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Previous Article | Next Article 
Journal of Virology, April 2007, p. 3130-3141, Vol. 81, No. 7
0022-538X/07/$08.00+0 doi:10.1128/JVI.02464-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.
Department of Infectious Diseases, St. Jude Children's Research Hospital, 332 N. Lauderdale, Memphis, Tennessee 38105-2794
Received 8 November 2006/ Accepted 9 January 2007
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ABSTRACT |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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HRB-derived peptides usually inhibit membrane fusion mediated only by the F protein from which they are derived (31). However, inhibition of closely related F proteins has been demonstrated (3), and an HRB-derived peptide from human parainfluenza virus type 3 (hPIV3) has been shown to inhibit Hendra virus fusion (39). The mechanism of cross-species inhibition is unresolved but may involve the topography of the nonpolar side chains on the hydrophobic face of the HRB-derived hPIV3 peptide allowing favorable packing interactions with the HRA coiled coil in the Hendra virus F protein. An implication of this finding is that the F protein hairpin may tolerate mutations, as long as the periodicity of nonpolar and polar residues is conserved and packing interactions are favorable. Conservative mutations to HR residues in the F protein of PIV5 (formerly known as simian virus 5) slightly reduce hairpin stability (17, 47) but more dramatically affect the stability and activation of the prefusion conformation of the F protein (47, 60). Thus, the tolerance of multiple conformations of the F protein to conservative mutations in the HR regions remains an unresolved issue, despite these residues being targeted in drug design efforts.
X-ray crystal structures
have been determined for two conformations of the F protein ectodomain:
an uncleaved prefusion form and a hairpin form (Fig.
1C and D). The prefusion
form of the PIV5 F protein has a mushroom-like shape formed by a large
globular head attached to a rod-like stalk
(64). The stalk consists
of a triple-stranded coiled coil formed by HRB, and the head is formed
by three domains: DI, DII, and DIII. At the base of the head, DI forms
a highly twisted ß-barrel-like assembly, and DII forms an
immunoglobulin-like ß-sandwich domain. The fusion peptide is
located on the side of the head and is held in place by interactions
with other residues in DIII and with residues in DII. At the top of the
head, DIII contains HRA in a crumpled, spring-loaded structure
consisting of 
-helices, ß-strands, and turns (Fig.
1C, inset). In the hairpin
structures of the F proteins of Newcastle disease virus (NDV)
(7) and hPIV3
(63), HRA forms a
triple-stranded coiled coil that is buttressed by antiparallel HRB
helices. DI and DII remain intact in both forms of the F protein,
undergoing changes in relative orientation to each other.
Many questions remain about how the paramyxovirus F protein regulates its irreversible structural change from a prefusion to a hairpin structure to cause membrane fusion at the right time and place. Mutations to isolated residues in the fusion peptide (27, 45), in HRA (28, 60), between DI and DIII (52), and in HRB (37, 47) of the F proteins of PIV5, NDV, and hPIV3 have been found to increase membrane fusion activity, presumably by destabilizing the prefusion conformation of the F protein. A retrospective analysis of the mutational studies, in light of the recently determined high-resolution structures of the F protein (7, 63, 64), showed that these residues are located in regions of the F protein that undergo significant conformational changes during the transition from the prefusion to the hairpin structure.
On the basis of these
observations, we hypothesized that F protein residues that undergo
dramatic changes in secondary and tertiary structure between the
prefusion and hairpin conformations are important in regulating the
membrane fusion activity of the F protein. To test this hypothesis, we
performed a mutational analysis on residues in a highly conserved
portion of the HRA domain of the Sendai virus (SeV) F protein (Fig.
1). The 10 residues in
this sequence form part of a ß-strand/turn/-helix
structure in the prefusion conformation and part of the triple-stranded
coiled coil in the hairpin conformation (Fig.
1C and D). Consistent with
this portion of the HRA region playing an important role in the folding
and stability of the prefusion F conformation, nine of the mutations
significantly disrupted F protein expression, processing,
and fusogenicity. Conversely, the other six
mutations caused the SeV F protein to have hyperactive membrane fusion
activity, most likely by destabilizing the spring-loaded conformation
of HRA. Overall, these HRA residues that undergo significant structural
changes during protein refolding were observed to be key regulators of
F protein folding and
fusogenicity.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The HRA mutants of SeV Fcl were generated in the background of pGEM3X plasmids and subcloned into the pCAGGS plasmid as described above. The nucleotide sequences of all the plasmid constructs were verified by DNA sequencing, which was performed in the Hartwell Center for Bioinformatics and Biotechnology at St. Jude Children's Research Hospital.
Cell culture. Monolayer cultures of Vero cells (ATCC CCL-81), BHK-21 cells (ATCC CCL-10), and BSR-T7/5 cells (4) were grown in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum, 1% glutamine, 1% penicillin, and 1% streptomycin. BSR-T7/5 cells were also grown in the presence of Geneticin (1 mg/ml, final concentration), which was added to the DMEM every other passage. BHK-21 cells were also supplemented with 10% tryptose phosphate broth. CV-1 cells (ATCC CCL-70) were grown in DMEM supplemented with 10% Nu-serum, 1% glutamine, 1% penicillin, and 1% streptomycin.
Transient expression of viral envelope glycoproteins. Monolayers of Vero cells in six-well dishes (85% to 95% confluence) were transiently transfected with 1 µg of pCAGGS F protein DNA, 1 µg pCAGGS HN protein DNA, or both by using the Lipofectamine Plus expression system (Invitrogen) according to the manufacturer's instructions. Transfected Vero cells were incubated for 4 h at 37°C. DMEM (containing 10% fetal bovine serum and 1% glutamine) was then added to the cells, which were subsequently incubated for 16 h at 37°C. Cells were then treated as indicated for each experiment.
Flow cytometry. At 16 h posttransfection, the Vero cells were washed five times with phosphate-buffered saline containing calcium and magnesium at 0.1 g/liter(PBS+) solution, overlaid with PBS+ solution containing primary antibody, and incubated at 4°C for 30 min. For quantification of the cell surface expression of the SeV F protein, the primary monoclonal antibody M16 (dilution, 1:200) (41) was used. The transfected cells were subsequently washed five times with PBS+, overlaid with PBS+ solution containing fluorescein isothiocyanate-conjugated goat anti-mouse secondary antibody (dilution, 1:100), incubated at 4°C for 30 min, and washed five times with PBS+. Cells were removed from six-well dishes with PBS deficient in calcium and magnesium (PBS–) and containing 50 mM EDTA and were fixed in suspension by adding methanol-free formaldehyde (final concentration, 0.5%). The cell surface fluorescence of 10,000 cells was measured using a FACSCalibur flow cytometer (Becton Dickinson). Mean fluorescence intensities (MFIs) were normalized to the MFI of the SeV F wild-type protein.
Generation of a monoclonal antibody against the cytoplasmic tail of the SeV F protein. B6 mice were immunized with 25 µg keyhole limpet hemocyanin (KLH)-conjugated peptide from the cytoplasmic tail of the SeV F protein (sequence, NPDDRIPRDTYTLEPKIR). One month later, mice received a 25-µg KLH-peptide boost with complete Freund's adjuvant. Three days prior to cell fusion, mice received a final 25-µg boost of KLH-peptide in PBS. Hybridomas were produced by fusion of spleen cells to P3x63-Ag8.653 mouse myeloma cells (ATCC CRL-158). Monoclonal antibodies were selected from tissue culture supernatant plated on 96-well plates coated with nonconjugated SeV F cytoplasmic tail peptide. Hybridoma SeV F tail 31705-1C12 was produced in larger quantities by Harlan Bioproducts for Science (Madison, WI).
Radioimmunoprecipitation. Vero cells expressing F proteins (wild type or mutant) were maintained in culture for 30 min in methionine- and cysteine-free medium and then labeled for 15 min with 50 µCi [35S]Promix (Amersham Pharmacia Biotech) in 0.5 ml of DMEM lacking methionine and cysteine and containing 20 mM HEPES buffer (pH 7.3). The cells were then washed once with PBS+ and chased with 3 ml of DMEM containing 2 mM methionine, 2 mM cysteine, and 20 mM HEPES buffer (pH 7.3) for the reported time. Samples were lysed with ice-cold radioimmunoprecipitation assay (RIPA) buffer containing 0.15 M NaCl, 9.25 mg/ml iodoacetamide, 1.7 mg/ml aprotinin, and 10 mM phenylmethylsulfonyl fluoride (36). The lysate was spun at 67,000 x g in an Optima TLX ultracentrifuge (Beckman Coulter).
The supernatant was incubated overnight (18 to 22 h) at 4°C with 5 µl mouse anti-F tail peptide monoclonal antibody (dilution, 1:200). Immune complexes were adsorbed to protein G-Sepharose 4 Fast Flow (GE Healthcare) for 1 h at 4°C. Samples were washed three times with RIPA buffer containing 0.3 M NaCl, three times with RIPA buffer containing 0.15 M NaCl, and once with 50 mM Tris buffer (0.25 mM EDTA, 0.15 M NaCl, pH 7.4). The samples were resuspended in 50 µl of sample dye buffer containing 200 mM Tris, 8% sodium dodecyl sulfate (SDS), 0.2% bromophenol blue, 40% glycerol, and 12% ß-mercaptoethanol. The samples were then boiled for 5 min, centrifuged at high speed for 1 min, and fractionated on 12% NuPAGE bis-Tris polyacrylamide-SDS gels (Invitrogen). Protein bands were visualized using a Typhoon 9200 phosphorimager (GE Healthcare) and quantified using ImageQuant 5.2 software (Molecular Dynamics).
Total F0 protein expression and cleavage. Five hours after transfection, the cells were washed twice with PBS+ solution. Cells were then chased for 0 or 180 min and radioimmunoprecipitated as described above. F0 bands were visualized as described above. For analysis of F protein cleavage, at 5 h posttransfection cells were pulsed for 15 min and chased for 3 h as described above. During the last hour of the chase, TPCK (tosylsulfonyl phenylalanyl chloromethyl ketone)-treated trypsin (5 µg/µl; Sigma) was added to each sample in 0.5 ml chase medium (DMEM, 1% penicillin, 1% streptomycin, 2 mM methionine, 2 mM cysteine, and 20 mM HEPES). The samples were then radioimmunoprecipitated as described above. F0 and F1 bands were visualized and quantified as described above.
Biotinylation of surface-expressed F protein. At 16 h posttransfection, the Vero cells were washed twice with PBS+ solution. Cells were then radiolabeled and chased for 180 min. The samples were subsequently biotinylated twice for 15 min at 4°C with 2 mg of EZ-Link Sulfo-NHS-SS-biotin (Pierce) in 1 ml of PBS at pH 8. Excess biotin was washed off with PBS containing 50 mM glycine. Samples were then radioimmunoprecipitated as described above. Following the wash with 50 mM Tris buffer (0.25 mM EDTA, 0.15 M NaCl, pH 7.4), the samples were resuspended in 100 µl of 50 mM Tris buffer (0.5% SDS, pH 7.4), boiled for 5 min, and centrifuged at high speed for 1 min. The supernatants were split into two 50-µl fractions. One fraction was saved for direct loading onto the SDS-polyacrylamide gel for polyacrylamide gel electrophoresis (PAGE). The other fraction was diluted to 1 ml with streptavidin buffer (20 mM Tris, 0.15 mM NaCl, 5 mM EDTA, 1% Triton X-100, 0.2% bovine serum albumin, pH 8) and incubated with streptavidin-agarose overnight at 4°C. The samples were washed as described above for radioimmunoprecipitation, resuspended in SDS-PAGE sample buffer containing 12% ß-mercaptoethanol, and fractionated on SDS-12% polyacrylamide gels. The F0 bands were visualized and quantified as described above.
Endo H digestion. To test for the conversion of N-linked carbohydrate chains from the high-mannose form to the complex form, we radioimmunoprecipitated the F protein from transfected cells at chase times of 0, 45, 90, 135, and 180 min. The immune complexes were dissociated by boiling in 50 mM Tris-HCl (pH 7.4) and 0.5% (wt/vol) SDS. To each sample, a solution of 0.1 M sodium citrate (pH 5.3) containing 1 mM phenylmethylsulfonyl fluoride and 2 mU of endoglycosidase H (endo H) (Roche Diagnostics) was added, and digestion was carried out for 18 to 24 h at 37°C. The reaction was terminated by the addition of sample dye buffer. Samples were fractionated in a 12% polyacrylamide-SDS gel under nonreducing conditions. Results were visualized as described above.
Luciferase reporter gene assay for cell-cell membrane fusion. To quantify membrane fusion, we performed a luciferase reporter gene assay as described previously (47). Briefly, six-well dishes containing Vero cells (70% to 80% confluence) were transfected with 1.0 µg luciferase control DNA (Promega), 1.0 µg pCAGGS F DNA, and 1.0 µg pCAGGS HN DNA. At 16 h posttransfection, Vero cells were treated with TPCK-treated trypsin (5 µg/ml) for 1 h at 37°C. Soybean trypsin inhibitor (20 µg/ml; Sigma) was then added, and the samples were incubated at 37°C for 30 min. BSR-T7/5 target cells (expressing T7 RNA polymerase) were overlaid onto the Vero cells expressing the SeV F and HN proteins. After an 8-h incubation at 37°C, the monolayers were washed, lysed in reporter lysis buffer (Promega), and clarified by centrifugation at 15,000 x g in a tabletop centrifuge (5417C; Eppendorf) at room temperature. From each clarified lysate, a 150-µl sample was transferred into a 96-well plate (Lumitrac 200; Promega). The luciferase activity resulting from fusion of the two cell populations was quantified with a Veritas luminometer (Promega); 100 µl luciferase assay substrate (Promega) was injected into each sample.
Dye transfer fusion assay for lipid mixing and content mixing. To measure whether any of the SeV F protein mutants were deficient in either lipid mixing or content mixing, we performed a standard dye transfer fusion assay as described previously (45), with a few modifications. Monolayers of CV-1 cells were grown in six-well plates and transfected with pGEM3X SeV F and HN plasmids by using the vaccinia virus-bacteriophage T7 RNA polymerase (vTF7.3) expression system (20). Before infection of the cells, the vaccinia virus was inactivated with psoralen (Trioxsalen, 4'-aminomethyl-, hydrochloride; 1 µl per well) (Calbiochem) and irradiated with long-wavelength UV light for 2 min. At 16 to 20 h posttransfection, CV-1 cells were incubated in the presence of TPCK-treated trypsin (5 µg/ml) for 30 min at 37°C. The Fcl mutants were tested for lipid mixing and content mixing in the absence of trypsin treatment. Human red blood cells (RBCs) were double labeled with octadecyl rhodamine B chloride (R18) and 6-carboxyfluorescein (6CF) (both from Molecular Probes) as described previously (45). Membrane fusion was quantified by counting 6CF or R18 dye transfer events from labeled RBCs to CV-1 cells. To measure the temperature dependence of cell-cell fusion by the different SeV F protein mutants, we labeled CV-1 effector cells with the red fluorescent nucleic acid stain SYTO 17 (Molecular Probes). RBCs were single labeled with 6CF. Fusion was promoted by incubation for 15 min at 25, 29, 33, or 37°C followed by washing with PBS+ at 4°C and a second incubation on ice. Dye transfer was visualized using a Nikon eclipse TE300 fluorescence microscope, and random fields were captured using the IPLab 3.7 software (BD Biosciences).
Syncytium assay for cell-cell fusion by Fcl mutants. Monolayers of BHK-21 cells grown in six-well plates were transfected with 1 µg pCAGGS SeV Fcl and 1 µg pCAGGS HN DNA as described above. At 24 h posttransfection, cells were fixed and stained with Hema 3 solution (Fisher) per the manufacturer's instructions. Representative fields were captured with a Nikon D70 digital camera attached to a Nikon Eclipse TS100 inverted microscope.
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-helix
formation, protein expression and processing, or membrane fusion
activity (60). Therefore,
we investigated the functional importance of the 10-residue HRA
sequence of the SeV F protein by mutating each residue individually to
the residues of Nipah virus, hPIV3, PIV5, and NDV (Fig.
1; Table
1). The 15 conservative mutations maintained the periodicity of hydrophobic
and polar residues in HRA and thus were not expected to disrupt
significantly the HRA coiled coil in the prehairpin intermediate or the
hairpin structure.
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Mutant F proteins that reach the cell surface are efficiently cleaved by trypsin. To become active for membrane fusion, the precursor F0 protein must first be cleaved into F1 and F2 subunits (31). Many paramyxovirus F0 proteins contain an R-X-R/K-R consensus sequence for intracellular cleavage by furin or furin-like proteases. However, most isolates of SeV contain F proteins that lack a multibasic cleavage site and are cleaved by exogenous proteases while replicating in the lungs of mice (49). When expressed on the surface of tissue culture cells, the SeV F0 protein was efficiently cleaved by exogenous trypsin (Fig. 3A). A densitometric analysis of the relative abundance of the wild-type F0 and F1 bands showed that the wild-type F protein was cleaved by exogenous trypsin with an efficiency of 90% (Fig. 3B; Table 1). Because mutations to HRA residues could potentially interfere with proteolysis, we measured the extents to which the 15 mutant F proteins were cleaved by exogenous trypsin (Fig. 3). All of the mutant F0 proteins that reached the cell surface were cleaved to an extent comparable to that of wild-type F0 protein, whereas the mutant F0 proteins that did not reach the cell surface were not efficiently cleaved.
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The dramatic degree to which the HRA mutations substantially increase, proportionally decrease, or completely eliminate membrane fusion activity confirms our original hypothesis that residues whose secondary and tertiary structures dramatically change during protein refolding are key regulators of the F protein's membrane fusion activity.
Wild-type and mutant F proteins with decreased fusogenicity do not have hemifusion phenotypes. Next, we further examined the mechanism by which the above-mentioned mutations regulate the fusogenicity of the F protein. The lipid bilayers of enveloped viruses are thought to rearrange during membrane fusion in a series of discrete steps that include dimpling, lipid stalk formation, hemifusion, transient pore formation, and pore enlargement (9, 18). In some cases, mutations in the fusion peptide of the influenza virus hemagglutinin (HA) protein cause a hemifusion phenotype (lipid mixing without content mixing) (42, 54). Therefore, we investigated whether the wild-type or mutant F proteins arrest membrane fusion at the hemifusion stage. We performed a fluorescent dye transfer assay with target RBCs colabeled with the lipidic probe R18 and the aqueous probe 6CF. For wild-type and mutant F proteins that had decreased fusogenicities in the luciferase assay (i.e., K180G, K180Q, T181A, and V186I), lipid mixing and content mixing were coincident (Fig. 6A). The mutant F proteins with hyperfusogenic phenotypes also had equivalent levels of lipid mixing and content mixing in the dye transfer assay (data not shown). The data were consistent with wild-type and mutant F proteins not arresting membrane fusion during late stages when a fusion pore opens and enlarges.
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The simplest mechanism by which the HRA mutations could increase fusogenicity is by destabilizing the spring-loaded conformation of HRA (Fig. 1C, inset) so that it has a lower activation energy barrier to form the coiled-coil structure (Fig. 1D, inset). To indirectly measure the activation energy of the F proteins containing hyperactive mutations (i.e., A178V, L179I, L179V, K180T, T181S, and F185Y), we compared their fusogenicities to that of the wild-type F protein as a function of incubation temperature (Fig. 6C). Wild-type F protein caused efficient dye transfer after incubation with target cells at 37°C but dropped to approximately 25% efficiency after incubation at 33°C. In contrast, the F proteins containing hyperfusogenic mutations caused dye transfer more efficiently at lower temperatures than did the wild-type F protein. In fact, the L179V and F185Y mutants caused dye transfer more efficiently at 25°C than the wild-type F protein did at 37°C.
In most cases, the results from the luciferase and dye transfer assays were roughly equivalent. However, the luciferase activity of the F185Y mutant was similar to that of wild-type F protein, whereas dye transfer caused by the F185Y mutant was almost twice as efficient. Target cells and effector cells were coincubated for 8 h in the luciferase assay and for 15 min in the dye transfer assay. This discrepancy between the phenotypes of the F185Y mutant may reflect fast activation kinetics that saturate over a longer period of time. However, further studies are needed to confirm this hypothesis.
Intracellular cleavage and delayed incubation with target cells do not inactivate F proteins carrying hyperfusogenic mutations. Unlike the SeV F0 protein, the PIV5 F0 protein is cleaved intracellularly by resident proteases in the secretory pathway (31). Mutation of residue L477 or I449 in the HRB region of the PIV5 F protein to an aromatic residue results in hyperfusogenic fusion activity in assays where the F protein comes into contact immediately with target cells after reaching the cell surface (e.g., syncytium and luciferase assays) (47). However, these same mutations also cause the PIV F protein to become quickly inactivated following intracellular cleavage and delayed incubation with target cells (i.e., in the dye transfer assay). To investigate further the "do-or-die" phenotypes, the L447 and I449 mutations were introduced into the background of a PIV5 F protein that had a modified cleavage site. As a result, the mutant PIV5 F proteins were not cleaved intracellularly but instead were cleaved by exogenous trypsin immediately before incubation with target cells. Extracellular cleavage of the mutant PIV5 F proteins resulted in hyperactive membrane fusion activity in the dye transfer assay similar to that found in the luciferase assay. Thus, if the hyperfusogenic HRA mutations in the SeV F protein studied here have an effect similar to that of the hyperfusogenic HRB mutations in the PIV5 F protein (destabilizing the native F protein structure before HN binds to target cells), then intracellular cleavage of the SeV F protein might also result in do-or-die phenotypes for the SeV F protein mutants in the dye transfer assay.
To investigate whether the mutational effects of the HRA residues were influenced by extracellular cleavage, we generated an SeV F protein in which its cleavage site was mutated to R-Q-K-R (designated Fcl) (see Materials and Methods), which is similar to another SeV F protein construct that is intracellularly cleaved (26). The 10 HRA mutations that did not previously eliminate cell surface expression (Fig. 2B) were then subcloned into the background of the Fcl construct. The Fcl/HRA mutants were cleaved intracellularly similarly to wild-type Fcl, and they were also expressed at levels similar to those when they were in the trypsin-cleavable F background (data not shown). When coexpressed with SeV HN, the Fcl/HRA mutants promoted cell-cell fusion in syncytium, luciferase, and dye transfer assays after intracellular cleavage (Fig. 7), at levels comparable to those obtained after extracellular cleavage (Fig. 5 and 6). The L179V, K180T, T181S, and F185Y mutants in the Fcl background had hyperfusogenic phenotypes, and the A178V and L179I mutants caused fusion at levels similar to or slightly above that caused by the wild-type Fcl. Conversely, the K180G, K180Q, T181A, and V186I mutants in the Fcl background were generally less fusogenic than wild-type Fcl in the three assays. Interestingly, the F185Y mutation in the Fcl background caused a much larger increase in membrane fusion in the dye transfer assay than in the luciferase assay, similar to the effects of the F185Y mutation in the background of the F protein having a wild-type cleavage site. The results from the fusion assays of the Fcl mutants showed that the means by which the SeV F protein is cleaved does not significantly alter the effects of the HRA mutations. Moreover, the hyperfusogenic HRA mutations do not inactivate the SeV F protein before the binding of HN to target cells. Although not conclusive, these results are consistent with the hypothesis that hyperactive mutations enhance the activation of the F protein only after the HN protein triggers F protein refolding, most likely at the step when the HRA region springs into its coiled coil form.
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DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To assess the importance of such residues in the functional mechanism of the F protein, we performed a mutagenesis analysis on a 10-residue sequence in HRA of the SeV F protein. This portion of HRA was selected because it is highly conserved, it undergoes a dramatic structural transformation during protein refolding, and it forms a prominent cavity that is both the nucleation site for HRB binding and a target in antiviral drug design. Nine of the 15 conservative mutations that were characterized either reduced or eliminated the expression and fusogenic activity of the SeV F protein. In contrast, the other six mutations induced hyperactive membrane fusion activity. Interestingly, various mutations made at residues K180 and T181, nonconserved residues that were not in a or d positions, either disrupted protein folding or led to hyperactive membrane fusion activity, depending on the substituted amino acid residue. This novel finding shows that residues not expected to be important in the formation of the coiled-coil structure may have crucial roles in regulating functional activity. Overall, these results demonstrate the importance of the entire 10-residue sequence in the folding, processing, and activation of the F protein. These results are also consistent with the hypothesis that residues undergoing significant structural changes during F protein refolding are important in regulating the functional activity of the F protein.
The available structural and biochemical data are consistent with the following sequence of events occurring after the F protein is triggered by the receptor-binding protein (48, 64): first, the HRB coiled coil melts, pulling the transmembrane domains apart; second, the crumpled HRA structure extends into a coiled coil, propelling the fusion peptide into the target membrane; and finally, the HRB segments swing around the base of the head formed by DI and DII, thereby binding to the HRA coiled coil and forming the fusogenic hairpin structure. The hyperfusogenic SeV F mutations studied here could, therefore, affect any or all of these structural transitions. The direct effects of the HRA mutations on F protein conformational changes have not yet been investigated. However, the location of the mutations and the low-temperature activation are consistent with their predominant effects occurring during the second step, the springing of the HRA coil.
After cleavage, the first step in F protein refolding is thought to involve a conformational change in HRB (46). Moreover, a peptide derived from HRB interacts physically with a peptide derived from the stalk region of the HN receptor-binding protein (22). Mutations to residues 447 and 449 in HRB cause the PIV5 F protein to become activated at lower temperatures, to be more easily triggered in the absence of the HN protein, and to become inactivated after delayed incubation with target cells in a dye transfer assay (47). In the prefusion structure of the PIV5 F protein, residues 447 and 449 interact with the F protein head at the nucleation site of the HRB coiled coil (64). Therefore, the mutations in HRB appear to affect the first step in F protein refolding, i.e., the melting of the HRB coil. Although the HRA mutations studied here also cause the SeV F protein to become activated at lower temperatures, they do not result in membrane fusion activity in the absence of HN coexpression, and they do not lead to F protein inactivation after intracellular cleavage and delayed incubation with target cells in the dye transfer assay. Thus, the hyperfusogenic HRA mutations in the SeV F protein do not appear to enhance F protein activation until after the HN protein is bound to target cells. The data are also inconsistent with hyperfusogenic HRA mutations alleviating a potential inefficiency by the wild-type SeV F protein during late steps in membrane fusion. Neither wild-type F protein nor any of the mutants studied here have a hemifusion phenotype. The hyperfusogenic HRA mutations could perhaps stabilize the hairpin structure, thereby coupling more energy to membrane fusion. Future biophysical studies directly measuring the effects of the HRA mutations on the thermostability of the coiled-coil hairpin could rule out this possibility. However, previous studies on HR-derived peptides from the PIV5 F protein have shown that mutations to HRA and HRB residues generally do not increase coiled-coil stability (17, 47).
An L161M mutation in HRA of the PIV5 F protein causes normal expression and fusogenicity at 37°C but results in enhanced fusogenicity at 30°C (60). Residue L161 is located three amino acid residues upstream of the 10-residue sequence in the SeV F protein studied here. In the prefusion structure of the PIV5 F protein (64), the L161 residue is located in a ß-strand that may also include SeV F protein residues A178 and L179, if analogous residues in the F proteins of PIV5 and SeV both form similar ß-strands in their prefusion forms. These results suggest that destabilizing the ß-strands in the prefusion structure of HRA decreases the activation energy barrier of the F protein, thereby resulting in increased membrane fusion activity at lower temperatures. Conversely, stabilizing the ß-strands and turns in HRA may perhaps favor the prefusion structure and consequently increase the activation energy barrier. Just as residues in the HRA region of the paramyxovirus F protein play an important role in regulating protein folding and membrane fusion, residues in and around the helix-loop-helix structure in the HRA region of the influenza virus HA protein also regulate its activation (13). Moreover, proline mutations predicted to stabilize the spring-loaded conformation of the HA protein prevent its conformational rearrangement and functional activity (24, 43).
Membrane fusion inhibitors that block virus replication can be clinically effective. For example, an HRB-derived peptide from HIV gp41 (T-20, Enfuvirtide) has been successfully used in drug combination therapies against HIV infection (23). HRB-derived peptides and small-molecule mimetics are also potent inhibitors of F protein-mediated membrane fusion and virus replication (3, 11, 33, 39, 46, 62). HRB-derived peptides block membrane fusion by binding to the HRA coiled coil in a prehairpin intermediate of the F protein, consequently preventing formation of the fusogenic hairpin (47). A small molecule designed to bind a prominent cavity in the HRA coiled coil inhibits replication of respiratory syncytial virus (RSV) (11). A similar potential drug target exists in the HRA regions of the parainfluenza viruses, including hPIV1, whose residues are identical to those in SeV that were mutated in this study. Some conservative mutations to HRA drug target residues might allow for efficient virus replication, just as they allowed efficient membrane fusion after expression from plasmid DNA in the present study. If such is the case, then resistance mutations could potentially arise in the HRA region of the F protein, analogous to those in the HRA region of HIV gp41 resulting from T-20 treatment (2, 34, 44). Previous studies on a small-molecule inhibitor that binds to the HRA region of the RSV F protein have shown that a K394R mutation in DII of the F protein can also lead to resistance (10). Although the mechanism of resistance has not yet been determined, we believe that the mutation most likely alters the kinetics of accessibility of the HRA coiled coil in the prehairpin intermediate form of the F protein. Future studies characterizing the effects of F protein mutations on virus replication by Sendai virus, RSV, and other paramyxoviruses will be needed to explore the importance of HRA residues and other F protein residues in virus pathogenesis and susceptibility to fusion inhibitors.
Prefusion structures have been determined for two
class I viral fusion proteins, the influenza virus HA protein
(61) and the
paramyxovirus F protein
(64). In the HA protein,
HRA forms a helix-loop-helix structure. In the F protein, HRA forms an
11-segment structure consisting of -helices,
ß-strands, and turns. In both structures, the HRA region is in
a spring-loaded conformation, which helps position the fusion peptide
on the side of the molecule until the fusion protein becomes activated.
Just as the HRA residues in the SeV F protein studied here play an
important role in regulating expression and fusogenicity, residues in
and around the helix-loop-helix structure in the HA protein also
regulate its activation
(13). Based on
similarities between paramyxovirus F and influenza virus HA, class I
viral fusion proteins may, in general, be expected to regulate protein
expression and activation with HRA residues that are spring loaded in
the prefusion form. Mutations to conserved HRA residues in HIV gp41,
analogous to those in SeV F studied here, are accommodated in the
hairpin structure of gp41
(29). However, these same
mutations eliminate membrane fusion activity by gp41
(5), a finding that
suggests that the HRA mutations may compromise other conformations of
the protein. The importance of HRA residues in the membrane fusion
activities of HIV gp160
(5,
8,
15,
59), Ebola virus GP
(58), and severe acute
respiratory syndrome coronavirus S
(38) may perhaps suggest
that the HRA regions of class I viral fusion proteins may, in general,
undergo significant conformational changes during protein refolding.
However, determinations of the prefusion structures of these proteins
are needed to confirm this hypothesis and reveal whether their HRA
regions also adopt spring-loaded conformations analogous to
those found in the prefusion structures of paramyxovirus F
and influenza virus HA. Ultimately, further structural, biochemical,
and virological studies will be needed to understand better how
regulated conformational changes by class I viral fusion proteins
contribute to virus replication and pathogenesis.
|
ACKNOWLEDGMENTS |
|---|
This work was supported by a Cancer Center support grant (CA 21765), the American Lebanese Syrian Associated Charities (ALSAC), and the Children's Infection Defense Center (CIDC) at St. Jude Children's Research Hospital.
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FOOTNOTES |
|---|
Published ahead of print on 24 January 2007.
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REFERENCES |
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2
x 106 relative light units). Error bars represent
two standard deviations from triplicate experiments. The solid line
represents 100% Fwt expression. Asterisks indicate a significant
difference as measured by Student's t test (P
< 0.05). (B) Cell-cell fusion versus cell surface
expression. The open squares connected by a solid line
represent a titration of
increasing amounts
of Fwt DNA. The open circles connected by a dotted line represent a
titration of increasing amounts of F L179V DNA. The cell surface
expression levels of the F proteins were determined by flow cytometry
using the M16 monoclonal antibody. The data were normalized to 100%
fusion and 100% expression, corresponding to the transfection of 1
µg of pCAGGS Fwt DNA. (C) Enlargement of panel
B.
