Biochemistry 1986, 25,4223-4232
Sellin, S., Eriksson, L. E. G., Aronsson, A X . , & Mannervik, B. (1983) J . Biol. Chem. 258, 2091-2093. Sutton, D. (1968) Electronic Spectra of Transition Metal Complexes, McGraw-Hill, New York.
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Vallee, B. L., & Williams, R. J. P. (1968) Proc. Natl. Acad. Sci. U.S.A. 59, 498-505. Vallee, B. L., & Galdes, A. (1984) Adv. Enzymol. Relat. Areas Mol. Biol. 56, 283-430.
Novel Preparation of Functional Sindbis Virosomest Ronald K. Scheule Department of Pharmacology, The University of Texas Medical School, Houston, Texas 77225 Received December IO, 1985; Revised Manuscript Received February IO, 1986
ABSTRACT: Lipid and protein factors important for the preparation and stability of reconstituted membranes prepared by the insertion of detergent-solubilized Sindbis virus glycoproteins into preformed lipid vesicles have been defined. I t was found that both the state of aggregation of the membrane protein and the phase of the lipid are critical for the insertion of proteins into preformed lipid vesicles. The membranes prepared with the insertion technique were characterized in terms of residual detergent, protein orientation, and whether or not they were sealed. Binding and fusion experiments have been carried out with the insertion membranes and virus. It was found that BHK-21 cells a t 4 OC bound one-fifth to one-tenth the number of insertion membranes as intact virus, and binding was saturable in both cases. Variation of the lipid/protein ratio did not result in significant differences in binding. The insertion membranes were found to fuse to a model lipid bilayer a t least as well as the virus. These results are discussed in terms of structural factors important for the biological functionalities of the viral spike glycoproteins.
S i n d b i s virus is an enveloped virus approximately 700 A in diameter (Harrison et al., 1971), composed of equimolar copies (presumably 240 per virion) of two glycoproteins embedded in its lipid coat and a third protein associated with the encapsulated viral RNA. Although the amino acid sequences and stoichiometries of the individual viral proteins are known, the molecular factors that are necessary for viral binding and fusion are understood only poorly. These two aspects of viral functionality are thought to have their origins in the “spike” glycoproteins of the viral envelope. Sindbis virus, like many other enveloped viruses (Marsh et al., 1982, 1983a), attaches to a receptor on the host cell surface and is taken up by endocytosis. The low pH of the endosome is thought to trigger a fusion between the viral envelope and the endosomal membrane, releasing the viral RNA into the cell cytoplasm. The cell-surface receptor of Sindbis virus is presumed to be a protein, and two different candidates have been identified. A 90000-dalton cell-surface protein has been identified, by cross-linking, as the receptor for Sindbis virus on JY and Daudi cells (Maassen & Terhorst, 1981), while a 45 000-dalton protein has been implicated as the viral receptor on BHK cells (Duda & Berencsi, 1980). Although another togavirus, Semliki Forest virus, appears to bind to HLA-A and HLA-B antigens (Helenius et al., 1978), it can also infect cells lacking these antigens (Oldstone et al., 1980). Thus, while some progress has been made in identifying the cell-surface receptors of these enveloped viruses, relatively little is known about the requirements for the receptor-virus binding interaction. For example, to what extent are the binding and fusion functions of the spike proteins dependent on the specific arrangement of the spikes in the viral envelope? Also, one may ask whether or not these biological functions are indeed properties of the individual components of the viral spike or This investigation was supported by Biomedical Research Support Grant 201-2-2526 from The University of Texas and Grant PCM 8303948 from the National Science Foundation.
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whether the two envelope proteins act in a concerted fashion to accomplish a given biological function. The structural and compositional similarity of Semliki Forest and Sindbis viruses suggests that results of reconstitution with envelope proteins of the former virus are likely to be applicable to those of the latter. Two forms of the spike proteins of Semliki Forest virus have been used previously to examine binding to cells, namely, protein micelles (Fries & Helenius, 1979) and reconstituted membranes (virosomes) prepared by cosolubilization (Marsh et al., 1983b). Lipid-free octamers comprised of the Semliki envelope proteins can be formed by detergent disruption of the virus, sedimentation, and dialysis to remove detergent (Helenius & von Bonsdorff, 1976; Helenius et al., 1977; Simons et al., 1978). The apparent binding constants to cells for the octameric complexes were 100-1000 times lower than those for intact virus; the number of particles bound was the basis for comparison (Fries & Helenius, 1979). Although the binding of SFV virosomes to cells was not studied at physiological pH, binding at pH values below 7.0 was generally much less than that of the virus (Marsh et al., 1983b). Both Sindbis virus and Semliki Forest virus have been shown to interact with lipid bilayers at low pH (Mooney et al., 1975; White & Helenius, 1980). The SFV’ interaction with target liposomes has been determined to result in actual membrane fusion (White & Helenius, 1980). Thus, the viral ability to fuse membranes is not dependent on the interaction of the virus with a receptor. The fact that polykaryon formation can be induced in cells expressing the envelope glycoproteins of SFV
’
Abbreviations: TX-100, Triton X-100;BSA, bovine serum albumin; SFV, Semliki Forest virus; DML, dimyristoyllecithin; DPL, dipalmitoyllecithin; EPE,egg phosphatidylethanolamine; LUV, large unilamellar vesicle; SUV, small unilamellar vesicle; MES, 2-(Nmorpho1ino)ethanesulfonic acid; R- 18, octadecylrhodamine; TLC, thinlayer chromatography; Hepes, N-(2-hydroxyethyl)piperazine-N’-2ethdnesulfonic acid; NP-40, Nonidet P-40; DBED, dibenzylethylenediamine.
0 1986 American Chemical Society
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B I ocH E M I sT R Y
(Kondor-Koch et al., 1983) implies that an intact viral structure is not necessary for membrane fusion. The fact that cells expressing envelope protein E2 only do not form polykaryons (Kondor-Koch et al., 1983) implies that the second envelope protein E l is necessary for fusion but does not prove that E l is sufficient for fusion. Some information regarding the ability of the purified viral proteins to induce fusion has been obtained from reconstitution studies. Virosomes containing SFV glycoproteins have been found to fuse to cells, but with one-fourth the efficiency of the intact virus (Marsh et al., 1983b). These virosomes also could not induce polykaryon formation. The reasons for these differences in fusion ability between virus and virosomes are not clear. Finally, there is some evidence that the water-soluble domain of a viral spike glycoprotein contains all the necessary information for its pH-dependent interaction with a bilayer. The water-soluble domain of influenza virus has been isolated by bromelain treatment (Brand & Skehel, 1972) and has been shown to interact with lipids in the same pH-dependent fashion as the intact virus (Skehel et al., 1982). Analogous subviral protein domains of a togavirus have not yet been isolated. In addition to Sindbis and Semliki Forest virus, reconstitution procedures have been described for the envelope proteins of Sendai virus (Shimizu et al., 1972; Hosaka & Shimizu, 1972a,b; Hsu et al., 1979), the bacteriophage PM2 (Schafer & Franklin, 1975), influenza virus (Almeida et al., 1975; Huang et al., 1980); and vesicular stomatitis virus (Petri & Wagner, 1979; Miller et al., 1979). These reconstitutions have all involved the cosolubilization of lipid and viral protein in detergent, followed by the removal of that detergent. Triton X-100 (TX-100)' is the detergent that has been employed most often for these reconstitutions. It has been demonstrated repeatedly that TX- 100 is very effective in separating viral membranes from nucleocapsids and is sufficiently mild that glycoproteins are not denatured. One unattractive feature of TX-100, however, is that it is difficult to remove entirely at the end of the reconstituted procedure. There are several solutions to this problem. One is to use a second detergent that is easier to remove to assist in separating TX-100 from reconstituted membranes. This has been done with cholate (Helenius et al., 1977; Hsu et al., 1979) or octyl glucoside (Helenius et al., 1977, 1981) as the second detergent. Another approach that was employed in some of the initial reconstitutions with Sindbis virus is to absorb TX-100 onto beads of a styrene-divinylbenzene copolymer (Scheule & Gaffney, 1981a,b). This approach has been used in reconstitution of fusogenic Sendai virus envelopes (Volsky & Loyter, 1978). It has also been reported that an improvement in removal of TX-100 from reconstituted membranes can be achieved if the middle-molecular-weight Triton molecules are chromatographically separated and used in the reconstitution (Scheule & Gaffney, 1981b). Because none of these procedures yields a reconstituted membrane preparation of homogeneous size and lipid/protein ratio, the possibility of inserting detergent-solubilized viral glycoproteins into preformed lipid vesicles was examined. If feasible, an insertion protocol should produce reconstituted structures of a size dictated by the size of the starting vesicles. In addition, since the solution containing the membrane protein in TX- 100 is added to sealed vesicles, such a protocol might reduce the final level of detergent while yielding a vectorial protein orientation in the reconstituted membranes. Large, unilamellar lipid vesicles of defined size can be prepared by treating smaller sonicated vesicles with deoxy-
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cholate for 10-15 min followed by removing detergent on Sephadex (Enoch & Strittmatter, 1979; Dufour et al., 1981). Vesicles prepared in this way are extremely homogeneous in size, with an average diameter of 1000 A. This preparation has been combined with an observation made earlier by Wickner (1976) that the correct orientation of a membrane protein in a dimyristoyllecithin (DML) membrane could be achieved by a cholate dilution technique at the lipid phase transition temperature (23 "C) but not at higher or lower temperatures. The Sindbis virus E l and E2 glycoproteins have been reconstituted into preformed DML macrovesicles at or above the lipid phase transition without loss of vesicle integrity. The resulting well-characterized insertion membranes2 were then used to ask questions about the ability of the inserted viral glycoproteins to mimic the biological functions of the intact virus, thereby gaining further insight into the molecular determinants of virus-cell binding and membrane fusion. MATERIALS AND METHODS Egg phosphatidylcholine (EPC) was prepared according to Bangham et al. (1974) and stored at -20 OC under inert gas. Its purity was checked by TLC on silica gel plates using an eluant of chloroform/methanol/water, 65/24/4 (v/v), and was shown to be greater than 99.9% with respect to other lipids. P,y-Dimyristoyl-L-a-lecithin (DML) was used as obtained from Calbiochem. y-Palmitoyl-P- [9,1O-3H(N)]palmitoyl-La-phosphatidylcholine ( [3H]DPL), 50 Ci/mmol, was used as obtained from Amersham. Egg phosphatidylethanolamine (EPE) was obtained from Avanti and gave a single ninhydrin and iodide positive spot on TLC in CHC13/CH30H/NH3/ HzO, 90/54/5.7/5.4. Triton X-100, deoxycholic acid, and ribonuclease A were from Sigma. The deoxycholic acid was recrystallized from ethanol prior to use. [3H(G)]Deoxycholic acid (f3H]DOC), 10 mCi/mg, and [phenyl-3H(N)]Triton X-100 ([3H]TX), 1.58 mCi/mg, were obtained from New England Nuclear. The radiochemical purities of 13H]DOCand [3H]TX were assayed prior to use by TLC in benzene/dioxane/acetic acid, 15/2/3 (v/v), and chloroform/methanol/water, 65/25/4 (v/v), respectively, and found to be greater than 99.9%. Octadecylrhodamine was obtained from Molecular Probes. Pronase (nuclease free) from Calbiochem was predigested for 1 h at 37 OC before use. Polystyrene beads were from Bio-Rad (Biobeads SM-2) and were washed prior to use according to the procedure of Holloway (1 973). Initial stocks of Sindbis virus were gifts from B. Sefton of the Salk Institute. The large quantities of virus needed for the preparation of membranes for the binding experiments were obtained from the MIT Cell Culture Center. Cells. BHK-21 cells (clone 13) were obtained from the American Type Culture Collection (Rockville, MD). Cells were maintained in Dulbecco's modified Eagle's medium (DME, Gibco) supplemented with 10% tryptose phosphate broth (Flow), 10% calf serum (HyClone), 100 units of penicillin/mL, and 100 pg of streptomycin/mL. For infection with virus, a confluent monolayer of cells was subcultured at a ratio of 1:2 and infected within 18-24 h. Growth and Purification of Virus. Small quantities (100-300 pg) of virus were grown in 100-mm plastic tissue culture dishes (Falcon), while large quantities (1-3 mg) were grown in 1750-cm plastic roller bottles (Falcon). For the The terms insertion membranes, virosomes, reconstituted membranes, and membranes are used interchangeably to mean the lipoprotein structures obtained by adding detergent-solubilized viral envelope glycoproteins to preformed unilamellar lipid vesicles.
FUNCTIONAL VIRAL INSERTION MEMBRANES
growth of nonradioactive virus, cells were infected at a multiplicity of 0.1 plaque-forming unit (PFTJ) per cell in a minimal volume of DME with 1.5% fetal calf serum. After 1 h at 37 OC with occasional redistribution of the inoculum, additional DME with 1% fetal calf serum was added and incubation at 37 OC continued. Virus was harvested after approximately 18 h by following the procedure of Welch and Sefton (1979). Harvesting of virus from roller bottles followed the same procedure, except for the use of a type 55.2 rotor (Beckman) to accommodate the larger supernatant volume. Virus was purified on continuous 15-3076 (w/w) sucrose gradients as described previously (Welch & Sefton, 1979). The virus band in the gradient was located visually, removed by aspiration, quick-frozen in liquid nitrogen, and stored at -70 OC. [3sS]Methionine-labeled virus was prepared by infecting confluent monolayers of cells in 100-mm dishes at high multiplicity (50-100 PFU per cell) in 1.0 mL of DME with 1% calf serum and 2% tryptose phosphate broth. After 1 h a t 37 "C with occasional redistribution of the inoculum, the medium was removed and the monolayer rinsed once with methionine-free DME with 1% calf serum. Labeling was for 10 h in 3.0 mL of methionine-free DME with 1% calf serum and 10 pCi/mL [3sS]methionine(Amersham, 600 Ci/mmol). Virus labeled in the oligosaccharide moiety was prepared similarly, with [6-3H]glucosamine (45 Ci/mmol, ICN) in medium that contained one-tenth the normal amount of glucose. Radiolabeled virus was purified as above (Welch & Sefton, 1979). Virus with labeled cholesterol was prepared as described previously (Sefton & Gaffney, 1979). Briefly, cells were grown for approximately 3 days in medium containing [I4C]mevalonate [ [(RS)-2-14C]mevalonic acid DBED salt, 50 mCi/mmol, New England Nuclear] a t 1.5 pCi/mL, after which time they were infected at low multiplicity and further incubated with [14C]meval~nate(1.5 pCi/mL) in DME without fetal calf serum. Virus was harvested after 18 h and purified as described above. Isolation of Delipidated Viral Glycoprotein. Sindbis virus envelope glycoproteins free of lipid and cholesterol were isolated by solubilization of the virus with TX-100 followed by equilibrium sucrose gradient centrifugation in the presence of TX-100. The gradient consisted of a 60% (w/w) sucrose cushion (0.800 mL), followed by a linear 30-15% sucrose gradient (3.0 mL) and topped with a 10% sucrose layer (0.800 mL), all in bicarbonate buffer (0.050 M bicarbonate, 0.15 M NaCl, pH 8.2) containing 0.2% (v/v) TX-100. Virus (0.30 mg) was dissociated with 200 pL of 0.2% TX-100 in bicarbonate buffer and washed onto the gradient with an additional 200 pL of this buffer. Centrifugation was at 200000gav for 24 h at 4 "C in a 5-mL Ultraclear (Beckman) tube. Fractions were collected from below into siliconized (Sigmacote, Sigma) glass tubes at 4 OC, quick-frozen in liquid nitrogen, and stored at -70 OC. Purifkd Sindbis virus glycoproteins can be isolated in at least two discrete forms, which for simplicity here will be called monomer and oligomer. Both forms contain equal proportions of the viral glycoproteins E l and E2 as determined by gel electrophoresis (data not shown). Glycoprotein monomer is the form isolated from the sucrose gradient described above, i.e., the isolation procedure used for the preparation of delipidated viral glycoprotein. The oligomeric form of the viral glycoproteins was produced by isolating 5 times as much viral protein on the same gradient, applying this purified protein to a Sephadex G50 column in 0.05% TX-100 to remove sucrose, and then incubating it with
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polystyrene beads to remove TX-100. Monomer and oligomer were characterized by their apparent densities on equilibrium sucrose gradients and by their apparent size on a calibrated gel filtration column (0.8 X 21 cm) of Sepharose CL-48. Preparation of Small Unilamellar Vesicles. Small unilamellar vesicles (SUV's) of DML and EPC were prepared by sonication. A dried film of lipid was resuspended to 10 mg/mL in 5 mM bicarbonate buffer and sonicated under inert gas above the phase transition of the lipid for 20-30 min with the cup-horn attachment to a Heat Systems ultrasonifier cell disrupter. This solution was centrifuged for 1 h at 25 OC at 1000OOgav,and the upper three-fourths of the supernatant was removed and stored under inert gas until use. Preparation of DML Macrovesicles. Large unilamellar vesicles of DML (DML-LUV's) were prepared by using method I of Enoch and Strittmatter (1979) as modified by Dufour et al. (1981). The bML-LUV's were characterized by both gel filtration (Sepharose 2B) and electron microscopy (results not shown). Consistent with the results of Enoch and Strittmatter (1979), they were found to be very homogeneous and had an average diameter of about 1000 A. Aqueous solutes were entrapped by including them either in the lipid suspension during sonication or just before the addition of detergent, or both. Preparation of Reconstituted Membranes by Insertion into Preformed D M L - L W s . Reconstitutions of small amounts of purified viral glycoprotein (8-10 pg in 40 pL of bicarbonate buffer containing 0.2% TX-100) were accomplished by adding it to a rapidly mixed solution of DML-LUV's in 760 pL of bicarbonate buffer at 23.0 OC. The reaction mixture was incubated for an additional 1 h at 23.0 "C. Larger amounts (- 100 pg) of purified viral protein were reconstituted by using a scaled-up version of this protocol. After the incubation step, the insertion membranes were concentrated at 4 OC to a few hundred microliters by using an Amicon Model 8MC ultrafiltration apparatus with a YM30 membrane. This concentrated sample was applied to a 10-mL, 15-45% linear sucrose gradient in bicarbonate buffer and centrifuged to , equilibrium at 200O00gavat 4 OC in an SW41 rotor. Fractions were collected from below into siliconized glass tubes. Fractions containing the insertion membranes were pooled, dialyzed, and concentrated by ultrafiltration as above, applied to a 5-mL, 1545% sucrose gradient in 5 mM bicarbonate, and centrifuged to equilibrium. The purified insertion membranes were isolated from this second gradient and assayed for lipid and protein. Overall yields of protein for this procedure were 30-50%, based on protein recovered in the purified reconstituted membranes from the second gradient compared to viral protein initially solubilized. Most losses were found to be due to irreversible sticking of the glycoproteins to the ultrafiltration membranes, even in the presence of BSA included to minimize nonspecific adsorption. Protein-Transfer Experiments. The ability of the protein of the insertion membranes to undergo transfer to target membranes was investigated by incubating the reconstituted membranes (of either DML or EPC) with sonicated vesicles (of either DML or EPC) and then applying the mixture to a calibrated column (0.9 X 60 cm) of Sepharose CL-4B at 4 OC. Binding Experiments. Binding isotherms of the insertion membranes, virus, oligomeric viral glycoproteins, and LUV's to BHK fibroblasts at pH 7.4 were performed at 4 OC to minimize endocytosis. Bifiding medium was DME with 50 mM Hepes adjusted to pH 7.4 at 4 OC containing 5 mg of
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BSA/mL to minimize nonspecific binding. Confluent monolayers of cells in a 24-well dish (Falcon) were rinsed 3 times with 0.5 mL of binding medium at 4 "C; 250 /.tL of binding medium was then added to each well. Variable amounts of ligand in 5 mM bicarbonate buffer plus a volume of buffer to give 100 pL were added to the well to give a total volume of 350 pL. The dishes were rocked gently at 4 "C for 3 h, after which time the medium was removed and the pH checked. Wells were rinsed 3 times with 1 mL of a buffer made up of 0.15 M NaCl, 20 mM Hepes, and 5 mg of BSA/mL, which was adjusted to pH 7.4 at 4 "C. Next, they were incubated twice with 1 mL of this buffer for 2 min and then rinsed quickly twice with 1 mL of this same buffer lacking BSA. Cells were removed by trypsinization and assayed for radioactivity. Duplicate wells were trypsinized for cell counting. The kinetics of ligand binding were studied by removing aliquots from the incubation supernatant at various time points and assayed for radioactivity to determine the number of ligands unbound. Bound ligand was calculated from the difference between the ligand initially added and the unbound ligand. Cell counts were obtained at the end of the experiment as described above. Fusion Assay. The fusion activity of virus and insertion membranes was measured quantitatively by using the R-18 fluorescence assay described by Hoekstra et al. (1984). This assay is based on the relief of fluorescence quenching that occurs as a result of membrane fusion between a target membrane and a protein-containing membrane that bears the fluorescent probe. The R-18 probe has been shown to be nontransferable between membranes, and the fluorescence increase is directly related to the degree of fusion (Hoekstra et al., 1984). Control studies with R-18-labeled Sindbis virus confirm that there is no pH-dependent change in the fluorescence of the R-18 probe in the absence of fusion; Le., there are no pH-dependent changes in the structure or charge state of the probe that result in fluorescence changes. To label virus with the probe, an aliquot of a concentrated solution of R-18 in ethanol was added to an aqueous solution of virus or membranes and incubated for 1 h at room temperature. Unincorporated probe was removed by gel filtration on Sephadex G75 (Hoekstra et al., 1984). The extent of fusion was measured at 37 "C by adding target vesicles to a stirred fluorescence cuvette containing a 5 mM MES buffer at pH 7.4, followed by the addition of the R-18-labeled virus or membranes, and then adding acid to decrease the pH to 5.4. A value corresponding to 100% fusion was obtained by adding TX- 100 to solubilize completely the R- 18 containing membrane. Corrections were made for dilution and for the light scattering caused by the LUV's. The concentration of LUV's was varied to confirm that they were not limiting the amount of fusion. The target vesicles for the fusion assay were LUV's composed of EPE. LUV's were found t o be superior to multilamellar vesicles (MLV's) as fusion targets due to their minimal scattering background. The LUV's were prepared by extrusion of a solution of EPE-MLV's in a 5 mM MES buffer at pH 7.4 in a LUVET device (Hope et al., 1985) and had a nominal diameter of 1000 A. A small correction was made for the fusion ability of R- 18-labeled DML-LUV's that did not contain viral glycoprotein. Thus, the fusion reported for the insertion membranes is a specific fusion ability, attributable to the presence of the viral glycoproteins. Other Methods. Protein concentrations were determined by the method of Lowry et al. (1951), using bovine serum
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albumin (Sigma, 3 times recrystallized) as a standard. Phospholipid analyses were accomplished by using an adaptation of the phosphate assay of Rouser et al. (1970). All radioactivity determinations were corrected for background and channel spillover. Virus was precipitated before use by dilution with cold water and centrifugation as described by Pfefferkorn and Clifford (1 963). Ribonuclease A (RNase) was iodinated in 0.10 M NaC1, pH 7.0, with Iodobeads (Pierce) and NaIZSI(17 Ci/mg; New England Nuclear). Free iodide was removed by gel filtration on Sephadex G25. RESULTS Isolation of Purified Spike Glycoprotein. Triton X-100 and the related NP-40 have been used often to solubilize and separate by sedimentation viral envelope proteins from nucleocapsids. The phospholipids are also removed from the glycoproteins in this step (Helenius & von Bonsdorff, 1976). Because cholesterol is thought to be necessary for the fusion activity of glycoproteins from several viruses (White & Helenius, 1980), it seemed possible that its presence might also influence the course of reconstitution. In order to examine whether gradient-purified TX- 100-solubilized Sindbis glycoproteins were free of viral cholesterol, the viral cholesterol was labeled biosynthetically by growth of cells on ['4C]mevalonate (Sefton & Gaffney, 1979). Cholesterol is well resolved from the envelope glycoproteins when sedimented into a sucrose gradient containing 0.2% (v/v) TX- 100; the viral phospholipids were found at the same position as cholesterol while the viral core protein sedimented to the bottom (data not shown). Hence, this purification protocol separates cholesterol and phospholipids as well as the capsid protein from the viral spike glycoproteins. Aggregation State of Membrane Protein. In order to evaluate the importance of the aggregation state of the purified membrane protein on its interaction with lipid vesicles, the envelope glycoproteins of Sindbis virus were isolated under conditions designed to produce oligomeric (micellar) or monomeric protein (see Materials and Methods). The oligomeric form of the envelope protein appears as a peak on a Sepharose CL-4B gel filtration column eluted in the absence of detergent with an apparent diameter of greater than 250 A. This, presumably micellar, form of the viral glycoprotein has an apparent density of 1.174 f 0.006 g/cm3, as determined by isopycnic centrifugation at 4 "C in the absence of detergent. On the same gel filtration column, the monomeric form of the viral glycoprotein has an apparent diameter of about 100 A. In addition, it is characterized by an apparent density of 1.097 f 0.003 g/cm3 in an equilibrium sucrose gradient containing 0.2% TX-100. The insertion characteristics of the monomeric and oligomeric glycoprotein were compared by incubating each in the presence of DML-LUV's at 23 OC. Figure 1A shows that the oligomeric form of the glycoprotein inserts very little, if at all, into preformed lipid vesicles. By contrast, Figure 1B shows that the monomeric form of the protein inserts virtually quantitatively into the same preformed vesicles at the same temperature. Phase of Lipid. Most reconstitutions are carried out at either room temperature or 4 "C and use a natural lipid such as egg lecithin. Thus, such reconstitutions are performed above the phase transition of the lipid. By using a synthetic lipid, such as DML, which has a gel-liquid crystalline phase transition at 23.0 OC, one can ask whether or not the phase of the lipid can influence the course of a reconstitution involving an insertion mechanism. The interaction of the monomeric glycoprotein with DML-LUV's was characterized therefore at
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FRACTION 1: Comparison of the interaction between oligomeric and monomeric forms of the viral glycoprotein and DML-LUV's by gel exclusion chromatographyat 4 OC on Sepharose CL-4B. (A) Elution profile of 23 O C reaction product between oligomeric protein and LUV's. (B) Elution profile of 23 O C reaction product between monomeric protein and LUV's. Viral glycoprotein is labeled with [35S]methionineand LUV's are labeled with ['HIDPL. Void and total volumes occur at fractions 9 and 29, respectively. FIGURE
temperatures below, at, and above the DML phase transition. Figure 2A shows that the addition of monomeric protein to DML-LUV's at a temperature below the phase transition temperature results in virtually no insertion. Figure 2a indicates that incubation of protein by itself results in some aggregation of the monomeric glycoprotein, presumably in response to diluting the detergent to a concentration below its
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critical micelle concentration. Figure 2B shows that at the phase transition, incubating monomeric viral protein with lipid vesicles results in virtually quantitative insertion of the protein into the lipid component. Figure 2b is a control incubation, showing that the protein by itself undergoes some aggregation under these conditions, as in (2a). It is clear, however, that the protein eluting in the void volume of the column in Figure 2B is the result of its association with the lipid vesicles at this temperature. Figure 2C indicates that incubating the monomeric form of the membrane protein with lipid vesicles at a temperature aboue the phase transition of the lipid also appears to lead to quantitative insertion. It is also apparent from Figure 2c, however, that at 41 OC the protein itself appears to aggregate into structures that are even larger than the protein micelles, which elute at about fraction 14 on this column (data not shown). The results of these insertion experiments with DML were confirmed in part by analogous experiments using EPC-LUV's. Insertion was found to occur at both room temperature and 4 O C , Le., at temperatures above the phase transition of EPC (data not shown). Since reconstitution occurs at 4 "C with EPC, but not with DML, it would appear that the phase of the lipid is critical in determining whether the monomeric protein can insert into lipid. Purification of Reconstituted Membranes. The reconstituted membranes prepared by the insertion technique were purified by using two sequential equilibrium sucrose gradients. The gradient profile shown in Figure 3A shows that virtually all the viral protein is associated with lipid in a structure that is more dense than the bulk of the lipid. When the protein peak from this first gradient is isolated, ultrafiltered to remove sucrose, and applied to a second gradient, the profile shown in Figure 3B is obtained. In this second gradient, the purified insertion membranes occur at fractions 15-2 1, while the residual, unreacted DML-LUV's appear as a very small shoulder of lipid at fractions 22-25. Although the protein and lipid coequilibrate on this gradient, it should be noted that the peaks of lipid and protein do not coincide exactly. Thus, there
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FRACTION FIGURE2: Temperature dependenceof the interaction between monomeric viral protein and DML-LUV's as assayed by gel exclusion chromatography at 4 O C on Sepharose CL-4B. Incubations were carried out at 0 O C with both protein and LUV's (A) or protein only (a), at 23 O C with both protein and LUV's (B) or protein only (b), and at 41 O C with both protein and LW's (C) or protein only (c). Identical amounts and concentrations of protein and LUV's were employed in all incubations. Lipid and protein are labeled with [3H]DPLand [35S]methionine,respectively. Column void and total volumes occur at fractions 9 and 29, respectively.
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FIGURE 3: Isolation of purified insertion membranes by consecutive equilibrium sucrose gradients. (A) Gradient profile of 23 O C reaction product between monomeric viral glycoprotein and DML-LUV’s on a IO-mL, 15-45% linear gradient run at 200000gavat 4 O C in an SW41 rotor. (B) Gradient profile of pooled fractions 16-24 from (A)which were dialyzed, concentrated, and applied to a SmL, 15-45% linear gradient run at 200000g,, at 4 O C in an SW50.1 rotor. (C) Gradient profile of reconstituted membranes prepared by the cosolubilization method on a gradient identical with that in (B). Lipid and protein are labeled with [3H]DPL and [35S]methionine,respectively, and density decreases from left to right.
appears to be a distribution of lipid to protein (L/P) through the peak, with the more protein rich membranes appearing at higher density. Gel filtration of this peak on a calibrated Sepharose 2B column (1.8 X 60 cm) reveals a homogeneous population of membranes with an average diameter of about 1000 A, consistent with the size of the starting LUV’s. It should be noted also that the average apparent density of the membranes obtained here is approximately 1.1 1 g/cm3, which is well resolved on this gradient from that of the purified oligomer ( p = 1.17 g/cm3; data not shown). For comparison with the insertion membranes, Figure 3C shows a sucrose density gradient profile of Sindbis virus membranes that were prepared by using a cosolubilization technique (Scheule & Gaffney, 1981a,b). In contrast to the insertion membranes, several protein-containing species are generated when reconstitution is accomplished by cosolubilization [see also Helenius et al. (1977)l. Gel filtration of the individual protein peaks in this gradient shows that the peak of highest density is composed of relatively small structures (-350-A diameter) with a L / P of - 2 2 , while the peak of lowest density is composed of larger vesicles (>900-A diameter) with a L/P of 1000. The complex and virtually unknown kinetic details of reconstitution by cosolubilization are reflected in the wide variety of lipoprotein structures observed. An additional aspect of the reconstitution procedure is demonstrated in Figure 3A, namely, that the interaction of monomeric protein with LUV’s does not appear to be a random process. The initial mole ratio of viral protein monomer to LUV’s in this particular reaction mixture was approximately 20. However, the fraction of LUV’s containing protein at the end of the incubation represents only about 1% of the total lipid present initially. The preferential interaction of protein with such a small fraction of the vesicles implies that this interaction is probably cooperative; Le., the initial interaction of viral protein with a particular vesicle makes the subsequent interaction of additional protein with that vesicle even more favorable. The net result is a population of insertion membranes in which the average molar lipid to protein ratio (L/P) is much lower than the proportion of the lipid to protein in
-
Table I: Characterization of Purified Reconstituted Membranes Prepared by Insertion into DML-LUV’s‘ lipid/protein (mol/mol) d PZO
initialb
1500 4000 8000 1 1400
finalc 70
130 440 1050
(g/cm’) 1.134 1.114 1.106
1.097 For comparison with these data, Sindbis virus has a L/P of 3 1 and a density of 1.21 g/cm3 (Pfefferkorn & Hunter, 1963). bRatio in initial reaction mixture. Ratio determined in purified membranes from second sucrose gradient. dDensities of peak fractions from second sucrose gradient, determined by refractive index measurement.
the initial reaction mixture. In the particular preparation of the insertion membranes illustrated in parts A and B of Figure 3, the final average number of lipids per protein (as El or E2) is approximately 130, with a variation through the peak of f 3 0 . Despite the observed cooperativity of the insertion process, the data in Table I demonstrate that it has been possible to prepare virosomes in which the number of lipids per protein are varied systematically from less than 100 to greater than 1000 simply by increasing the L / P of the initial reaction mixture. Orientation of Inserted Proteins. The orientation of viral glycoproteins inserted into preformed vesicles, as well as the integrity of the bilayer, was investigated by protease treatment of purified insertion membranes that contained trapped RNase. Figure 4A shows that reconstituted membranes prepared by using [3H]glucosamine-labeled viral protein with trapped [‘251]RNa~e are stable in the absence of protease under the conditions of a proteolysis incubation, viz. 37 OC for 1 h. Figure 4B demonstrates that proteolysis results in the removal of greater than 93%of the viral glycoprotein from the membranes while they retain roughly 85% of their internal RNase marker. Thus, membranes prepared by insertion are sealed, maintain their structural integrity at 37 OC, and are characterized by a vectorial protein orientation. Residual Detergent. The preparation of the DML-LUV’s used in these studies makes use of DOC; the subsequent in-
VOL. 2 5 , N O . 1 5 , 1986
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I
0
2
I
3
4
TIME(Hours)
20
40
60
FRACTION
FIGURE 5: Kinetics of binding of Sindbis virus and reconstituted membranes to BHK cells at pH 7.4 and 4 OC. (A) Virus (3.4 X loLo) labeled with [3sS]methioninewas added to 1 X lo6 cells at zero time and binding followed for 3 h as described under Materials and Methods. The medium then was replaced with new medium lacking virus, and the amount of virus remainin bound was determined. (B) Reconstituted membranes (1.5 X lo1{; L/P = 190) labeled with [3H]DPL were added to cells and the kinetics of binding followed as in (A).
4: Pronase treatment of insertion membranes with trapped [12sI]RNase.Viral protein is labeled with [3H]glucosamine.Membranes were incubated for 1 h at 37 OC either in (A) the absence or (B) the presence of 2 mg of Pronase/mL and then applied to a Sepharose CL-4B column. Void and total volumes occur at fractions 14 and 45, respectively. FIGURE
I
I
h
x
sertion of detergent-purified protein with these vesicles introduces TX-100 into the reconstitution. In principle, residual detergent in reconstituted membranes could result in altered membrane properties, such as permeability, receptor binding ability, and fusogenic potential. Due to the differences of the topology of preparation of insertion membranes compared to those prepared by cosolubilization, one might predict that any residual detergent effects would be minimized in the insertion membranes. Radiolabeled detergents were used to evaluateresidual levels of DOC and TX-100 in both the reconstituted membranes and DML-LUV’s to which TX-100 but no protein had been added, i.e., a mock reconstitution. For both DOC and TX-100, the final levels of detergent were found to be independent of protein and were 1-2 molecules of detergent per 1000 lipids. For comparison, membranes prepared by cosolubilization of lipid and protein with TX-100 were found to have greater than 100 molecules of TX-100 per 10oO molecules of lipid (Scheule & Gaffney, 1981b). Binding of Insertion Membranes to BHK Cells. The kinetics of the binding of the insertion membranes and virus to BHK cells were determined at pH 7.4 and 4 OC. Figure 5 shows that the kinetics are similar for both the membranes and virus, requiring several hours to reach maximal values. Therefore, to obtain binding isotherms of virus or membranes with cells, incubations were carried out for 3 h. Control incubations of virosomes (prepared by using DML-LUV’s) with SUV’s at 4 O C for 3 h showed no evidence for the exchange of viral proteins between membranes (data not shown).
-1 W -1
. 0
d
3 2 0
m
2 0 2
a K
m
EH
O Y 0
I
2
I
MEMBRANES ADDEDICELL
X
16’
Competition between Sindbis virus and insertion membranes for binding to BHK cells at pH 7.4 and 4 O C : membranes by themselves (0); varying amounts of membranes added together at zero time with a 5-fold excess of virus ( 0 ) . Membranes are labeled with [3sS]methionineviral proteins. Bound membranes were determined after 3 h. FIGURE 6:
Figure 5 demonstrates an additional feature that is common to the binding of virus and insertion membranes. That is, if virus or membranes are allowed to bind for several hours and then the medium is replaced by medium that is free of virus, very little bound virus or membranes are seen to reequilibrate with the medium. Thus, it appears from these data that the binding of both virus and insertion membranes to cells is virtually irreversible. Figure 6 illustrates a further similarity between virus and membrane binding; that is, they both appear to compete for
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5.0
6.0
7.0
PH
I
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0
10 PARTICLES ADDED/CELL
I
20 X
FIGURE 7: BHK cell binding isotherms of virus (O), reconstituted membranes (A) L/P = 190, (A) L/P = 270, and (V)L/P = 84, viral glycoprotein oligomers (O), and DML-LUV's ( O ) , at pH 7.4 and 4 OC. Error bars reflect the magnitude of uncertainties arising from the determination of particle numbers.
some of the same cellular binding sites. In this experiment, both virus and membranes were added to the cells at zero time and allowed to bind for 3 h. An isotherm was constructed by varying the total number of virosomes and virus added initially; the ratio of virus to virosomes was kept constant at 5: 1. Figure 6 shows that virus competes with the virosomes for cell binding over the entire range of concentrations studied. If one assumes identical binding sites and binding affinities, one would expect fewer virosomes to bind in the presence of virus than are shown in Figure 6, given the relative numbers of virus and reconstituted membranes added in this competition experiment. One interpretation of these data is that the virosomes bind to both viral and other sites; i.e., they have two modes of binding. Figure 7 compares binding isotherms for virus, insertion membranes (made by using DML), viral protein oligomers, and pure lipid vesicles of DML. At equivalent numbers of particles added per cell, these data indicate that the cells bind 5-10 times more virus than insertion membranes, which in turn bind about as well as the viral protein oligomers. Further, the cells bind an order of magnitude fewer DML-LUV's than reconstituted membranes. Two additional points can be made regarding these binding isotherms. First, given the error limits involved, there does not appear to be any simple correlation between the lipid/ protein ratio of the insertion membranes and their ability to bind. For example, insertion membranes with a L/P = 190 bind better than membranes with either larger or smaller L/P ratios. Second, although not obvious due to the semilogarithmic nature of Figure I , the binding of virus and insertion membranes exhibits saturability at roughly the same range of particles added per cell. In contrast, the binding of viral glycoprotein oligomers and DML-LUV's does not exhibit saturability, even beyond the range of particles added that is depicted in this figure. Fusion of Insertion Membranes. Togaviruses are able to fuse their membranes with that of the endosomal membrane of the host cell (Helenius et al., 1980). This process is de-
FIGURE 8: pH dependence of the fusion of Sindbis virus ( 0 ) and insertion membranes (0) with target EPE-LUV's at 37 O C . The percent fusion was obtained as described under Materials and Methods by using the fluorescent R-18 probe.
pendent on pH and is probably a result of a conformational change in the viral spike. The fusion competence of the insertion membranes was compared to that of the virus by using a quantitative fluorescence assay. Figure 8 shows the results of a comparative study of the pH dependence of the fusion of intact virus and insertion membranes with target lipid vesicles. Virus and virosomes promote fusion with the same pH dependence, exhibiting a midpoint for fusion at about pH 5.8. In addition, the insertion membranes appear to fuse to an even greater extent than the virus in this model system. DISCUSSION The results presented here demonstrate that there are several important molecular considerations that should be taken into account in order to assure a successful reconstitution of a detergent-solubilized amphipathic protein by insertion. The aggregation state of the protein that is added to the preformed lipid vesicles is crucial in determining the outcome of the interaction between protein and lipid. Thus, monomeric, detergent-solubilized proteins insert readily into preformed lipid vesicles, while water-soluble, detergent-free protein micelles are totally refractory toward insertion. It would appear that, by diluting a detergent-solubilized protein into a solution containing lipid vesicles, the protein satisfies the hydrophobic requirements of its membranespanning domain by inserting into the hydrocarbon region of the lipid bilayer. Protein micelles, on the other hand, are stable in the absence of detergent by virtue of their amphipathic construction. Reconstitutions of membrane proteins that present a detergent-solubilized, monomeric form of the protein to lipid vesicles should therefore increase the likelihood that a successful protein-lipid interaction will ensue. In addition to the aggregation state of the protein, these studies have also demonstrated the critical importance of the lipid phase of the vesicles into which the protein is to be inserted. Even detergent-solubilized monomeric protein does not insert into a lipid that is in the gel phase, i.e., below its gel-liquid crystalline phase transition temperature. Since both the lipid head groups and acyl chains are more disordered in the liquid crystalline phase than in the gel phase, this obser-
FUNCTIONAL VIRAL INSERTION MEMBRANES
vation is consistent with the hypothesis that the insertion of a protein into a lipid bilayer requires a certain degree of disorder in the lipid bilayer; perhaps some minimal degree of lateral compressibility of the lipid (Linden et al., 1973) is required for insertion to occur. It is interesting to note that once a protein has inserted into a fluid-phase lipid bilayer, it can remain associated with the lipid even below the phase transition temperature. This aspect of the reconstituted membranes is illustrated by the insertion of the Sindbis virus glycoproteins into DML-LUV’s and their subsequent incubation at 4 OC-conditions under which they remained stable. It is also instructive to compare the present findings with other examples of the insertion of amphipathic proteins into lipid bilayers. The insertion of cytochrome b5 into lipid vesicles has been well studied, but exclusively above the phase transition of the lipid (Enoch et al., 1979). The details of the orientation of the protein in the bilayer have been elucidated with photoactive lipids and indicate that a hydrophobic hairpin of the protein inserts into the outer lamella of the bilayer (Takagaki et al., 1983). In another example, the procoat glycoprotein of bacteriophage M13 inserts entirely across a target lipid bilayer of Escherichia coli lipids (Wickner et al., 1978). Together with the present results, these experiments indicate that the potential topological interactions between a lipid bilayer and an added amphipathic protein are several and depend on the molecular details of both the lipid bilayer and the protein. The insertion membranes prepared here have several characteristics that make them ideal structures with which to probe the biological functionality of the viral glycoproteins. They are well characterized and homogeneous, in both their size and L/P ratio. Residual detergent levels are exceedingly low, thus minimizing any artifacts arising from this source. Finally, the inserted proteins have a vectorial orientation, ensuring that all the protein in the membranes has the potential of being functional. With regard to the biological activities of the virus, these studies show that viral glycoprotein which has been inserted into preformed LUV’s is fully fusion competent but does not bind as well as intact virus. It would appear that an intact viral structure is not necessary for fusion and that the ability to fuse membranes is a property that can be associated with an individual spike. Furthermore, the fusion potential of the spike proteins does not appear to be impaired as a result of the manner in which they have been inserted into the preformed LUV’s. The process of insertion is likely to involve the interaction of a hydrophobic hairpin of the membranespanning domain of the spike glycoproteins with the outer lamella of the LUV, as has been shown to be the case for cytochrome b5 (Takagaki et al., 1983). This conformation would result in the carboxyl termini of the spike glycoproteins residing on the outside of the insertion membrane. Assuming this to be the case, it does not appear to affect fusion adversely, which can be interpreted to mean that the information necessary for fusion resides entirely in the domain of the viral spike that lies outside the viral bilayer. This result is consistent with the finding that the aqueous domain of the influenza hemagglutinin is capable of interacting with lipid vesicles in the same pH-dependent fashion as the intact virus (Skehel et al., 1982). The preparation of a well-characterized, homogeneous population of insertion membranes also has permitted meaningful comparisons of the cellular binding abilities of virosomes and virus, since actual “particles” can be counted. The present
VOL. 2 5 , NO. 15, 1986
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studies demonstrate that insertion membranes can bind to a host cell of the virus at physiological pH. At equivalent concentrations of ligands, BHK cells bind fewer reconstituted membranes than intact virus in the concentrated range studied. The reconstituted membranes bind about equally as well on a particle basis as viral protein oligomers and an order of magnitude better than pure lipid vesicles, which do not contain the viral glycoprotein. For comparison, protein micelles of SFV have been found to exhibit apparent binding affinities that are 100-1000-fold lower than that of the intact virus (Fries & Helenius, 1979). The insertion membranes also appear to possess some of the binding properties that characterize the intact virus. Thus, the binding of the virosomes, like the virus, is both saturable and irreversible. In contrast, the binding of viral glycoprotein oligomers does not appear to be saturable in the concentration range studied. This latter observation also has been made with glycoprotein micelles derived from Semliki Forest virus, where saturation was achieved only at very high concentrations of the protein complexes (Fries & Helenius, 1979). While the origins of these differences in saturability are not known, it is clear that the reconstituted membranes resemble the virus more closely than do the protein oligomers with regard to this aspect of their interaction with the cell-surface receptor. Although the ability of the reconstituted membranes to saturate cellular binding sites resembles that of the virus, it is also apparent that cells bind significantly fewer of these membranes compared to virus, despite the fact that the size and lipid/protein ratio of these membranes approaches that of the virus. This finding does not appear to be related to the details of preparation of the reconstituted membranes, since membranes of Sindbis virus (R. K. Scheule, unpublished observations) and Semliki Forest virus (Marsh et al., 1983b) prepared by cosolubilization techniques also do not bind as well as intact virus. Thus, it would appear that some other aspect of the viral structure enables the virus to bind more efficiently to its receptor. The most obvious remaining structural difference between the intact virus and the reconstituted membranes is the viral capsid. It has been proposed that in togaviruses there are specific interactions between at least one of the glycoproteins and the viral capsid (Garoff & Simons, 1974). Such interactions no doubt would stabilize a particular geometric configuration of the envelope proteins. Topological stabilizing interactions of this kind are not preserved in the reconstituted membranes and may contribute to the differences in binding between the reconstituted membranes and the virus for the cellular receptor. With regard to the biological functions of the viral spike glycoproteins, these studies lead to the conclusions that (i) correct, viruslike binding appears to be a function that is critically dependent on the sum of inter xtions between the components of several glycoprotein spikes and the cellular receptor(s) and (ii) the ability to fuse membranes resides in the aqueous domain of the spike. These results should provide a starting point for additional studies that seek to elucidate the molecular details of the biological functionalities of the spike glycoproteins. ACKNOWLEDGMENTS I thank Dr. B. J. Gaffney for her support and encouragement during the course of this work and for her critical reading of the manuscript. I acknowledge the technical assistance of Leah Goldberg in the binding experiments. In addition, I thank Dr. Peter Davies for the use of a fluorometer and Amicon concentrator and Drs. Alan Schroit and Rajiv Nayar for the use of the LUVET device and for helpful discussions.
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Registry No. DML, 18194-24-6.
REFERENCES Almeida, J. B., Edwards, D. C., Brand, C. M., & Heath, T. D. (1975) Lancet, 899-901. Bangham, A. D., Hill, M. W., & Miller, N. G. A. (1974) Methods Membr. Biol. I , 1-68. Brand, C. M., & Skehel, J. J. (1972) Nature (London),New B i d . 238, 145-147. Duda, E., & Berencsi, K. (1980) Acta Virol. 24, 149-152. Dufour, J.-P., Nunally, R., Buhle, L., Jr., & Tsong, T. Y. (1981) Biochemistry 20, 5576-5586. Enoch, H. G., & Strittmatter, P. (1979) Proc. Natl. Acad. Sci. U.S.A. 76, 145-149. Enoch, H. G., Fleming, P. J., & Strittmatter, P. (1979) J.Biol. Chem. 254, 6483-6488. Fries, E., & Helenius, A. (1979) Eur. J . Biochem. 97, 2 13-220. Garoff, J., & Simons, K. (1974) Proc. Natl. Acad. Sci. U.S.A. 71, 3988-3992. Harrison, S . C., David, A,, Jumblatt, J., & Darnell, J. E. (1971) J . Mol. Biol. 60, 523-528. Helenius, A., & von Bonsdorff, C. H. (1976) Biochim. Biophys. Acta 436, 895-899. Helenius, A., Fries, E., & Kartenbeck, J. (1977) J . Cell Biol. 75, 866-880. Helenius, A., Morein, B., Fries, E., Simons, K., Robinson, P., Schirrmacher, V., Terhorst, C., & Strominger, J. L. (1978) Proc. Natl. Acad. Sci. U.S.A. 75, 3846-3850. Helenius, A., Kartenbeck, J., Simons, K., & Fries, E. (1980) J . Cell Biol. 84, 404-420. Helenius, A., Sarvas, M., & Simmons, K. (1981) Eur. J. Biochem. 116, 27-35. Hoekstra, D., deBoer, T., Klappe, K., & Wilshut, J. (1984) Biochemistry 23, 5675-568 1 . Holloway, P. 0. (1973) Anal. Biochem. 53, 304-308. Hope, M. J., Bally, M. B., Webb, G., & Cullis, P. R. (1985) Biochim. Biophys. Acta 812, 55-65. Hosaka, Y., & Shimizu, Y. K. (1972a) Virology 49, 627-639. Hosaka, Y., & Shimizu, Y. K. (1972b) Virology 49,640-646. Hsu, M.-C., Scheid, A., & Choppin, P. W. (1979) Virology 95, 476-49 1 . Huadg, R. T. C., Wahn, K., Klenk, H.-D., & Rott, R. (1980) Virology 104, 294-302. Kondor-Koch, C., Burke, B., & Garoff, H. (1983) J . Cell Biol. 97, 644-651. Linden, C. D., Wright, K. L., McConnell, H. M., & Fox, C. F. (1973) Proc. Natl. Acad. Sci. U.S.A. 70, 2271-2275. Lowry, 0. H., Rosebrough, A., Farr, L., & Randall, R. J. (1951) J . Biol. Chem. 193, 265-275.
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Maassen, J. A., & Terhorst, C. (1981) Eur. J . Biochem. 115, 153-158. Marsh, M., M a t h , K., Simons, K., Reggio, H., White, J., Kartenbeck, J., & Helenius, A. (1982) Cold Spring Harbor Symp. Qunnt. Biol. 46, 835-843. Marsh, M., Bolzau, E., & Helenius, A. (1983a) Cell (Cambridge, Mass.) 32, 931-940. Marsh, M., Bolzau, E., White, J., & Helenius, A. (1983b) J. Cell Biol. 96, 455-46 1 . Miller, D. K., Feuer, B. I., Vanderoef, R., & Lenard, J. (1979) J . Cell Biol. 84, 421-429. Mooney, J. J., Dalrymple, J. M., Alving, C. R., & Russell, P. K. (1975) J. Virol. 15, 225-231. Oldstone, M. B. A., Tishon, A., Dutko, F. Y., Kennedy, S . I. T.,Holland, J. J., & Lampert, P. W. (1980) J. Virol. 34, 256-265. Petri, W. A., Jr., & Wagner, R. R. (1 979) J.Biol. Chem. 254, 43 13-43 16. Pfefferkorn, E. R., & Clifford, R. L. (1963) Virology 21, 273-274. Pfefferkorn, E. R., & Hunter, H. S. (1963) Virology 20, 433-445. Rouser, G., Fleischer, S., & Yamamoto, A. (1970) Lipids 5, 494-496. Schafer, R., & Franklin, R. M. (1975) J . Mol. Biol. 97, 21-34. Scheule, R. K., & Gaffney, B. J. (1981a) in Liposomes, Drugs and Immunocompetent Cell Functions (Nicolau, C., & Paraf, A., Eds.) pp 79-93, Academic, London. Scheule, R. K., & Gaffney, B. J. (1 98 1 b) Anal. Biochem. I 1 7 , 61-66. Sefton, B. M., & Gaffney, B. J. (1979) Biochemistry 18, 436-442. Shimizu, K., Hosaka, Y., & Shimizu, Y. K. (1972) J . Virol. 9, 842-850. Simons, K., Garoff, H., Helenius, A,, & Ziemiecki, A. (1 978) in Frontiers in Physicochemical Biology (Pullman, B., Ed.) pp 387-407, Academic, New York. Skehel, J. J., Bayley, P. M., Brown, E. B., Martin, S . R., Waterfield, M. D., White, J. M., Wilson I. A., & Wiley, D. C. (1982) Proc. Natl. Acad. Sci. U.S.A. 79, 968-972. Takagaki, Y., Radhakrishnan, R., Gupta, C. M., & Khorana, H. G. (1983) J . Biol. Chem. 258, 9128-9135. Volsky, D. J., & Loyter, A. (1978) FEBS Lett. 92, 190-194. Welch, W . J., & Sefton, B. M. (1979) J. Virol. 29, 1186-1 195. White, J., & Helenius, A. (1980) Proc. Natl. Acad. Sci. U.S.A. 77, 3273-3277. Wickner, W. (1976) Proc. Natl. Acad. Sci. U.S.A. 73, 1159-1 163. Wickner, W., Mandel, G., Zwizinski, C., Bates, M., & Killick, T. (1978) Proc. Natl. Acad. Sci. U.S.A. 75, 1754-1758.
