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Journal of Virology, February 2002, p. 1194-1205, Vol. 76, No. 3
0022-538X/01/$04.00+0     DOI: 10.1128/JVI.76.3.1194-1205.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Molecular Dissection of the Semliki Forest Virus Homotrimer Reveals Two Functionally Distinct Regions of the Fusion Protein

Don L. Gibbons and Margaret Kielian*

Department of Cell Biology, Albert Einstein College of Medicine, Bronx, New York 10461

Received 31 July 2001/ Accepted 23 October 2001


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ABSTRACT
 
Semliki Forest virus (SFV) is an enveloped alphavirus that infects cells via a membrane fusion reaction triggered by the acidic pH of endosomes. In response to low pH, the E1 proteins on the virus membrane undergo a series of conformational changes, resulting in the formation of a stable E1 homotrimer. Little is known about the structural basis of either the E1 conformational changes or the resulting homotrimer or about the mechanism of action of the homotrimer in fusion. Here, the E1 homotrimer was formed in vitro from either virus or soluble E1 ectodomain and then probed by various perturbants, proteases, or glycosidase. The preformed homotrimer was extremely stable to moderately harsh conditions and proteases. By contrast, mild reducing conditions selectively disrupted the N-terminal region of trimeric E1, making it accessible to proteolytic cleavage and producing E1 fragments that retained trimer interactions. Trypsin digestion produced a fragment missing a portion of the N terminus just proximal to the putative fusion peptide. Digestion with elastase produced several fragments with cleavage sites between residues 78 and 102, resulting in the loss of the putative fusion peptide and the release of membrane-bound E1 ectodomain as a soluble trimer. Elastase also cleaved the homotrimer within an E1 loop located near the fusion peptide in the native E1 structure. Mass spectrometry was used to map the C termini of several differentially produced and fully functional E1 ectodomains. Together, our data identify two separate regions of the SFV E1 ectodomain, one responsible for target membrane association and one necessary for trimer interactions.


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INTRODUCTION
 
Enveloped viruses must circumvent the double barrier of their own membranes and the membrane of the host cell in order to deliver their genomes into the cytoplasm and cause infection. They bypass both membrane barriers simultaneously by means of a membrane fusion event, either at the cell surface or in the endocytic pathway (18). The well-characterized alphavirus Semliki Forest virus (SFV) is a positive-strand RNA virus that enters cells by receptor-mediated endocytosis and then fuses with the endosome membrane to infect the cell (see references 23 and 47 for reviews). SFV fusion is mediated by the viral E1 protein, a type I membrane glycoprotein of about 50 kDa. Upon exposure to the low pH of the endosomes, E1 dissociates from its stable dimeric interaction with the companion E2 protein. E1 then undergoes a series of conformational changes resulting in the exposure of previously hidden acid-conformation-specific epitopes and the formation of a highly stable E1 homotrimer. The protein associates with the target membrane, presumably via the putative fusion peptide (approximately residues 83 to 100), and the dual interaction of E1 with the target and virus membranes drives the fusion reaction. SFV fusion is strongly dependent on the presence of cholesterol and sphingolipid in the target membrane (reviewed in reference 24). Although treatment of virus at low pH in the absence of target membranes induces acid-specific epitope exposure and homotrimer formation, these conformational changes are clearly promoted by the presence of cholesterol- and sphingolipid-containing membranes (5, 6). Several virus mutants with reduced cholesterol dependence have been isolated, and srf-3, the best-characterized such mutant, has a single-amino-acid change of E1 proline 226 to serine (48). This mutation specifically reduces the cholesterol dependence of both membrane fusion and E1 homotrimer formation (5), and the 226 region of E1 similarly affects the cholesterol dependence of Sindbis virus, a related alphavirus (34).

Several lines of evidence indicate that the formation of a highly stable E1 homotrimer is critical for SFV fusion. The E1 homotrimer was originally discovered because it is highly resistant to trypsin digestion and because the trimer interaction is stable to gradient sedimentation in the presence of the nonionic detergent NP-40 and to mild sodium dodecyl sulfate (SDS) treatment followed by SDS-polyacrylamide gel electrophoresis (PAGE) (49, 50). In vivo, formation of the homotrimer occurs during virus entry into cells, is blocked by agents that raise the pH of endosomes, and is coincident with virus fusion in the endosome (29, 43, 50). In vitro, homotrimer formation occurs with kinetics, pH dependence, and lipid preferences similar to those of fusion (4, 5, 16, 20, 26, 43). Homotrimer formation and fusion are specifically induced by exposure to low pH rather than by general destabilizing treatments of E1 (15). Importantly, formation of the E1 homotrimer is blocked by a mutation within the E1 fusion peptide, glycine 91 -> aspartate (G91D), that also blocks virus fusion and infection (26). In addition, both E1 homotrimer formation and virus-liposome fusion are inhibited in the presence of Zn2+ (7). Thus, these data suggest that the formation of the trimeric structure of the fusion protein is a critical intermediate in fusion and may be regulated by the E1 fusion peptide.

A number of enveloped animal viruses mediate fusion by the formation of a highly stable trimer structure based on a six-helix bundle with a central {alpha}-helical coiled coil (reviewed in references 19, 46, and 52). Viral fusion proteins using this mechanism, termed class I fusion proteins (32), are exemplified by influenza virus hemagglutinin and currently include members of the orthomyxovirus, paramyxovirus, filovirus, and retrovirus families. The present model for fusion of these viruses is that the virus fusion peptide first interacts with the target membrane and then the energy released by the transition from the native, metastable structure to the highly stable six-helix bundle acts to drive the fusion reaction (reviewed in references 18, 37, 42, and 52). Although an alternative structural motif has not yet been identified, mounting evidence suggests that members of .the alphaviruses and flaviviruses mediate membrane fusion via a novel, non-coiled-coil-based structure (see references 15, 17, 24, and 32 for reviews). The existence of a separate group, termed class II fusion proteins (32), would not be surprising, given some of the obvious differences between these fusion proteins and those of the coiled-coil type, including the lack of extensive {alpha}-helical structure, the dimeric rather than trimeric state of the native alphavirus and flavivirus fusion proteins, and their proteolytic activation by cleavage of a companion subunit rather than the fusogenic subunit.

What is currently known about the formation of the SFV E1 homotrimer? We recently reported that, unlike the coiled-coil class of fusion proteins, neither SFV fusion nor E1 homotrimer formation are induced by heat or treatment with denaturants (15). Instead, low-pH treatment appears to be required as a specific trigger for both fusion and E1 conformational changes. Although little has been published regarding the structural properties of the homotrimer or the portions of the protein that account for its biological roles, a purified E1 ectodomain monomer termed E1* has been extensively characterized (25, 30). E1* associates with liposomes in a low-pH-, cholesterol-, and sphingolipid-dependent manner. Under these conditions, E1* concomitantly undergoes acid-dependent conformational changes, including acid-specific epitope exposure and formation of the trypsin- and SDS-resistant homotrimer. The E1* homotrimer shows induction properties and stability similar to those of the full-length protein, but its formation is more strictly dependent on the presence of cholesterol- and sphingolipid-containing membranes (15, 30).

Recently, an ectodomain preparation of E1 termed E1{Delta}S was crystallized, and the crystal structure was determined at a resolution of 3.5 Å (32, 53). Strikingly, the structure demonstrates that the neutral-pH form of the SFV E1 protein has an overall fold similar to that of E protein, the fusion protein from the flavivirus tick-borne encephalitis (TBE) virus. The SFV E1{Delta}S structure is composed of three domains made up primarily of antiparallel ß-sheet secondary structure, with the transmembrane anchor at one end of the molecule and the putative fusion peptide at the opposite end (Fig. 1A shows a schematic). The protein structure was fitted into the previously published 9-Å cryoelectron microscopy (cryo-EM) reconstruction of the SFV particle (36). These analyses show that, similar to the TBE virus E protein, the elongated E1 protein forms a protein shell covering the virus membrane surface rather than a vertical spike (12, 32). The crystal structure and cryo-EM fitting, along with the recent cryo-EM analyses of glycoprotein mutants from Sindbis virus and an SFV capsid assembly mutant (13, 40), have established the idea that E1 is not only the fusogenic protein of the virus but also forms an icosahedral protein lattice structure on the outer membrane surface that helps to drive budding and to maintain the T=4 symmetry of the virion. The structure supports the model that the E1-E2 dimer interaction is released at low pH, freeing E1 to form the stable homotrimer and drive fusion of the two membranes. Thus, the mechanism of fusion protein trimerization is crucial to our understanding of the fusion of both alphaviruses and flaviviruses, as well as other viruses that may fall into this fusion protein class.



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  		FIG. 1. Effect of ß-ME on the trypsin digestion of the E1 or E1* homotrimer. (A) Linear diagram of domains I (solid), II (hatched), and III (stippled) within the primary sequence of SFV E1 plus the putative stem region (shaded) and transmembrane domain (checkered). (Data are from reference 32). (B) [35S]methionine-labeled virus was mixed with liposomes (0.8 mM), treated at pH 5.5 for 3 min at 20°C to form the E1 homotrimer, and adjusted to neutral pH. Samples were then incubated in the indicated concentrations of ß-ME for 30 min at 37°C, followed by digestion with (+) trypsin (125 µg/ml) in 0.5% TX-100 for 1 h at 37°C. The digestion was stopped by addition of PMSF to a final concentration of 5 mM, and the samples were incubated in SDS sample buffer for 3 min at 30 or 95°C as indicated and analyzed by SDS-PAGE on an 11% acrylamide gel using the Tris-glycine buffer system. The positions of the E1 homotrimer (HT), E2, E1, capsid (C), and trypsin-cleaved E1 (E1-T1) are indicated. Higher-order oligomers of the E1 protein are indicated by the asterisk. (C) [35S]methionine-labeled ectodomains were mixed with liposomes (1 mM), treated at pH 5.5 for 10 min at 37°C to form the E1* homotrimer, and adjusted to neutral pH. Samples were then incubated in 0.25% ß-ME for 30 min at 37°C, digested with trypsin as for panel B for the indicated times (0 to 2.5 h), and analyzed by SDS-PAGE as for panel B, using a 10% polyacrylamide gel with the Tris-Tricine buffer system. The E1* and E2* ectodomains comigrate under these conditions. The positions of the E1* truncated homotrimer (HT{Delta}T) and the T1, T2, and T3 fragments are indicated, and the arrowheads on the left denote the electrophoretic positions of molecular mass standards in kilodaltons.

The goal of the present study was to generate a minimal core fragment of the SFV E1 protein that would account for the interaction of the three subunits of the homotrimer and allow for further biochemical characterization of the mechanism involved. The data reveal that the remarkable stability of homotrimeric E1 is maintained in part by its complex disulfide organization. Treatment under mild reducing conditions allowed proteolytic access specifically to the end of the E1 molecule containing the fusion peptide. The putative fusion peptide region could be removed by proteolysis without disruption of the homotrimer, while the remaining large core of the homotrimer was inaccessible to further proteolysis or to deglycosylation, even under fairly harsh conditions. Removal of the fusion peptide region released the E1 ectodomain from its association with target membranes, producing a soluble trimer.

(The data in this paper are from a thesis to be submitted by D.L.G. in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Sue Golding Graduate Division of Medical Sciences, Albert Einstein College of Medicine, Yeshiva University.)


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MATERIALS AND METHODS
 
Virus and cells. BHK-21 cells were cultured at 37°C in Dulbecco’s modified Eagle medium containing 5% fetal calf serum, 100 U of penicillin per ml, 100 µg of streptomycin per ml, and 10% tryptose phosphate broth (39). The SFV used in these experiments was a well-characterized plaque-purified isolate propagated in BHK-21 cells (16). The virus was radiolabeled with [35S]methionine and purified as previously described (28) or propagated in the absence of label and purified by banding on tartrate gradients (21).

Preparation of SFV E1 ectodomains. Soluble forms of the E1 and E2 proteins (E1* and E2*) were prepared as previously described (25, 30). In brief, for the radiolabeled preparations, a mixture of 35S-labeled SFV (~107 cpm) and purified unlabeled SFV (100 µg of protein) was digested with proteinase K or subtilisin (100 µg/ml) in phosphate-buffered saline containing 0.9 mM CaCl2, 0.5 mM MgCl2, and 0.5% Triton X-114 (TX-114) for 60 to 90 min on ice. The ectodomains were then purified by TX-114 detergent phase separation (3) and concanavalin A chromatography, dialyzed against morpholine ethanesulfonic acid (MES) buffer (20 mM MES [pH 7.0], 130 mM NaCl), and stored at -140°C until they were used. For preparation of unlabeled ectodomains, the same procedure was followed, starting with 1 mg of purified virus and using a protein-to-protease ratio of ~3:1 (wt/wt).

Liposomes. Liposomes were prepared by extrusion, as previously described (5), with phosphatidylcholine (egg yolk)-phosphatidylethanolamine (derived from egg phosphatidylcholine by transphosphatidylation)-sphingomyelin (bovine brain)-cholesterol at a molar ratio of 1:1:1:1.5 for experiments with virus (27). For experiments with ectodomains, equimolar amounts of total phospholipids and cholesterol were used for liposome preparation, yielding a molar ratio of 1:1:1:3 (30). All phospholipids were purchased from Avanti Polar Lipids (Alabaster, Ala.), and cholesterol was from Steraloids (Wilton, N.H.). For assays, liposomes were mixed with ectodomains or SFV at a final concentration of 1 or 0.8 mM, respectively, unless otherwise noted.

Formation and enzymatic digestion of the E1 and E1* homotrimer. [35S]methionine-labeled virus or ectodomains were preincubated with liposomes for 5 min at 20 or 37°C, respectively, adjusted to pH 5.5 by addition of a precalibrated volume of 0.5 N acetic acid, incubated further as indicated in each figure legend, and then neutralized by addition of 0.5 N NaOH. Samples were then incubated in ß-mercaptoethanol (ß-ME) at the indicated concentrations for 30 min at 37°C. Digestions with trypsin (N-tosyl-l-phenylalanine chloromethyl ketone treated; type XIII; Sigma, St. Louis, Mo.) were at a concentration of 125 µg/ml, and those with elastase (grade II; Boehringer Mannheim, Indianapolis, Ind.) were at either 125 or 250 µg/ml as noted in the figure legends. Protease digestions were carried out in the presence of 0.5% TX-100 unless otherwise indicated. Proteolysis was terminated prior to further sample analysis by addition of phenylmethylsulfonyl fluoride (PMSF) to a final concentration of 5 mM. Digestion with peptide N-glycosidase F (PNGase F; New England Biolabs, Beverly, Mass.) was performed at an enzyme concentration of 10,000 U/ml in MES-saline buffer, pH 7.0, for 3 h at 37°C.

SDS-PAGE analysis. Two different SDS gel electrophoresis systems were utilized in this study. Standard SDS-polyacrylamide gels with a Tris-glycine buffer system were used to analyze most samples (31). For samples with smaller polypeptides (<=20 kDa) a Tris-Tricine buffer system was utilized instead (44). For the former, samples were heated for 3 min in SDS sample buffer (200 mM Tris-Cl [pH 8.8], 4% SDS, 10% glycerol, 0.02% bromophenol blue) at either 30°C, to preserve the homotrimer, or 95°C (15) prior to loading, as indicated in the figure legends. For the Tris-Tricine gels, samples were heated for 3 min at the indicated temperature in SDS sample buffer (100 mM Tris-Cl [pH 6.8], 4% SDS, 12% glycerol, 0.01% Coomassie blue G-250). Proteins were further reduced (beyond the ß-ME preincubation) and alkylated where indicated. Quantitation of protein radioactivity was performed by PhosphorImager analysis with Image Quant version 1.2 software (Molecular Dynamics, Sunnyvale, Calif.)

Gel filtration chromatography. A 3-h elastase digestion containing ~350 kcpm of 35S-labeled ectodomains and ~90 kcpm of 3H-labeled liposomes was terminated by the addition of PMSF. The sample was then chromatographed on a 35- by 1.2-cm Superdex 200 column at room temperature in buffer (20 mM Tris-HCl [pH 7.5], 150 mM NaCl, 0.2 mM PMSF, 37.5 µg of N-tosyl-L-phenylalanine chloromethyl ketone/ml, 50 µg of bovine serum albumin/ml, and 0.02% NaN3) at a flow rate of 0.3 ml/min. Fractions (~0.45 ml) were collected, and the radioactivity in a 50-µl aliquot of each fraction was measured by liquid scintillation counting. The column was calibrated with the protein standards ferritin (440 kDa), catalase (232 kDa), aldolase (158 kDa), albumin (67 kDa), ovalbumin (43 kDa), and chymotrypsinogen (25 kDa), and their elution was determined by absorbance at 280 nm. In separate column runs, the elution of 3H-labeled liposomes alone was followed by liquid scintillation counting, and the elution of blue dextran was followed by absorbance at 280 nm.

Sucrose density gradient flotation analysis. Binding of ectodomains to liposomes was detected by coflotation of protein with liposomes on sucrose step gradients as previously described (30). Briefly, low-pH-treated samples were adjusted to neutral pH and a final concentration of 40% sucrose, layered into the bottom of TLS55 ultracentrifuge tubes, and then overlaid with 1.4 ml of 25% sucrose in TN buffer (50 mM Tris-HCl[pH 7.4], 100 mM NaCl) and 0.2 ml of 5% sucrose. The samples were centrifuged for 3 h at 54,000 rpm at 4°C. Seven fractions of 300 µl each were then collected from the gradients. The top four fractions and the bottom three fractions from each gradient were pooled for subsequent analysis. In control experiments, 3H-labeled liposomes were recovered in the top three fractions.

Sequence and mass analysis. Matrix-assisted laser desorption ionization-time of flight (MALDI-TOF) mass spectrometry was used to determine the mass of E1* prepared by either proteinase K or subtilisin digestion. To separate the E1* from the E2* and p62* proteins, purified ectodomains were further purified by reversed-phase high-performance liquid chromatography (HPLC) on a Vydac C18 column at 50°C. The sample was bound to the column and washed in 5% CH3CN-0.1% trifluoroacetic acid buffer and then eluted from the column with an increasing gradient of CH3CN. Fractions from the peaks eluted from the column were concentrated and analyzed by SDS-PAGE and Western blotting with a polyclonal rabbit antiserum against SFV to assign identities to the proteins in each fraction. Fractions containing exclusively E1* were pooled and concentrated. The samples were then mixed with purified bovine serum albumin as an internal standard and the matrix {alpha}-cyano-4-hydroxycinnamic acid before being spotted for MALDI-TOF mass spectrometry analysis, which was performed on a PerSeptive Biosystems Voyager machine in the Einstein Proteomics Facility.

For Edman sequencing, samples were electrophoresed on standard SDS-containing polyacrylamide gels with either the Tris-glycine or Tris-Tricine buffer system and then transferred to polyvinylidene difluoride membranes (Millipore Corp., Bedford, Mass.) by the tank method in a buffer containing 192 mM glycine, 25 mM Tris base, 0.05% lithium dodecyl sulfate, 20% methanol, and 2 mM dithiothreitol. Bands were identified by staining with Coomassie blue R-250, excised, and sequenced in the Einstein Proteomics Facility on an Applied Biosystems Procise 494 automated sequencer.


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RESULTS
 
Mass spectrometry and functional analysis of various E1 ectodomains. In previous work, we have described the generation and characterization of soluble monomeric ectodomains of the E1 and E2 proteins, termed E1* and E2*, by digestion of native SFV with proteinase K in nonionic detergent in the cold (15, 25, 30). The initial report of these ectodomains noted that they are also produced by digestion of the virus with other proteases, such as subtilisin or pronase. Recently, an alternative protocol for production of an E1 ectodomain (termed E1{Delta}S) by subtilisin digestion of purified SFV E1 and E2 protein octamers was published (53) and was the basis for the determination of the protein’s structure (32). Although there is limited biological information for the E1{Delta}S ectodomain, it forms a stable heterodimer with E2{Delta}S in solution and acts as a pH-dependent hemagglutinin, presumably correlating with exposure of the fusion peptide and membrane binding. To gain a better understanding of the E1* ectodomains that have been studied previously and to allow for their comparison with the E1{Delta}S ectodomain used for the X-ray crystal structure, we sought to identify the cleavage point at the C terminus of E1* by mass spectrometry and to compare the biological properties of the various forms of truncated E1.

We produced two forms of the E1* ectodomain using either digestion with proteinase K (E1*pk) or subtilisin (E1*su) and purified them by detergent phase separation and concanavalin A chromatography as previously described (25, 30). The purified E1*su ectodomain was tested for acid-specific epitope exposure, coflotation with target liposomes of the appropriate lipid composition, and homotrimer formation to compare it with E1*pk, which has been characterized by those assays in a number of earlier studies (15, 25, 30). As listed in Table 1, E1*su and E1*pk were functionally indistinguishable from one another. To accurately assign an experimental molecular mass value to the E1*pk or E1*su ectodomain, E1* was separated from E2* and p62* by reversed-phase HPLC and analyzed by MALDI-TOF mass spectrometry. As shown in Table 1, the mass of E1* produced by either proteinase K or subtilisin digestion in our protocol is 44,290 Da. In comparison, the mass reported for E1{Delta}S was 44,075 Da, and the protein’s C terminus was concluded to be at Y390 (53) (Table 1). Our virus was propagated and purified similarly to the virus used to produce E1{Delta}S, allowing direct comparison of the mass results. Although Wengler et al. (53) calculated the C terminus of E1{Delta}S by comparing its mass to that of full-length E1, we have been unable to measure the mass of full-length E1 or of deglycosylated E1*, which would afford a more precise interpretation of the C terminus. However, the most reasonable interpretation of our data, based on comparison with those of Wengler et al. (53), is that E1*pk and E1*su are the same molecule, that their C terminus is A392, and that they differ, at most, by only a few residues from the E1{Delta}S used for crystallization. The functional studies indicate that all three forms of E1 (E1*pk, E1*su, and E1{Delta}S) undergo conformational changes in response to low pH and interact with target membranes. Since E1*su and E1*pk appear to be identical molecules structurally and functionally, they have been used interchangeably in our studies and will subsequently be referred to as E1*. The protein yield of ectodomains with the subtilisin protocol seems to be slightly better than with proteinase K, and the subtilisin protocol is therefore our preferred method for most routine preparations.


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  		TABLE 1. Analysis of E1 ectodomains by MALDI-TOF mass spectrometry

A region of domain II in trimeric E1 is stabilized by intramolecular disulfide bonds. We next sought to define the regions of the E1 ectodomain important in trimerization. The homotrimers of viral E1 or the ectodomain E1*, like the "triggered" conformations of fusion proteins from many viruses, are extremely stable oligomeric structures resistant to proteolysis, heat, and denaturants (15, 25, 49). Two simple assays for homotrimer formation and stability are the molecule’s resistance to digestion by trypsin and its resistance to dissociation by SDS sample buffer at 30°C. When boiled in SDS sample buffer, the homotrimer dissociates into monomers, and the two forms can be readily distinguished in SDS-PAGE (49). We took advantage of these properties to screen for conditions that would allow proteolytic cleavage of trimeric E1 to a core oligomer. The E1 homotrimer was generated by treatment of virus at low pH in the presence of liposomes, followed by solubilization in 0.5% TX-100 and digestion with a wide variety of proteases (data not shown). Most proteases, including trypsin, elastase, bromelain, endoproteinase Glu-C (V8), papain, and thermolysin, showed no detectable digestion of the homotrimer. Chymotrypsin showed no effect early in digestion but completely proteolyzed the homotrimer after incubation for several hours. Pretreatment of the homotrimer with various detergents and denaturants or heat did not alter its general resistance to protease digestion, in keeping with previous results with trypsin (15).

It has been known for many years that reducing agents abrogate the infectivity of SFV and change the electrophoretic mobility of the E1 protein in SDS-PAGE (22). The recently solved crystal structure of E1{Delta}S directly demonstrates the presence of intramolecular disulfide bonds stabilizing the E1 structure, with five disulfide bonds in domain II, the dimerization domain containing the fusion peptide, and three disulfide bonds in domain III, the immunoglobulin-like domain that connects to the stem and transmembrane regions (32) (Fig. 1A shows a schematic of E1 domain organization). We therefore reasoned that perturbation of trimeric E1 with a reducing agent might allow access by proteases where other perturbants had failed. The preformed E1 homotrimer was incubated with increasing concentrations of ß-ME (0.06 to 0.25%) for 30 min at 37°C and then digested with trypsin. Samples were solubilized in SDS gel buffer at either 30 or 95°C and analyzed by SDS-PAGE (Fig. 1B). Lane 1 demonstrates the presence of the homotrimer after 30°C solubilization, while lane 2 shows that the homotrimer band was converted to monomeric E1 by solubilization at 95°C. Pretreatment with ß-ME caused the E2 and E1 subunits to comigrate in SDS-PAGE (lane 9), confirming that this mild reducing treatment accessed at least some of the disulfide bonds in E1. Trypsin completely digested capsid and monomeric E1 and E2, while the E1 homotrimer became selectively sensitive to trypsin following treatment with increasing concentrations of reducing agent (lanes 3 to 8). The homotrimer band migrated as a slightly smaller species (see also Fig. 1C) but retained trimer interactions (lanes 3, 5, and 7). Dissociation of this truncated trimer by heating it at 95°C in sample buffer demonstrated the presence of both intact E1 and increasing amounts of a smaller species, termed E1-T1, migrating at approximately 38 kDa (lanes 4, 6, and 8). The production and size of this digestion product was unchanged by increasing ß-ME concentrations up to at least 1% (130 mM) or by treatment at a fixed reductant concentration (e.g., 0.25%) with digestion times up to 18 h (data not shown). In addition, although the ß-ME-treated homotrimer was more accessible to trypsin, its resistance to dissociation by urea (up to 5 M) or by detergents such as TX-100 and SDS appeared unchanged (data not shown). Thus, ß-ME reduction selectively disrupted local structure in trimeric E1, resulting in protease access, while the remainder of E1 was resistant to further proteolysis and remained trimeric.

N-terminal sequencing of the E1-T1 trypsin digest fragment from an unlabeled virus sample treated as in Fig. 1B, lane 8, identified the site of trypsin cleavage as K79, producing a fragment with Val 80 at the N terminus (Table 2). The calculated peptide mass for an E1 fragment containing residues 80 to 438 is 38.5 kDa, in approximate agreement with the observed E1-T1 mass of approximately 38 kDa in SDS-PAGE (including the single carbohydrate chain). TX-114 phase separation experiments demonstrated that the E1-T1 product, like full-length E1, partitioned into the detergent phase, consistent with the presence of the transmembrane segment (data not shown). Thus, the E1-T1 fragment begins at residue 80, and its properties are consistent with an intact C terminus. We then addressed whether the digested homotrimer might still contain any fragments derived from the region of E1 N-terminal to the K79 cleavage site. Trypsin digestion products similar to those in Fig. 1B, lane 8, were analyzed by Tris-Tricine gel electrophoresis. While smaller fragments were observed, the pattern was complicated by the presence of capsid digestion products (data not shown). We therefore performed the same analysis with homotrimers derived from the E1* ectodomain.


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  		TABLE 2. Trypsin fragments of E1 or E1* homotrimer

Radiolabeled ectodomains were treated at low pH in the presence of liposomes, and the resultant mixture containing E1*, E2*, and the E1* homotrimer was treated with 0.25% ß-ME, digested with trypsin, and analyzed by SDS-PAGE using a Tris-Tricine buffer system (Fig. 1C). A prominent band (E1*-T1) migrating below the position of intact E1* was observed in samples heated in sample buffer at 95°C (lanes 4, 6, and 8), and N-terminal sequence analysis confirmed that this fragment resulted from cleavage after K79, analogous to the E1-T1 fragment (Table 2). In addition, two smaller fragments were visualized, one of ~22 kDa (E1*-T2) and another of ~6 kDa (E1*-T3). These fragments were observed only when the samples were heated in sample buffer at 95°C (e.g., Fig. 1C, lane 7 versus lane 8) and thus are components of the digested homotrimer. The 22-kDa band was not analyzed further due to its overlapping migration with trypsin. A similar product was produced by elastase digestion and will be discussed below. The ~6-kDa fragment was difficult to visualize using radiolabeled protein, presumably because of the limited number of methionine residues in a fragment of this size. However, digestion of unlabeled ectodomains readily allowed E1*-T3 to be visualized by Coomassie blue staining and permitted analysis of its N-terminal sequence (Table 2). The fragment has the same amino terminus as the intact E1 protein. Based on its apparent molecular mass of 6 kDa, E1*-T3 is probably not the entire N-terminal 79 residues of the protein (which has a calculated molecular mass of ~8,950 Da) but has probably been cleaved after one or each of the three lysine residues present between amino acids 52 and 75. The retention of this fragment in the homotrimer suggests that it is either still connected to the E1*-T1 fragment by an intact disulfide bond (perhaps due to the cysteine at residue 49) or that it remains associated via the extended ß-sheet secondary structure of the subunits or the quaternary structure of the oligomer. Together, these results indicate that trimeric E1 is disrupted by mild reduction, allowing protease cleavage at a site(s) adjacent to the fusion peptide. The native E1{Delta}S crystal structure shows that three stabilizing disulfide bonds flank the fusion peptide loop (32), and it is likely that reduction of one or more of these disulfides would destabilize the trimeric E1 sufficiently to allow protease access.

Elastase digestion of the E1 homotrimer. A variety of proteases were next tested for the ability to digest the ß-ME-reduced homotrimer. Several proteases (papain, subtilisin, and thermolysin) completely digested the reduced protein, although they had minimal or no effect on the unreduced protein (data not shown). Strikingly, several nonspecific or relatively nonspecific proteases (elastase, endoproteinase Glu-C, and bromelain) showed no digestion in the absence of reduction but produced fragments similar to trypsin following ß-ME treatment of the homotrimer. The elastase digestion products were characterized in more detail (Fig. 2A). Elastase completely digested monomeric E1, E2, and capsid and produced a faster-migrating form of the E1 homotrimer (HT{Delta}L [Fig. 2A, lanes 2, 4, and 6]). Treatment of these samples with SDS gel buffer at 95°C followed by complete reduction and alkylation resolved the E1 cleavage products into at least three bands, termed E1-L1, E1-L2, and E1-L3, which migrate faster than intact E1 and which progressively arise over the time course of the digestion (Fig. 2A, lanes 3, 5, and 7). Sequence analysis of E1-L1 and E1-L2 revealed that E1-L1 is a heterogeneous mixture of at least three polypeptides with amino termini at E1 K79, F95, and D97, while E1-L2 was composed of a single digestion product beginning at Q102 (Table 3). We were unable to obtain enough E1-L3 for N-terminal sequence analysis.



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  		FIG. 2. Effect of ß-ME on the elastase digestion of the E1 or E1* homotrimer. [35S]methionine-labeled virus (A) or ectodomains (B) were low-pH treated in the presence of liposomes as for 1B and C. Samples were then incubated in 0.25% ß-ME for 30 min at 37°C and digested with elastase (125 µg/ml) in 0.5% TX-100 for the indicated times at 37°C, and PMSF was added to a final concentration of 5 mM. The samples were then incubated in SDS sample buffer at 30°C for 3 min or reduced at 95°C and alkylated prior to SDS-PAGE on a 13% acrylamide gel with the Tris-glycine buffer system (A) or an 11% acrylamide gel with the Tris-Tricine buffer system (B). The labels refer to the elastase-truncated homotrimer (HT{Delta}L), E1 fragments (E1-L1, E1-L2, and E1-L3), and E1* fragments (E1*-L1 to -L6). # (B) denotes an unidentified fragment in lanes 2 and 3 that is seen in some samples early in digestion but is not part of the stable homotrimer. * indicates higher-order oligomers of the E1 protein, and the arrowheads on the left indicate molecular mass standards as in Fig. 1C.


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  		TABLE 3. Elastase fragments of E1 or E1* homotrimer

The elastase digestion studies were repeated with the E1* homotrimer, and the results are shown using the Tris-Tricine gel system to better resolve lower-molecular-mass digestion products (Fig. 2B). Fragments comparable to the E1-L1 and E1-L2 products were observed (E1*-L1 and E1*-L2 [Fig. 2B, lanes 4, 6, and 8 and data not shown]), along with additional bands of approximately 18.5 and 6 kDa, labeled E1*-L4-5 and E1*-L6. N-terminal sequencing of the E1*-L1 fragment revealed a set of fragments similar to those found in E1-L1, with amino termini at E1 residues 81, 95, and 97 (Table 3). E1*-L2 contained a single species with an amino terminus at Q102, identical to that of the E1-L2 product. The ~18.5-kDa fragment could be partially resolved into two bands by SDS-PAGE, and sequence analysis identified their N termini as Q102 and H230 (Table 3). The cleavage sites and molecular masses of these two fragments are consistent with their being generated by cleavage of E1*-L2 approximately in half at residue 230. Although we were unable to obtain sufficient E1*-L6 for sequence analysis, based on its similarity to the 6-kDa product of trypsin digestion, we believe that it is also derived from the E1 region N-terminal to the fusion peptide. Based on the similarities between the trypsin and elastase fragmentation patterns, the two proteases cleave at approximately the same sites in the reduced E1 homotrimer, even though one protease is very specific and the other is nonspecific. Strikingly, the data indicate that the only two regions in the homotrimer accessible to proteolysis are the fusion peptide loop and the region around residue 230.

Target membrane-independent E1 homotrimer. Data from several groups indicate that treatment of SFV at low pH in the absence of target membranes produces an E1 homotrimer but that the pH-treated virus loses the ability to carry out subsequent fusion with target membranes (4, 49). Two alternative explanations for this "fusion-incompetent" homotrimer are, first, that the homotrimer is nonfusogenic because it undergoes different conformational changes in the absence and in the presence of target membranes and, second, that the energy released upon conversion of the native E1 to the stable trimer is utilized in the fusion reaction and formation of the trimer in the absence of target membranes uncouples the two reactions. To address this issue, the homotrimer was formed by acid treatment of virus in the absence of liposomes and then subsequently assayed for trypsin sensitivity with and without ß-ME treatment. As shown in Fig. 3, the E1 homotrimer formed without target membranes was resistant to proteolysis without prior reduction (lanes 3 to 6). Treatment with ß-ME allowed digestion of the trimer to produce a product (labeled E1-T1 [Fig. 3, lanes 8 and 10]) that had the same electrophoretic mobility as the E1-T1 produced in the presence of target membranes (e.g., Fig. 1B, lanes 4, 6, and 8, and data not shown). Homotrimer produced in the absence of liposomes retained its trimeric association after trypsin cleavage (Fig. 3, lanes 7 and 9). Thus, the E1 homotrimer formed in the absence of target membranes had the same trypsin digestion properties as the homotrimer formed in the presence of target membranes from either E1 or E1*, suggesting that the overall homotrimer structure is the same. Even though E1* produces a homogeneous homotrimer population that is uniformly inserted into the target bilayer via the fusion peptide, such an E1* homotrimer was similar to homotrimers produced from virus in the absence of target membranes. These data support the notion that low-pH inactivation of SFV fusion occurs because the irreversible protein conformational changes that normally drive fusion have been triggered without the requisite insertion of the fusion peptide into the target membrane.



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  		FIG. 3. Trypsin digestion of E1 homotrimer (HT) formed in the absence of target membranes. [35S]methionine-labeled virus was acid treated at pH 5.5 for 3 min at 20°C in the absence of liposomes. After neutralization, samples were incubated in the presence (+) or absence (-) of 0.25% ß-ME for 30 min at 37°C and then digested with trypsin (125 µg/ml) in 0.5% TX-100 at 37°C as indicated. Control samples were incubated without trypsin and analyzed by SDS-PAGE as for Fig 1B. * indicates higher-order oligomers of the E1 protein.

The E1 subunit forms higher-order oligomers. In gel samples that were heated to 30°C in SDS buffer, we frequently observed both the homotrimer band and a series of bands that migrated above the position of the homotrimer (1B, 2A, and 3). These species have been observed in homotrimer preparations from radiolabeled virus, from unlabeled virus, and from E1* ectodomains (e.g., see Fig. 6B, lane 1 [15]). Their detergent, heat, and denaturant stability, resistance to proteolysis, and sensitivity to ß-ME all resemble the properties of the E1 homotrimer (1B, 2A, and 3 and data not shown). Similar to the homotrimer, boiling in sample buffer dissociated these higher-molecular-mass species and caused them to migrate in the position of the monomeric E1-E1* or as the proteolytically truncated species produced by trypsin or elastase. Given the large sizes of these oligomers, they did not resolve well on the standard 11% acrylamide gels shown in 1B, 2A, and 3. We therefore treated virus at low pH in the presence of liposomes and removed the other viral subunits by trypsin digestion as shown in 1B and C. The remaining E1 homotrimer products were analyzed by electrophoresis on a 4 to 15% gradient acrylamide gel (data not shown). Under these conditions, the higher-molecular-mass oligomers resolved as discrete bands. The migration of each of these bands was plotted versus the log of the theoretical molecular masses for multimers of the E1 homotrimer (300, 450, 600, and 750 kDa). The plot showed a linear relationship, supporting the idea that the higher-molecular-mass E1 species represent higher-order oligomers of the E1 trimer. An alternative plot using the theoretical masses of multiples of an E1 monomer (50, 150, 200, 250, 300, and 350 kDa) gave a reasonable fit to the data, but the best fit came from the assumption that the bands are higher-order trimers. It was difficult to analyze these species further given the relatively small amounts of the protein population found in each of the oligomer bands, but it is intriguing that the E1 homotrimer may form oligomers that could function similarly to the multiple trimers involved in influenza virus fusion (8).



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  		FIG. 6. Gel filtration analysis of the elastase-truncated E1* homotrimer. (A) [35S]methionine-labeled ectodomains were mixed with [3H]-labeled liposomes (1 mM) and treated at pH 5.5 for 10 min at 37°C to form the E1* homotrimer. Samples were then incubated in 0.25% ß-ME for 30 min at 37°C and digested with elastase (250 µg/ml) for 3 h at 37°C in the absence of detergent. PMSF was added to a final concentration of 5 mM, and the sample was chromatographed on a Superdex 200 column. The elution of radioactivity representing [35S]-labeled protein and [3H]-labeled liposomes was determined by liquid scintillation counting of a 50-µl aliquot of each of the fractions collected. The elution volumes of 3H-labeled liposomes alone or the protein standard albumin (67 kDa), aldolase (158 kDa), or catalase (232 kDa) (positions marked by long arrows) were determined by separate column runs. The short arrows on the abscissa mark the void (determined by blue dextran elution) and total elution volumes of the column. (B) The fractions of the three radioactive peaks shown in panel A were pooled, acid precipitated, and analyzed by SDS-PAGE with a Tris-glycine buffer system after reduction and alkylation of the samples (lanes 3 to 5). Lane 2 contains a sample of the digest mixture prior to chromatography and represents ~7% of the sample loaded onto the column. Lane 1 shows a control of ectodomains treated at low pH without digestion to indicate the positions of E1*, E2*, and the homotrimer (HT).

PNGase F treatment of the E1* homotrimer. The mass studies of the E1* ectodomains reveal that the C-terminal region of native E1 is accessible to proteolysis, resulting in ectodomain generation. Proteolysis of the homotrimers demonstrates that under mild reducing conditions the region around the fusion peptide becomes accessible to proteolysis. In contrast, save for the cleavage site at residue 230, trimeric E1 between residues Q102 and A392 is remarkably stable, even after treatment with reducing agents, detergents, and denaturants. As an additional probe for the stability and accessibility of this central homotrimer region, we followed the deglycosylation of the preformed homotrimer by PNGase F. The SFV E1 protein has a single glycosylation site C-terminal to the fusion peptide region at N141 in domain I. The carbohydrate chain in this position is at least partially accessible to PNGase F in the monomeric form of the E1{Delta}S ectodomain (53). We mixed E1* and E2* with liposomes, treated them at low pH to form E1* homotrimers, digested them with PNGase F, and compared the results to those obtained with parallel reduction and trypsin digestion (Fig. 4). In the absence of prior reduction, most of the monomeric E1* and E2* in the reaction was deglycosylated and comigrated as a band running below the positions of undigested E1* and E2* (Fig. 4, lanes 2 to 3). In contrast, the E1* subunits in the homotrimer were resistant to deglycosylation, as shown by their migration at the control E1* position when the homotrimer was dissociated by being boiled in sample buffer (lane 3). Prior treatment with ß-ME and trypsin did not increase the accessibility of the core E1* homotrimer to PNGase F digestion. A double digest with PNGase F and trypsin (lanes 7 and 8) gave a protein fragment migrating at the position of E1*-T1 indistinguishable from the product produced by digestion of the homotrimer with trypsin alone (lanes 5 and 6). Thus, trypsin was able to digest the reduced E1* homotrimer at residue K79, while the PNGase F site at residue 141 was inaccessible. In a separate experiment, we found that PNGase F digestion of untreated E1* gave partial cleavage (data not shown), similar to the results of Wengler et al. (53). However, given the incomplete nature of the digest, it was unclear if the N141 site switches from an accessible position in monomeric E1* to an inaccessible position in the homotrimer or if PNGase F was only able to cleave a biologically inactive population of E1* monomers that had been somehow denatured. In any event, the results clearly support the general inaccessibility of the core residues of the E1 homotrimer to both proteolysis and deglycosylation.



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  		FIG. 4. PNGase F treatment of the E1* homotrimer. [35S]methionine-labeled ectodomains were mixed with liposomes (1 mM) and treated at pH 5.5 for 10 min at 37°C to form the E1* homotrimer (HT). Samples were then incubated in the absence (-) or presence (+) of 0.25% ß-ME for 30 min at 37°C, followed by treatment with PNGase F (10,000 U/ml) for 3 h at 37°C and/or trypsin (125 µg/ml) for 1 h at 37°C. Samples were reduced and analyzed as for Fig. 1B. The open arrow marks the position of deglycosylated E1* and E2*. The solid arrow marks the position of the E1*-T1 fragment.

Elastase-truncated E1* dissociates from liposomes as a soluble trimer. The elastase-cleaved form of the homotrimer remained trimeric and resistant to further proteolysis even after 6 h of digestion (Fig. 2A, lane 6). The E1*-L2 and E1*-L6 peptides were retained in the stable trimer core, suggesting that removal of the fusion peptide loop by cleavage at sites within and flanking this region (residues 83 to 100 in the crystal structure [32]) did not affect the stability of the trimer. We next explored the effect of elastase digestion on the membrane association of the E1* homotrimer. Low-pH treatment triggers the interaction of the E1* ectodomain with cholesterol- and sphingomyelin-containing liposomes via the fusion peptide region (30; A. Ahn, D. L. Gibbons, and M. Kielian, submitted for publication). Membrane-associated E1* is trimeric and can be separated from E2* and monomeric E1* by flotation on a sucrose step gradient (30). To assess the effect of elastase digestion on this stable E1*-membrane interaction, ectodomains were mixed with liposomes, low-pH treated, adjusted to neutral pH, reduced with ß-ME, digested with elastase in the absence of any detergent, and analyzed by flotation on sucrose gradients. Control experiments showed that the liposomes were unaffected by these treatments and consistently migrated to the top fractions of the sucrose gradients (data not shown). Also, association of the E1* homotrimer with liposomes was unchanged by ß-ME treatment alone; about 60% of the low-pH-treated E1* cofloated with the liposomes with or without reduction by ß-ME (data not shown). Elastase digestion of E1* homotrimer in the absence of detergent produced the same pattern of fragments as that observed in the presence of detergent, including E1*-L1, -2, -4, and -5 (Fig. 5A). All of these cleavage products were found in the bottom fractions of the gradients (lanes 2, 4, and 6). A small amount of undigested E1* remained early in digestion and still cofloated with the liposomes at the top of the gradient (lane 1). These results suggested that elastase digestion released the E1* homotrimer from the liposomes. To confirm that the fragments in the bottom of the gradient were derived from liposome-associated E1* homotrimer, membrane-bound E1* homotrimers were first separated from E2* and free E1* by flotation on a density gradient (Fig. 5B, lane 1). The membrane-bound E1* was then either refloated without further treatment (Fig. 5B, lanes 2 and 3), or treated with ß-ME and elastase, followed by gradient flotation (Fig. 5B, lanes 4 and 5). In the absence of elastase digestion, E1* refloated at the top of the gradient (lane 2), as did a small amount of undigested E1* in the elastase-treated sample (lane 4). In contrast, the elastase-digested homotrimers dissociated from the liposomes and were found at the bottom of the gradient (lane 5). Thus, removal of the fusion peptide region by elastase digestion led to loss of homotrimer-membrane association.



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  		FIG. 5. Gradient flotation analysis of the elastase-truncated E1* homotrimer. (A) [35S]methionine-labeled ectodomains were used to generate E1* homotrimers, and the preparations were digested with elastase for the indicated times, all as for Fig. 2B but in the absence of any detergent. The samples were then analyzed for their association with liposomes by flotation on sucrose step gradients (see Materials and Methods). The top (fractions 1 to 4; T) and bottom (fractions 5 to 7; B) fractions of each gradient were pooled, concentrated by acid precipitation, and then analyzed on 11% acrylamide gels after reduction and alkylation of the samples. (B) [35S]methionine-labeled ectodomains were mixed with liposomes (1 mM), treated at pH 5.5 for 10 min at 37°C, and neutralized, and the E1* homotrimer was isolated by flotation on a sucrose step gradient. The top fractions of the gradient (fractions 1 to 4) were pooled and then split into three aliquots. One sample was electrophoresed without further treatment (lane 1). One sample was reanalyzed in another sucrose step gradient without any other treatment (Refloat; lanes 2 and 3). One sample was incubated in 0.25% ß-ME for 30 min at 37°C, digested with elastase (250 µg/ml) for 2 h at 37°C in the absence of detergent, and then analyzed by flotation on another sucrose step gradient (Elastase; lanes 4 and 5). The gradients were fractionated, and the samples were analyzed as for panel A.

Because trypsin cleaves only on the N-terminal side of the fusion peptide, it would be interesting to determine the membrane association of E1* after trypsin digestion in the absence of detergent. Unfortunately, unlike elastase, trypsin did not efficiently digest the homotrimer without a nonionic detergent in the reaction (data not shown). It is possible that trypsin and elastase have somewhat different accessibilities to the N terminus of the fusion peptide region or that elastase cleavage at the C-terminal side of the fusion peptide of one E1* subunit might enhance cleavage at the N-terminal sites on another subunit in the trimer.

To assay the solubility and oligomeric nature of the elastase-released E1*, radiolabeled ectodomain trimers were digested with elastase in the absence of detergent and chromatographed on a Superdex 200 gel filtration column. The reaction mixture resolved into three peaks of radioactivity (Fig. 6A). Peak I corresponded to the measured void volume of the column (indicated on the abscissa at ~10.2 ml) and was also the elution volume for a 3H-labeled liposome control (data not shown). We estimate that at least 60% of the radioactivity in peak I is accounted for by the presence of 3H-labeled liposomes in the reaction mixture. Peak III corresponded to the total elution volume of this column (indicated on the abscissa at ~27.5 ml). Peak II eluted between albumin (67 kDa) and catalase (232 kDa) and slightly faster than aldolase (158 kDa), three of the molecular mass markers used to calibrate the column. The fractions corresponding to peaks I, II, and III were pooled, concentrated by acid precipitation, and analyzed by SDS-PAGE (Fig. 6B). Lane 2 shows an aliquot of the complete digest mixture before separation on the column (lane 2; ~7% of total). Peak I (lane 3; 9.6 to 12 ml) contains a small amount of full-length E1*, which was presumably still associated with liposomes and therefore eluted in the void volume. Peak II (lane 4; 14.4 to 18.4 ml) contains the fragments (E1*L-1, -2, -4-5, and -6) described above. These fragments coelute as a soluble oligomer in a position consistent with a truncated trimer of E1*. These data are thus in agreement with the electrophoretic migration of elastase-cleaved E1* as a trimer (Fig. 2). The elastase-cleaved molecule does not chromatograph ideally versus standard globular proteins of known molecular mass, perhaps suggesting that it has a nonglobular shape. Peak III (Lane 5; 23.2 to 32 ml) shows no discrete bands, suggesting that it contains primarily nonprecipitable and/or very small peptide products of the digestion, consistent with an elution position in the total volume of the column. The three peaks represent 13, 16, and 71% of the radioactivity recovered from the column, respectively, and together correspond to a yield of ~89% of the total radioactivity loaded, of which ~21% was liposomes and ~79% was protein. A substantial amount of the protein in the reaction mixture was thus completely degraded by elastase, as would be predicted by the presence of the elastase-sensitive E1* and E2* monomers, while a small amount of E1* was uncleaved and remained associated with liposomes. Cleavage by elastase released E1* from the target membrane as a soluble trimer containing a discrete set of polypeptides.


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DISCUSSION
 
The kinetics of formation of the SFV E1 homotrimer are similar to those of virus fusion in vitro and in vivo (4, 20, 50), and the homotrimer constitutes an extremely stable structure that, similar to the coiled-coil-based trimers of the class I fusion proteins, is believed to play a key role in virus-membrane fusion (for reviews, see references 15 and 23). Here, we have used proteolytic cleavage as a tool to define the regions of E1 important in the maintenance of the stable homotrimer. The data revealed that E1 in the preformed homotrimer could be destabilized by mild reducing conditions, selectively allowing protease access to sites flanking the fusion peptide. Cleavage within this region removed the fusion peptide without disruption of the homotrimer and released the trimer from its association with target membranes. The proteolytically released homotrimer was inaccessible to further proteolysis or to deglycosylation, even under fairly harsh conditions. Homotrimers formed in the presence or absence of target membranes were remarkably similar in stability and in sensitivity to reduction and proteases, suggesting that their overall structures were similar. The proteolytically released, soluble homotrimer was composed of a large core of the E1 protein, including most of domains I, II, and III. The truncated homotrimer contained a peptide N-terminal to the fusion peptide and residues 102 to 392 C-terminal to the fusion peptide. Within the latter region, only a site at position 230 was accessible to cleavage. These data collectively indicate that different regions of the SFV E1 subunit are responsible for target membrane association and trimer interactions.

E1* and E1{Delta}S were produced by cleavage within a few residues of each other (~390 to 392) and have similar properties. The one difference appeared to be that the E1{Delta}S ectodomains are present as stable E1{Delta}S-E2{Delta}S dimers in solution (53), while radiolabeled E1* is present as a soluble monomer (25). This may reflect differences in preparation or concentration, or differences in the E2 ectodomain in the two preparations, which we did not characterize to the same degree. In separate studies, we have found that the N-terminal sequence of the proteinase K-cleaved E1 "tail" begins at residue A406, some 7 residues before the start of the transmembrane domain (X. Zhang and M. Kielian, unpublished results). The difference between the C terminus of the ectodomain (A392) and the N terminus of the tail (A406) suggests that further proteolysis occurs in this stem or hinge region, presumably due to its increased accessibility compared to the rest of E1, which is uncleaved during ectodomain generation. It is interesting to compare the role of the stem region of E1 and that of the TBE virus E protein in light of the remarkable similarity in overall fold between the two proteins. Because the membrane-proximal stems are missing in the crystal structures of SFV E1 and TBE virus E, their structures cannot be directly compared. However, the residues from ~392 to 414 constitute one of the least conserved regions of the alphavirus E1, while the larger stem region of flavivirus E protein is conserved and appears to play an important role in trimerization of the E protein in the absence of target membranes (2, 17). The many shared properties of E1* and full-length E1 suggest that the stem region and the transmembrane domain are probably not involved in the early stages of the SFV fusion reaction, including acid-specific E1 conformational changes, trimerization, and target membrane association. However, since the ectodomains cannot be assayed for fusion activity, we cannot comment on whether this region may be important for subsequent steps in membrane fusion.

The internal fusion peptide regions of several viral fusion proteins, including those of SFV, the flaviviruses TBE virus and West Nile virus, the retrovirus avian sarcoma and leukosis virus, and the filovirus Ebola virus, are anchored at either end by disulfide bonds that help to hold the fusion peptide in a loop configuration (9, 14, 35, 38, 41, 51). This loop conformation is presumably important for the insertion of the fusion protein into the target bilayer, and such a loop insertion would differ from the insertion of the amino-terminal fusion peptides found on many, although not all, viral fusion proteins of the influenza virus class. In the case of SFV, the disulfides visualized in the crystal structure of E1{Delta}S fall into two groups, each completely conserved: one stabilizes the ß-strands and loops of domain II, including the fusion peptide, and the other group stabilizes the immunoglobulin-like domain III (32). Domain I contains no disulfide bonds. There is a striking contrast between the protease accessibility of domain II and that of domains I and III. While our data do not directly demonstrate the reduction of specific disulfide bonds, it is clear that the only regions of trimeric E1 that were cleaved by trypsin and elastase following mild reduction were the domain II area containing the fusion peptide and another domain II region containing residue 230.

The evidence presented here indicates that elastase digestion removed the E1 fusion peptide region from the preformed homotrimer without detectable disruption of the trimer structure or stability. Given the cleavage sites and the size of the E1*-L6 fragment, we estimate that the region removed extended from at least residue 79 to 102, including the entire fusion peptide loop. The portion of E1 removed may in fact be somewhat larger (from approximately residue 55 to 102), but this cannot be definitively determined from the current evidence. Since its removal produces a soluble trimer, the fusion peptide is either exclusively responsible for the interaction of the E1 ectodomain with target membrane or plays a dominant role.

The biological role of the SFV fusion peptide region has been explored in a series of mutagenesis studies (10, 26, 33, 45). The G91D mutation, which falls in the middle of the 83-to-100 loop observed in the crystal structure, completely blocks homotrimer formation, virus fusion, and infection, although the mutant protein still shows at least some activity in binding target membranes and acid-specific monoclonal antibodies. Thus, although the fusion peptide region does not appear to be required for the stable trimer once it has formed, this protein region clearly can have functional effects on homotrimer formation. Trimerization of the SFV E1 ectodomain has a strict requirement for target membranes containing specific lipids (30). Recent studies with the E1 ectodomain have demonstrated that membrane-inserted E1* is associated with cholesterol-rich domains in the target membrane via the fusion peptide (Ahn et al., submitted). This may reflect the need to anchor the fusion peptide in the target membrane and permit proper E1* orientation and/or local protein concentration for the conformational changes to occur. In contrast, the stable viral E1 homotrimer can be formed by low-pH treatment in the absence of target membranes. It is possible that in this situation the viral fusion peptide is inserting into its own membrane, obviating the need for an exogenous target membrane. In this regard, however, it is interesting that a mutation within the fusion peptide of the TBE virus E protein, leucine 107->aspartate, completely blocks membrane insertion but still permits homotrimer formation (1). Future studies should address the possibility that the fusion peptide drives fusion via simple anchoring in the target membrane versus a more specific role that might include association with specific lipids and/or regulation of homotrimer formation.

It is striking that, aside from the fusion peptide region, only the E1 region containing residue 230 was accessible to cleavage in the reduced homotrimer. This domain II residue falls within a loop that is closely associated with the fusion peptide in the native E1 structure (F. Rey, personal communication). In addition, the cleavage site is close to the site of the srf-3 mutation P226S, which confers decreased cholesterol dependence for the fusion of both SFV and Sindbis virus (34, 48). It is not yet clear how this region might interact with membranes or with other portions of the E1 protein to affect the cholesterol phenotype. The fact that both the fusion peptide loop and the srf-3-containing loop are selectively proteolyzed could mean that they retain their native close association once the homotrimer has formed, or it could be an independent property of each loop. It will be important to determine if and how the srf-3 loop might affect the activity or lipid requirement of the fusion peptide during membrane insertion.

How might the E1 homotrimer function in mediating membrane fusion? The ability to form the E1 homotrimer is clearly built into the E1 protein, which likely exists in a metastable state on the virion surface (15). The available mutant, inhibitor, and kinetic data suggest that upon exposure to low pH, the E1-E2 dimer dissociates and the E1 subunit then interacts with the target membrane via the fusion peptide (4, 7, 26). However, following E1 membrane insertion, the next step in fusion is not clear. One scenario would be that the energy derived from the formation of the E1 homotrimer would drive the membrane fusion reaction, similar to findings with virus fusion proteins of the coiled-coil type (37, 42). This would imply that the formation of the homotrimer would occur concomitantly with membrane fusion. This model is appealing, given its utilization of the energy generated from stable homotrimer formation and its logical similarity to the fusion mechanisms of other virus fusion proteins. The key event would be the conformational transition, which would function similarly for different viruses, even though the structural basis of the conformation would differ between the class I and class II fusion proteins. However, the available kinetic data suggest that SFV homotrimer formation occurs before the actual membrane-mixing event, not concurrently (4). Thus, an alternative model would be that the homotrimer forms with the fusion peptide productively inserted in the target membrane and then subsequent events and/or E1 protein rearrangements act to drive the fusion reaction. For example, a role for the observed higher-order oligomers of E1 could be envisioned. Given the lag time between low-pH treatment and SFV fusion (4), it is tempting to speculate that these higher-order oligomers might reflect the formation of a fusion protein complex between the two merging membranes.

It may be difficult to differentiate between these two models using only kinetic data. The current measurements do not distinguish between virus homotrimers that involve E1 inserted in the target membrane and nonproductive homotrimers that involve E1 subunits not interacting with the target membrane. This question is also complicated by the rapidity and efficiency of both SFV homotrimer formation and SFV fusion, as well as the fact that the number of homotrimers required to mediate a fusion event is unknown. The availability of information on the structure of the six-helix bundle has made it possible to design peptide inhibitors of the conformational change for the class I fusion proteins (reviewed in references 11 and 52). Such inhibitors were instrumental in determining the role of the conformational change in driving fusion by the class I proteins (e.g., references 37 and 42). Further information on the structure of the class II fusion proteins, including SFV E1, will help in designing inhibitors of the transition to homotrimer, which may address this issue. In addition, it will be important to determine if the class II proteins, similar to the class I fusion proteins, adopt a final conformation that places the fusion peptide and the transmembrane domain at the same end of the molecule (52).


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ACKNOWLEDGMENTS
 
We thank Félix Rey for discussion of results prior to publication and for helpful comments on the manuscript. We thank Anna Ahn for experiments on the activity of E1*su. We also thank the members of our laboratory for their technical expertise, helpful discussions and suggestions, and critical reading of the manuscript. We thank Linda Sinconolfi-Baez and Ruth Angeletti in the Einstein Laboratory for Macromolecular Analysis and Proteomics for expert assistance with HPLC purification, mass spectrometry, and protein sequencing.

This work was supported by a grant to M.K. from the Public Health Service (R01 GM52929) and by the Jack K. and Helen B. Lazar Fellowship in Cell Biology. D.L.G. was supported through the Medical Scientist Training Program (NIH T32 GM 07288). The Einstein Laboratory for Macromolecular Analysis and Proteomics is supported in part by the Albert Einstein Comprehensive Cancer Center (CA13330) and the Diabetes Research and Training Center (DK20541).


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Cell Biology, Albert Einstein College of Medicine, 1300 Morris Park Ave., Bronx, NY 10461. Phone: (718) 430-3638. Fax: (718) 430-8574. E-mail: kielian{at}aecom.yu.edu. Back


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REFERENCES
 
    1
  1. Allison, S. L., J. Schalich, K. Stiasny, C. W. Mandl, and F. X. Heinz. 2001. Mutational evidence for an internal fusion peptide in flavivirus envelope protein E. J. Virol. 75:4268–4275.[Abstract/Free Full Text]
    2. 2
  2. Allison, S. L., K. Stiasny, K. Stadler, C. W. Mandl, and F. X. Heinz. 1999. Mapping of functional elements in the stem-anchor region of tick-borne encephalitis virus envelope protein E. J. Virol. 73:5605–5612.[Abstract/Free Full Text]
    3. 3
  3. Bordier, C. 1981. Phase separation of integral membrane proteins in Triton X-114 solution. J. Biol. Chem. 256:1604–1607.[Abstract/Free Full Text]
    4. 4
  4. Bron, R., J. M. Wahlberg, H. Garoff, and J. Wilschut. 1993. Membrane fusion of Semliki Forest virus in a model system: correlation between fusion kinetics and structural changes in the envelope glycoprotein. EMBO J. 12:693–701.[Medline]
    5. 5
  5. Chatterjee, P. K., M. Vashishtha, and M. Kielian. 2000. Biochemical consequences of a mutation that controls the cholesterol dependence of Semliki Forest virus fusion. J. Virol. 74:1623–1631.[Abstract/Free Full Text]
    6. 6
  6. Corver, J. 1998. Membrane fusion activity of Semliki Forest virus. Ph.D. thesis. Groningen University, Groningen, The Netherlands.
    7. 7
  7. Corver, J., R. Bron, H. Snippe, C. Kraaijeveld, and J. Wilschut. 1997. Membrane fusion activity of Semliki Forest virus in a liposomal model system: specific inhibition by Zn2+ ions. Virology 238:14–21.[CrossRef][Medline]
    8. 8
  8. Danieli, T., S. L. Pelletier, Y. I. Henis, and J. M. White. 1996. Membrane fusion mediated by the influenza virus hemagglutinin requires the concerted action of at least three hemagglutinin trimers. J. Cell Biol. 133:559–569.[Abstract/Free Full Text]
    9. 9
  9. Delos, S. E., and J. M. White. 2000. Critical role for the cysteines flanking the internal fusion peptide of avian sarcoma/leukosis virus envelope glycoprotein. J. Virol. 74:9738–9741.[Abstract/Free Full Text]
    10. 10
  10. Duffus, W. A., P. Levy-Mintz, M. R. Klimjack, and M. Kielian. 1995. Mutations in the putative fusion peptide of Semliki Forest virus affect spike protein oligomerization and virus assembly. J. Virol. 69:2471–2479.[Abstract]
    11. 11
  11. Eckert, D. M., and P. S. Kim. 2001. Design of potent inhibitors of HIV-1 entry from the gp41 N-peptide region. Proc. Natl. Acad. Sci. USA 98:11187–11192.[Abstract/Free Full Text]
    12. 12
  12. Ferlenghi, I., M. Clarke, T. Ruttan, S. L. Allison, J. Schalich, F. X. Heinz, S. C. Harrison, F. A. Rey, and S. D. Fuller. 2001. Molecular organization of a recombinant subviral particle from tick-borne encephalitis virus. Mol. Cell 7:593–602.[CrossRef][Medline]
    13. 13
  13. Forsell, K., L. Xing, T. Kozlovska, R. H. Cheng, and H. Garoff. 2000. Membrane proteins organize a symmetrical virus. EMBO J. 19:5081–5091.[CrossRef][Medline]
    14. 14
  14. Gallaher, W. R. 1996. Similar structural models of the transmembrane proteins of Ebola and avian sarcoma viruses. Cell 85:477–478.[CrossRef][Medline]
    15. 15
  15. Gibbons, D. L., A. Ahn, P. K. Chatterjee, and M. Kielian. 2000. Formation and characterization of the trimeric form of the fusion protein of Semliki Forest virus. J. Virol. 74:7772–7780.[Abstract/Free Full Text]
    16. 16
  16. Glomb-Reinmund, S., and M. Kielian. 1998. fus-1, a pH-shift mutant of Semliki Forest virus, acts by altering spike subunit interactions via a mutation in the E2 subunit. J. Virol. 72:4281–4287.[Abstract/Free Full Text]
    17. 17
  17. Heinz, F. X., and S. L. Allison. 2000. Structures and mechanisms in flavivirus fusion. Adv. Virus Res. 55:231–269.[CrossRef][Medline]
    18. 18
  18. Hernandez, L. D., L. R. Hoffman, T. G. Wolfsberg, and J. M. White. 1996. Virus-cell and cell-cell fusion. Annu. Rev. Cell Dev. Biol. 12:627–661.[CrossRef][Medline]
    19. 19
  19. Hughson, F. M. 1997. Enveloped viruses: a common mode of membrane fusion? Curr. Biol. 7:R565–R569.[CrossRef][Medline]
    20. 20
  20. Justman, J., M. R. Klimjack, and M. Kielian. 1993. Role of spike protein conformational changes in fusion of Semliki Forest virus. J. Virol. 67:7597–7607.[Abstract/Free Full Text]
    21. 21
  21. Kääariäinen, L., K. Simons, and C.-H. von Bonsdorff. 1969. Studies of Semliki Forest virus subviral components. Ann. Med. Exp. Biol. Fenn. 47:235–248.[Medline]
    22. 22
  22. Kaluza, G., and G. Pauli. 1980. The influence of intramolecular disulfide bonds on the structure and function of Semliki Forest virus membrane glycoproteins. Virology 102:300–309.[CrossRef][Medline]
    23. 23
  23. Kielian, M. 1995. Membrane fusion and the alphavirus life cycle. Adv. Virus Res. 45:113–151.[Medline]
    24. 24
  24. Kielian, M., P. K. Chatterjee, D. L. Gibbons, and Y. E. Lu. 2000. Specific roles for lipids in virus fusion and exit: examples from the alphaviruses., p.409–455. In H. Hilderson and S. Fuller (ed.), Subcellular biochemistry, vol. 34. Fusion of biological membranes and related problems. Plenum Publishers, New York, N.Y.
    25. 25
  25. Kielian, M., and A. Helenius. 1985. pH-induced alterations in the fusogenic spike protein of Semliki Forest virus. J. Cell Biol. 101:2284–2291.[Abstract/Free Full Text]
    26. 26
  26. Kielian, M., M. R. Klimjack, S. Ghosh, and W. A. Duffus. 1996. Mechanisms of mutations inhibiting fusion and infection by Semliki Forest virus. J. Cell Biol. 134:863–872.[Abstract/Free Full Text]
    27. 27
  27. Kielian, M. C., and A. Helenius. 1984. The role of cholesterol in the fusion of Semliki Forest virus with membranes. J. Virol. 52:281–283.[Abstract/Free Full Text]
    28. 28
  28. Kielian, M. C., S. Keränen, L. Kääariäinen, and A. Helenius. 1984. Membrane fusion mutants of Semliki Forest virus. J. Cell Biol. 98:139–145.[Abstract/Free Full Text]
    29. 29
  29. Kielian, M. C., M. Marsh, and A. Helenius. 1986. Kinetics of endosome acidification detected by mutant and wild-type Semliki Forest virus. EMBO J. 5:3103–3109.[Medline]
    30. 30
  30. Klimjack, M. R., S. Jeffrey, and M. Kielian. 1994. Membrane and protein interactions of a soluble form of the Semliki Forest virus fusion protein. J. Virol. 68:6940–6946.[Abstract/Free Full Text]
    31. 31
  31. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680–685.[CrossRef][Medline]
    32. 32
  32. Lescar, J., A. Roussel, M. W. Wien, J. Navaza, S. D. Fuller, G. Wengler, and F. A. Rey. 2001. The fusion glycoprotein shell of Semliki Forest virus: an icosahedral assembly primed for fusogenic activation at endosomal pH. Cell 105:137–148.[CrossRef][Medline]
    33. 33
  33. Levy-Mintz, P., and M. Kielian. 1991. Mutagenesis of the putative fusion domain of the Semliki Forest virus spike protein. J. Virol. 65:4292–4300.[Abstract/Free Full Text]
    34. 34
  34. Lu, Y. E., T. Cassese, and M. Kielian. 1999. The cholesterol requirement for Sindbis virus entry and exit and characterization of a spike protein region involved in cholesterol dependence. J. Virol. 73:4272–4278.[Abstract/Free Full Text]
    35. 35
  35. Malashkevich, V. N., B. J. Schneider, M. L. McNally, M. A. Milhollen, J. X. Pang, and P. S. Kim. 1999. Core structure of the envelope glycoprotein GP2 from Ebola virus at 1.9-Å resolution. Proc. Natl. Acad. Sci. USA 96:2662–2667.[Abstract/Free Full Text]
    36. 36
  36. Mancini, E. J., M. Clarke, B. E. Gowen, T. Rutten, and S. D. Fuller. 2000. Cryo-electron microscopy reveals the functional organization of an enveloped virus, Semliki Forest virus. Mol. Cell 5:255–266.[CrossRef][Medline]
    37. 37
  37. Melikyan, G. B., R. M. Markosyan, H. Hemmati, M. K. Delmedico, D. M. Lambert, and F. S. Cohen. 2000. Evidence that the transition of HIV-1 gp41 into a six-helix bundle, not the bundle configuration, induces membrane fusion. J. Cell Biol. 151:413–423.[Abstract/Free Full Text]
    38. 38
  38. Nowak, T., and G. Wengler. 1987. Analysis of disulfides present in the membrane proteins of the West Nile flavivirus. Virology 156:127–137.[CrossRef][Medline]
    39. 39
  39. Phalen, T., and M. Kielian. 1991. Cholesterol is required for infection by Semliki Forest virus. J. Cell Biol. 112:615–623.[Abstract/Free Full Text]
    40. 40
  40. Pletnev, S. V., W. Zhang, S. Mukhopadhyay, B. R. Fisher, R. Hernandez, D. T. Brown, T. S. Baker, M. G. Rossmann, and R. J. Kuhn. 2001. Locations of carbohydrate sites on alphavirus glycoproteins show that E1 forms an icosahedral scaffold. Cell 105:127–136.[CrossRef][Medline]
    41. 41
  41. Rey, F. A., F. X. Heinz, C. Mandl, C. Kunz, and S. C. Harrison. 1995. The envelope glycoprotein from tick-borne encephalitis virus at 2A resolution. Nature 375:291–298.[CrossRef][Medline]
    42. 42
  42. Russell, C. J., T. S. Jardetzky, and R. A. Lamb. 2001. Membrane fusion machines of paramyxoviruses: capture of intermediates of fusion. EMBO J. 20:4024–4034.[CrossRef][Medline]
    43. 43
  43. Salminen, A., J. M. Wahlberg, M. Lobigs, P. Liljeström, and H. Garoff. 1992. Membrane fusion process of Semliki Forest virus. II. Cleavage-dependent reorganization of the spike protein complex controls virus entry. J. Cell Biol. 116:349–357.[Abstract/Free Full Text]
    44. 44
  44. Schagger, H., and G. von Jagow. 1987. Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal. Biochem. 166:368–379.[CrossRef][Medline]
    45. 45
  45. Shome, S. G., and M. Kielian. 2001. Differential roles of two conserved glycine residues in the fusion peptide of Semliki Forest virus. Virology 279:146–160.[CrossRef][Medline]
    46. 46
  46. Skehel, J. J., and D. C. Wiley. 1998. Coiled coils in both intracellular vesicle and viral membrane fusion. Cell 95:871–874.[CrossRef][Medline]
    47. 47
  47. Strauss, J. H., and E. G. Strauss. 1994. The alphaviruses: gene expression, replication, and evolution. Microbiol. Rev. 58:491–562.[Abstract/Free Full Text]
    48. 48
  48. Vashishtha, M., T. Phalen, M. T. Marquardt, J. S. Ryu, A. C. Ng, and M. Kielian. 1998. A single point mutation controls the cholesterol dependence of Semliki Forest virus entry and exit. J. Cell Biol. 140:91–99.[Abstract/Free Full Text]
    49. 49
  49. Wahlberg, J. M., R. Bron, J. Wilschut, and H. Garoff. 1992. Membrane fusion of Semliki Forest virus involves homotrimers of the fusion protein. J. Virol. 66:7309–7318.[Abstract/Free Full Text]
    50. 50
  50. Wahlberg, J. M., and H. Garoff. 1992. Membrane fusion process of Semliki Forest virus. I. Low pH-induced rearrangement in spike protein quaternary structure precedes virus penetration into cells. J. Cell Biol. 116:339–348.[Abstract/Free Full Text]
    51. 51
  51. Weissenhorn, W., A. Carfi, K. H. Lee, J. J. Skehel, and D. C. Wiley. 1998. Crystal structure of the Ebola virus membrane fusion subunit, GP2, from the envelope glycoprotein ectodomain. Mol. Cell 2:605–616.[CrossRef][Medline]
    52. 52
  52. Weissenhorn, W., A. Dessen, L. J. Calder, S. C. Harrison, J. J. Skehel, and D. C. Wiley. 1999. Structural basis for membrane fusion by enveloped viruses. Mol. Membrane Biol. 16:3–9.[CrossRef][Medline]
    53. 53
  53. Wengler, G., G. Wengler, and F. A. Rey. 1999. The isolation of the ectodomain of the alphavirus E1 protein as a soluble hemagglutinin and its crystallization. Virology 257:472–482.[CrossRef][Medline]

Journal of Virology, February 2002, p. 1194-1205, Vol. 76, No. 3
0022-538X/01/$04.00+0     DOI: 10.1128/JVI.76.3.1194-1205.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



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