Cell-Cell Fusion Induced by the Avian Reovirus Membrane Fusion Protein Is Regulated by Protein Degradation

The p10 fusion-associated small transmembrane protein of avianreovirus induces extensive syncytium formation in transfectedcells. Here we show that p10-induced cell-cell fusion is restrictedby rapid degradation of the majority of newly synthesized p10.The small ectodomain of p10 targets the protein for degradationfollowing p10 insertion into an early membrane compartment.Paradoxically, conservative amino acid substitutions in thep10 ectodomain hydrophobic patch that eliminate fusion activityalso increase p10 stability. The small amount of p10 that escapesintracellular degradation accumulates at the cell surface ina relatively stable form, where it mediates cell-cell fusionas a late-stage event in the virus replication cycle. The unusualrelationship between a nonstructural viral membrane fusion proteinand the replication cycle of a nonenveloped virus has apparentlycontributed to the evolution of a novel mechanism for restrictingthe extent of virus-induced cell-cell fusion.

INTRODUCTION

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Introduction

Membrane fusion is essential to a wide array of biological processes(21, 25, 54, 56). Since inappropriate fusion of membranes wouldhave severe adverse consequences, the process of membrane fusionmust by tightly regulated. For vesicle transport within cells,the proteins involved in mediating vesicle fusion are ubiquitouslyand stably expressed, and the timing and the specificity ofvesicle fusion are controlled by complex interactions amonga diversity of soluble and membrane-bound proteins (4, 34, 38).Conversely, extracellular fusion of cell membranes, such asthat which occurs during the formation of multinucleated myotubes,syncytiotrophoblasts, and osteoclasts (16, 50, 54), is regulatedin large part by the differentiation- or tissue-specific expressionof candidate cellular fusion proteins (24, 48, 35).

Enveloped virus entry into cells is also dependent on membranefusion mediated by multimeric membrane fusion proteins embeddedin the virus envelope (21, 55). The fusion activity of theseviral fusion proteins must be regulated within cells duringvirus assembly to prevent aberrant fusion of cell membranes.In addition, the activity of the multimeric fusion proteinsin the context of the mature, extracellular virus particle isregulated to ensure the correct timing and location of fusionof the virus envelope with a cell membrane during the entryprocess. Regulation is achieved through a series of complexconformational changes in the fusion proteins. Viral fusionproteins are frequently synthesized as precursors that mustbe proteolytically cleaved to allow assembly into fusion-incompetent,multimeric complexes during virus assembly (39). In addition,structural rearrangements of metastable, multimeric fusion proteinsinduced by environmental triggers, such as low endosomal pHand/or receptor binding, regulate fusion activity during virusentry (12, 19, 46, 49).

The fusogenic reoviruses are one of the few groups of nonenvelopedviruses capable of inducing cell-cell fusion and polykaryonformation (10). The nonenveloped avian reovirus (ARV) and NelsonBay reovirus (NBV) encode homologous 10-kDa proteins (p10) thatinduce syncytium formation when expressed in transfected orvirus-infected cells (41). The p10 fusion-associated small transmembrane(FAST) protein is the smallest known viral membrane fusion proteinand contains a central transmembrane (TM) domain that separatessmall, approximately equally sized (about 40 residues) ectodomainsand endodomains (41). Since nonenveloped viruses lack a lipidmembrane and do not enter cells through membrane fusion, p10is not a component of virus particles; therefore, unlike thatof enveloped viruses, the p10 membrane fusion protein has norole in reovirus entry. Rather, p10 is expressed in virus-infectedcells, where it is trafficked to the cell surface in a typeI topology to mediate cell-cell fusion and the induction ofsyncytium formation, a relatively late-stage event in the virusreplication cycle (11, 41). In addition to providing directcell-cell transmission of the virus, syncytium formation contributesto a more rapid lytic response and a more rapid rate of virusrelease, possibly due to an inherent membrane-destabilizingactivity of p10 (2, 11).

In view of the toxic nature of p10-induced syncytium formation(i.e., the membrane integrity of syncytial cells is compromised),it seems clear that the rate and the extent of syncytium formationmust be restricted in order to allow sufficient time for progenyvirus assembly. However, p10 expression is not temporally regulatedin virus-infected cells and follows the same general time courseas the expression of other reovirus proteins (42). Furthermore,p10 is a promiscuous fusogen (i.e., it induces cell-cell fusionin a wide range of cell types) and functions at neutral pH;therefore, there is no evidence for the activation of p10 fusionactivity by a specific trigger, such as low pH or receptor binding.Here we show that p10-induced syncytium formation is restrictedby the rapid degradation of p10, thereby delaying the stableaccumulation of p10 at the cell surface in quantities sufficientto initiate and propagate polykaryon formation. It would appearthat the unusual role of p10 in the replication cycle of a nonenvelopedvirus has contributed to a novel means of restricting cell-cellfusion.

MATERIALS AND METHODS

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Materials and Methods

Cells. Continuous quail cell line QM5 was cultured as previously described(9). Mouse hybridoma cells producing 12CA5 monoclonal antibodiesto hemagglutinin (HA) were grown in RPMI 1640 containing 10%fetal bovine serum and penicillin-streptomycin.

Materials. The anti-HA immunoglobulin G2b kappa-chain [IgG2b( ${kappa}$ )] 12CA5 antibodieswere produced in-house from mouse hybridoma cells, and the cellsupernatant (approximately 0.12 µg of anti-HA immunoglobulin/µl)was used either directly for surface staining or following concentration(to approximately 1.6 µg/µl) by precipitation with35% ammonium sulfate. Fluorescein isothiocyanate (FITC)-conjugatedgoat anti-rabbit IgG antibodies and protein G-agarose were obtainedfrom Life Technologies Inc. Streptavidin-conjugated Texas red,biotin-conjugated rat anti-mouse IgG, and mouse anticalnexinantibodies were purchased from Sigma-Aldrich, St. Louis, Mo.A rabbit polyclonal anti-p10 antiserum specific for residues19 to 34 of p10 was a kind gift from Richard Mulligan (HarvardUniversity). A rabbit polyclonal antiserum raised against theC-terminal endodomain was previously described (41).

Cloning and transfection. The eukaryotic expression vector pcDNA3 (Invitrogen) was usedfor the expression of p10 and its derivatives. The constructionof the amino-terminal double-HA-tagged p10 expression construct(p10-2HAN) and the generation of all p10 substitutions by three-primerPCR were previously described (41). The p10 ectodomain construct(p10e) was amplified by a single "touchdown" PCR with a forwardprimer corresponding to the N terminus of HA-tagged p10. Theprimer contained the sequence encoding the signal peptide ofinfluenza virus HA with a signal peptidase cleavage site (MLTIIALSYIFCLALGQ).The sequences of all constructs were confirmed. Plasmids weretransfected into cells by using Lipofectamine (Life Technologies)as previously described (41).

Immunofluorescence staining. For double labeling of p10-transfected cells with anti-p10 antiserumand anti-calnexin antibodies, cells were fixed with methanoland preblocked with normal goat serum and whole rat IgG beforebeing incubated with rabbit anti-p10 antiserum (1:400) and mouseanti-calnexin antibodies (1:1,000). Bound antibodies were visualizedby using FITC-conjugated goat anti-rabbit antibodies (1:400)and biotin-conjugated rat anti-mouse IgG (1:1,000) plus streptavidin-conjugatedTexas red (1:2,000). Stained cells were washed extensively andvisualized by using a Zeiss LSM510 scanning argon laser confocalmicroscope and a x100 objective.

To examine the stability of surface-localized p10, cells transfectedwith p10-2HAN were incubated on ice for 30 min at 36 h posttransfectionto slow cell metabolism and then were incubated for 30 min onice with a 1:20 dilution of concentrated anti-HA antibodies.Cells were washed extensively with cold Hanks' balanced saltsolution and then incubated at 37°C in complete medium 199for various times (0, 45, or 90 min). Cells were reacted withFITC-labeled anti-mouse antibodies on ice for 30 min to detectthe anti-HA antibodies that remained on the surface of cells.Cells were washed as described above and fixed with 4% paraformaldehydeprior to visualization and photography with appropriate filtersets. Low levels of surface-localized p10 confounded the quantificationof p10 surface fluorescence by fluorescence-activated cell sortinganalysis. Therefore, p10 stability at the cell surface was qualitativelyassessed by fluorescence microscopy by capturing fluorescenceimages over time under identical capture parameters.

Immunoprecipitation. Radiolabeled cells were lysed in 1x final lysis (radioimmunoprecipitation[RIPA]) buffer (50 mM Tris-HCl [pH 7.5], 150 mM NaCl, 1% NonidetP-40, 0.5% sodium deoxycholate, 1 µg of aprotinin/ml,1 µg of leupeptin/ml, 1 µg of pepstatin/ml). Forimmunoprecipitation of cell lysates under harsh lysis conditions(6), 2x final lysis buffer containing 0.1% sodium dodecyl sulfate(SDS) was used. For every 2 x 10⁶ cells, 4 µl of concentratedanti-HA antibodies was preincubated with 15 µl of proteinG-agarose for 1 h with shaking at room temperature. Antibody-proteinG-agarose complexes were washed once with 1x lysis buffer andincubated with the cell lysates for 1 h at room temperature;these steps were followed by two stringent washes with eachof the following: 1x lysis buffer, high-salt buffer (50 mM Tris-HCl[pH 7.5], 500 mM NaCl, 0.1% Nonidet P-40, 0.05% sodium deoxycholate),and low-salt buffer (50 mM Tris-HCl [pH 7.5], 0.1% Nonidet P-40,0.05% sodium deoxycholate). Immune complexes were analyzed bySDS-polyacrylamide gel electrophoresis (PAGE) and fluorographywith 15% acrylamide gels as previously described (8).

Pulse-chase analysis. Transfected cells were prelabeled for 30 min with methionine-freeDulbecco minimal essential medium at 30 h posttransfection,pulse-labeled for 15 min with the same medium containing 75µCi of [³⁵S]methionine/ml, and chased for different timeswith complete medium 199 containing 10% fetal bovine serum.Samples were analyzed by immunoprecipitation, SDS-PAGE, andfluorography. Captured images were scanned, and pixels correspondingto p10 were quantified by using Scion software to determinethe levels of p10 after various chase times relative to thosein the pulse-labeled p10 sample. For assessment of the effectsof proteasome or lysosome inhibitors, E64C, chloroquine, methylamine,and MG132 were added at final concentrations of 200 µM,0.5 mM, 15 mM, and 100 µM, respectively, to all incubations.For assessment of the stability of p10 proteins resident inmembranes, cells were pulse-chased as described above and thenfractionated into membrane and soluble fractions. The membranefraction was stripped of peripheral membrane proteins by treatmentwith sodium carbonate as previously described (41) prior tosolubilization with 1x final lysis buffer and immunoprecipitation.

RESULTS

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Results

The ectodomain targets p10 for rapid and extensive degradation. Nonsyncytial constructs of p10 were previously created by site-specificsubstitutions of the ectodomains or the endodomains (41). Whiledetermining whether the loss of fusion activity displayed bythese modified p10 constructs reflected increased p10 instability,we made the surprising discovery that functional, HA-taggedp10 was rapidly degraded inside cells. Pulse-chase analysisdemonstrated that the vast majority of newly synthesized p10was degraded within 30 min of synthesis (Fig. 1A). Sigma C,a soluble structural protein of ARV, remained stable under identicalconditions. The levels of p10 over time showed similar patternsunder both mild and stringent solubilization conditions (Fig.1A), suggesting that the loss of p10 reflected degradation andnot aggresome formation (26). Rapid p10 degradation was notdue to overexpression, since the levels of p10 expression attainedin transfected cells were comparable to the levels attainedin virus-infected cells (41). Since both N- and C-terminallytagged p10 constructs were functional and displayed the samedegradation profiles (data not shown), rapid degradation wasnot attributable to the epitope tag but appeared to be an inherentfeature of p10.

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FIG. 1. The ectodomain targets p10 for rapid degradation. (A) (Left) Transfected cells expressing full-length p10-2HAN were pulse-labeled and chased for the indicated times in minutes. Radiolabeled cells were solubilized in RIPA buffer or double-strength RIPA buffer, and p10 was immunoprecipitated with anti-p10 antiserum ( ${alpha}$ p10), resolved by SDS-PAGE, and visualized by fluorography. (Right) Same analysis with a control protein, the soluble sigma C protein of avian reovirus. (B) Full-length p10-2HAN (p10), p10ecto^– (ecto^–), or p10e, with two N-terminal HA epitope tags (HA) and the cleavable HA signal peptide (SP), were expressed in transfected cells. The stability of the various p10 constructs was assessed by pulse-chase analysis as described for panel A. The nature of the various constructs is diagrammed at the bottom.

The p10 protein is an integral membrane protein and residesexclusively in the membrane fraction of virus-infected and p10-transfectedcells (41, 43, 44). Although p10 lacks an N-terminal signalpeptide for membrane insertion, it utilizes a central TM domainas an internal signal-anchor sequence to direct an N exoplasmic/Ccytoplasmic membrane topology (41). Accordingly, deletions inthe p10 ectodomain do not alter p10 localization to the membranefraction of cells (41). However, deletion of the majority ofthe p10 ectodomain (p10ecto^–) stabilized the protein,eliminating the rapid degradation profile observed for the full-lengthprotein (Fig. 1B). The p10ecto^– protein still residedexclusively in the membrane fraction of cells (data not shown).These results suggested that the ectodomain contributes to p10instability directly rather than indirectly by altering p10membrane localization. To determine whether the ectodomain alonewas responsible for the short half-life of p10, we expressedthe soluble HA-tagged p10e construct in transfected cells. Sincep10e lacked the natural p10 signal-anchor sequence (i.e., theTM domain), a cleavable signal sequence of influenza virus HAwas added to the N terminus of p10e for translocation of theectodomain into the lumen of the endoplasmic reticulum (ER),where it normally resides. Pulse-chase analysis revealed thatthe soluble, and presumably luminal, p10 ectodomain displayedthe same rapid degradation profile as authentic p10 (Fig. 1B),further indicating that determinants in the p10 ectodomain targetthe protein for rapid degradation.

Hydrophobic and thiol residues in the ectodomain influence p10 degradation. Shmulevitz and Duncan previously identified in the ectodomainof p10 a small, moderately hydrophobic patch (HP) that is flankedby two conserved cysteine residues (residues 9 to 24) (Fig.2A) and that is important for p10-induced polykaryon formation(41). Since exposed hydrophobic or thiol residues can serveas recognition signals for the quality control degradation machinery(14, 15, 57), we examined the stability of p10 constructs containingpoint substitutions in the p10 HP (Fig. 2A).

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FIG. 2. Residues in the p10 HP influence p10 degradation. (A) Sequence and schematic representation of the p10 ectodomain. Numbers refer to amino acid residues. The locations of the HP, the CR in the ARV and NBV p10 proteins, and the start of the downstream TM are indicated. The locations of various site-specific substitutions in the ARV p10-2HAN construct are indicated. Constructs are named by using the single-letter amino acid code to indicate the identity of the authentic amino acid, its position, and the identity of the substitution. A/S indicates a substitution with either Ala or Ser. (B) Degradation rates for authentic p10-2HAN (p10) and constructs containing single amino acid substitutions, assessed by pulse-chase analysis (chase times are indicated in minutes) as described in the legend to Fig. 1. (C) Time course analysis of p10 degradation performed as described for panel B but with more precise time points (in minutes) to compare authentic p10-2HAN (p10) to two p10 constructs containing single residue substitutions; one of these constructs retains fusion activity (T13M), and one is nonfusogenic (V15M). Numbers below the gel lanes indicate the percentages of p10 detected after various chase times relative to that in the pulse-labeled sample with no chase, as determined by image analysis of the fluorogram. (D) Degradation rates for p10-2HAN (p10) and constructs containing single amino acid substitutions in the endodomain dicysteine (C63/64A) or polybasic (K67M) motifs or in the ectodomain conserved region (L32A), assessed as described for panel B.

Previous results indicated that both conserved ectodomain cysteineresidues are essential for p10 fusion activity (41). Replacementof either cysteine residue with alanine or serine (C9A/S orC21A/S) eliminated both p10 fusion activity and rapid p10 degradation(Fig. 2A and B). To examine possible hydrophobic effects, bothvaline residues in the p10 HP were individually replaced withmethionine residues (other than tryptophan, with its much bulkierside group, methionine introduced the smallest incremental decreasein the hydrophobicity of the p10 HP, from 0.285 to 0.256) (13).In spite of the subtle change in hydrophobicity and the relativelyminor alteration in side group volumes imparted by these aminoacid changes, the V15M and V19M substitutions had the same effectas the cysteine substitutions, eliminating p10-induced syncytiumformation and rapid p10 degradation (Fig. 2A and B). The nonfusogenicproperty and increased stability of the p10 proteins containingthe cysteine and valine substitutions did not reflect alteredp10 membrane interactions, since all constructs assumed thecorrect surface-localized N exoplasmic/C cytoplasmic membranetopology (44).

A more detailed quantitative analysis indicated that less than30% of authentic HA-tagged p10 remained 30 min after synthesis,and only 5 to 10% remained following a 60-min chase, yieldinga p10 half-life of approximately 20 to 25 min (Fig. 2C). Longerchases indicated that the residual 5 to 10% of p10 that escapedrapid degradation persisted for an additional 2 h or more (Fig.2D). In contrast, decreased degradation of the V15M constructresulted in approximately 70% of the protein persisting formore than 1 to 2 h (Fig. 2C). A functional, fusogenic threonine-methioninesubstitution construct, T13M, was also examined to probe thepotential influence of altered hydrophobicity versus alteredside chain volume on the degradation properties of the p10 HP.The degradation profile of T13M was generally the same as thatof authentic p10, with the possible exception of a slightlymore rapid initial rate of p10 degradation (Fig. 2C). Therefore,methionine substitutions of either of the two aliphatic residuesin the p10 HP eliminated p10 degradation.

The ability of substitutions to alter p10 stability was restrictedto the HP. As expected from the results obtained with the p10ecto^–and p10e constructs, which suggested that the ectodomain aloneis responsible for p10 degradation, alterations in the dicysteinemotif (C63/64A) or the conserved basic region (K67M) presentin the p10 endodomain had no effect on p10 stability (Fig. 2D).On the membrane-proximal side of the p10 HP resides a nine-residueregion that is completely conserved between the ARV and theNBV p10 proteins (Fig. 2A, CR [conserved region]) The importanceof this relatively polar region for p10 function has not beendetermined. A substitution (L32A) that significantly decreasedthe hydrophobicity of the CR (from 0.146 to 0.093) also decreasedthe rate of p10 degradation (Fig. 2D), although this effectwas far less pronounced than that observed with the more subtlealterations in the aliphatic residues in the p10 HP. Therefore,changes in the structure and/or relative hydrophobicity of thep10 ectodomain have profound effects on p10 degradation.

Degradation of p10 occurs following p10 insertion into an early membrane compartment. Proteins can be targeted for degradation by misfolding in thecytoplasm, by failure to properly translocate across the ERmembrane, or following successful ER translocation (14, 15,57). To determine which of these pathways might contribute top10 degradation, confocal microscopy was used to examine thep10 subcellular distribution. Immunofluorescence staining withanti-p10 antisera revealed a punctate, reticular staining pattern(Fig. 3) similar to that observed when cells were stained withantisera specific for calnexin, an ER membrane-localized chaperone(51). An overlay of the two images revealed significant colocalizationof p10 and calnexin (Fig. 3), indicating that the majority ofp10 localizes, at least transiently, in the ER. Along with previousresults indicating that p10-induced syncytium formation is inhibitedby the vesicle transport inhibitor brefeldin A (11) and thatp10 resides exclusively as an integral membrane protein (41),the colocalization data suggested that p10 is targeted to theER membrane and trafficked through the secretory pathway tothe cell surface.

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FIG. 3. p10 localizes to the ER. (Left) Cells transfected with a p10 expression plasmid were fixed and permeabilized, and the subcellular distribution of p10 was examined by confocal microscopy with anti-p10 polyclonal antiserum ( ${alpha}$ p10) and FITC-conjugated secondary antibody. (Middle) A similar analysis was performed with anticalnexin antibody ( ${alpha}$ calnexin) and Texas red-conjugated secondary antibody to reveal the distribution of an ER-localized protein. (Right) Merge of the p10 and calnexin panels, showing extensive colocalization of the two proteins. Scale bars, 10 µm.

To determine whether p10 degradation reflected inefficient signal-anchor-mediatedinsertion of p10 into membranes, the stability of membrane-associatedp10 was assessed by pulse-chase analysis. Following the pulseor each chase period, the membrane fraction was isolated andtreated with high-pH carbonate buffer to removal peripheralmembrane proteins. Subsequent gel analysis therefore would monitorthe fate and relative stability of p10 molecules already presentas integral membrane proteins. The p10 proteins resident inthe membrane displayed the characteristic rapid degradationprofile (Fig. 4A), indicating that p10 is degraded followinginsertion into the membrane.

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FIG. 4. Degradation of p10 occurs following membrane insertion and is altered by a proteasome inhibitor. (A) Pulse-chase analysis for the indicated times (in minutes) was performed with p10-transfected cells as described in the legend to Fig. 1. After the pulse or each chase, the membrane fraction was isolated, and the presence of membrane-associated p10 was determined by immunoprecipitation, SDS-PAGE, and fluorography. (B) Pulse-chase analysis for the indicated times (in minutes) was performed with p10-transfected cells as described in the legend to Fig. 1, except that cells were incubated during the pulse-chase in the presence of inhibitors of lysosomal proteases (E64C) or lysosome acidification (chloroquine [chlor] or methylamine [MA]) or a proteasome inhibitor (MG132).

Protein degradation following successful membrane translocationcan occur through trafficking to the lysosome, retrieval fromthe ER, and targeting to the proteasome or through retrievalfrom the plasma membrane and targeting to the lysosome or proteasome(22, 47). The membrane-permeable proteasome inhibitor MG132(31) significantly reduced the rate of p10 degradation (Fig.4B). Since peptide aldehyde inhibitors of the proteasome mayalso inhibit certain lysosomal proteases (31, 37), we examinedthe effects of inhibitors of lysosome acidification (chloroquineand methylamine) or lysosomal proteases (E64C) on p10 degradation.None of these lysosome inhibitors reversed the rapid degradationof p10 (Fig. 4B), suggesting that p10 degradation might be proteasomemediated.

To determine whether p10 is retrieved for degradation beforeor after transport to the plasma membrane, the stability ofsurface-localized HA-tagged p10 was assessed. Relatively lowlevels of p10 surface expression confounded attempts to quantifyp10 surface expression by flow cytometry (data not shown). Therefore,p10 surface expression was qualitatively assessed by immunofluorescencemicroscopy. HA-specific antiserum was added to live p10-transfectedcells, which then were incubated for various times under optimalcell growth conditions prior to fixation as described in Materialsand Methods. Fluorescence images were captured under identicalparameters such that the extent of fluorescence was a relativeindicator of p10 surface expression. In contrast to the extremelyrapid degradation observed for membrane-associated p10 (Fig.4A), the quantities of surface-localized, antibody-bound p10remained relatively constant over the 90-min chase period followingantibody binding (Fig. 5). Consequently, p10 appears to be relativelystable once expressed at the cell surface. In conjunction withthe fact that p10 degradation begins within 5 min of its synthesis(Fig. 2C), these results were consistent with p10 degradationoccurring following retrieval from an early membrane compartment.

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FIG. 5. Surface-localized p10 is relatively stable. The stabilities of surface-localized p10-2HAN (p10) and the V19M construct were assessed by surface staining of live transfected cells with anti-HA monoclonal antibody. Following antibody addition, the presence of surface-bound antibody specifically attached to the HA-tagged, surface-localized p10 ectodomain was monitored over time (indicated in minutes on the right). Fluorescence images were captured under identical parameters for comparison of the intensities of fluorescence.

Increased p10 stability leads to increased polykaryon formation. It seemed likely that the degradation of p10 would serve torestrict the rate of syncytium formation. If this hypothesiswere correct, then increasing p10 stability should lead to anincreased rate of syncytium formation. All of the substitutionsthat prolonged the p10 half-life also eliminated the fusogenicactivity of the protein (Fig. 2A). We therefore used a proteasomeinhibitor to decrease the extent of p10 degradation and monitoredthe effects on the rate and extent of cell-cell fusion overtime by microscopic examination of Giemsa-stained monolayers(Fig. 6). The total number of nuclei present in syncytia inrandom microscopic fields was increased by approximately 70%following 6 h of treatment with the inhibitor (Fig. 6A). Thiseffect was manifested by a modest increase in both the numberof syncytial foci (Fig. 6B) and the relative size of individualfoci (Fig. 6C). The correlation between decreased p10 degradationand increased syncytium formation suggested that rapid and extensivedegradation of p10 might represent a significant factor regulatingp10-induced cell-cell fusion.

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FIG. 6. Inhibition of proteasome activity leads to enhanced p10-mediated syncytium formation. Cell monolayers were transfected with a p10 expression plasmid. At 15 h posttransfection, as syncytia were beginning to appear, the culture medium was removed and replaced with medium lacking or containing the proteasome inhibitor MG132. Treated monolayers were incubated for a further 4 to 6 h prior to Giemsa staining to reveal syncytial foci. Microscopic examination was used to quantify the average number of syncytial nuclei per field at 6 h posttreatment (A) or the average number of syncytia per field at 4 and 6 h posttreatment (B). Results are presented as the mean and standard deviation of triplicate samples from a representative experiment. The numbers and sizes of the syncytia present in Giemsa-stained monolayers at 6 h posttreatment were captured by light microscopy at a magnification of x194 (C).

DISCUSSION

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Discussion

Extensive degradation restricts p10-induced syncytium formation. Previous studies indicated that the time course of p10 expressionin ARV-infected cells follows a pattern similar to that of otherreovirus proteins, suggesting little, if any, temporal controlof syncytium formation (11, 41). However, syncytium formationis regulated, in part, by inefficient translation of p10 froma suboptimal translation start site (30, 42). Here we show thatthe majority of p10 is rapidly degraded shortly after synthesisand before reaching the plasma membrane. The net effect of extensivep10 degradation would be a delay in the rate of p10-inducedsyncytium formation. We assume that the small percentage ofp10 that escapes degradation following membrane insertion successivelytransits the ER-Golgi apparatus transport pathway to slowlyaccumulate in a relatively stable state at the plasma membraneuntil some minimal threshold level is exceeded and cell-cellfusion ensues. Although protein degradation influences the activitiesof numerous cellular regulatory proteins (5, 27), as far aswe are aware, this is the only example in which the cellulardegradation machinery is exploited to regulate cell-cell fusioninduced by a membrane fusion protein.

p10 degradation occurs following membrane insertion. The ER-associated degradation (ERAD) pathway is an essentialcomponent of the cellular quality control machinery involvedin monitoring protein folding and maturation (14, 15, 23, 32,47, 57). Our present results suggest the possible involvementof the ERAD pathway in mediating p10 degradation. This speculationis supported by the following evidence: (i) degradation of p10follows insertion of the protein into the membrane fractionof cells (Fig. 4A); (ii) colocalization studies indicated thatp10 associates, at least transiently, with the ER (Fig. 3);(iii) targeting of the soluble p10 ectodomain to the lumen ofthe ER by using a cleavable HA signal sequence resulted in rapiddegradation of this ectodomain construct (Fig. 1); (iv) pulse-chaseanalysis indicated that p10 degradation begins within 5 minof its synthesis (Fig. 2C); and (v) p10 molecules in the plasmamembrane are relatively stable (Fig. 5). All of these resultsimply that p10 degradation involves an early membrane compartment,most likely the ER. According to this model, p10 degradationwould reflect retrieval of p10 from the ER membrane and targetingto the proteasome, a hypothesis consistent with the abilityof a proteasome inhibitor to decrease p10 degradation (Fig.4B) and increase p10-induced syncytium formation (Fig. 6). Sincethe soluble, and presumably luminal, p10e construct was alsorapidly degraded, retrieval of this polypeptide from the ERlumen would have to occur, possibly by retrotranslocation, assuggested for other luminal proteins degraded by the ERAD pathway(47). Additional studies are required to conclusively determinethe roles, if any, of ER retrieval, ubiquitination, and theproteasome in p10 degradation.

Ectodomain signals in p10 target the protein for degradation. Although many of the steps in the ERAD pathway have been elucidated,the specific signals that target proteins for rapid degradationare less clear. Two such signals are exposed hydrophobic orthiol residues, which may serve as general indicators of aberrantprotein folding or failure to achieve a final native structure(15). Exposed hydrophobic or thiol residues are recognized byER-resident molecular chaperones or protein disulfide isomerase(3, 17, 18, 20, 36). Interactions with these luminal ER proteinsinvolved in facilitating protein folding can lead to proteinretention in the ER and subsequent degradation.

Mutational analysis clearly revealed that the ectodomain ofp10 serves as a signal for rapid degradation, somewhat analogousto a degron (53), specifically imparting a short half-life tothe protein. The basis for this activity is still unclear, althoughthere was a correlation between p10 degradation and the relativehydrophobicity of the ectodomain, primarily in the HP and toa lesser extent in the adjacent CR. For example, slight decreasesin the relative hydrophobicity of the HP (e.g., V15M and V19M)decreased p10 degradation, while increasing hydrophobicity (T13M)had no such effect on preventing p10 degradation and, if anything,resulted in a slightly more rapid initial rate of p10 degradation.Since the methionine substitutions could exert their influencethrough means other than changes in hydrophobicity, more extensivemutagenic and structural analyses are required to more clearlydefine the roles of specific residues in p10 degradation.

The two ectodomain cysteine residues that are essential forp10 fusion activity are also important determinants of the p10half-life. Similar to the situation with the valine substitutions,substitution of either C9 or C21 eliminates p10 fusion activity(41) while, as we have now shown, simultaneously extending thep10 half-life. These cysteines do not participate in the formationof an intermolecular disulfide bond, since p10 exhibits thesame relative gel mobilities under reducing and nonreducingconditions (44). The cysteines may, however, contribute to theformation of a disulfide-stabilized loop structure through theformation of an intramolecular disulfide bond. If the p10 HPdoes exist as a cysteine noose structure, then substitutionof either cysteine residue would be expected to significantlyalter the structure of the p10 ectodomain, thereby affectingprotein degradation.

How does the ectodomain contribute to p10 degradation? Generally, mutations causing a loss of the native structurewould be expected to increase degradation by the cellular qualitycontrol machinery (14, 15, 57). Paradoxically, ectodomain mutationsthat eliminate p10 fusion activity lead to a decrease in p10degradation. What, then, might be the basis for p10 degradationand the effects of ectodomain substitutions on this process?Since misfolded proteins are targeted for degradation, the majorityof p10 may fail to achieve the native structure and be targetedfor degradation. More than 50% of proteins carrying N-terminaldegrons are degraded cotranslationally, suggesting that nascentprotein folding may occur in kinetic competition with cotranslationaldegradation pathways (52). This is not the case with p10, sincefull-length p10 is synthesized and inserted into the ER membraneprior to degradation, indicating that p10 degradation is a posttranslationalevent.

Inefficient formation of the predicted intramolecular disulfidebond is one conceivable cause of posttranslational misfoldingof p10 (Fig. 7, model 1). Disulfide bond formation is knownto be a slow process (40). Delayed formation of the predictedp10 cysteine noose structure would lead to exposed thiols andpossibly hydrophobic residues in the nonnative structure, targetingp10 for ER retention and subsequent export to the cytoplasmfor degradation (17). Inhibition of the proteasome would providesufficient time for increased formation of the predicted cysteinenoose and native p10 structure, a prediction in accord withthe observed enhanced syncytium formation in the presence ofproteasome inhibitors (Fig. 6). According to the slow disulfidebond formation model, the V15M and V19M substitutions shouldpromote more efficient disulfide bond formation, thereby decreasingp10 degradation. Hydrophobic forces driving the valine residuesto the interior of the loop structure could interfere with localresidue packing and constrain disulfide bond formation (18).The less hydrophobic methionine residue more easily orientstoward the solvent phase, a conformation which might allow tighterpacking of the loop structure, increased disulfide bond formation,and decreased p10 degradation. However, the stabilizing effectof the cysteine substitutions appears to be at odds with thismodel, since the removal of one cysteine would guarantee thepresence of a free thiol group. Possible explanations for thisanomaly include sequestration of the free thiol within the nonnativestructure or removal of the thiol by aberrant intermoleculardisulfide bond formation between p10 molecules or with luminalER proteins. However, such aberrant disulfide bond formationgenerally leads to increased protein degradation (7, 33, 36),which is not the case with the p10 cysteine substitutions.

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FIG. 7. Models of p10 degradation. Three possible models to explain the influence of the p10 ectodomain and the HP on p10 degradation by the ERAD pathway are shown. The first model implies that the slow formation of an intramolecular disulfide bond (SS) leads to exposed thiol (SH) or hydrophobic residues (filled circles) in the nonnative structure which target p10 for degradation. The second model implies that it is not the rate of folding of the p10 ectodomain but rather a low thermodynamic stability of the native structure that leads to transient and repeated exposure of thiol or hydrophobic residues. The third model implies that hydrophobic residues are naturally surface exposed in the native p10 ectodomain structure, leading to recognition and degradation of the majority of p10.

A thermodynamic model of p10 degradation predicts that the p10ectodomain may have a low free-energy barrier between nativeand nonnative conformations, with structural fluctuations inthe p10 HP leading to reversible disulfide bond formation orexposure of hydrophobic residues and p10 degradation (Fig. 7,model 2). This model of protein degradation was derived fromstudies of bovine pancreatic trypsin inhibitor and single-chainT-cell receptor (28, 29, 45), where thermodynamic stabilityrather than the rate of protein folding correlates with proteinsecretion efficiency. The small size of the p10 ectodomain suggeststhat there may be minimal differences in free energy betweenthe native and the nonnative states. In this scenario, the V15Mand V19M substitutions would either increase the stability ofthe "native" ectodomain structure (while paradoxically eliminatingthe fusion activity of p10) or decrease the hydrophobic signalthat targets p10 for degradation. Once again, one would haveto explain how the cysteine substitutions promote a more stable"native" structure and decrease p10 degradation while eliminatingp10-induced syncytium formation.

A third possibility is that native p10, rather than a misfoldedpopulation of p10, is targeted for degradation. While the exposureof hydrophobic residues may serve as a general indicator ofaberrant protein folding, the relative surface hydrophobicityof the native protein correlates with the protein half-lifeand may be necessary for ubiquitination (1, 3). In this model,the stable native structure of the p10 ectodomain would containsolvent-exposed hydrophobic residues leading to recognitionand degradation of the majority of p10 by the ERAD pathway (Fig.7, model 3). The lack of complete fidelity of the quality controlmachinery or the extent of p10 surface hydrophobicity wouldallow a small percentage of p10 to escape degradation and slowlyaccumulate at the cell surface to induce cell-cell fusion. Methioninesubstitutions would reduce the surface hydrophobicity to belowthe level required for recognition by the cellular quality controlmachinery, while cysteine substitutions, by removing the predictedloop structure, could allow exposed hydrophobic residues tomore easily bury themselves within a more compact or aggregatednonnative structure. As with the other models, how the freethiol created by a single cysteine substitution escapes recognitionis not clear.

Further studies of p10 degradation and a structural determinationof the p10 ectodomain are required to distinguish among theseand other possible models of the basis for the ectodomain-mediateddegradation of p10 and the relationship, if any, between p10degradation and the mechanism of p10-induced membrane fusion.

Why would p10 evolve to be rapidly degraded? The ARV p10 protein contributes to a loss of membrane integritylate in the virus replication cycle and is therefore toxic tocells (2). The p10 ectodomain may have evolved purely as a meansto limit the rate of accumulation of p10, thereby delaying syncytiumformation and membrane leakiness to allow sufficient time forprogeny virus assembly.

A more intriguing possibility to explain the degradation propertiesof the p10 HP was derived from recent studies indicating thatsynthetic peptides based on the HP induce lipid mixing in liposomefusion assays (44). These results suggested that the p10 HPmay function as a fusion peptide, analogous to the fusion peptidespresent in enveloped virus fusion proteins. Reversible or constitutivesolvent exposure of hydrophobic residues may be a requirementfor the fusion peptide activity of the p10 HP. Therefore, hydrophobiceffects may serve a dual function in p10: as an essential featureof the membrane fusion reaction and as a means to delay syncytiumformation by targeting the majority of p10 for degradation.The inherent or readily reversible exposure of hydrophobic residuesin the predicted p10 fusion peptide is not observed with theenveloped virus fusion peptides that are stably sequesteredwithin the tertiary structure of the multimeric fusion proteinuntil triggered conformational changes lead to their exposure.Differences in the structure and solvent exposure propertiesof the enveloped and nonenveloped viral fusion peptides mayreflect significant differences in the mechanisms by which thesetwo groups of fusion proteins regulate and promote the fusionof biological membranes.

ACKNOWLEDGMENTS

We thank Jingyun Shou for expert technical assistance and RichardMulligan (Harvard University) for the donation of anti-p10 antiserum.

This research was funded by a grant from the Canadian Institutesof Health Research (CIHR). R.D. is the recipient of a CIHR-RPPinvestigator award. M.S. was funded by scholarships from theNatural Sciences and Engineering Research Council of Canadaand from the Killam Foundation. J.S. was funded by a scholarshipfrom Cancer Care Nova Scotia.

FOOTNOTES

* Corresponding author. Mailing address: Department of Microbiology and Immunology, Faculty of Medicine, Dalhousie University, Halifax, Nova Scotia, Canada B3H 4H7. Phone: (902) 494-6770. Fax: (902) 494-5125. E-mail: roy.duncan{at}dal.ca.

REFERENCES

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