayers discussed already. The apoproteins that have been studied cover a range of sizes. This is important because interactions between multilamellar vesicles of DMPC and various apoproteins apparently depend on the size of the proteins. Also, the rates of formation of

49 solubilized phospholipid-apoprotein complexes, for any apoprotein, have maxima at 24°C for vesicles of DMPC [284,285]. The temperature-dependence of these rates appears to depend critically on the size of the apoprotein [239]. Significant rates for interactions between apoA 1 (28.4 kDa) and vesicles of DMPC are limited to conditions existing at the temperature for the phase transition of DMPC. The rate increases 40-fold when T is raised from 22.5°C to 24°C and then decreases by 30-fold when T is raised to 27 ° C. By contrast, RCM-A-II (8700 Da) interacts relatively rapidly with vesicles of DMPC either above or below 24 ° C. There is even a smaller effect of temperature on rates of interaction between DMPC and the 20-amino-acid model peptide referred to as LAP 20 [288]. In addition to a strong dependence of rates of solubilization on temperature, rates of solubilization at any temperature are much smaller for large vs. small apoproteins. Rates of solubilization of vesicles of DMPC at 24 o C are about 103 times greater for LAP as compared with apoA 1. It seems, therefore, that whatever happens at 24°C to allow insertion of apoproteins and other hydrophobic proteins is important only for the insertion of relatively large apoproteins into vesicles of DMPC. Conditions for insertion of relatively small apoproteins are present at 24 ° C, as well as above and below this temperature. On the other hand, the rate of interaction between apoproteins and bilayers is enhanced in the presence of guanidine hydrochloride, which suggests that the rate-limiting step in the interaction is the accessibility of the hydrophobic regions of the proteins (Epand, R., personal communication). This accessibility could depend in some way on the size of the apoprotein; but Epand's data suggest that size per se is not the only factor determining interactions between apoproteins and lipid bilayers. Given the usual methods for studying interactions between apoproteins and vesicles, one must be cautious in equating solubilization of phospholipids by the apoproteins (the measured property) with the insertion (the inferred event). The methods used in the work by Pownall et al. [284,285,288] and Swaney et al. [286,287], as quoted above, have shortcomings in this regard. Rates of solubilization reported in Ref. 288, for

example, are quite slow as compared with spontaneous rates of insertion into ULV of DMPC of proteins larger than apoA 1. This may reflect only that solubilization occurs after a critical level of insertion is exceeded. Solubilization, therefore, is unlikely to be a direct index of insertion. The data in Ref. 290 support this conclusion. Measurements of the effects of apoproteins on the turbidity of large multilamellar vesicles (i.e. solubilization), suggested that the apoproteins did not solubilize pure vesicles of DPPC and DSPC even at the Tm for these vesicles [286-288]. On the other hand, experiments not based on measurement of turbidity suggested that apoA 1 did interact with ULV's of pure DPPC and DSPC (as well as DMPC) [290]. Interaction was measured in the latter studies by the sudden, rapid leakage (phase transition release) of the internal contents of ULV's, which was presumed to be secondary to protein-bilayer interactions. The initial ratio of mol phospholipid to mol protein was quite high in these experiments (about 3400). So it was presumed that there was insufficient protein to solubilize all the phospholipid, and that lipid remained in a bilayer with interdigitated apoA 1. Not surprisingly, the condition required for interaction between apoA 1 and ULV's of DPPC or DSPC, at low ratios of protein to phospholipid, was that the bilayers be at Tm for the phospholipid comprising them. Unfortunately, there was no direct evidence to support the idea that the apoprotein formed irreversible complexes with the bilayer. Nevertheless, the data in Ref. 290 do demonstrate that the apoprotein must have interacted with bilayers of DPPC and DSPC. Although the serum apoproteins interact with vesicles of pure, single species of phospholipid, the data for the larger apoproteins can be considered a variation of the theme displayed by cytochrome b5, cytochrome c oxidase, bacteriorhodopsin and UDPglucuronosyltransferase, which is that a mixture of phospholipids or impurities in bilayers of single phospholipid facilitate insertion by producing an 'activated' state of the bilayer. The mixtures of phospholipids required for protein-lipid interactions in the work in Refs. 285-291 comprised co-existing gel and liquid crystalline phases of the same species of phospholipid. Thus, insertion was facilitated under conditions of co-existing gel and fluid phase - the bilayer at the midpoint

50 of the phase transition. This idea is reinforced by observations that other kinds of mixtures of phospholipids also facilitated insertion of apoproteins into preformed bilayers of phospholipid. Bilayers of DMPC containing cholesterol reacted more rapidly with apoA 1 than did bilayers of pure DMPC [282,283,287]; 13.7 mol% cholesterol stimulated this rate optimally [287]. The rate of interaction declined at higher concentrations of cholesterol. Lysophosphatidylcholine also stimulated the rate of interaction between bilayers and apolipoproteins [288].

Reconstitution of the glucose transporting protein The glucose transporter from erythrocyte membranes was reconstituted by Kasahara and Hinkle [292] by adding the protein to ULV's prepared by sonication above Tm. The mixture then was frozen rapidly and thawed. Lipid-protein complexes were formed and consisted of large unilamellar vesicles. These were sonicated briefly (20 to 30 s) to reduce the size of the vesicles. The sonication step, however, was not the one that induced insertion, which was completed prior to sonication. The success of the freeze-thaw method may be another example of the relationship between the barrier to the spontaneous insertion of membrane-bound proteins into vesicles and the presence of mixtures of phospholipid species in the preformed vesicles. The mixture in this case did not comprise different species of phospholipid, but different arrays of phospholipids separated by defects in the packing order. Defects in packing are induced by rapid cooling of vesicles prepared above Tm [293,294]. This probably is the mechanism for the reconstitution method developed by Kasahara and Hinkle. An important piece of evidence in support of this conclusion is that the freeze-thaw cycle in [292] produced vesicles that fused with each other, which was obvious from the very large vesicles of lipid and protein produced by the freeze-thaw method of reconstitution. The method has been applied to proteins other than the glucose transporter [295-298]. The reconstituted proteins in these cases span the membranes of the reconstituted systems, as reflected by transport activity.

Hepatic asialoglycoprotein receptor Plasma membranes of hepatocytes contain a

receptor for desialylated glycoproteins [299]. The receptor has been solubilized and purified by detergent treatment of an acetone powder of liver [299]. The detergent-depleted pure protein, which aggregates to a significant extent, contains five polypeptides ranging in size from 37 to 270 kDa. The pure, lipid-free proteins do not bind ligands, but they do function as a receptor for asialoglycoproteins after reconstitution by addition to sonicated ULV's of DPPC [32]. Several lines of evidence suggest that the proteins inserted spontaneously into bilayers of DPPC at room temperature. Lipid-protein complexes were isolated by density gradient centrifugation. There were lipidinduced changes in the physical properties of the proteins; and there were protein-dependent changes in the physical properties of the lipids. It was not determined whether all proteins of the purified fraction inserted into the ULV. Interestingly, formation of the lipid-protein complexes did not cause leakage of the internal content of ULV, which contrasts with results for formation of complexes between DPPC and apoA 1. Several important aspects of the interaction between ULV's and the receptor for asialoglycoproteins remain to be established. For example, if the receptor for asialoglycoproteins inserts spontaneously into pure ULV of DPPC in a gel phase, then this protein might have a mechanism for insertion different from those directing the insertion of other membrane-bound proteins of similar or smaller size. This mechanism might not be a property of the phospholipids, but depends on special properties of the receptor for asialoglycoproteins, which is in contrast with the data for UDPglucuronosyltransferase, bacteriorhodopsin, cytochrome c oxidase, apoproteins of serum lipoproteins and several other large hydrophobic or amphipathic proteins. On the other hand, it is possible that the DPPC used in Ref. 32 was contaminated with impurities that promote spontaneous insertion of membrane-proteins. This idea is not just a theoretical possibility. Zakim and Scotto (unpublished data) have found that commercial preparations of PC from all suppliers can be contaminated with uncharacterized fusogenic compounds. They have evidence too that contaminating compounds that do not promote fusion when present alone may do so when present in the same

51 ULV. Commercial preparations of PC have been found to be contaminated with fusogens of this second type. Finally, Zakim and Scotto have also found that some membrane proteins inserted spontaneously into vesicles of 'pure' DMPC in a gel phase, but not into vesicles made from the same lots of DMPC that were repurified by the method in Ref. 300. These observations underline an important problem of reconstitution studies. It is critical to define carefully the conditions for reconstitution in order to develop a rational basis for proposing ideas about mechanism(s) for spontaneous reconstitution. Apocytochrome c Cytochrome c is attached to membranes by electrostatic interactions with acidic phospholipids [301]. It is not an integral membrane protein. The protein nevertheless is interesting in the context of the spontaneous assembly of proteins into membranes because apocytochrome c is an integral membrane protein as it is transported across the outer mitochondrial membrane [302,303]. Although it has been suggested that the transport of apocytochrome c across the outer mitochondrial [304] membrane is via pores in this membrane, there is considerable evidence that the protein spontaneously can span synthetic phospholipid bilayers [63,305-307]. Cytochrome c is present in cells in the inner mitochondrial membrane. But apocytochrome c is synthesized in the cytoplasm [308,309]. The apoprotein is transported intact across the outer mitochondrial membrane; the heme moiety is added to it at some point in this sequence [310,311]. The holo protein does not penetrate the outer mitochondrial membrane, and in vitro its interactions with phospholipid bilayers differ markedly with those for apocytochrome c. Therefore, addition of heme to apocytochrome c, within the cell, causes the cytochrome c to be 'trapped' in the space between the outer and inner mitochondrial membranes. Its attraction for acidic phospholipids will cause the protein to become associated with the outer leaflet of the inner membrane, which is the site of its biologic function. The process by which apocytochrome c incorporates into membranes in vitro is extremely interesting. It is possible now to explain a great deal of

the process by which it enters mitochondria simply on the basis of spontaneous interactions between it and phospholipids. The first step in the transport of apocytochrome c across membranes is binding to acidic phospholipids. Avidity for these phospholipids is better for PG and DPG versus PS [301,305]. Subsequent to binding, apocytochrome c penetrates the apolar span of the bilayer to reach the inside of the vesicles. Proteolytic enzymes trapped within vesicles will partly hydrolyze apocytochrome c added to the outside of vesicles [63,305,312]. Penetration of the protein to the inside of vesicles of egg PC and PS from bovine brain depended, however, on the concentration of PS in the vesicles [305]. Thus, although binding of apocytochrome c to vesicles with less than 15 mol% PS could be demonstrated, penetration of protein to the inside of the vesicles occurred only for higher concentrations of PS. These results suggest that penetration depended not only on binding but also on some structural disorganization within the bilayer. Of possible relevance to this aspect of the problem of entry of apocytochrome c into the apolar region of the membrane was that penetration depended not only on the amount of acidic phospholipid but also on its 'quality'. Penetration into the apolar region of the membrane was facilitated by increasing unsaturation of PS [305]. Also, mixed acidic phospholipids from mitochondria were more effective in this regard than equal amounts of PS. Dumont and Richards found that penetration of protein into the apolar region of vesicles was temperature-dependent. There was no apparent insertion at 0 o C vs. 37 o C [63]. Unfortunately it is difficult to discern in this work the exact compositions of vesicles used in different experiments. In addition to the importance of the physical state of the acyl chains of bilayer phospholipids, clustering of complexes of PS-apocytochrome c could be important for transfer of the protein across the bilayer, possibly via effects on the bilayer on protein. This idea is based on evidence for clustering of protein in the vesicles. Thus oligomers of apocytochrome c form in vesicles containing greater than 30 mol% PS [305]. These clusters might delineate defect structures in the bilayer and/or they could provide the stimulus for unfolding/refolding of the apocytochrome c to

52 provide a conformational isomer that can interact favorably with the apolar interior of the bilayer. Apocytochrome c does not have a cleavable signal sequence, and it has the same amino-acid sequence as cytochrome c [313,314]. Nevertheless, the conformations of apo- and holocytochrome c are quite different. The CD spectrum of the apoprotein is featureless, whereas the holoprotein contains considerable amounts of a-helix [307]. It is suggested from studies of mixtures of apocytochrome c and negatively charged or neutral detergents, that binding to negatively charged lipids induces the C-terminal region to form an a-helix that inserts into and spans the bilayer [63,307]. It seems that the C-terminal end of apocytochrome c is the portion of the molecule that crosses the membrane [63]. It is suggested too, however, that this is the portion of the protein that recognizes the mitochondrial binding site [303]. Obviously, a considerable amount of important detail must still be elucidated before these processes will be understood completely. For example, apocytochrome c binds with high avidity to acidic phospholipids. This then leads by a combination of electrostatic and presumably hydrophobic interactions to penetration of the protein into the apolar region of the membrane. Yet, binding is easily displaced by addition of more protein [305]. This is difficult to understand in the case that the protein spans the bilayer. Perhaps, however, the presumed hydrophobic interactions (the C-terminal end of the protein within the membrane) do not add to the overall lowering of free energy in the transfer of protein from water to membrane. It may be that the energy of the electrostatic interactions between apocytochrome c and acidic phospholipids is used to drive the protein into the apolar region of the membrane. Binding of apocytochrome c to suitable membranes does not occur in the presence of salt [63,306], which is not unexpected. It would be useful to know whether binding is reversed by salt. In addition to these questions it is not clear how the apocytochrome c eventually is 'pulled through' the outer mitochondrial membrane to reach the inter-membrane space. Nevertheless, there are sufficient data to show that the protein can enter membranes in the absence of mechanisms other than the consequences of lipid-protein interactions.

Coat protein of coliphage M13 The coat protein of coliphage M13 spans the inner membrane of Escherichia coli [315]. The mature protein is 6500 Da. It is synthesized on soluble polysomes as a procoat that contains a 23-amino-acid leader sequence on the N-terminus, which is cleaved after assembly into the membrane [316,317]. It has been thought, as for apocytochrome c, that insertion of procoat into membranes might require a translocase [318]. It has been shown recently, however, that the procoat protein inserts spontaneously into vesicles made from E. coli phospholipids [9]. Whether insertion depends on the types of phospholipid comprising the bilayer remains an open question. Nevertheless, protein inserted into membranes made from E. coli phospholipids spans the membrane. Entrapped chymotrypsin cleaves the procoat at some site beyond the 23-amino-acid leader sequence [9], i.e., the fragment remaining after cleavage is smaller than coat protein. Tubulin It seems to us (and this point will be elaborated on in following sections) that a better understanding of the events associated with the spontaneous insertion of proteins into preformed bilayers will depend in part on studies of the kinetics of the process. Such studies can provide information on the heights of energy barriers to insertion, whether a variety of different 'impurities' in membranes have identical effects on these barriers, and the extent to which the barrier heights depend on the nature of the proteins being inserted. These kinds of data will be useful in providing insight into detailed mechanisms, both from the point of view of membrane and protein dynamics. A major difficulty in approaching this problem experimentally is that most of the membrane proteins of interest, e.g., bacteriorhodopsin, UDPglucuronosyltransferase, etc., are aggregates of uncertain size and dispersity. There is reason to believe, however, that tubulin may be a suitable protein for beginning to study the detailed kinetics of the spontaneous insertion of proteins into preformed membranes. This is true even though tubulin is considered to be a soluble protein. Tubulin has been reported, however, to be an integral component of some membranes [319,320]. We include a brief discussion of this protein because it is a

53 soluble protein that can have the properties of an embedded, membrane protein [320,321]. Tubulin, consisting of A and B subunits of 50 kDa each, induces a rapid leakage of the contents of ULV's of DPPC at Tm [321]. This observation suggests that tubulin can insert into bilayers of DPPC. More important, vesicles of tubulin incorporated into DPPC have been isolated by density-gradient centrifugation [321]. Tubulin appears to form disc-like complexes when mixed with vesicles of DMPC [320]. Protein bound to membranes of DPPC was protected partially from digestion by trypsin. There was a minimal protection of the A subunit, but substantial protection of the B subunit in the bound form [321]. (Eventually both subunits were degraded completely by trypsin.) The fluorescence of tubulin and the content of a-helix increase for bound as compared with soluble tubulin [321]. Hence the protein has optical properties that can be used to follow the timecourse of its insertion into bilayers.

III. The phenomenon of spontaneous incorporation of embedded proteins into preformed bilayers One of the key features for successful incorporation of delipidated proteins into preformed bilayers appears to be the state of organization of the bilayer. Bilayers conducive to spontaneous incorporation of large membrane proteins are achieved by incorporating 'impurities', such as fatty acids, lysophospholipids, mixtures of structurally different phospholipids, cholesterol, detergents and membrane-bound proteins. Obviously, there are many other ways to introduce defects in the uniform packing of lipids in bilayers, including the presence of non-bilayer phases [6]. In the following section we explore certain mechanistic aspects of the phenomenon of spontaneous incorporation of proteins into bilayers, without specific regard for the exact membrane defects that promote this event.

account for spontaneous incorporation of all proteins into bilayers. Nevertheless, to a first approximation, the kinetic and thermodynamic constraints of lipid-protein interactions can be viewed in the context of the general problem of solute-bilayer interactions. Consider the interaction of a solute, S, in the biphasic bilayer-water system. The overall equilibrium distribution of the solute between the two phases will be determined by the free energies of interaction of the solute with itself, with water, and with the bilayer. The overall process for incorporation of S into a preformed bilayer can be viewed at several levels of complexity. Some, but not all possible alternative pathways, are illustrated in Figs. 1 and 2. For small solutes and monomeric proteins in water, partitioning or binding equilibria are dominated generally by hydrophobic effects on the solute. A variety of hydrophobic solutes, including small peptides and proteins, incorporate preferentially into bilayers because of an overall gain in the free energy of the system, which arises from desolvation, conformational changes, and perturbation of the organization of the bilayer. The rate of incorporation of small solutes into bilayers appears to be diffusion limited. This implies that interfacial factors, like the hydration layer, do not offer a significant kinetic barrier. Breaking of hydrogen bonds between a solute and water does contribute 2-3 kcal/mol to activation energy [322,323]. For larger solutes the rate of incorporation will be determined by conformational changes that may be necessary to accommodate the protein in the bilayer. Thus, the

A,

~D

it

III.A. The mechanism of incorporation of proteins into preformed bilayers M+S

The literature reviewed in Section II substantiates that a single mechanistic description will not

"

M'S

S n '~-

S + M--,,~S •M

Fig. 1. Reaction coordinate diagrams for incorporation of a monomeric solute (S) into bilayer when the solute exists as a monomer (A) or as an aggregate (S,,) in the aqueous phase (B).

54 A.

g.

LL

M+Sn

. M'Sn

M

l, M ' + S n ,

p M'Sn

Fig. 2. Reaction coordinate diagram for spontaneous incorporation of aggregated solute (S,,) into a bilayer without defects (M), (A), or with defects (M*), (B). The ordinates in this figure are not meant to be comparable to those in Fig. 1.

activation energy barrier for incorporation of small solutes into bilayers (S--, MS) is relatively small, and the energy profile of the reaction coordinates would be as shown by Fig. la. Many hydrophobic solutes (and membrane proteins) aggregate in the aqueous phase. The ground state energy of the solute in the aggregated state, or within the bilayer, is lower than for monomers in the aqueous phase. Transfer of monomers of solute (S) from aggregates in water (Sn --, nS) to a bilayer (Sn + M ---, MS) could be a relatively slow process that is limited by the concentration of monomeric solute in the aqueous phase. The reaction coordinate for this process is given in Fig. lb, which is different from Fig. l a only in the sense that the initial state of S is aggregated in lb (Sn) but not in la. Here the activation energy for incorporation of the solute into the bilayer (MS) is apparently high because the ground state for the solute molecules, present as aggregates in water (S,), is lower than that for monomers (S). For example, the apparent rate of transfer of carcinogenic polycyclic hydrocarbons from water to vesicles can be very slow because the hydrocarbons are dispersed as microcrystals in the aqueous phase [324,325]. For similar reasons, the transfer of proteins from aggregates to monomers in water to bilayers also could be slow. Other processes for which the energy profile of the reaction coordinate is expected to be dominated by a relatively high activation energy, due to lower ground state energy of the reactants, include fusion of vesicles, intervesicle transfer, and trans-

membrane movement of protein and phospholipids. The ground state energy of a bilayer can be increased by introducing impurities. The putative effect of impurities is the formation of 'organizational defects' that act as sites for fusion of vesicles, not only with other vesicles, but also with aggregates of protein. In such cases an aggregate of protein molecules can be transferred en masse to a host vesicle as is observed for sheets of bacteriorhodopsin and aggregates of UDPglucuronosyltransferase [240,271]. The rate of spontaneous insertion of this type is too low to measure when the vesicles are annealed and free of impurities. As shown in Fig. 2, the activation energy for the process of spontaneous incorporation of aggregated proteins apparently is lowered by defects because the ground state energy of the vesicles is increased in the energy profiles of the reaction coordinate. M in Fig. 2a is the ground state of the membrane without defects. M* in Fig. 2b is the ground state of the membrane with defects. The energy coordinates are not meant to be comparable in Figs. 1 and 2. It should be noted that, according to the scheme outlined in Figs. 1 and 2, defects would increase the free energy gain for the overall process of insertion. Theoretically, defects or impurities could lower the activation energy for insertion without influencing the ground-state energy. Such a mechanism requires the formation of a unique transition state in which defects and proteins participate. It might apply in the case of a protein like apocytochrome c in which entry into the apolar region of the bilayer depends on prior binding to the bilayer interface. It would be a highly unlikely mechanism, however, for insertion into the bilayer of proteins like bacteriorhodopsin. An increase in the ground state energy of vesicles secondary to the presence of defects has several interesting consequences. From the vantage of this review, organizational defects in a bilayer effectively will decrease the apparent activation energy for insertion of proteins and increase the free energy change associated with incorporation of proteins. Aggregated proteins will be incorporated into preformed vesicles by a fusion type of mechanism. Incorporation of monomeric proteins also could be promoted by defects because the

55 transfer of protein from water to bilayer would lower the overall energy of the system. Similarly, an anchored protein like phospholipase A 2, which does not penetrate into the acyl chain region of the bilayer, also appears to require defects in order to bind to a bilayer. The defects in this specific case appear to dislodge a molecule of substrate from association with other phospholipids. By binding at the active site, this molecule of phospholipid anchors the protein to the interface. The significance of defects for increasing the ground state energy of vesicles is implicated too in the observation that the same state promotes fusion of vesicles, leakage of polar solutes, enhanced rates of transbilayer and intervesicle transfer of phospholipids [51,59,170,184,325a], as well as the spontaneous incorporation of proteins. Thus, a state of a membrane with a higher than minimal energy in the ground state facilitates incorporation of proteins and promotes other processes that require a certain type of modulation of bilayer organization. In order to elaborate the possible molecular basis of the phenomenon of incorporation of proteins into preformed bilayers, we discuss some of the relevant energetic, structural, organizational, and motional features of bilayer organization that bear on the formation and the role of defects.

III.B. Defects vs. general disorder Organizational defects in a bilayer can be viewed in the context of two extreme assumptions. First, if the average behavior of all the lipid molecules in a bilayer is considered, the phase properties of the bilayer can be described in terms of bulk averaging parameters like fluidity [326-328], hydration layer at the interface [329], interfacial tension, surface free energy, compressibility, bending elasticity or elastic deformation [330,331]. On the other hand, if the motion and organizational parameters of the components of the bilayer are not averaged on the time scale of the functions under consideration, a distinct local environment will be sampled by interacting molecules. The properties of this microenvironment can be significantly different from the average properties of the bilayer. This microenvironment is thus a 'defect' in the bulk average behavior [332,333].

If the properties of molecules involved in a given process are similar to the average properties of the system, a description in terms of the bulk properties is a reasonable approximation. Such an attempt to describe the space and time-averaged behavior of all the molecules in a system provides a biophysical rationale for terms like fluidity and elasticity. Of necessity, this treatment does not consider and hence underestimates the contribution of local environments (defects) that depart significantly from the average. All attempts to distinguish local disorganization and general disorder, as embodied in the above discussion, are vague abstractions. Nevertheless, like all generalizations, they serve the useful purpose of polarizing dialectics.

The hydrophobic effect Formation of bilayers and insertion of small solutes into a bilayer are believed to be driven by entropic factors that give rise to the hydrophobic effect [334,335]. The basis of the hydrophobic effect is probably quantum mechanical [336]. Solute-water interactions on a descriptive level are of the van der Waals type. Interactions are repulsive at small separations; they are attractive at large separations because of dipole-induced dipoles and dispersion interactions [335]. At a qualitative level, hydrophobic groups exposed to water tend to arrange water molecules in their vicinity, yet permitting a solvent exchange on a picosecond scale, as happens in bulk water. The free energy for transfer of hydrophobic groups away from water is favorable because it increases the disorder of water molecules. In other words, a sufficiently attractive solvent-induced interaction causes solutes to form a second phase. In thermodynamic terms, enthalpic contributions promote solubility of hydrophobic solutes in water. However, favorable (negative) enthalpy is counterbalanced by unfavorable (negative) entropic contributions. The negative entropy of solution also implies that liquid water becomes a less favorable solvent for hydrophobic solutes as temperature is increased. The free energies for transfer of -CH 2- groups or hydrophobic amino-acid side-chains from water to a hydrocarbon-like phase are respectively - 2 . 5 to - 3 . 5 k J / m o l and - 5 to - 1 5 k J / m o l [232].

56 For amino acids, only the transfer of aliphatic and aromatic side-chains from water to apolar regions of a bilayer is favored. Transfer of all "other functional groups is energetically unfavorable, i.e., there is a positive free energy of transfer. Thus, the entropy-driven hydrophobic effect is the major thermodynamic force for forming and stabilizing the folded structure of proteins as well as for aggregation of delipidated membrane proteins. Other manifestations of the hydrophobic effect include low solubility of hydrocarbons in water, the partitioning of hydrocarbon chains into a nonpolar medium, adhesion, wetting, and fusion properties of surfaces, and aggregation of phospholipids to form micelles, monolayers, and bilayers. Under most experimental conditions the kinetic barrier for the formation of aggregated phospholipid structures, or for incorporation of small solutes into bilayers, is relatively insignificant, but the overall change in the free energy is substantial due to the hydrophobic effect.

Motional properties in aggregates of phospholipids Within the constraints of the hydrophobic effect, a bilayer is only one mode of organization that can maximize the total entropy of lipid-water systems. The most stable organization of the monomers in aggregates of phospholipid depends on the shapes and sizes of the molecules. The native and induced (by temperature, p H and salt concentration) molecular geometry of a phospholipid molecule is the single most important factor determining the state of aggregation. Highly curved structures like micelles are the preferred state for cone-shaped molecules with a polar group that is larger than the cross-sectional area of the acyl chains, e.g., salts of fatty acids and lysophospholipids. On the other hand, the hexagonal II phase is the preferred phase for wedgeshaped molecules because the polar group is smaller than the cross-section of the acyl chains, as in the case of highly unsaturated PE and Ca. D P G [6]. Several weak interactions and perturbations can be accomodated within the gross thermodynamic and geometric constraints of the bilayer. Based on the characteristics of the thermotropic phase transition, for example, an increasing degree of thermally induced motion, within the bilayer, is

accommodated on going from the sub-gel to gel to fluid phases. All-trans acyl chains can rotate about their long axis without significantly disrupting the lateral organization in the hexagonal lattice of the gel phase; but this rotation is not possible in the sub-gel phase in which the chains are close-packed in a pseudo-crystalline lattice. Introduction of gauche conformers, due to rotation about C--C bonds, leads to a cooperative gel to fluid transition because formation of gauche conformers in the hexagonally-packed acyl chains must be associated with similar changes in adjacent chains. The distortion caused by a gauche conformer is accomodated by formation of kinks, i.e.~ the simultaneous appearance of gauche conformers in neighboring C - C bonds [337]. The consequences of rotation and kink formation in acyl chains on the organizational order and disorder within the bilayer can be discussed at several levels. The following discussion relates largely to the induction of locally disordered regions in the plane of a bilayer [332,333]. The thermally and geometrically induced disorder due to formation of gauche conformers, in a bitayer of a single species of phospholipid molecules, will be distributed uniformly in the plane of the bilayer on a space-time averaged basis. However, at any given instant the disorder may be 'squeezed' away into regions of relative disorder. These then give rise to line defects and grain boundaries that separate domains of relatively organized structure. Coalescence of the disorganized conformers into a single region of a bilayer maximizes the extent of domains with hexagonal packing and minimizes the number of acyl chains exposed to water. Hence, concentrating the disorganization at a few points in the plane of a bilayer is favored energetically. Such considerations suggest that there will be lateral phase separation in the plane of the bilayer in the presence of solutes or co-dispersions of structurally different phospholipids. Again, the coalescence of disordered molecules can form phaseboundaries. Yet another factor could operate in bilayers containing two or more components. The transgauche motion of acyl chains in a bilayer occurs on a time scale of about 10-100 ps. The lifetime of the collated defects in such a bilayer will be in

57 this range because on a time-averaged basis all the molecules within the bilayer have equal probability of assuming any of the thermally induced conformational states. The lateral interactions between structurally different molecules in a bilayer are different in this context. The organizational disorder at the interface between different molecules in a bilayer would be time-averaged on the basis of the time-scale for diffusion of molecules across the organized domains, which is more than 10 ns. Such considerations suggest that the lifetimes of organizational defects in a bilayer are enhanced significantly in mixed bilayers or in bilayers containing impurities versus pure single species of phospholipid. It seems clear from the above that packing defects will exist in membranes composed only of a single pure lipid and that the lifetimes of true defects can be modulated by the presence of impurities. Even excluding the presence of tens to hundreds of species of proteins and lipids other than phospholipids, biological membranes still are complex mixtures. The presence of mixtures of phospholipids increases the potential for defect structures simply because of possible lateral phase separation. But, in addition, lipid-water systems exhibit organizational polymorphism. Two or more organizational forms can coexist under certain conditions. They can be induced thermotropically or isothermally by addition of proteins, ions and amphipaths [6]. For example, besides the gel and fluid phases, fluid bilayers can co-exist with regions of hexagonal II phase [338]. Nonrandom lateral organization in a bilayer can be induced by geometrical constraints, specific intermolecular interactions between the apolar or polar groups [332,336], by hydrogen bonding [178], by calcium bridging [339], or by interdigitation of acyl chains in the two monolayer halves of the bilayer [340,341]. Such phases appear to coexist over narrow but well-defined ranges of conditions. All these factors are known to contribute to disorder in bilayer organization. The lifetime of defects Questions related to the density, size and lifetime of defects in bilayers can be posed at this stage of our understanding, but cannot be answered. Defects may be considered localized if

they are long-lived as compared with the time constant for the molecular processes occurring at such sites. Fluctuations occurring in the nanosecond range would barely influence processes related to permeability. If, however, a significant fraction of the defects have life times of more than a few microseconds, incorporation of proteins and permeability of solutes could be enhanced. Stabilization of defects to the range of seconds may be required for enhancement of fusion and transmembrane movement. IlL C. The state of proteins and bilayers in reconstituted systems With regard to the energetics for spontaneous incorporation of proteins into vesicles, it is worthwhile to consider the effect of proteins on the structure of bilayers. Embedding a protein molecule into a bilayer perturbs the organization of the lipid molecules. Such perturbations, by themselves, would be energetically unfavorable as locally they induce trans to gauche (free energy change AG = +0.4 kcal/mol) conformational changes. On the other hand, defects and intrinsic instabilities (latent or expressed) in the organization of the bilayer could promote lipid-protein interactions because defect sites or regions can accommodate a protein molecule without inducing additional energetically unfavorable general disorder, desolvation, and lateral compression. Integral membrane proteins, like myelin proteolipid and Ca 2+-Ca-ATPase, for example, disorder chains and prevent chain ordering in the bulk gel phase [69]. Lower enthalpies for gel to fluid transitions in the presence of proteins also suggest that some lipid molecules are prevented from participating in a cooperative phase transition. These observations imply that most proteins and peptides remain in a relatively disordered region, irrespective of the state of the bulk phase [342]. Hence, it is not unexpected that freeze-fracture electron micrography shows that myelin lipoprotein [122] and glycophorin [94] are clustered along phase boundaries in bilayers. Incorporation of the hydrophobic helices of an integral membrane protein into bilayers of phospholipids leads to leaky vesicles, as occurs with bacteriorhodopsin and glycophorin. It is believed that the leakage pathways

58

arise from the mismatch between a transmembrane hydrophobic helix and neighboring acyl chains. Apparently, unsaturated lipids or mixtures of lipids fit better into the rugged edges of a helix as compared with saturated acyl daains [92]. A precise matching of the length of a hydrophobic helix (each amino-acid residue contributes 1.5 to the length of the helix) with the thickness of the bilayer also is important in regulating the phase state of the reconstituted bilayer [131].

Desolvation of the interface and possible role of ionic interactions As has been pointed out by others (and in Section II), there is a striking correlation between conditions that promote spontaneous insertion of proteins into preformed vesicles and conditions that promote fusion between vesicles [4,278,274]. Moreover, in both processes there must be exposure of hydrophobic surfaces on both 'partners' in the fusion event and removal of water from the surfaces of each'partner. It has been proposed in fact that the close approach of the partners in intervesicle fusion is limited by removal of surface water [329]. Intrusion into, or exposure of a hydrophobic surface, or specific ionic interactions on the bilayer interface could lead to desolvation. However, desolvation is a secondary process, and other manifestations of such a desolvation may include stabilization of ionic interactions. Although desolvation has obvious importance in determining the overall thermodynamic stability of ionic interactions, desolvation cannot be considered a kinetically rate-limiting step. Thus, the rate of exchange of water molecules at a solvated interface is of the same order of magnitude as in bulk water [322-324] and its activation energy is less than 5 kcal/mol. An array of cationic groups on polymyxin could bind tightly to the polar region of an anionic phospholipid. The resulting complex would remain anchored in the bilayer through the acyl chains. The ability of polymyxin to lower significantly the phase transition temperature of a bilayer can be accounted for by the possibility that the molecule of phospholipid bound to polymyxin is displaced, leading to a loss of overlap between its acyl chains and those of its neighbors. As discussed already, anchoring of phospholipase A 2

to the substrate interface appears to be dominated by ionic interactions; yet, hydrophobic interactions within the bilayer are also modulated [133,136]. In such cases as diphtheria toxin, signal peptide, apocytochrome c and factor Va, the initial interaction between the peptide and the bilayer is ionic, which may perturb the bilayer and expose thereby the hydrophobic surface of the protein to the hydrophobic regions of the bilayer. Thus initial ionic interactions could stabilize hydrophobic interactions between a peptide and a bilayer, probably by desolvating the intervening region. An increase in helical content secondary to ionic binding of proteins to lipids suggests formation of the appropriate elements of secondary structure required for stable protein-lipid interactions. Functionally, such interactions could lead to translocation, catalytically active conformation, and aggregation. Interaction of polymeric ionic arrays on a protein with complementary ionic arrays on a bilayer interface has interesting consequences, especially if such an interaction has the character of binding to a site. Ion-site interactions involve desolvation of ion and the site [340a]. Binding of Ca 2+ to anionic phospholipids shows considerable specificity for the head group. Similarly, intermolecular repulsion in bilayers of anionic phospholipids could be overcome by cationic groups on a protein. Under appropriate conditions such factors could contribute to the overall binding energy.

Integrity of the bilayer containing proteins The organization of bilayers is modified by incorporating a solute, which can modulate the free energy change for the overall process of insertion of the solute. In some cases this effect is subtle and can be quantitated by changes in the organizational and motional parameters of phospholipids [3]. At the other extreme, some solutes cause extensive reorganization of bilayers so that the integrity of the bilayer is lost [343]. The consequences of protein incorporation for the properties of the bilayer sometimes are not considered. For example, a leaky reconstituted system may be adequate for enzymatic or spectroscopic studies. By contrast, the osmotic integrity of a bilayer is critical for evaluating transport in reconstituted systems in which parallel leakage pathways could

59 appear as protein-mediated translocation. Leakage due to mismatch between proteins and lipids in a bilayer could account, at least in part, for the apparent lipid specificity of reconstituted translocators. Leakage pathways due to lateral mismatch of the acyl chains with the surface of the protein can, as mentioned, be compensated for by mixtures of lipids as opposed to a single species of phospholipid. In any case, interactions of integral membrane proteins with bilayers, which are dominated by insertion of hydrophobic helices, retain the gross integrity of bilayers. Transmembrane channels formed by hexameric a-toxin of Staphylococcus aureus and several other multimeric proteins (gap junction connection, acetylcholine receptor, complement attack complex, porin) appear to be predominantly in the r-sheet conformation [153]. The detailed organizational significance of this is not clear. It is possible, however, that the polar face of the r-sheet forms the aqueous side of the transmembrane channel. Similarly, several toxins undergo a conformational change that exposes hydrophobic domains for the entry of the protein itself into the cell [151] or for the formation of channels by aggregation in the membrane [152]. Certain cationic peptides solubilize anionic phospholipids from vesicles. A similar phenomenon is observed with zwitterionic vesicles at their phase-transition temperatures (cf. Table I). The particles produced by calcitonin, glucagon, apolipoproteins and related model peptides appear to be disks of 100 to 600 ~, diameter with a thickness corresponding to that of a bilayer. One of the most interesting structural features of these proteins is segments of primary sequence in which every third or fourth residue is hydrophobic [28,29]. Helices of such segments will have one hydrophobic face and one hydrophilic face. That is, the helix will have amphipathic faces, which is a structural organization well-suited for stabilizing a disk of bilayer because the hydrophobic face of the amphipathic helix binds to the acyl chains exposed at the edges of the disk. In this context it is interesting to note that peptides like mellitin, which form a hydrophobic helix and probably insert through the bilayer, do not solubilize zwitterionic vesicles. The mellitin containing vesicles are leaky and fuse readily [110], presumably due

to the phase separation induced by mellitin. Interaction of apocytochrome c with a lipid-hke environment apparently leads to a significant increase in the helical content of the protein. Hydrophobic regions in these helices could play a role in the translocation of apocytochrome c across membranes. The bilayer-protein interactions involving the polar and the glycerol backbone regions induce not only leakage but also fusion of vesicles. Such interactions predominate with anchored proteins. Hydrophobic interactions between proteins and bilayers can also come into play following ionic interaction of a solute with the charges on the bilayer surface. The molecular basis for such an 'ion-lock' mechanism for lipid-protein interaction [137] is implicit in the structural element of the amphipathic helix. In an amphipathic helix the charged residues will exhibit a specific distribution [26-28]. These charged residues also form topographically close complementary ion pairs, and interact appropriately with the anionic and cationic groups on a zwitterionic lipid. Such interactions can contribute significantly to the stability of a lipid-protein complex.

Thermotropic phase-transition properties of bilayers containing proteins The gel to fluid phase transition in bilayers of saturated phospholipids has been used to investigate the effects of lipid-protein interactions on phase properties and organizational defects. A change in the transition profile indicates a modification of the environment of the acyl chains. Alamethicin [98], gramicidin, Ca2+-ATPase, and bacteriorhodopsin [344] increase the gauche content in the gel phase, and lower it in the fluid phase. In general it is very difficult to interpret the thermotropic transition data. A direct contribution to the heat capacity from free or bound proteins is generally irreversible, and its not observed in the cooling runs or in repeated heating runs. Transition profiles are modified not only by modulating the local environment of the acyl chains but also by long-range factors. These include any one of the numerous solute-induced changes in the macroscopic organization of the bilayer such as formation of interdigitated bilayer,

6O

solubilization leading to a disk of bilayer structure stabilized by proteins, and induction of hexagonal and cubic phases. In cases in which it can be demonstrated clearly that the gross organization of the bilayer is retained in the reconstituted system, on a macroscopic level, the thermotropic transition data can be used for quantifying the extent of binding. However, caution is needed in extrapolating such corroborative information, which may be obtained at low concentrations and high lipid-to-protein ratios. The conditions usually are exactly opposite for the calorimetry runs. A detailed phenomenological theory explaining all aspects of the gel to fluid transition in bilayers is not available yet. Some insights may be obtained, however, from the phase rules developed for transitions from bulk solid to bulk liquid. These extrapolations have been quite successful in providing an understanding of the phase diagrams of mixed lipid systems [44,132,338]; however, no mechanistic insights are available, and overextension of these rules to protein-bilayer systems can be misleading. The effect of additives on the freezing point of bulk liquids usually is interpreted in terms of Raoult's law or the Clausius-Claperon equation. In its simplest form the change in the transition temperature induced by a solute is given by: ~H

AHm

T

rm

RT In Xg/Xf

(4)

Tm and T are the transition temperatures and A H m and A H are the associated enthalpies in the absence and presence of the solute. R is the gas constant, Xg and Xf are the mole fractions of the solute in the gel and the fluid phases, respectively. This relationship will be valid for ideally mixed solutes that do not perturb the organization of the bilayer. Obviously, these conditions are not satisfied in reconstituted bilayers. Therefore, interpretation of the phase transition data, in terms of Eqn. 4, does not provide information on the relative affinities of a protein for gel and the fluid phases. Moreover, most of the approximations described in the literature assume that A H = A H m" This creates a major difficulty because A H / T is small and highly sensitive, therefore, to small changes in A H.

IV. Biologic implications of spontaneous incorporation of proteins into membranes Review of the literature provides many examples of the spontaneous assembly of proteins into preformed membranes. Although such events have been observed under conditions that might not occur in vivo, it is important to emphasize that natural components of biologic membranes, such as proteins, fatty acids and cholesterol, will promote spontaneous insertion of proteins into preformed membranes. Moreover, the complexity of membranes, with attendant possibilities for lateral phase separation and the presence of non-bilayer phases, indicates that natural membranes have the properties required in vitro for 'activating' membranes so that insertion of proteins (i.e., membrane assembly) is a facile process. Given the abundance of evidence for the spontaneous assembly of proteins into synthetic membranes and the conditions required in vitro, we think it likely that nature makes use of these mechanisms in vivo. These mechanisms may be especially important for the assembly into membranes of proteins that loop across a membrane several times. The solution that has been proposed for this problem is co-translational insertion of proteins into membranes, as an adaptation of the 'signal hypothesis' for secretion of proteins [232,345]. There are many uncertainties, however, as to whether proteins that span a bilayer several times can be inserted into bilayers by some variation of the co-transitional theme [232,345,347]. Nevertheless, the hypothesis is popular probably because there has been no other reasonable alternative to co-translational insertion of proteins into membranes, and many membrane-bound proteins have signal sequences, are synthesized on membrane-bound ribosomes, and synthesis is arrested in the absence of signal recognition particles and a suitable receptor for signal recognition particle-bound ribosomes [10]. With regard to the first point, it has not been appreciated that the energy barrier to insertion can be lowered drastically in fairly simple ways, as reflected by the data discussed above. With regard to the second point, the linking of translation and insertion into membranes of integral membrane proteins may confound understanding of the mechanism of the latter phenomena and the bio-

61 logic significance of the former. For example, translation of a membrane bound enzyme on ribosomes bound to the endoplasmic reticulum might lead simply to the secretion of the protein into the lumen of the endoplasmic reticulum. The secreted, translated protein within the lumen of the endoplasmic reticulum could fold and aggregate perhaps with other like molecules and then insert spontaneously into the endoplasmic reticulum. One result of this process would be to alter the orientation of the protein in the endoplasmic reticulum versus what it would have if it had inserted from the cytosolic surface. The advantage to the cell of co-transitional secretion into the lumen of the endoplasmic reticulum followed by secondary insertion into the membranes of this organelle, therefore, could be proper orientation of some proteins, which could determine in turn the modes of function and regulation of a given protein as well as how the protein will undergo post-translational glycosylation. Purified membrane-proteins are aggregated, even in the presence of detergents. Spontaneous reconstitution in vitro, as reviewed in subsection II.C, therefore, often reflects interactions between bilayers and aggregates of membrane-proteins. The only data relating to this question, of which we are aware, indicate that a membrane-protein synthesized on cytosolic ribosomes is aggregated in the cytosol [348]. Mitochondrial ATP/ADP transport protein, which is synthesized on cytosolic ribosomes is aggregated prior to entry into mitochondria [348].

V. Epilog The main goal of this review has been to examine the evidence for spontaneous incorporation of membrane proteins into preformed bilayers. The experimental evidence in this regard demonstrates that monomeric as well as aggregated proteins can be incorporated readily into suitably prepared preformed bilayers. The proteins of interest may be absorbed, anchored, or embedded within the bilayer. Proteins with a relatively small extent of hydrophobic surface probably are transferred into bilayers as monomers. Aggregates of delipidated, integral membrane proteins probably are transferred en masse from water to mem-

branes by a fusion type of mechanism. It is interesting in this regard that conditions favoring fusion between bilayers also promote spontaneous incorporation of aggregated proteins as well as monomeric proteins. In those instances in which conditions were controlled carefully, the data indicate that traces of impurities (detergents, lipids, other proteins, amphipathic solutes), in bilayers of pure phospholipids, are necessary for spontaneous incorporation of aggregated membrane proteins. We have proposed, therefore, that impurities induce or increase the lifetime of defects in the organization of bilayers and that these defects are the loci for spontaneous incorporation of proteins into preformed bilayers. The defects also are probably the sites for processes like fusion, intervesicle exchange, transbilayer movement of phospholipids, and anomalous leakage of polar solutes. A feature common to all these processes is the requirement that hydrophobic regions of the bilayer become exposed to the aqueous phase, which we propose is the rate-limiting step in each of these processes. According to the scheme shown in Fig. 2b, local defects in the organization of bilayers increase the energy of the bilayers in the ground state, leading thereby to increased rates for processes like spontaneous incorporation of membrane proteins and fusion between bilayers. A mechanism for increasing the energy in the ground state is implicit in the metastability of defects, as well as in the local disorder that could accomodate the aminoacid side-chains of a protein. The nature of the proposed defects in packing within bilayers is uncertain. It is not clear whether such defects pre-exist in bilayers containing impurities, or whether they are latent, i.e., they are expressed only after an encounter with an appropriate protein. A possible role for regions of pre-existing hexagonal II phase can be excluded because in most cases there is no evidence in the data reviewed for the presence of such nonbilayer structures (cf., however, Ref. 6). On the other hand, within the constraints of bilayer organization, defects can be viewed as local features of organization required to accommodate mismatch arising from transbilayer asymmetry, lateral immiscibility, thermal motion, or geometrical constraints. Obviously, a confluence of all such fac-

62

tors is possible. A change in parameters of bilayers that are measured as time-averaged properties for the whole bilayer (e.g., fluidity, desolvation, elasticity) would be the macroscopic manifestation of defects. Unfortunately, there are no methods for measuring directly the density, location and lifetime of defects in a bilayer. We believe, nevertheless, that most defects pre-exist in the bilayer organization. Impurities affect membrane processes because they increase the lifetimes of defects. That is, long-lived defects in the organization of bilayers are sites for processes with time constants that are less than those for the lifetimes of defects. That many purified integral membrane proteins insert spontaneously into preformed vesicles has obvious practical value for the reconstitution of such proteins. In addition, observations of these systems may apply to the problem of assembly of membranes in cells. Finally, the protocols for spontaneous incorporation of proteins into lipid bilayers provide an opportunity for studying this process per se, which cannot be accomplished with other methods used to reconstitute lipid-protein complexes from purified delipidated proteins and pure phospholipids.

8 9 l0 11

12 13

14 15 16 17 18 19 20 21

22

Acknowledgements The authors thank Professors Karl Koehler and Richard Epand for their critical reading of the manuscript. Work from the authors' laboratories was supported by grants NIH GM29703 (M.K.J.) and NSF PCM 8311967 (D.Z.).

References 1 Levitzki, A. (1985) Biochim. Biophys. Acta 822, 127-153 2 Etemadi, A. (1985) Adv. Lipid Res. 21, 281-428 3 Devaux, P.F. and Seigneuret, M. (1985) Biochim. Biophys. Acta 822, 63-125 4 Eytan, G.D. (1982) Biochim. Biophys. Acta 694, 185-202 5 Semenza, G., Kessler, M., Mosang, M., Weber, J. and Schmidt, U. (1984) Biochim. Biophys. Acta 779, 343-379 6 DeKruijff, B., Cullis, P.R., Verkleij, A.J., Hope, M.J., van Echteld, C.J.A., Taraschi, T.F., van Hoogevest, P., Killian, J.A., Rietveld, A. and van der Steen, A.T.M. (1985) Progress in Protein-Lipid Interactions (Watts, A. and DePont, J.J.H.H.M., eds.), pp. 89-142, Elsevier, Amsterdam 7 Unwin, P.N.T. and Henderson, R. (1984) Sci. Am. 250, 78-94

23 24 25 26 27

28

29

30 31 32

33

Wickner, W. (1979) Annu. Rev. Biochern. 49, 23-45 Geller, B.L. and Wickner, W. (1985) J. Biol. Chem. 260. 13281-13285 Rapoport, T.A. and Wiedmann, M. (1985) Curr. Top. Membranes Transp. 24, 1-63 Scherphof, G., Damen, J. and Hoekstra, D. (1981) in Liposomes: From Physical Structure to Therapeutic Applications (Knight, E., ed.), pp. 299-322, Elsevier, Amsterdam Nicholls, P. (1981) Int. Rev. Cytol. Suppl. 12, 327 388 Hebdon, G.M., Levine, H., Shayoun, N.E., Smitges, C.J. and Cuatrecasas, P. (1981) Proc. Natl. Acad. Sci. USA 78, 120 3 Klein, J., Moore, L. and Pastan, I. (1978) Biochim. Biophys. Acta 506, 42-53 Sinha, A.K., Shattik S.J. and Colman, R.W. (1976) Fed. Proc. 35, 1714 Verhallen, P.F.J., Demel, R.A., Zwiers, H. and Gispen, W.H. (1984) Biochim. Biophys. Acta 775, 246-254 Garcia, L.A.M., Araujo, P.S. and Chaimovich, H. (1984) Biochim. Biophys. Acta 772, 231-234 Christiansen, K. and Carlsen, J. (1983) Biochim. Biophys. Acta 735, 225-233 Kohne, W., Deuticke, B. and Heast, C.W.M. (1983) Biochim. Biophys. Acta 730, 139-150 Wetterau, J.R. and Jonas, A. (1982) J. Biol. Chem. 257, 10961-10966 Andrews, A.L., Atkinson, D., Barratt, M.D., Finer, E.G., Hauser, H., Heury, R., Leslie, R.G., Owens, N.L., Phillips, M.C. and Robertson, R.N. (1976) Eur. J. Biochem. 64, 549-563 Massey, J.B., Rohde, M.F., Van Winkle, W.B., Gotto, A.M. and Pownalk H.J. (1981) Biochemistry 20, 1569-1574 Swaney, J.B. (1980) J. Biol. Chem. 255, 8791-7, 8798-8803 Swaney, J.B. and Chang, B.C. (1980) Biochemistry 19, 5637-44 Pownall, H.J. and Massey, J.B. (1982) Biophys. J. 37, 177-179 Segrest, J.P, Chung, B.H.M., Brouillette, C.G., Kanellis, P. and McGahan, R. (1983) J. Biol. Chem. 258, 2290-2295 Weinstein, J.N., Klausner, R.D., Innerarity, Y., Ralston, E. and Blumenthal, R. (1981) Biochim. Biophys. Acta 647, 270-284 Anatharamaiah, G.M., Jone, J.L., Brouillette, C.G., Schmidt, C,F., Chung, B.H., Hughes, T.A., Bhown, A.S. and Segrest, J.P. (1985) J. Biol. Chem. 260, 10248-10256 Chung, B.H., Anatharamaiah, G.M., Brouillette, C.G., Nishida, T. and Segrest, J.P. (1985) J. Biol. Chem. 260, 10256-10262 Vaughan, D.J., Breckenridge, W.C. and Stanacev, N.Z. (1980) Can. J. Biochem. 58, 592-598 Cardin, A.D., Jackson, R.L. and Johnson, J.D. (1982) FEBS Lett. 141,193-196 Klausner, R.D., Bridges, K., Tsunoo, H., Blumenthal, R., Weinstein, J.N. and Ashwell, G. (1980) Proc. Natl. Acad. Sci. USA 77, 5087-91 East, J.M., Jones, O.T., Simmonds, A.C. and Lee, A.G.

63 (1984) J. Biol. Chem. 259, 8070-1 34 Lentz, B.R., Clubb, K.W., Alford, D.R., Hochil, M. and Meissner, G. (1985) Biochemistry 24, 433-442 35 McIntosh, D.B. and Ross, D.C. (1985) Biochemistry 24, 1244-1251 36 Reirthardt, R., Lindemann, B. and Anner, B.M. (1984) Biochim. Biophys. Acta 774, 147-150 36a Jain, M.K., White, F.P., Williams, E., Strickholm, A. and Cordes, E.H. (1972) J. Membrane Biol. 8, 363-374 37 Zachowski, A. and Devaux, P.F. (1985) FEBS Lett. 163, 245-249 38 Esman, M., Watts, A. and Marsh, D. (1985) Biochemistry 24, 1386-1393 39 George, R., Lewis, N.A.H. and McElhaney, R.N. (1985) Biochim. Biophys. Acta 821,253-258 40 Ramirez, F., Okazaki, H. and Tu, S. (1981) FEBS Lett. 135, 123-126 41 Huang, K., Bayley, H. and Khorana, H.G. (1980) Proc. Natl. Acad. Sci. 77, 323-7 42 Ovchirmikov, Y.A. (1982) FEBS Lett. 148, 179-189 43 Lewis, B.A., Harbison, G.S., Herzfeld, J. and Griffin, R.G. (1985) Biochemistry 24, 4671-79 44 Alonso, A., Restall, C.J., Turner, M., Gomez-Fernandez, J.C., Goni, F.M. and Chapman, D. (1982) Biochim. Biophys. Acta 689, 283-289 45 Rehorek, M., Dencher, N.A. and Heyn, M.P. (1985) Biochemistry 24, 5980-5988 46 Keniry, M.A., Gutowsky, H.S. and Oldfield, E. (1984) Nature 307, 383-386 47 Grant, S.R., Babbitt, B.P., West, L.K. and Huang, L. (1982) Biochemistry 21, 1274-1279 48 Epand, R.M., Epand, R.F., Orlowski, R.C., Schlueter, R.J., Boni, L.T. and Hui, S.W. (1983) Biochemistry 22, 5074-5084 49 Vincent, J.S., Steer, C.J. and Levin, I.W. (1984) Biochemistry 23, 625-631 50 Steer, C.J., Vincent, J.S. and Levin, I.W. (1984) J. Biol. Chem. 259, 8052-5 51 Hong, K., Yoshimura, T. and Papahadjopoulos, D. (1985) FEBS Lett. 191, 17-20 52 Henderson, R. (1981) Membranes and Intercellular Communications (Balian, R., Chabre, M. and DeVaux, P.F., eds.), pp. 229-249, Elsevier, Amsterdam 53 Kimmelman, D., Tecoma, E.S., Wolber, P.K., Hudson, B.S., Wickner, W. and Simoni, R.D. (1979) Biochemistry 18, 5874-5880 54 Hagen, D.S., Weiner, J.H. and Sykes, B.D. (1978) Biochemistry 17, 3860-3866 55 Wiener, J.H., Lemire, B.D., Jones, R.W., Anderson, W. and Scraba, D.G. (1984) J. Cell. Biochem. 24, 207-216 56 Davidson, V.L., Brunden, K.R., Cramer, W.A. and Cohen, F.S. (1984) J. Membrane Biol. 79, 105-118 57 Akhrem, Adrianov, V.T., Bokut, S.B., Luka, Z.A., Kissel, M.A., Skornyakova, T.G. and Kisselev, P.A. (1982) Biochim. Biophys. Acta 692, 287-295 58 Freire, E., Markello, T., Rigell, C. and Holloway, P.W. (1983) Biochemistry 22, 1675-80 59 Greenhut, S.F. and Roseman, M.A. (1985) Biochemistry 24, 1252-1260

60 61 62 63 64

65 66 67 68 69

70 71 72 73 74 75 76 77 78 79 80 81 82 83 84 85 86 87 88 89

Papahadjopoulos, D., Moscarello, M.A., Eylar, E.H. and Isac, T. (1975) Biochim. Biophys. Acta 401, 317-335 Brown, L.R. and Wuthrich, K. (1977) Biochim. Biophys. Acta 468, 389-410 DeKruijff, B. and Cullis, P.R. (1980) Biochim. Biophys. Acta 602, 477-490 Dumont, M.E. and Richards, F.M. (1984) J. Biol. Chem. 261, 4147-4156 Rietveld, A., Ponjee, G.A.E., Schiffers, P., Jordi, W., Coolwijk, P.J.F.M., Demel, R.A., Marsh, D. and DeKruijff, B. (1985) Biochim. Biophys. Acta 818, 398-409 Knowles, P.F., Watts, A. and Marsh, D. (1979) Biochemistry 18, 4480-4487 Semin, B.K. and Wickstrom, M. (1984) Biochim. Biophys. Acta 769, 15-22 Powell, G.L., Knowles, P.F. and Marsh, D. (1985) Bioclaim. Biophys. Acta 816, 191-194 Rice, D.M., Hsung, J.C., King, T.E. and Oldfield, E. (1979) Biochemistry 18, 5885-5892 Rice, D.M., Meadow, M.D., Scheinman, A.O., Goni, F.M., Gomez-Fernandez, J.C., Moscarello, M.A., Chapman, D. and Oldfield, E. (1979) Biochemistry 18, 5893-5903 Young, M., Dinda, M. and Singer, M.A. (1983) Biochim. Biophys. Acta 735, 429-432 B~Ssterling, B., Trndell, J.R. and Galla, H.J. (1981) Biochim. Biophys. Acta 643, 547-556 Yang, C.S. (1977) J. Biol. Chem. 252, 293-298 Navarro, J., Chabot, J., Sherill, K., Aneja, R., Zahler, S.A. and Racker, E. (1985) Biochemistry 24, 4645-4650 Erne, D., Sargent, D.F. and Schwyzer, R. (1985) Biochemistry 24, 4261-63 Myers, M. and Freire, E. (1985) Biochemistry 24, 4076-4082 Loh, H.H. and Law, P.Y. (1980) Annu. Rev. Pharmacol. Toxicol. 20, 201-234 Van der Waart, P., Bruls, H., Hemker, H.C. and Lindhout, T. (1983) Biochemistry 22, 2427-2432 Pusey, M.L. and Nelsestuen, G.L. (1984) Biochemistry 23, 6202-6210 Jones, A.J.S., Epand, R.M., Lin, K.F., Walton, D. and Vail, W.J. (1978) Biochemistry 17, 2301-7 Ernandes, J.R., Epand, R.M. and Schreir, S. (1983) Biochirn. Biophys. Acta 733, 75-86 Epand, R.M. and Sturtevant, J.M. (1981) Biochemistry 20, 4603-4606 Epand, R.M. (1980) Trends Biochem. Sci. 8, 205-207 Carruthers, A. and Melchiors, D.L. (1984) Biochemistry 23, 2712-2718 Hah, J., Hui, S.W. and Jang, C.Y. (1983) Biochemistry 22, 4763-4769 Deleted Kapitza, H.G., Ruppel, D.A., Galla, H.J. and Sackman, E. (1984) Biophys. J. 45, 577-587 Dluhy, R.A., Mendelsohn, R., Casel, H.L. and Mantsch, H.H. (1983) Biochem. 22, 1170-1177 Rothschild, K.J. and Morrero, H. (1982) Proc. Natl. Acad. Sci. USA 79, 4045-4049 Nakanishi, M. (1984) FEBS Lett. 176, 385-389

64 90 Taraschi, T.F., DeKruijff, B., Verkleij, A.J. and Van Echteld, C.J.A. (1982) Biochim. Biophys. Acta 685, 153-161 91 Taraschi, T.F., van der Steen, A.T.M., De Kruijff, B., Tellier, C. and Verkleij, A.J. (1982) Biochemistry 21, 5756-5764 92 Van Hoogevest, P., Du Maine, A.P.M., De Kruijff, B. and De Gier, J. (1984) Biochim. Biophys. Acta 771, 119-126 93 Van der Steen, A.T.M., Taraschi, T.F., Voorhout, W.F. and De Kruijff, B. (1983) Biochim. Biophys. Acta 733, 51-64 94 Ruppel, D., Kapitka, H.G., Gala, H.J., Sixl, F. and Sackman, E. (1982) Biochim. Biophys. Acta 692, 1-17 95 Pal, R., Wiener, J.R., Barenholz, Y. and Wagner, R.R. (1983) Biochemistry 22, 3624-3630 96 Urry, D.W., Trapane, T.L. and Prasad, W.U. (1983) Science 221, 1064-1067 97 Weinstein, S., Durkin, J.T., Veatch, W.R. and Blout, E.R. (1985) Biochemistry 24, 4374-82 98 Lee, D.C., Durrani, A.A. and Chapman, D. (1984) Biochim. Biophys. Acta 769, 49-56 99 Cornell, B.A., Davenport, J.B. and Separovie, F. (1982) Biochim. Biophys. Acta 689, 337-45 100 Feigenson, G.W. (1983) Biochemistry 22, 3106-3112 101 Van Echteld, C.J.A., De Kruijff, B., Mandersloot, J.G. and De Gier, J. (1981) Biochim. Biophys. Acta 649, 211-220 102 Van Echteld, C.J.M., De Kruijff, B., Leunissen-Bijvelt, J., Verkleij, A.J. and De Gier, J. (1982) Biochim. Biophys. Acta 692, 126-138 103 Killian, J.A., De Kruijff, B., Van Echteld, C.J.A., Verkleij, A.J., Leunissen-Bijvelt, J. and De Gier, J. (1983) Biochim. Biophys. Acta 728, 141-144 104 Bamberg, E., Hageman, P. and Oesterhelt, D. (1984) Biochim. Biophys. Acta 773, 53-60 105 Kimura, H. and Futai, M. (1978) J. Biol. Chem. 252, 1095-1100 106 Hoekstra, D., Dtizgtines, N. and Wilschut, J. (1985) Biochemistry 24, 565-572 107 Grant, C.W.M. and Peters, M.W. (1984) Biochim. Biophys. Acta 779, 403-422 108 Higashijima, T., Wakamatsu, K., Takemitsu, M., Fujino, M., Nakajima, T. and Miyazawa, T. (1983) FEBS Lett. 152, 227-231 109 Habermann, R. (1980) in Natural Toxins (Eaher, A., and Wadstrom, B., eds.), p. 173-185, Pergamon Press, London 110 Morgan, C.G., Thomas, E.W., Moras, T.S. and Yianni, Y.P. (1982) Biochim. Biophys. Acta 692, 196-201 111 Eytan, G.D. and Almary, T. (1983) FEBS Lett. 156, 29-32 112 Dessux, J., Faucon, J., Lafleur, M , Pezolet, M. and Dufourcq, J. (1984) Biochim. Biophys. Acta 775, 37-50 113 Dufton, M.J., Hilder, R.C. and CheRry, R.J. (1984) J. Membrane Biol. 11, 17-24 114 Georghiou, S.M., Thompson, A.K., and Mukhopadhyay (1982) Biochim. Biophys. Acta 688, 441-452 115 Epand, R.M. and Moscarello, M.A. (1982) Biochim. Bio-

116 117 118 119 120 121 122 123 124 125 126 127 128 129 130 131 132 133

134

135 136 137 138 139 140 141 142 143

phys. Acta 685, 230-232 Bach, D. and Sela, B. (1985) Biochim. Biophys. Acta 819, 225-230 Stollery, J.G., Boggs, J.M. and Moscarello, M.A. (1980) Biochemistry 19, 1219-1226 Boggs, J.M., Wood, D.D. and Moscarello, M.A. (1981) Biochemistry 20, 1065-1073 Dufourcq, J., Faucon, J.F., Maget-Dana, R., Pileni, M.P. and Helene, C. (1981) Biochim. Biophys. Acta 649, 67-75 Sixl, F., Brophy, P.J. and Watts, S.A. (1984) Biochemistry 23, 2032-39 Curatolo, W., Sakura, J.D., Small, D.M. and Shipley, G.G. (1977) Biochemistry 16, 2313-19 Boggs, J.M., Clement, I.R. and Moscarello, M.A. (1980) Biochim. Biophys. Acta 601, 134-151 Boggs, J.M., Wood, D.D., Moscarello, M.A. and Papahadjopoulos, D. (1977) Biochemistry 16, 2325-29 Verma, S.P., Wallach, D.F.H. and Sakura, J.D. (1980) Biochemistry 19, 574-579 Lavialle, F. and Levin, I.W. (1980) Biochemistry 23, 6044-6050 Brophy, P.J., Horvath, L.I. and Marsh, D. (1984) Biochemistry 23, 860-865 Epand, R.M. (1985) Biochemistry 24, 7092-7095 Epand, R.M. and Sturtevant, J.M. (1984) Biophys. Chem. 19, 355-362 Surewicz, W.K. and Epand, R.M. (1985) Biochemistry 24, 3135-3144 Davis, J.H., Clare, D.M., Hodges, R.S. and Bloom, M. (1983) Biochemistry 22, 5298-5305 Huschilt, J.C., Hodges, R.H. and Davis, J.H. (1985) Biochemistry 24, 1377-1386 Moorow, M.R., Huschilt, J.C. and Davis, J.H. (1985) Biochemistry 24, 5396-5406 Jain, M.K., Egmond, M.R., Verheij, H.M., Apitz-Castro, R.J., Dijkman, R. and De Haas, G.H. (1982) Biochim. Biophys. Acta 688, 341-348 Jain, M.K., Rogers, J., Jahagirdar, D.V., Marecek, J.R. and Ramirez, F. (1986) Biochim. Biophys. Acta 860, 435-447 Jain, M.K., Maliwal, B., Slotboom, A.J. and De Haas, G.H. (1986) Biochim. Biophys. Acta 860, 448-461 Jain, M.K., Rogers, J., Marecek, J.F., Ramirez, F. and Eibl, H. (1986) Biochim. Biophys. Acta 860, 462-474 Jain, M.K., De Haas, G.H., Marecek, J.F. and Ramirez, F. (1986) Biochim. Biophys. Acta 860, 475-483 Xu, Y., Gietzen, K., Galla, H. and Sackman, E. (1983) Biochem. J. 213, 21-24 Wang, C.Y. and Huang, L. (1984) Biochemistry 23, 4409-16 Uster, P.S. and Deamer, D.W. (1985) Biochemistry 24, 1-8 Boggs, J.M., Rangaraj, G., Moscarello, M.A. and Koshy, K.M. (1985) Biochim. Biophys. Acta 816, 208-220 De Kruijff, B. and Cullis, P.R. (1980) Biochim. Biophys. Acta 601, 235-240 Sixl, F. and Galla, H.J. (1982) Biochim. Biophys. Acta 693, 466-478

65 144 Boggs, J.M. and Rangaraj, G. (1985) Biochim. Biophys. Acta 816, 221-233 145 Parrish, R.F., Strauss, J.W., Polakowski, K.I. and Dombrose, F.A. (1978) Proc. Natl. Acad. Sci. USA 75, 149-152 146 Davoust, J., Schoot, B.M. and Devaux, P.F. (1979) Proc. Natl. Acad. Sci. USA 76, 2755-59 147 Briggs, M.S. and Oierasch, L.M. (1986) Adv. Protein Chem. 38, 109-179 148 Mombers, C.P.W.M., Van Dijs, L.L.M., Van Deenen, J., De Gier, A.J. and Verkleij, A. (1977) Biochim. Biophys. Acta 470, 152-160 149 Deleted. 150 Hong, K., Duzgunes, N., Ekerdt, R. and Papahadjopoulos, D. (1982) Proc. Natl. Acad. Sci. USA 79, 4642-4644 151 Olsnes, S., Sandvig, M., Madshus, J.H. and Sundan, A. (1985) Biochem. Soc. Symp. 50, 171-191 152 Bhakdi, S. and Tranum-Jensen, J. (1985) Biochem. Soc. Symp. 50, 221-233 153 Tobkes, N., Wallace, B.A. and Bayley, H. (1985) Biochemistry 24, 1915-1920 154 Dufourcq, C.J., Faucon, J.F., Bernard, E., Perolet, M., Tessier, M., Bougis, P., van Rietschoten, J., Delori, P. and Rochat, H. (1982) Toxicon 20, 165-180 155 Gulick-Krzywicki, T., Baerna, M., Vincent, J.P. and Lazdunski, M. (1981) Biochim. Biophys. Acta 643, 101-114 156 Blumenthal, K.M. (1982) Biochemistry 21, 4229-4233 157 Tomasi, M., D'Agnolo, G. and Montecucco, C. (1982) Biochim. Biophys. Acta 692, 339-344 158 Tomasi, M., Battistini, A., Cardelli, M., Sonnino, S. and D'Agnolo, G. (1984) Biochemistry 23, 2520-2526 159 Richards, R.L., Moss, J., Alving, C.R., Fishman, P.H. and Brady, R.O. (1979) Proc. Natl. Acad. Sci. USA 76, 1673-77 160 Gorin, B. and Freire, E. (1985) Biochemistry 24, 1791-97 161 Zalman, L.S. and Wisnieski, B.J. (1984) Proc. Natl. Acad. Sci. USA 81, 3341-3345 162 Blewitt, M.G., Chung, L.A. and London, E. (1985) Biochemistry 24, 5458-64 163 Cabiaux, V., Van den Branden, M., Falmagne, P. and Ruysschaert, J. (1984) Biochim. Biophys. Acta 775, 31-36 164 Boquet, P. (1979) Eur. J. Biochem. 100, 483-487 165 Alving, C.R., Iglewski, B.H., Urban, K.A., Moss, I., Richards, R.L. and Sandhoff, J.C. (1979) Proc. Natl. Acad. Sci. USA 77, 1986-1990 166 Thomas, W.E. and Ellar, D.J. (1983) FEBS Lett. 154, 362-365 167 Shin, M.L., Michaels, D.W. and Mayers, M.M., (1979) Biochim. Biophys. Acta 555, 79-88 168 Linder, R. and Bernheimer, A.W. (1984) Toxicon 22, 641-651 169 Fehrenbach, F.J. and Eibl, M. (1977) Z. Naturforsch. 32, 101-106 170 Duncan, J.L. and Buckingham, L. (1981) Biochim. Biophys. Acta 648, 6-12 171 Morris, D.A.N., McNeil, R., CasteUino, F.J. and Thomas, J.K. (1980) Biochim. Biophys. Acta 599, 380-390 172 Morris, N.P., Consiglio, E., Kohn, L.D., Habig, W.H.,

173 174 175 176 177 178 179 180

181 182 183 184 185 186 187 188 189 190 191 192 193 194 195

196 197 198 199 200 201

Hardegree, M.C. and Helting, T.B. (1980) J. Biol. Chem. 255, 6071-76 Kumar, N., Klausner, R.D., Weinstein, J.N., Blumenthal, R. and Flavin, M. (1981) J. Biol. Chem. 256, 5886-5889 Olson, E.N., Towler, D.A. and Glaser, L. (1985) J. Biol. Chem. 260, 3784-3790 Magee, A.I. and Schlesinger, M.J. (1982) Biockim. Biophys. Acta 694, 279-289 K_im,B.H. and Rosenberry, T.L. (1985) Biochemistry 24, 3586-3592 Low, M.G. and Kincade, P.W. (1985) Nature 318, 62-64 Boggs, J.M. (1983) Membrane Fluidity Biol. 2, 89-130 Kitawaga, T., Inoue, K. and Nojima, S. (1976) J. Biochem. 79, 1135-1145 Rainier, S., Jain, M.K., Ramirez, F., Iannou, P.V., Marecek, J.F. and Wagner, R. (1979) Biochim. Biophys. Acta 558, 187-198 Verger, R. and De Haas, G.H. (1977) Annu. Rev. Biophys. Bioeng. 5, 77-117 Jain, M.K., Crecely, R., Hille, J.D.R., De Haas, G.H. and Grunner, S. (1985) Biochim. Biophys. Acta 813, 68-76 Apitz, R.C., Jain, M.K. and De Haas, G.H. (1982) Biochim. Biophys. Acta 688, 349-356 Jain, M.K., Streb, M., Rogers, J. and De Haas, G.H. (1984) Biochem. Pharm. 33, 2541-2553 Jain, M.K. and Jahagirdar, D.V. (1985) Biochim. Biophys. Acta 814, 319-326 Jain, M.K. and Maliwal, B.P. (1985) Biochim. Biophys. Acta 814, 135-140 Jurtchuk, P., Sekuzu, I. and Green, D.E. (1963) J. Biol. Chem. 238, 3595-3605 Vidal, J.C., Guglielmucci, E.A. and Stoppani, A.D.M. (1970) Arch. Biochem. Biophys. 187, 138-152 Bock, H.-G. and Fleischer, S. (1974) Methods Enzymol. 31, 374-391 McIntyre, J.O., Brock, H.-G. and Fleischer, S. (1978) Biochim. Biophys. Acta 513, 255-267 Miyahara, M., Nishihara, Y., Morimizato, Y. and Utsumi, K. (1981) Biochim. Biophys. Acta 641,232-241 Gotterer, G.S. (1967) Biochemistry 6, 2147-2152 Grover, A.K., Slotboom, A.J., De Haas, G.H. and Hammes, G.G. (1975) J. Biol. Chem. 250, 31-38 Gazzotti, P., Bock, H.-G. and Fleischer, S. (1975) J. Biol. Chem. 250, 5782-5790 Isaacson, Y.A., Deroo, P.W., Rosenthal, A.F., Bittman R., McIntyre, J.O., Bock, H.-G., Gazzotti, P. and Fleischer, J. (1979) J. Biol. Chem. 254, 117-126 McIntyre, J.O., Wang, C.T. and Fleischer, S. (1979) J. Biol. Chem. 254, 5199-5207 Cortese, J.D., Vidal, J.C., Churchill, P., McIntyre, J.O. and Fleischer, S. (1982) Biochemistry 16, 3899-3908 Clancy, R.M., McPherson, L.H. and Glaser, M. (1983) Biochemistry 22, 2350-2364 Berrez, J.M., Pattus, F. and Latruffe (1985) Arch. Biochem. Biophys. 243, 62-69 Cortese, J.D. and Vidal, J.C. (1983) Arch. Biochem. Biophys. 224, 351-357 Mclntyre, J.O., Churchill, P., Maurer, A., Berenski, C.J., Jung, C.Y. and Fleischer, S. (1983) J. Biol. Chem. 258, 953-959

66 202 203 204 205 206 207 208 209 210 211 212 213 214 215 216

217 218

219 220 221

222

223 224 225 226 227 228 229 230

E1 Kebbaj, M.S. and Latruffe, N. (1986) Arch. Biochem. Biophys. 244, 662-670 Mclntyre, J.O., Holladay, L.A., Smigel, H., Puett, D. and Fleischer, S. (1978) Biochemistry 17, 4169-4177 E1 Kebbaj, M.S., Berrez, J.M., Lakhlifi, T., Morpaln, C. and Latruffe, N. (1985) FEBS Lett. 182, 176-180 Eibl, H.J, Churchill, P., Mclntyre, J.O. and Fleischer, S. (1982) Biochem. Int. 4, 551-558 Churchill, P., Mclntyre, J.O., Eibl, H.J. and Fleischer, S. (1983) J. Biol. Chem. 258, 208-214 Fleischer, B., Casu, A. and Fleischer, S. (1966) Biochem. Biophys. Res. Commun. 24, 189-194 Nielson, N.C. and Fleischer, S. (1979) J. Biol. Chem. 248, 2549-2555 Miyahara, M., Utsumi, K. and Deamer, D.W. (1981) Biochim. Biophys. Acta 641, 222-231 Mombers, C., De Gier, J., Demel, R.A. and Van Deenen, L.L.M. (1980) Biochim. Biophys. Acta 604, 52-62 Sixl, F. and Galla, H.J. (1979) Biochim. Biophys. Acta 557, 320-330 Ranck, J.L. and Tocanne, J.F. (1982) FEBS Lett. 143, 171-174 Epand, R.M., Epand, R.F., Orlowski, R.C., Flanigan, E. and Stahl, G.L. (1985) Biophys. Chem. 23, 39-48 Dufourcq, J. and Faucon, J.F. (1978) Biochemistry 17, 1170-1176 Faucon, J.F., Dufourcq, J., Couraud, F. and Rochat, H. (1979) Biochim. Biophys. Acta 554, 332-339 Yogeeswaran, G. (1980) In Cancer Markers: Diagnostic and Developmental Significance, (Sell, S., ed.), pp. 371-401, Humana Press, Clifton, NJ Marcus, D.M. and Kundu, D.M. (1980) Adv. Exp. Med. Biol. 125, 321-326 Maget-Dana, R., Veh, R., Sander, M., Roche, A.C., Schauer, R. and Monsigny, M. (1981) Eur. J. Biochem. 114, 11-16 Abrahamson, M.B., Yu, R.K. and Zaby, V. (1972) Biochim. Biophys. Acta 280, 365-372 Hayashi, K. and Katagiri, A. (1974) Biochim. Biophys. Acta 337, 107-117 Ankel, H., Krishnamurti, Ch., Besanqon, F., Stefanos, S., and Falcoff, E. (1980) Proc. Natl. Acad. Sci. USA 77, 2528-2532 Hakamori, S.I., Young, W.W., Patt, L.M., Yoshino, T., Halfpap, L. and Lingwood, C.A. (1980) Adv. Exp. Med. Biol. 125, 247-261 Markwell, M.A.K. and Paulson, J.C. (1980) Proc. Natl. Acad. Sci. USA 77, 5693-5697 Markwell, M.A.K., Svennerholm, L. and Paulson, J.C. (1981) Proc. Natl. Acad. Sci. USA 78, 5406-5410 Van Heyningen, W.E. and Mellanby, J. (1968) J. Gen. Microbiol. 52, 447-454 Cuatrecasas, P. (1973) Biochemistry 12, 3547-3557; 3567-3576 Simpson, L.L. and Rapport, M.M. (1971) J. Neurochem. 18, 1341-1343 Wiegandt, H. (1979) Adv. Cytopharmacol. 3, 17-25 Fishman, P.H. (1982) J. Membrane Biol. 69, 85-97 Kundrot, C.E., Spangler, E.A., Kendall, D.A., Mac-

231 232 233 234 235 236 237 238 239 240 241 242 243 244 245 246 247 248 249 250 251 252 253 254 255 256 257 258 259

Donald, R.C. and MacDonald, R.I. (1983) Proc. Natl. Acad. Sci. USA 80, 1608-1612 Von Schulthers, G.K., Cohen, R.K., Sakato, N. and Benedek, G.B. (1976) Immunochemistry 13, 955-962 Von Heijne, G. (1985) Curr. Top. Membranes Transp. 24, 151-179 Strittmatter, P., Rogers, M.J. and Spatz, L. (1972) J. Biol. Chem. 247, 7188-7194 Endomoto, K. and Sato, R. (1973) Biochem. Biophys. Res. Commun. 51, 1-7 Sullivan, M.R. and Holloway, P.W. (1973) Biochem. Biophys. Res. Commun. 54, 808-815 Dufourcq, J, Faucon, J.F., Lusssan, C. and Bernon, R. (1975) FEBS Lett. 57, 112-116 Spatz, L. and Strittmatter, P. (1971) Proc. Natl. Acad. Sci. USA 68, 1042-1046 Greenhut, S.F. and Rosenman, M.A. (1985) J. Biol. Chem. 260, 5883-5886 Eytan, G., Matheson, M.J. and Racker, E. (1976) J. Biol. Chem. 251, 6831-6837 Scotto, A. and Zakim, D. (1985) Biochemistry 24, 4066-4075 Enoch, H.G., Fleming, P.J. and Strittmatter, P. (1979) J. Biol. Chem. 252, 5656-5660 Roseman, M.A., Holloway, P.W., Calabro, M.A. and Thompson, T.E. (1977) J. Biol. Chem. 252, 4842-4849 Leto, T.L., Roseman, M.A. and Holloway, P.W. (1980) Biochemistry 19, 1911-1916 Enoch, H.G., Fleming, P.J. and Strittmatter, P. (1979) J. Biol. Chem. 254, 6483-6488 Takagaki, Y., Radhakrishan, R,, Wirtz, K.W.A. and Khorana, H.G. (1983) J. Biol. Chem. 258, 9136-9142 Christiansen, K. and Carlsen, J. (1983) Biochim. Biophys. Acta 735, 225-233 Christiansen, K. and Carlsen, J. (1985) Biochim. Biophys. Acta 815, 215-222 Gogol, E.P. and Engelman, D.M. (1984) Biophys. J. 46, 491-495 Fleming, P.J. and Strittmatter, P. (1978) J. Biol. Chem. 253, 8198-8202 Hochman, Y., Kelley, M. and Zakim, D. (1983) J. Biol. Chem. 258, 6509-6516 Erickson, R.H., Zakim, D. and Vessey, D.A. (1978)Biochemistry 17, 3706-3711 Hochman, Y., Zakim, D. and Vessey, D.A. (1981) J. Biol. Chem. 256, 4783-4788 Singh, O.M.D., Graham, A.B. and Wood, G.C. (1981) Eur. J. Biochem. 116, 311-316 Cullis, P.R. and De Kruijff, B. (1979) Biochim. Biophys. Acta 559, 399-420 Alonso, A., Villena, A. and Goni, F.M. (1981) FEBS Lett. 123, 200-204 Hunt, G.R.A. (1980) FEBS Lett. 119, 132-136 Kantor, H.L. and Prestegard, J.H. (1975) Biochemistry 14, 1790-1795 Kantor, H.L. and Prestegard, J.H. (1978) Biochemistry 17, 3592-3597 Kantor, H.L., Mabrey, J., Prestegard, J.H. and Sturtevant, J.M. (1977) Biochim. Biophys. Acta 466, 402-410

67 260 Zakim, D. and Vessey, D.A. (1982) in Membranes and Transport (Martonosi, A., ed.), pp. 269-273, Plenum Press, New York 261 Zakirn, D., Hochman, Y. and Vessey, D.A. (1985) Biochem. Pharm. Toxicol. 1, 161-227 262 Hochrnan, Y. and Zakim, D. (1983) J. Biol. Chem. 258, 4143-4146 263 Oesterhelt, D. and Stoeckenius, W. (1974) Methods Enzymol. 31,667-678 264 Heyn, M.P., Cherry, R.J. and Denclier, N.A. (1981) Biochemistry 20, 840-849 265 Heyn, M.D., Blume, A., Rehorek, M. and Dencher, N.A. (1981) Biochemistry 20, 7109-7115 266 Oesterhelt, D. and Stoeckenius, W. (1971) Nature 233, 149-152 267 Blaurock, A.E. and Stoeckenius, W. (1971) Nature 223, 152-155 268 Oesterhelt, D. and Stoeckenius, W. (1973) Proc. Natl. Acad. Sci. USA 70, 2853-2857 269 Racker, E. and Stoeckenius, W. (1974) J. Biol. Chem. 249, 662-663 270 Gerber, G.E., Gray, C.P., Wildenauer, D. and Khorana, H.G. (1979) Proc. Natl. Acad. Sci. USA 74, 5426-5430 271 Scotto, A.W. and Zakim, D. (1986) Biochemistry 25, 1555-1561 272 Yonetani, T. (1961) J. Biol. Chem. 236, 1680-1688 273 Mason, T.L., Poyton, R.O., Wharton, D.C. and Shatz, G. (1973) J. Biol. Chem. 248, 1346-1354 274 Downer, N.W., Robinson, N.C. and Capaldi, R.A. (1976) Biochemistry 15, 2930-2936 275 Capaldi, R.A., Malatesta, F. and Darley-Usmar, V.M. (1983) Biochim. Biophys. Acta 726, 135-148 276 Kandrach, A. and Racker, E. (1973) J. Biol. Chem. 248, 5841-5847 277 Eytan, G.D., Matheson, M.J. and Racker, E. (1975) FEBS Lett. 57, 121-125 278 Eytan, G.D. and Broza, R. (1978) J. Biol. Chem. 253, 3196-3202 279 Eytan, G.D. and Racker, E. (1977) J. Biol. Chem. 252, 3208-3213 280 Kagawa, Y., Kandrach, A. and Racker, E. (1973) J. Biol. Chem. 248, 676-684 281 Poole, A.R., Howell, J.L. and Lucy, J.A. (1970) Nature 227, 6630-6635 282 Tall, A., Shipley, G.G. and Small, D.M. (1976) J. Biol. Chem. 251, 3749-3755 283 Jonas, A. and Krajnovich, D. (1977) J. Biol. Chem. 252, 2194-2199 284 Pownall, H.J., Massey, J.B., Kusserow, J.K. and Gotto, A.M., Jr. (1978) Biochemistry 17, 1183-1188 285 Pownall, H.J., Massey, J.B., Kusserow, S.K. and Gotto, A.M., Jr. (1979) Biochemistry 18, 574-579 286 Swaney, J.B. and Chang, B.C. (1980) Biochemistry 19, 5637-5644 287 Swaney, J.B. (1980) J. Biol. Chem. 255, 879-887 288 Pownall, H.J., Pao, Q., Hickson, D., Sparrow, J.T., Kusserow, S.K. and Massey, J.B. (1981) Biochemistry 20, 6630-6635 289 Smith, L.C., Pownall, H.J. and Gotto, A.M. (1978) Annu. Rev. Biochem. 47, 751-771

290

291 292 293 294 295 296 297 298 299

300 301 302 303

304 305 306 307

308 309 310 311 312 313 314 315 316 317 318

Weinstein, J.N., Klausner, R.D., Irmerarity, T., Ralston, E. and Blumenthal, R. (1981) Biochim. Biophys. Acta 647, 270-284 Wetterau, J.R. and Jonas, A. (1982) J. Biol. Chem. 257, 10961-10966 Kasahara, M. and Hinkle, P.C. (1977) J. Biol. Chem. 252, 7384-7390 Lawaczeck, R., Kainosho, M. and Chan, S.I. (1976) Biochim. Biophys. Acta 443, 313-330 Salsbury, N.J. and Chapman, D. (1968) Biochim. Biophys. Acta 163, 314-320 Haaker, H. and Racker, E. (1979) J. Biol. Chem. 254, 6598-6602 Carter-Su, C., Pillion, D.J. and Czech, M.P. (1980) Biochemistry 19, 2374-2385 Oku, N. and MacDonald, R.C. (1983) Biochemistry 22, 855-863 Navarro, J. and Essig, A. (1984) Biophys. J. 46, 709-717 Hudgin, R.L., Pricer, W.E., Jr., AshweU, G., Stockert, R.J. and Morell, A.G. (1974) J. Biol. Chem. 249, 5536-5544 Colacicco, G. (1972) Biochim. Biophys. Acta 266, 313-319 Rietveld, A., Sijens, P., Verkleij, A.J. and De Kruijff, B. (1983) EMBO J. 2, 907-913 Korb, H. and Neupert, W. (1978) Eur. J. Biochem. 91, 609-620 Matsuura, S., Aprin, M., Margoliash, E., Sabatini, D.D. and Morimoto, T. (1981) Proc. Natl. Acad. Sci. USA 78, 4368-4372 Walter, P. and Blobel, G. (1981) J. Cell. Biol. 91, 551-556 Rietveld, A., Jordi, W. and De Kruijff, B. (1986) J. Biol. Chem. 261, 3846-3856 Gorrissen, H., Marsh, D., Rietveld, A. and De Kruijff, B. (1986) Biochemistry 25, 2904-2910 Rietveld, A., Ponjee, G.A., Schiffers, P., Jordi, W., Van de Coolwijk, P.J.F.M., Demel, R.A., Marsh, D. and DeKruijff, B. (1985) Biochim. Biophys. Acta 818, 390-409 Hallermayer, G., Zimmerman, R. and Neupert, W. (1977) Eur. J. Biochem. 81,523-532 Harmey, M.A., Hallermayer, G., Korb, H. and Neupert, W. (1977) Eur. J. Biochem. 81, 533-544 Basile, G., DiBello, C. and Taniuchi, H. (1980) J. Biol. Chem. 255, 7181-7191 Hennig, B. and Neupert, W. (1981) Eur. J. Biochem. 121, 203-212 Rietveld, A. and De Kruijff, B. (1984) J. Biol. Chem. 259, 6704-6707 Smith, M., Leung, D.W., Gillian, S., AsteU, C.R., Montgomery, D.L. and Hall, B.D. (1979) Cell 16, 753-761 Zimmerman, R., Paluch, V. and Neupert, W. (1979) FEBS Lett. 108, 141-145 Wickner, W. (1976) Proc. Natl. Acad. Sci. USA 73, 1159-1163 Ohno-Iwashita, Y. and Wickner, W. (1983) J. Biol. Chem. 250, 1895-1900 Mandell, G. and Wickner, W. (1979) Proc. Natl. Acad. Sci. USA 76, 236-240 Zimmerman, R., Watts, C. and Wickner, W. (1982) J. Biol. Chem. 257, 6529-6536

68 319 Bhattacharya, B. and Wolf, J. (1975) J. Biol. Chem. 250, 7639-7646 320 Caron, J.M. and Berlin, R.D. (1979) J. Cell Biol. 81, 665 -671 321 Klausner, R.D., Kumar, N., Weinstein, J.N., Blumenthal, R. and Flavin, M. (1981) J. Biol. Chem. 256, 5079-5885 322 Cohen, B.E. (1975) J. Membrane Biol. 20, 205-234 323 Nachliel, E. and Gutman, M. (1984) Eur. J. Biochem. 143, 83-88 324 Lackowicz, J.R. and Hylden, J.L. (1978) Nature 275, 446-448 325 Li, K.-P., Li, Y.-Y., Boley, L. (1983) Biochem. Biophys. Res. Commun. 112, 1069-1076 325a Young, T.M. and Young, J.D. (1984) Biochim. Biophys. Acta 775,441-445 326 Singer, S.I. and Nicholson, G.L. (1972) Science 175, 720-731 327 Shinitzky, M. and Barenholz, Y. (1978) Biochim. Biophys. Acta 515, 367-394 328 Stubbs, C.D. and Smith, A.D. (1984) Biochim. Biophys. Acta 779, 89-137 329 Rand, R.P. and Parsegian, V.A. (1984) Can. J. Biochem. Cell Biol. 62, 752-759 330 Petrov, A.G., Selezner, S.A. and Derzanski, A. (1979) Acta Physica. Pol. A55, 385-405 331 Sackman, E. (1984) Biol. Membranes 5, 105-143 332 Jain, M.K. and White, H.B. (1977) Adv. Lipid Res. 15, 1-60 333 Jain, M.K. (1983) Membrane Fluidity Biol. 1, 1-37 334 Kauzman, W. (1961) Adv. Protein Chem. 65, 1071335 Pratt, L.R. (1985) Annu. Rev. Phys. Chem. 36, 433-449

336 337 338

339

340 340a 341 342 343 344 345 346 347 348

Finney, J.L. Gellatly, B.J., Golton, I.C. and Goodfellow, I. (1980) Biophys. J. 32, 17-33 Lee, A.G. (1977) Biochim. Biophys. Acta 472, 285-344 De Kruijff, B., Cullis, P.R., Verkleij, A.J., Hope, M.J., Van Echteld, C.J.A., Taraschi, T.F., Van Hoogevest, P., Killian, J.A., Rietveld, A. and Vandersteen, A.T.M. (1978) in Progress in Protein-Lipid Interactions (Watts, A. and DePont, J.J.H.H.M., eds.), pp. 89-142, Elsevier, Amsterdam Eibl, H.J. (1983) in (Alioa, R.C., ed.), Membrane Fluidity in Biology, Vol. 2, pp. 217-236, Academic Press, New York McDaniel, R.V., Mclntosh, T.J. and Simon, S.A. (1983) Biochim. Biophys. Acta 731, 97-108 Wright, E.M. and Diamond, G.M. (1977) Physiol. Rev. 57, 109-156 Mclntosh, T.J., McDaniel, R.V. and Simon, S.A. (1983) Biochim. Biophys. Acta 731, 109-114 Jain, M.K., Rogers, J., Simpson, L. and Gierasch, L. (1985) Biochim. Biophys. Acta 816, 153-162 Epand, R.M. and Surewicz, W.K. (1984) Can. J. Biochem. Cell Biol. 62, 1167-1176 Cortijo, M., Alonso, A., Gomez-Fernandez, J.C. and Chapman, D. (1982) J. Mol. Biol. 157, 597-618 Blobel, G. (1980) Proc. Natl. Acad. Sci. USA 77, 1496-1500 Von Heijne, G. (1980) Eur. J. Biochem. 103, 431-438 Randall, L.L. (1983) Cell 33, 231-240 Zimmerman, R. and Neupert, W. (1980) Eur. J. Biochem. 109, 217-229