Efficient Reovirus- and Measles Virus-Mediated Pore

Efficient Reovirus- and Measles Virus-Mediated Pore Expansion during Syncytium Formation Is Dependent on Annexin A1 and Intracellular Calcium Department of Microbiology & Immunology,a Department of Biochemistry & Molecular Biology,b and Department of Pediatrics,c Dalhousie University, Halifax, Nova Scotia, Canada

ABSTRACT

Orthoreovirus fusion-associated small transmembrane (FAST) proteins are dedicated cell-cell fusogens responsible for multinucleated syncytium formation and are virulence determinants of the fusogenic reoviruses. While numerous studies on the FAST proteins and enveloped-virus fusogens have delineated steps involved in membrane fusion and pore formation, little is known about the mechanics of pore expansion needed for syncytiogenesis. We now report that RNA interference (RNAi) knockdown of annexin A1 (AX1) expression dramatically reduced both reptilian reovirus p14 and measles virus F and H protein-mediated pore expansion during syncytiogenesis but had no effect on pore formation. A similar effect was obtained by chelating intracellular calcium, which dramatically decreased syncytiogenesis in the absence of detectable effects on p14-induced pore formation. Coimmunoprecipitation revealed calcium-dependent interaction between AX1 and p14 or measles virus F and H proteins, and fluorescence resonance energy transfer (FRET) demonstrated calcium-dependent p14-AX1 interactions in cellulo. Furthermore, antibody inhibition of extracellular AX1 had no effect on p14-induced syncytium formation but did impair cell-cell fusion mediated by the endogenous muscle cell fusion machinery in C2C12 mouse myoblasts. AX1 can therefore exert diverse, fusogen-specific effects on cell-cell fusion, functioning as an extracellular mediator of differentiation-dependent membrane fusion or as an intracellular promoter of postfusion pore expansion and syncytium formation following virus-mediated cell-cell fusion. IMPORTANCE

Numerous enveloped viruses and nonenveloped fusogenic orthoreoviruses encode membrane fusion proteins that induce syncytium formation, which has been linked to viral pathogenicity. Considerable insights into the mechanisms of membrane fusion have been obtained, but processes that drive postfusion expansion of fusion pores to generate syncytia are poorly understood. This study identifies intracellular calcium and annexin A1 (AX1) as key factors required for efficient pore expansion during syncytium formation mediated by the reptilian reovirus p14 and measles virus F and H fusion protein complexes. Involvement of intracellular AX1 in syncytiogenesis directly correlates with a requirement for intracellular calcium in p14-AX1 interactions and pore expansion but not membrane fusion and pore formation. This is the first demonstration that intracellular AX1 is involved in pore expansion, which suggests that the AX1 pathway may be a common host cell response needed to resolve virus-induced cell-cell fusion pores.

V

irus-induced syncytium formation is a common cytopathic effect associated with replication of numerous enveloped viruses in the families Paramyxoviridae, Herpesviridae, and Retroviridae. Syncytiogenesis is caused by viral membrane fusion proteins that evolved to mediate virus-cell membrane fusion during virus entry; when trafficked to the plasma membrane of virusinfected cells, these fusogens can promote cell-cell membrane fusion (1, 2). The specific functional role of syncytium formation in virus replication is unclear although generation of polykaryons is linked to viral pathogenesis (3–6). Additionally, pseudotyping oncolytic viruses with viral fusogens greatly increases viral spread and cancer cell killing (7, 8). The molecular mechanisms responsible for virus-induced syncytium formation are still unclear. Protein-mediated cell-cell membrane fusion followed by syncytium formation is a multistage process. The prefusion stage involves cell-cell attachment and close membrane apposition. In the case of enveloped-virus-induced syncytium formation, the prefusion stage is mediated by receptor-binding components of the fusion protein complex and formation of a trimeric hairpin structure that may promote close membrane apposition (2). During

June 2014 Volume 88 Number 11

physiological cell-cell fusion events, as during muscle myoblast fusion into multinucleated myotubes, prefusion events are not mediated by the fusogen per se but by several protein partners involved in cell recognition and adhesion (9). The membrane fusion stage of syncytium formation involves sequential steps of hemifusion (i.e., merger of proximal lipid leaflets), pore formation, and pore expansion to form a stable micropore (10). For the well-described enveloped-virus fusogens, this process is believed to be driven by energy released from dramatic conformational rearrangements of their large, complex, multimeric ectodomains (11). During the postfusion stage, expansion of stable micropores

Received 15 January 2014 Accepted 11 March 2014 Published ahead of print 19 March 2014 Editor: S. López Address correspondence to Roy Duncan, [email protected]. Copyright © 2014, American Society for Microbiology. All Rights Reserved. doi:10.1128/JVI.00121-14

Journal of Virology

p. 6137– 6147

jvi.asm.org

6137

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

Marta Ciechonska,a Tim Key,a Roy Duncana,b,c

Ciechonska et al.

6138

jvi.asm.org

expansion stage of syncytium formation. We also show that this process is not virus specific as AX1 also interacts with the measles F and H proteins and is necessary for efficient pore expansion mediated by this enveloped-virus fusion complex. This is the first demonstration that intracellular AX1 is involved in pore expansion during syncytium formation, which suggests that the AX1 pathway may be a common host cell response to resolve virusinduced cell-cell fusion pores. MATERIALS AND METHODS Cells and antibodies. Quail muscle fibroblast (QM5) and Vero cells were maintained in medium 199, as previously described (31). C2C12 mouse myoblast, HT1080, and HEK cells were grown in Dulbecco’s modified Eagle’s medium (DMEM) containing 10% fetal bovine serum (FBS) and 25 mM HEPES. C2C12 cells were differentiated using DMEM supplemented with 5% horse serum. Polyclonal rabbit anti-p14 and anti-p14 ectodomain (residues 5 to 31) antisera were described previously (30, 31). Monoclonal AX1 antibodies were used for immunoprecipitation and Western blotting (Santa Cruz Biotechnology) and in antibody blocking experiments (Abcam). Monoclonal anti-FLAG (Invitrogen), polyclonal anti-protein disulfide isomerase (PDI; Abcam), polyclonal anti-green fluorescent protein (GFP; Clonetech), and horseradish peroxidase (HRP)conjugated anti-rabbit and anti-mouse secondary antibodies (Santa Cruz Biotechnologies) were from the suppliers indicated here. Polyclonal rabbit anti-F and anti-H antibodies were provided by Chris Richardson (Dalhousie University). Polyclonal rabbit anti-HER2 antibody was obtained from Graham Dellaire (Dalhousie University). Plasmid cDNAs. Plasmid constructs expressing p14 or p14G2A were previously described (31). RRV p14 was tagged at the C terminus with mCherry. AX1 cDNA was prepared from a PC3 human prostate cancer cell RNA library, cloned into pcDNA3, and N-terminally tagged with a triple FLAG epitope or C-terminally tagged with enhanced GFP (EGFP). Plasmids expressing measles virus Edmonston strain F or H proteins were obtained from Chris Richardson (Dalhousie University), and the HER2expressing plasmid was obtained from Graham Dellaire (Dalhousie University). Transfections and syncytial indexing. Subconfluent cell monolayers in 12-well plates were transfected with 1 ␮g of DNA and 1.5 ␮l of Lipofectamine LTX (Invitrogen) (HT1080 cells) or 3 ␮l of Lipofectamine (Invitrogen) (QM5 cells) or polyethyleneimine (PEI) (HEK cells) per well. C2C12 cell monolayers in 48-well plates were transfected with 0.25 ␮g of DNA and 0.75 ␮l of Lipofectamine LTX in Opti-MEM medium (Invitrogen). Cells were fixed at various times posttransfection, as indicated in the figure legends, and WrightGiemsa stained, and the average number of syncytial nuclei per microscopic field was quantified as previously described (31). AX1 On-TargetPlus SMARTpool small interfering RNA (siRNA) and nontargeting control siRNA sequences were obtained from Dharmacon. For siRNA transfection, HT1080 cells in 12-well plates were transfected with 25 nM siRNA and 2.5 ␮l of DharmaFECT1 in Opti-MEM and then transfected with plasmid DNA at 24 to 36 h post-siRNA transfection. FRET. HT1080 cells on coverslips were cotransfected with EGFPtagged AX1 and mCherry-tagged p14, fixed with paraformaldehyde, and imaged at ⫻100 using a Zeiss LSM 510 Meta confocal microscope. The PixFRET ImageJ plug-in was used to calculate donor and acceptor spectral bleed-through (SBT) values and normalized fluorescence resonance energy transfer (NFRET) levels in each pixel. Donor and acceptor SBT values were determined by acquiring two images (the FRET and the donor/acceptor images) from cells expressing only EGFP or mCherry, and donor and acceptor SBT ratios were modeled using exponential relationships with fluorophore intensity after exclusion of aberrant background values at low intensities and the application of a Gaussian blur. To determine NFRET values for the p14-annexin interaction, three images were acquired from 10 cells in duplicate experiments: sensitized emission FRET (donor excitation and acceptor emission), donor (donor excitation and donor emission), and acceptor (acceptor excitation and acceptor emis-

Journal of Virology

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

into lumen-sized macropores large enough to accommodate migration of nuclei results in syncytium formation. Relatively little is known about the mechanism of this postfusion, pore expansion stage of syncytium formation or the players involved. While enveloped-virus fusogens evolved to mediate virus-cell fusion and virus entry, the reovirus fusion-associated small transmembrane (FAST) proteins are nonstructural viral proteins that evolved specifically to induce cell-cell, rather than virus-cell, membrane fusion (12). The FAST proteins and syncytium formation are virulence determinants of the fusogenic reoviruses (13, 14), and syncytiogenesis promotes localized cell-cell virus transmission, increased cytopathic effects, and enhanced progeny virus release in cell culture (15, 16). Members of the FAST protein family differ markedly from enveloped-virus fusogens in their size and distribution across membranes. At 95 to 198 residues in size, the FAST proteins are the smallest known viral fusogens. They assume a bitopic, Nexoplasmic/Ccytoplasmic topology in membranes, positioning very small (⬃20 to 40 residues) fusion peptide-containing domains external to the plasma membrane (17–20) and equalsized or considerably larger (⬃36 to 141 residues) domains in the cytoplasm (21). The ecto-, endo-, and transmembrane domains all function as fusion modules and play an active role in the membrane fusion process (22, 23). The mechanism of action of these unique viral fusogens also differs in several respects from enveloped-virus fusogens. The FAST protein ectodomains lack receptor binding activity and do not form trimeric hairpins (24), suggesting that they have little, if any, role in mediating prefusion cell attachment and membrane apposition. As with myoblast fusion, FAST proteins rely on separate adhesion factors to mediate the prefusion stage of syncytium formation, using cadherins to mediate cell attachment and actin remodeling to promote close membrane apposition (24). The rudimentary size of the FAST protein ectodomains is also incompatible with a membrane fusion reaction based on energy released from dramatic ectodomain structural remodeling, and the mechanism of membrane merger does not display the same sensitivity to membrane curvature agents as enveloped-virus fusogens (25). Lastly, the FAST protein endodomains are essential for cell-cell fusion, while the generally short endodomains of enveloped-virus fusogens are frequently dispensable or inhibit syncytium formation (26–29). The disproportionate size of their endodomains suggests that FAST proteins may be more reliant on interactions on the cytosolic side of the plasma membrane than envelopedvirus fusogens. A recent study revealed that the soluble endodomain of the reptilian reovirus (RRV) p14 FAST protein promotes syncytium formation mediated by FAST proteins, by enveloped-virus fusogens, and by the unidentified cellular fusogen(s) responsible for muscle cell fusion (30). The promiscuous nature of enhanced pore expansion mediated by the p14 endodomain suggests the involvement of a common cellular pathway involved in converting micropores into the macropores needed for syncytium formation. RRV p14 is the most robust fusogen in the FAST protein family (16) and has a 36-residue, myristoylated, N-terminal ectodomain and a 68-residue C-terminal endodomain (31). To identify cellular partners of the p14 endodomain, we analyzed the interaction profile of the p14 endodomain in a yeast two-hybrid screen and identified cellular annexin A1 (AX1) as a potential interaction candidate. We now show that intracellular AX1 interacts with p14 in a Ca2⫹-dependent manner and promotes the postfusion, pore

Annexin A1 and Syncytium Formation

June 2014 Volume 88 Number 11

treated with a 1:10 (100 ␮g/ml), 1:25 (40 ␮g/ml), or 1:50 (20 ␮g/ml) dilution of anti-AX1 or anti-PDI (control) antibody in growth medium. Cells were fixed at 12 to 14 h posttransfection, and syncytium formation was quantified as described above. Antibody inhibition of differentiating C2C12 cells was as described previously (32). Briefly, at 24 h postseeding, cells were induced to differentiate using DMEM containing 5% horse serum, medium was supplemented with 100 ␮g/ml of anti-AX1 or antiPDI antibody at 51 h postdifferentiation (prior to the appearance of myotubes), and cells were incubated for an additional 16 h before syncytium formation was quantified as described above.

RESULTS

AX1 is necessary for efficient RRV p14-mediated syncytium formation. To investigate whether annexins exert any effect on p14mediated syncytiogenesis, plasmid DNA expressing p14 was transfected into HT1080 cells previously treated with control or AX1 siRNA. Monolayers of cells containing the control siRNA fused at an appreciable rate, as indicated by the presence of multinucleated syncytia observed in Giemsa-stained monolayers, while treatment of cells with AX1 siRNA substantially impaired syncytium formation (Fig. 1A). Quantifying the number of syncytial nuclei per microscopic field revealed that AX1 siRNA treatment decreased p14-mediated syncytium formation by ⬃93% (Fig. 1B). Western blotting confirmed that AX1 expression was reduced by the pooled siRNAs (Fig. 1B, inset). Since AX1 has been implicated in membrane protein trafficking and endocytosis (33, 34), plasma membrane localization of a nonfusogenic p14G2A construct that displays normal surface localization levels (31) was quantified by flow cytometry following immunostaining of nonpermeabilized cells using anti-p14 antiserum. FACS analysis revealed no observable difference in p14 plasma membrane localization between cells expressing AX1 siRNA and cells expressing nontargeting siRNA (Fig. 1C). AX1 is therefore a positive effector of p14-induced syncytium formation. RRV p14 interacts with AX1 in cellulo. Fluorescence resonance energy transfer (FRET) was employed to determine whether p14 interacts with AX1 under physiologically relevant conditions (i.e., inside vertebrate cells). This approach detects in vivo protein-protein interactions that occur over distances of ⬍5 to 10 nm (35) and was applied to HT1080 cells cotransfected with C-terminally EGFPtagged p14 and N-terminally mCherry-tagged AX1. Donor and acceptor spectral bleed-through (SBT) values and normalized FRET (NFRET) intensities were calculated using the PixFRET ImageJ plug-in (36), and mean NFRET (mNFRET) values were determined from best-fit Gaussian distributions using images acquired from 10 cells in each of two separate experiments. Positive controls were cells cotransfected with EGFP-tagged- and mCherry-tagged-p14 constructs (Fig. 2A, top row) since coimmunoprecipitation indicated that p14 forms homomultimers (37). The negative controls included cells cotransfected with soluble EGFP and mCherry-tagged p14, which indicated no interaction between the fluorophores (Fig. 2A, bottom row), and cells cotransfected with soluble mCherry and EGFP-tagged AX1 that displayed a similar lack of FRET (data not shown). FRET between p14 and AX1 was clearly detected, as shown in the fluorescence images (Fig. 2A, middle row), and NFRET quantification indicated that p14-AX1 FRET intensity was ⬃50% of that obtained during p14 homomultimerization (Fig. 2B). Both yeast two-hybrid results and FRET analysis therefore indicated that p14 interacts with AX1. RRV p14 interaction with AX1 and syncytium formation is calcium dependent. To further confirm p14 interaction with

jvi.asm.org 6139

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

sion) images. Background subtraction and Gaussian blur of the donor, acceptor, and FRET channels were performed on each image. FRET signals were normalized to the square root of the product of the donor and acceptor fluorescence intensities to control for variations in fluorophore expression levels between cells and provided a quantification of FRET intensity comparable between different samples. Pixel amplitude distributions of the eight-bit NFRET images generated by the PixFRET software were summarized as histograms using a bin width of 0.03906 NFRET units; histograms were fit to four Gaussian distributions, and the distribution with the highest calculated R2 value was used to determine mean NFRET (mNFRET) values and average pixel amplitudes from each condition. Homotypic pore formation. Donor HT1080 cells (15% confluent) in six-well plates were cotransfected with plasmids expressing EGFP or p14 and incubated for 4 h or cotransfected with EGFP and measles F and H proteins and incubated for 6 h. Confluent T-175 flasks of HT1080 target cells were labeled with 12.5 ␮g/ml CellTrace calcein red-orange acetoxymethyl (AM) (Invitrogen) for 30 min at 37°C according to manufacturer’s specifications, incubated for 1 to 4 h at 37°C in growth medium, resuspended with trypsin, overseeded on the subconfluent donor cells, and cocultured until small syncytia (4 to 6 nuclei) appeared, ⬃ 6 h in the case of p14 and ⬃10 h in the case of measles F and H. Cells were resuspended with trypsin and fixed with 7.4% formaldehyde, and 10,000 cells were analyzed by flow cytometry (FACSCalibur; Becton, Dickinson). The percent cofluorescent cells signifying transfer of soluble contents was determined using Cell Quest and FCS Express, version 3, software from triplicate samples (n ⫽ 3 experiments). Intracellular Ca2ⴙ chelation. Transfected cells were treated with 20 to 100 ␮M BAPTA-AM [1,2-bis(o-aminophenoxy)ethane-N,N,N=,N=-tetraacetic acid acetoxymethyl ester] (Invitrogen) in Hanks’ balanced salt solution (HBSS) for 30 min at 37°C, washed with HBSS, and incubated in growth medium for 4 h (p14) or 20 h (measles virus F and H proteins), and syncytium formation was quantified as described above. For FRET analysis, BAPTA AM-treated HT1080 cells were fixed and processed at 24 h posttransfection. For pore formation, donor and target cells were separately treated with BAPTA-AM before being cocultured for 5 h and processed by fluorescence-activated cell sorter (FACS) analysis, as described above. Surface expression. HT1080 cells sequentially transfected with AX1 siRNA and p14G2A were incubated for 24 h at 37°C, washed and incubated in cold HBSS containing 1% bovine serum albumin (BSA; blocking buffer) for 30 min at 4°C, and then incubated with a 1:1,000 dilution of anti-p14-ectodomain antibody diluted in blocking buffer for 1 h at 4°C. Cells were extensively washed and then incubated with Alexa 647-conjugated goat anti-rabbit secondary antibody for 1 h at 4°C, resuspended with 10 mM EDTA in phosphate-buffered saline (PBS), and fixed with 7.4% formaldehyde; surface expression was then quantified by flow cytometry, as described above, for triplicate samples (n ⫽ 3 experiments). Coimmunoprecipitation. HEK cells in 10-cm dishes cotransfected with FLAG-tagged AX1 and p14 or measles H and F plasmid constructs were harvested by scraping at 12 to 16 h posttransfection, pelleted by centrifugation, resuspended in 1 ml of lysis buffer (20 mM Tris-HCl, 137 mM NaCl, 10% glycerol, 1% NP-40, protease inhibitor cocktail [SigmaAldrich]) supplemented with 0 to 5 mM CaCl2 or MgCl2, lysed by 10 passages through a 30-gauge needle, and incubated at 4°C for 30 min with agitation. Lysates were cleared by centrifugation at 16,000 ⫻ g at 4°C, and supernatants were incubated overnight with 2 ␮l of anti-FLAG antibody at 4°C. A 10-␮l aliquot of a 50% slurry of washed Dynabeads (Invitrogen) in lysis buffer was added to each sample and incubated at 4°C with agitation for 30 min. Beads were extensively washed with cold lysis buffer and transferred to new tubes, and immunoprecipitates were eluted with 60 ␮l of boiling protein sample buffer and analyzed by SDS-PAGE and Western blotting using the antibodies indicated in the figure legends. AX1 antibody inhibition of syncytium formation. HT1080 or C2C12 cells in 48-well plates were transfected with p14 and at 2 h posttransfection

Ciechonska et al.

AX1, cell lysates coexpressing full-length p14 and N-terminally FLAG-tagged AX1 were immunoprecipitated with anti-FLAG antibody followed by Western blotting using anti-p14 antiserum. As shown in Fig. 3A, p14 coprecipitated with FLAG-tagged AX1. Anti-FLAG antibody did not precipitate p14 in the absence of FLAG-tagged AX1, and AX1 did not coprecipitate the integral membrane protein HER2 (Fig. 3A), indicating specificity of the p14-AX1 coprecipitation. Since AX1 is Ca2⫹ regulated, coimmunoprecipitations were repeated in the presence of a range of Ca2⫹ concentrations. Decreasing Ca2⫹ concentrations from 100 ␮M to 25 ␮M led to corresponding decreases in p14 coprecipitation, with no detectible p14-AX1 interaction at Ca2⫹ concentrations of 10 ␮M or less (Fig. 3A). Similar experiments using a range of Mg2⫹ concentrations indicated only low levels of p14-AX1 interaction and only in the presence of 1 mM Mg2⫹ (Fig. 3B), indicating that

6140

jvi.asm.org

Journal of Virology

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

FIG 1 AX1 is necessary for efficient p14-mediated syncytium formation. (A) HT1080 cells were transfected with control (left) or AX1 (right) siRNA, cotransfected with p14 plasmid DNA at 36 h post-siRNA transfection, fixed at 12 h post-DNA transfection, and Giemsa stained to visualize syncytia. (B) HT1080 cells transfected with AX1 or control (ctrl) were cotransfected with p14 plasmid DNA and fixed and stained as described for panel A, and the extent of syncytium formation was quantified by determining the average number of syncytial nuclei per field. Results are mean ⫾ standard deviation of a representative experiment in triplicate (n ⫽ 3). AX1 siRNA knockdown was verified by SDS-PAGE and immunoblotting with monoclonal anti-annexin A1 antibody (inset). (C) HT1080 cells transfected with AX1 siRNA and then cotransfected with plasmid DNA expressing fusion-incompetent p14G2A were surface stained at 24 h post-DNA transfection with p14 anti-ectodomain antiserum and Alexa 647-conjugated secondary antibody. Cells were fixed and analyzed for cell surface fluorescence by flow cytometry. au, arbitrary units; ctrl, control.

p14 interaction with AX1 is Ca2⫹ specific. Intracellular Ca2⫹ concentrations are maintained at ⬃200 nM under normal physiological conditions but increase to low-micromolar concentrations following triggered release of intracellular Ca2⫹ stores (38). Thus, p14-AX1 interactions occur at calcium levels anticipated in cells with activated calcium signaling pathways. To examine the Ca2⫹ dependence of p14-AX1 interaction in the more physiologically relevant context of a whole cell, intracellular Ca2⫹ was sequestered using BAPTA-AM, a membrane-permeable Ca2⫹ chelator widely used in studies of intracellular Ca2⫹ release and neurotransmitter signaling (39, 40). Repeating the FRET assay in the presence of 0 to 40 ␮M BAPTA-AM revealed a dose-dependent decrease in FRET signal with increasing concentrations of BAPTA-AM, with near complete abolition of signal at 40 ␮M (Fig. 4). Coimmunoprecipitation and FRET results therefore indicate that RRV p14 interacts with AX1 in a Ca2⫹-dependent manner, both in vitro and in cellulo. To determine whether inhibiting p14-AX1 interactions by depleting intracellular Ca2⫹ had a corresponding inhibitory effect on p14-induced syncytium formation, p14-transfected QM5 cells were treated with a range of BAPTA-AM concentrations. Syncytiogenesis was reduced by ⬃60% following treatment with 20 ␮M BAPTA-AM and by ⬃80% in the presence of 30 to 40 ␮M BAPTA-AM (Fig. 5A), concentrations that nearly eliminated p14AX1 FRET interactions (Fig. 4). Moreover, the requirement for intracellular Ca2⫹ during p14-mediated syncytium formation was not cell type specific, as similar results were obtained using p14transfected HT1080 cells (Fig. 5B). AX1 interactions with p14 and p14-induced syncytium formation are therefore both dependent on intracellular Ca2⫹. AX1 interacts with measles F and H proteins and is necessary for efficient F protein-mediated syncytiogenesis. To determine whether AX1 might also play a role in syncytium formation mediated by other viral fusogens, we used HT1080 cells cotransfected with measles virus F and H proteins. The F and H proteins of measles virus form heterodimers, and both are required for fusion activity; H is responsible for receptor binding and activation of F while F mediates membrane fusion (41). The measles virus F-H fusion complex was chosen since these proteins contain slightly longer cytoplasmic tails than most enveloped-virus fusogens (33 residues for F and 34 residues for H) and since they promote fusion at neutral pH (42), two attributes shared with FAST proteins. HT1080 cells were sequentially transfected with nontargeting control or AX1 siRNA followed by cotransfection with measles virus F and H proteins. Decreasing AX1 expression had an obvious inhibitory effect on F protein-mediated syncytium formation (Fig. 6A). When quantified, AX1 knockdown inhibited F proteinmediated syncytiogenesis by ⬃50% at multiple time points throughout the course of syncytium formation (Fig. 6B). Presumably, residual levels of AX1 following siRNA (Fig. 7D) are sufficient to allow gradual syncytium formation over the extended 22-h time course of F protein-mediated syncytiogenesis. Furthermore, chelation of intracellular Ca2⫹ using 50 to 100 ␮M BAPTA-AM decreased F-mediated syncytium formation by ⬃60% (Fig. 6C). Coimmunoprecipitation assays also indicated that both F and H coprecipitated with AX1 when F and H were expressed either together or individually (Fig. 6D), indicating that F and H can independently interact with AX1. F or H was not precipitated by anti-FLAG antibody in the absence of FLAGtagged AX1 (Fig. 6D), implying that the coprecipitations were

Annexin A1 and Syncytium Formation

specific. Thus, AX1 and intracellular Ca2⫹ are important for syncytiogenesis mediated by diverse viral fusogens. AX1 is necessary for efficient pore expansion during syncytium formation mediated by RRV p14 and measles virus F and H proteins. To determine which stage of syncytiogenesis is dependent on AX1, a fluorescent, dual-color, homotypic pore formation assay was employed. Sparsely seeded HT1080 cells were transfected with control or AX1 siRNA and 36 h later cotransfected with plasmids expressing EGFP and empty vector or p14-expressing vector; the EGFP plasmid served as a surrogate marker for p14-expressing cells. Target HT1080 cells labeled with the soluble

cytoplasmic fluor calcein red-orange AM were overseeded on donor cell monolayers at 6 h after plasmid transfection, and donor and target cells were cocultured for another 6 h. Cells expressing EGFP and p14 that also contained calcein, indicative of pore formation between donor and target cells, were quantified by flow cytometry. As shown in Fig. 7A, p14 induced robust pore formation in the presence of both control and AX1 siRNAs. When pore formation was quantified over repeat experiments, there was no detectable difference in the extent of pore formation between cells treated with control or AX1 siRNA (Fig. 7B). Similarly, knockdown of AX1 had no effect on the extent of membrane fusion and

FIG 3 AX1 and p14 coimmunoprecipitate at physiological Ca2⫹ concentrations. (A) HEK cells cotransfected with FLAG-tagged AX1 and p14 were lysed

at 14 h posttransfection in buffer containing the following Ca2⫹ concentrations: 5 mM (lanes 1, 2, and 10 to 12), 100 ␮M (lane 3), 50 ␮M (lane 4), 25 ␮M (lane 5), 10 ␮M (lane 6), 1 ␮M (lane 7), 0.5 ␮M (lane 8), and 0 ␮M (lane 9). Lysates were immunoprecipitated (IP) with anti-FLAG antibody and analyzed by Western blotting (WB) using anti-p14 antiserum (lanes 1 to 9) or anti-HER2 antiserum (lanes 10 to 12) (top). Lysates without immunoprecipitation were immunoblotted using anti-FLAG (AX1) or anti-p14 antibodies (bottom). (B) The experiment is as described for panel A, except that cells were lysed in buffer containing the indicated concentrations of Mg2⫹ or Ca2⫹: Mg2⫹ at 5 mM (lanes 1 and 2), 1 mM (lane 4), 100 ␮M (lane 5), 25 ␮M (lane 6), 10 ␮M (lane 7), and 0 ␮M (lane 8) and Ca2⫹ at 100 ␮M (lane 9), 50 ␮M (lane 10), and 25 ␮M (lane 11). Lysates were immunoprecipitated and analyzed by Western blotting as described for panel A.

June 2014 Volume 88 Number 11

jvi.asm.org 6141

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

FIG 2 FRET analysis of AX1-interactions in cellulo. HT1080 cells were cotransfected with EGFP-tagged AX1 and mCherry-tagged p14 and fixed at 10 h posttransfection. (A) Representative images of sensitized emission FRET, showing the donor channel, acceptor channel, and the calculated normalized FRET (NFRET) image. Control images were obtained from cells expressing p14 linked to EGFP and mCherry (top row) as a positive FRET control for a known multimeric protein or expressing free EGFP and mCherry-tagged p14 (bottom row) as a negative FRET control. Images acquired from cells coexpressing EGFP-tagged AX1 and mCherry-tagged p14 were used to detect AX1-p14 interactions. The NFRET range is denoted by color gradations. Scale bar, 10 ␮m. (B) Fitted Gaussian distributions of 20 calculated NFRET images from two separate experiments were used to calculate the mNFRET (top) from cells transfected or cotransfected with the indicated fluorescent probes, as described for panel A. The box highlights the standard deviation of mNFRETs, the plus sign (⫹) is the mean mNFRET, the line is the median mNFRET, and the whiskers indicate minimum and maximum mNFRETs. The fitted NFRET distributions were also used to calculate the mean pixel amplitude (bottom) from each image. Error bars represent standard errors propagated within and across experiments.

Ciechonska et al.

FIG 5 Chelation of intracellular calcium inhibits p14-mediated syncytium formation. (A) QM5 cells transfected with p14 were treated with increasing concentrations of BAPTA-AM at 4 h posttransfection, fixed, and Giemsa stained at 8 h posttransfection to visualize syncytia. (B) The experiment is as described for panel A except that the extent of syncytium formation was quantified by determining the average number of syncytial nuclei per field. Results are mean ⫾ standard deviation of a representative experiment in triplicate (n ⫽ 2). (C) The experiment is as described in panel B except that HT1080 cells were used. Results are mean ⫾ standard deviation of a representative experiment in triplicate (n ⫽ 3).

6142

jvi.asm.org

pore formation mediated by the measles virus fusion complex (Fig. 7C). In addition, treatment of cell monolayers with 20 or 40 ␮M BAPTA-AM after target cells were overseeded on donor cells had no effect on p14-induced pore formation (Fig. 7E). Since both AX1 siRNA and BAPTA-AM profoundly inhibited p14-induced and measles virus F-H-induced syncytium formation but had no effect on pore formation, AX1 and intracellular Ca2⫹ function downstream of membrane fusion and pore formation to promote efficient pore expansion leading to syncytiogenesis. Antibody inhibition of extracellular AX1 does not inhibit RRV p14-mediated syncytium formation. To determine whether extracellular AX1 plays any role in p14-mediated cellcell fusion, extracellular AX1 was inhibited using anti-AX1 antibody, as recently reported in C2C12 mouse myoblasts (32). Blocking antibody was added to p14-transfected HT1080 cells at 2 h posttransfection, i.e., before the onset of syncytium formation (which first appears at ⬃8 h posttransfection). As previously reported (32), anti-AX1 blocking antibody inhibited myotube formation by ⬃50 to 60% following induction of differentiation-dependent myoblast fusion (Fig. 8A). The same concentration of antibody had no effect on p14-induced syncytiogenesis, and increasing concentrations of antibody were similarly ineffective (Fig. 8B). To ensure that the different effects of anti-AX1 antibody on p14 fusion of HT1080 versus differentiation-dependent fusion of C2C12 mouse myoblasts was not due to cell type, p14-mediated fusion of C2C12 cells was examined in the absence and presence of blocking antibody. As in HT1080 cells, p14-mediated C2C12 syncytium formation was insensitive to treatment of cells with extracellular

Journal of Virology

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

FIG 4 Chelation of intracellular Ca2⫹ abolishes AX1-p14 FRET interactions. (A) HT1080 cells cotransfected with EGFP-tagged AX1 and mCherry-tagged p14 were treated with 0, 20, or 40 ␮M BAPTA-AM and processed for FRET analysis as described in the legend of Fig. 2. The NFRET range is denoted by color gradations. Scale bar, 10 ␮m. (B) Fitted Gaussian distributions of 20 calculated NFRET images from two separate experiments were used to calculate the mNFRET values (top) and mean pixel amplitudes (bottom) from 20 cells cotransfected with the indicated fluorescent probes as described in the legend of Fig. 2. In the top panel, the box highlights the standard deviation of mNFRETs, the plus sign (⫹) is the mean mNFRET, the line is the median mNFRET, and the whiskers indicate minimum and maximum mNFRETs. Error bars in the bottom panel represent standard errors propagated within and across experiments.

Annexin A1 and Syncytium Formation

FIG 6 AX1 and intracellular Ca2⫹ are necessary for efficient measles virus F and H protein-mediated syncytium formation. (A) HT1080 cells transfected with control (ctrl) or AX1 siRNA were cotransfected with measles virus F and H plasmid DNA at 36 h post-siRNA transfection, fixed at 22 h post-DNA transfection, and Giemsa stained to visualize syncytia. (B) HT1080 cells transfected with control or AX1 siRNA and cotransfected with measles virus F and H plasmid DNA as described for panel A were fixed and stained at the indicated times post-DNA transfection, and the average number of syncytial nuclei per field was quantified. Results are the mean ⫾ standard deviation of a representative experiment in triplicate (n ⫽ 3). (C) HT1080 cells cotransfected with measles virus F and H proteins were treated with 0, 50, or 100 ␮M BAPTA-AM at 4 h posttransfection and fixed at 24 h posttransfection, and the extent of syncytium formation was quantified as described for panel B. Results are the mean ⫾ standard deviation of a representative experiment in triplicate (n ⫽ 3). (D) HEK cells cotransfected with FLAG-tagged AX1 and measles virus F and/or H proteins were lysed in buffer supplemented with 5 mM Ca2⫹; lysates were immunoprecipitated with anti-FLAG antibody, and immunoprecipitates were analyzed by Western blotting using polyclonal anti-F or anti-H antibody. Lysates were immunoblotted using anti-FLAG (AX1), anti-F, or anti-H antibody.

anti-AX1 antibody (Fig. 8C). Thus, while extracellular AX1 contributes to myotube formation, it is not involved in RRV p14-mediated syncytiogenesis. DISCUSSION

The disproportionately long FAST protein cytoplasmic tails and their predicted intrinsic disorder (21) suggested that these endodomains might interact with and recruit intracellular fusion cofactors. A yeast two-hybrid approach, using p14 endodomain as bait, yielded AX1 as a genetic p14 interaction partner. AX1-p14 interactions were verified in vitro by coimmunoprecipitation and in cellulo by FRET analysis, and these interactions were sensitive to physiological intracellular Ca2⫹ concentrations. Knockdown of AX1 expression and chelation of intracellular Ca2⫹ substantially inhibited p14-mediated syncytio-

June 2014 Volume 88 Number 11

formation. (A) A homotypic, dual-fluorescence pore formation assay was used to quantify pore formation. HT1080 cells transfected with control or AX1 siRNA were cotransfected at 36 h post-siRNA transfection with EGFP plasmid DNA and either empty vector or p14 plasmid DNA. At 4 h post-DNA transfection, donor cells were overlaid with target HT1080 cells labeled with CellTrace calcein red-orange AM; cells were fixed at 8 h postaddition of target cells, and transfer of fluorescent markers was quantified by flow cytometry. The percentage of EGFP-positive cells that acquired the aqueous calcein red-orange fluor in arbitrary fluorescence units (indicated above the horizontal gating line) was quantified, and values are shown relative to the forward scatter in arbitrary units. Results are for 10,000 cells from a representative experiment. (B) As in panel A, fluorescence intensity was quantified as mean ⫾ standard error for triplicate samples from n ⫽ 3 experiments. vec, vector. (C) The experiment is as described in panel B, except that measles virus F and H proteins were substituted for RRV p14. Cells cotransfected with EGFP and F and H were incubated for 6 h and subsequently cocultured with calcium-labeled target cells for 10 h prior to fixation. (D) AX1 siRNA knockdown in F- and H-expressing cells was verified by SDS-PAGE and immunoblotting with monoclonal anti-AX1 antibody at 40 h post-siRNA transfection. (E) The experiment is as described in panel B, using donor and target cells treated with the indicated concentrations of BAPTA-AM at 4 h posttransfection. Results are mean ⫾ standard deviation of a representative experiment (n ⫽ 2).

genesis with no adverse effects on pore formation, and p14-induced syncytiogenesis was not impaired by treatment with extracellular anti-AX1 blocking antibody. Taken together, these results identify intracellular annexins as Ca2⫹-dependent positive effectors of fusion pore expansion and syncytiogenesis. Notably, a similar involvement of AX1 in cell-cell fusion mediated by the measles virus fusion protein complex suggests the existence of a generic, AX1-dependent cellular pore expansion pathway applicable to diverse viral fusogens. The present study provides the first direct evidence for host factor interactions with a FAST protein. While FAST proteins can function as autonomous fusogens to induce the merger of artificial lipid bilayers (43), their rudimentary structure suggested that cellular cofactors might be involved in the more complex process of syncytiogenesis. Among such cofactors are cadherins. While there is no evidence of a physical interaction between p14 and cadherins, p14 does colocalize at adherens junctions and exploits these adhesive structures to mediate the prefusion, cell attachment

jvi.asm.org 6143

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

FIG 7 AX1 is not required for p14- or measles virus F- and H-induced pore

Ciechonska et al.

stage of syncytium formation (24). We now identify annexin as a p14 fusion cofactor involved in postfusion pore expansion. Results indicate that p14 physically associates with AX1 in a Ca2⫹dependent manner both in vitro and in cellulo. Since FRET requires the two fluorophores be separated by ⬍5 to 10 nm, a distance reflective of direct protein-protein interactions (44), p14 appears to directly interact with AX1. Furthermore, chelating intracellular Ca2⫹ inhibited both p14-AX1 interactions and p14induced syncytium formation, suggesting that p14 association with AX1 is biologically relevant. AX1 and intracellular Ca2⫹ also promoted pore expansion leading to syncytium formation induced by the measles virus membrane fusion complex, and the F and H components of this complex expressed together or individually in cells coprecipitated with AX1 (Fig. 6). Whether AX1 interacts directly or indirectly with measles virus F and H proteins is unclear, but the interaction appears to be specific since AX1 did not coimmunoprecipitate an unrelated integral membrane protein (HER2 receptor). As with AX1, measles virus M protein also interacts with the cytoplasmic tails of both the F and H proteins, where competitive M protein interactions with actin regulate the balance between syncytium formation and virus assembly (45–47). Nearly the entire 33-residue cytoplasmic tail of measles virus F protein can be deleted without compromising syncytium formation, while deleting more than 14 amino acids of the 34-residue H protein cytoplasmic tail greatly reduces syncytium formation (3, 42). These results suggest that AX1 interactions with measles virus H protein may be more biologically relevant to syncytium formation than AX1-F protein interactions. However, the F protein cytoplasmic tail of simian virus 5 (SV5), another member of the Paramyxoviridae, is dispensable for pore formation but required for pore expansion (48), suggesting that AX1-F protein interactions may be more relevant in this different virus system. Additional studies with measles virus F and H protein tail truncation mutants, using both virus-

6144

jvi.asm.org

infected and transfected cells, may provide further insights into how AX1 promotes measles virus-induced syncytium formation. Annexins are a diverse family of Ca2⫹-dependent, membraneinteracting proteins implicated in a remarkable range of cellular processes, including inflammation, coagulation, endocytosis, apoptosis, actin rearrangement, and plasma membrane repair (49–53). The conserved, C-terminal core domain comprises four repeats arranged to form a curved disc shape, with the convex surface responsible for Ca2⫹-binding and interaction of annexins with phospholipid bilayers and cellular partners. The nonconserved short N-terminal domain of annexins is thought to interact with the cytoplasm-facing concave surface of the core domain and contributes to functional variation between different members of this family (49). Members of the annexin family function both from inside and outside cells. Lacking a conventional signal peptide for secretion, annexins can be exported by ATP-binding cassette (ABC) transporters, in gelatinase granules, or in extracellular vesicles from where they can modulate anti-inflammatory responses and bind to signaling receptors, such as formyl peptide receptors (54, 55). Annexins are implicated in numerous aspects of virus-cell interactions. For example, AX2 and AX5 present in virions or extracellularly promote virus-cell attachment and infectivity of human papillomavirus (56), cytomegalovirus (57), enterovirus type 71 (58), hepatitis B virus (59), and HIV (60). Intracellularly, AX2 promotes formation of hepatitis C virus replication complexes and assembly of infectious HIV virions (61, 62), while AX6 interacts with the influenza virus M2 protein and inhibits viral release (63). Our present results define a new role for intracellular AX1 in regulating virus-cell interactions as a cofactor for virus-induced pore expansion and subsequent syncytium formation. Recent studies with mouse myoblasts provide a precedent for AX1 involvement in cell-cell fusion although in this instance AX1 functions extracellularly to promote cell migration, membrane apposition, and/or membrane merger (32, 64). While extracellular

Journal of Virology

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

FIG 8 Antibody inhibition of AX1 inhibits C2C12 differentiation-dependent myoblast fusion but not p14-mediated cell-cell fusion. (A) C2C12 mouse myoblasts induced to differentiate and fuse using horse serum were incubated for 51 h before addition of a 1:10 dilution of AX1 or PDI (control) antibody (Ab). Cells were incubated for a further 16 h, fixed, and Giemsa-stained; images were acquired by bright-field microscopy (top), and the numbers of syncytial nuclei per microscopic field were quantified (bottom). Results are mean ⫾ standard error from n ⫽ 3 experiments. (B) HT1080 cells transfected with p14 were treated with a 1:50, 1:25, or 1:10 dilution of AX1 antibody at 2 h posttransfection, fixed and Giemsa stained at 14 h posttransfection, and imaged (top), and the numbers of syncytial nuclei per microscopic field were quantified (bottom). Results are mean ⫾ standard error from n ⫽ 3 experiments. (C) C2C12 cells transfected with p14 were treated with a 1:10 dilution of AX1 antibody at 2 h posttransfection, fixed and Giemsa stained at 12 h posttransfection, and imaged (top), and the numbers of syncytial nuclei per microscopic field were quantified (bottom). Results are mean ⫾ standard error from n ⫽ 3 experiments.

Annexin A1 and Syncytium Formation

June 2014 Volume 88 Number 11

leads to transient Ca2⫹ influx. If so, then cell-cell membrane fusion would activate the Ca2⫹-triggered cellular membrane repair response, which, interestingly, is also dependent on AX1 (77). It is also possible that p14 activates endoplasmic reticulum store-operated Ca2⫹ channels, as recently shown for rotavirus NSP4 (78), a viroporin that shares structural and functional similarities to FAST proteins. The FAST proteins are the smallest known membrane fusion proteins and the only viral fusogens that specifically evolved to induce cell-cell, rather than virus-cell, membrane fusion. These structural and functional attributes are reflected in distinct differences between the mechanisms employed by FAST proteins and enveloped-virus fusogens to mediate membrane fusion (12, 25). However, our current findings indicate that enveloped-virus fusogens and FAST proteins may share an AX1- and Ca2⫹-dependent pathway for converting micropores into the macropores needed for syncytium formation. Recent studies also indicate that influenza virus HA-, baculovirus gp64-, and parainfluenza virus 5-induced syncytiogenesis are all dependent on ATP, actin remodeling, and/or membrane curvature for pore expansion (66– 69). Taken together, these studies suggest that the pore expansion stage of syncytium formation reflects generalized cellular responses to the presence of cell-cell fusion pores that serve to resolve these thermodynamically unfavorable membrane structures by converting them into lumen-sized openings and syncytia. ACKNOWLEDGMENTS This research was supported by a grant from the Canadian Institutes of Health Research. M.C. was funded by scholarships from the Natural Sciences and Engineering Research Council of Canada and the Nova Scotia Health Research Foundation. We thank Craig McCormick, Chris Richardson, and Graham Dellaire for reagents and advice on their use, Julie Boutilier for providing yeast two-hybrid results, and Cameron Landry for cloning AX1 cDNA.

REFERENCES 1. Harrison SC. 2008. Viral membrane fusion. Nat. Struct. Mol. Biol. 15: 690 – 698. http://dx.doi.org/10.1038/nsmb.1456. 2. White JM, Delos SE, Brecher M, Schornberg K. 2008. Structures and mechanisms of viral membrane fusion proteins: multiple variations on a common theme. Crit. Rev. Biochem. Mol. Biol. 43:189 –219. http://dx.doi .org/10.1080/10409230802058320. 3. Cathomen T, Naim HY, Cattaneo R. 1998. Measles viruses with altered envelope protein cytoplasmic tails gain cell fusion competence. J. Virol. 72:1224 –1234. 4. Takeda M, Ohno S, Seki F, Nakatsu Y, Tahara M, Yanagi Y. 2005. Long untranslated regions of the measles virus M and F genes control virus replication and cytopathogenicity. J. Virol. 79:14346 –14354. http://dx.doi .org/10.1128/JVI.79.22.14346-14354.2005. 5. Perfettini JL, Nardacci R, Seror C, Raza SQ, Sepe S, Saidi H, Brottes F, Amendola A, Subra F, Del Nonno F, Chessa L, D’Incecco A, Gougeon ML, Piacentini M, Kroemer G. 2010. 53BP1 represses mitotic catastrophe in syncytia elicited by the HIV-1 envelope. Cell Death Differ. 17:811– 820. http://dx.doi.org/10.1038/cdd.2009.159. 6. Watanabe S, Shirogane Y, Suzuki SO, Ikegame S, Koga R, Yanagi Y. 2013. Mutant fusion proteins with enhanced fusion activity promote measles virus spread in human neuronal cells and brains of suckling hamsters. J. Virol. 87:2648 –2659. http://dx.doi.org/10.1128/JVI.02632-12. 7. Guedan S, Grases D, Rojas JJ, Gros A, Vilardell F, Vile R, Mercade E, Cascallo M, Alemany R. 2012. GALV expression enhances the therapeutic efficacy of an oncolytic adenovirus by inducing cell fusion and enhancing virus distribution. Gene Ther. 19:1048 –1057. http://dx.doi.org/10 .1038/gt.2011.184. 8. Chen HH, Cawood R, El-Sherbini Y, Purdie L, Bazan-Peregrino M, Seymour LW, Carlisle RC. 2011. Active adenoviral vascular penetration

jvi.asm.org 6145

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

anti-AX1 antibody inhibited differentiation-dependent muscle cell fusion (Fig. 8), similar treatment had no effect on p14-induced syncytiogenesis in either HT1080 cells or undifferentiated C2C12 mouse myoblasts (Fig. 8). Furthermore, chelation of intracellular Ca2⫹ inhibited both p14-induced (Fig. 5) and measles virus-induced (Fig. 6C) syncytium formation. Thus, intracellular, not extracellular, AX1 is important for FAST protein- and measles virusinduced syncytium formation. AX1 can therefore function intracellularly or extracellularly to promote syncytium formation, depending on the nature of the fusogen. AX1 serves a similar dual role in maintaining endothelial blood-brain barrier integrity, interacting with the actin cytoskeleton and modulating Rho GTPase activity via signaling through formyl peptide receptor 2 to organize adherens junctions and tight junctions (65). While extracellular AX1 promotes the prefusion or membrane merger stages of myoblast fusion, intracellular AX1 functions at the postfusion, pore expansion stage of viral fusogen-induced syncytiogenesis. Pronounced inhibitory effects of AX1 knockdown and intracellular Ca2⫹ chelation on p14-induced syncytiogenesis (Fig. 1 and 5) in the absence of adverse effects on p14 and measles virus-mediated pore formation (Fig. 7) support this conclusion. Our current understanding of pore expansion is fragmentary at best. Studies with baculovirus gp64, influenza virus hemagglutinin (HA), and parainfluenza virus 5 indicate that pore expansion is an ATP-dependent event that is impeded by cortical actin and promoted by dynamin, phosphatidylinositol-4,5-bisphosphate (PIP2), and protein motifs such as ENTH and BAR domains that induce membrane curvature (66–69). The ability of annexins to integrate Ca2⫹ signaling with actin and plasma membrane dynamics may be relevant to a role in pore expansion (70). For example, AX2 and AX6 link PIP2 and cholesterol-rich membrane microdomains, respectively, to the actin cytoskeleton (71, 72), while AX1 is a regulator of Ca2⫹-dependent actin bundling and interacts with ceramide platforms to organize and internalize plasma membrane proteins (70, 73). PIP2 and actin dynamics are involved in myoblast fusion and virus-induced syncytium formation (32, 68), and p14-induced syncytiogenesis is influenced by membrane microdomains and actin remodeling (24, 74). Experiments examining annexin interactions with PIP2, actin, and/or lipid microdomains might therefore be informative with regard to how AX1 promotes pore expansion following p14- and measles virus-induced cell-cell fusion. The present results also revealed another parallel between muscle cell fusion and virus-mediated syncytium formation, both of which are dependent on intracellular Ca2⫹. Myoblast differentiation requires membrane hyperpolarization, Ca2⫹ influx, and Ca2⫹-mediated signaling through nuclear factor of activated T cells (NFAT) (40, 75, 76). Both measles virus- and p14-induced syncytium formation were inhibited by chelation of intracellular Ca2⫹ (Fig. 5 and 6), and BAPTA-AM treatment disrupted p14AX1 interaction (Fig. 4) but had no effect on pore formation (Fig. 7). Intracellular Ca2⫹ is therefore an essential factor for measles virus- and p14-induced syncytiogenesis, promoting the pore expansion stage of syncytium formation. Increased intracellular Ca2⫹ during myoblast fusion is provided by activation of T-type channels in the plasma membrane or store-operated Ca2⫹ entry channels in the endoplasmic reticulum (ER) (40, 75). The source of intracellular Ca2⫹ during p14- and measles virus-induced syncytium formation is presently unclear. One possibility is that disruption of membrane lamellar structure during the fusion process

Ciechonska et al.

9. 10. 11.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24. 25.

26.

27.

6146

jvi.asm.org

28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38. 39. 40.

41.

42.

43.

44. 45.

proteins of two subtypes differ in their ability to support assembly. J. Virol. 79:13673–13684. http://dx.doi.org/10.1128/JVI.79.21.13673-13684.2005. Long G, Pan X, Westenberg M, Vlak JM. 2006. Functional role of the cytoplasmic tail domain of the major envelope fusion protein of group II baculoviruses. J. Virol. 80:11226 –11234. http://dx.doi.org/10.1128/JVI .01178-06. Waning DL, Russell CJ, Jardetzky TS, Lamb RA. 2004. Activation of a paramyxovirus fusion protein is modulated by inside-out signaling from the cytoplasmic tail. Proc. Natl. Acad. Sci. U. S. A. 101:9217–9222. http: //dx.doi.org/10.1073/pnas.0403339101. Top D, Barry C, Racine T, Ellis CL, Duncan R. 2009. Enhanced fusion pore expansion mediated by the trans-acting endodomain of the reovirus FAST proteins. PLoS Pathog. 5:e1000331. http://dx.doi.org/10.1371 /journal.ppat.1000331. Corcoran JA, Duncan R. 2004. Reptilian reovirus utilizes a small type III protein with an external myristylated amino terminus to mediate cell-cell fusion. J. Virol. 78:4342– 4351. http://dx.doi.org/10.1128/JVI.78.8.4342-4351 .2004. Leikina E, Melikov K, Sanyal S, Verma SK, Eun B, Gebert C, Pfeifer K, Lizunov VA, Kozlov MM, Chernomordik LV. 2013. Extracellular annexins and dynamin are important for sequential steps in myoblast fusion. J. Cell Biol. 200:109 –123. http://dx.doi.org/10.1083/jcb.201207012. Futter CE, Felder S, Schlessinger J, Ullrich A, Hopkins CR. 1993. Annexin I is phosphorylated in the multivesicular body during the processing of the epidermal growth factor receptor. J. Cell Biol. 120:77– 83. http://dx.doi.org/10.1083/jcb.120.1.77. Tcatchoff L, Andersson S, Utskarpen A, Klokk TI, Skanland SS, Pust S, Gerke V, Sandvig K. 2012. Annexin A1 and A2: roles in retrograde trafficking of Shiga toxin. PLoS One 7:e40429. http://dx.doi.org/10.1371 /journal.pone.0040429. Schultz E, Jaryszak DL, Gibson MC, Albright DJ. 1986. Absence of exogenous satellite cell contribution to regeneration of frozen skeletal muscle. J. Muscle Res. Cell Motil. 7:361–367. http://dx.doi.org/10.1007 /BF01753657. Feige JN, Sage D, Wahli W, Desvergne B, Gelman L. 2005. PixFRET, an ImageJ plug-in for FRET calculation that can accommodate variations in spectral bleed-throughs. Microsc. Res. Tech. 68:51–58. http://dx.doi.org /10.1002/jemt.20215. Corcoran JA, Clancy EK, Duncan R. 2011. Homomultimerization of the reovirus p14 fusion-associated small transmembrane protein during transit through the ER-Golgi complex secretory pathway. J. Gen. Virol. 92: 162–166. http://dx.doi.org/10.1099/vir.0.026013-0. Marambaud P, Dreses-Werringloer U, Vingtdeux V. 2009. Calcium signaling in neurodegeneration. Mol. Neurodegen. 4:20. http://dx.doi.org /10.1186/1750-1326-4-20. Bird GS, DeHaven WI, Smyth JT, Putney JW, Jr. 2008. Methods for studying store-operated calcium entry. Methods 46:204 –212. http://dx .doi.org/10.1016/j.ymeth.2008.09.009. Bijlenga P, Liu JH, Espinos E, Haenggeli CA, Fischer-Lougheed J, Bader CR, Bernheim L. 2000. T-type alpha 1H Ca2⫹ channels are involved in Ca2⫹ signaling during terminal differentiation (fusion) of human myoblasts. Proc. Natl. Acad. Sci. U. S. A. 97:7627–7632. http://dx.doi.org/10 .1073/pnas.97.13.7627. Corey EA, Iorio RM. 2007. Mutations in the stalk of the measles virus hemagglutinin protein decrease fusion but do not interfere with virusspecific interaction with the homologous fusion protein. J. Virol. 81: 9900 –9910. http://dx.doi.org/10.1128/JVI.00909-07. Moll M, Klenk HD, Maisner A. 2002. Importance of the cytoplasmic tails of the measles virus glycoproteins for fusogenic activity and the generation of recombinant measles viruses. J. Virol. 76:7174 –7186. http://dx.doi.org /10.1128/JVI.76.14.7174-7186.2002. Top D, de Antueno R, Salsman J, Corcoran J, Mader J, Hoskin D, Touhami A, Jericho MH, Duncan R. 2005. Liposome reconstitution of a minimal protein-mediated membrane fusion machine. EMBO J. 24: 2980 –2988. http://dx.doi.org/10.1038/sj.emboj.7600767. Sekar RB, Periasamy A. 2003. Fluorescence resonance energy transfer (FRET) microscopy imaging of live cell protein localizations. J. Cell Biol. 160:629 – 633. http://dx.doi.org/10.1083/jcb.200210140. Wakimoto H, Shimodo M, Satoh Y, Kitagawa Y, Takeuchi K, Gotoh B, Itoh M. 2013. F-actin modulates measles virus cell-cell fusion and assembly by altering the interaction between the matrix protein and the cytoplasmic tail of hemagglutinin. J. Virol. 87:1974 –1984. http://dx.doi.org /10.1128/JVI.02371-12.

Journal of Virology

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

12.

by targeted formation of heterocellular endothelial-epithelial syncytia. Mol. Ther. 19:67–75. http://dx.doi.org/10.1038/mt.2010.209. Abmayr SM, Pavlath GK. 2012. Myoblast fusion: lessons from flies and mice. Development 139:641– 656. http://dx.doi.org/10.1242/dev.068353. Kozlov MM, McMahon HT, Chernomordik LV. 2010. Protein-driven membrane stresses in fusion and fission. Trends Biochem. Sci. 35:699 – 706. http://dx.doi.org/10.1016/j.tibs.2010.06.003. Melikyan GB. 2008. Common principles and intermediates of viral protein-mediated fusion: the HIV-1 paradigm. Retrovirology 5:111. http://dx .doi.org/10.1186/1742-4690-5-111. Boutilier J, Duncan R. 2011. The reovirus fusion-associated small transmembrane (FAST) proteins: virus-encoded cellular fusogens. Curr. Top. Membr. 68:107–140. http://dx.doi.org/10.1016/B978-0-12 -385891-7.00005-2. Brown CW, Stephenson KB, Hanson S, Kucharczyk M, Duncan R, Bell JC, Lichty BD. 2009. The p14 FAST protein of reptilian reovirus increases vesicular stomatitis virus neuropathogenesis. J. Virol. 83:552–561. http: //dx.doi.org/10.1128/JVI.01921-08. Duncan R, Sullivan K. 1998. Characterization of two avian reoviruses that exhibit strain-specific quantitative differences in their syncytiuminducing and pathogenic capabilities. Virology 250:263–272. http://dx.doi .org/10.1006/viro.1998.9371. Duncan R, Chen Z, Walsh S, Wu S. 1996. Avian reovirus-induced syncytium formation is independent of infectious progeny virus production and enhances the rate, but is not essential, for virus-induced cytopathology and virus egress. Virology 224:453– 464. http://dx.doi.org/10 .1006/viro.1996.0552. Salsman J, Top D, Boutilier J, Duncan R. 2005. Extensive syncytium formation mediated by the reovirus FAST proteins triggers apoptosisinduced membrane instability. J. Virol. 79:8090 – 8100. http://dx.doi.org /10.1128/JVI.79.13.8090-8100.2005. Barry C, Key T, Haddad R, Duncan R. 2010. Features of a spatially constrained cystine loop in the p10 FAST protein ectodomain define a new class of viral fusion peptides. J. Biol. Chem. 285:16424 –16433. http://dx .doi.org/10.1074/jbc.M110.118232. Corcoran JA, Syvitski R, Top D, Epand RM, Epand RF, Jakeman D, Duncan R. 2004. Myristoylation, a protruding loop, and structural plasticity are essential features of a nonenveloped virus fusion peptide motif. J. Biol. Chem. 279:51386– 51394. http://dx.doi.org/10.1074/jbc.M406990200. Dawe S, Corcoran JA, Clancy EK, Salsman J, Duncan R. 2005. Unusual topological arrangement of structural motifs in the baboon reovirus fusion-associated small transmembrane protein. J. Virol. 79:6216 – 6226. http://dx.doi.org/10.1128/JVI.79.10.6216-6226.2005. Top D, Read JA, Dawe SJ, Syvitski RT, Duncan R. 2012. Cell-cell membrane fusion induced by p15 fusion-associated small transmembrane (FAST) protein requires a novel fusion peptide motif containing a myristoylated polyproline type II helix. J. Biol. Chem. 287:3403–3414. http://dx .doi.org/10.1074/jbc.M111.305268. Barry C, Duncan R. 2009. Multifaceted sequence-dependent and -independent roles for reovirus FAST protein cytoplasmic tails in fusion pore formation and syncytiogenesis. J. Virol. 83:12185–12195. http://dx.doi .org/10.1128/JVI.01667-09. Clancy EK, Duncan R. 2009. Reovirus FAST protein transmembrane domains function in a modular, primary sequence-independent manner to mediate cell-cell membrane fusion. J. Virol. 83:2941–2950. http://dx .doi.org/10.1128/JVI.01869-08. Clancy EK, Duncan R. 2011. Helix-destabilizing, ␤-branched and polar residues in the baboon reovirus p15 transmembrane domain influence the modularity of the FAST proteins. J. Virol. 85:4707– 4719. http://dx.doi.org /10.1128/JVI.02223-10. Salsman J, Top D, Barry C, Duncan R. 2008. A virus-encoded cell-cell fusion machine dependent on surrogate adhesins. PLoS Pathog. 4:e1000016. http://dx.doi.org/10.1371/journal.ppat.1000016. Clancy EK, Barry C, Ciechonska M, Duncan R. 2010. Different activities of the reovirus FAST proteins and influenza hemagglutinin in cell-cell fusion assays and in response to membrane curvature agents. Virology 397:119 –129. http://dx.doi.org/10.1016/j.virol.2009.10.039. Broer R, Boson B, Spaan W, Cosset FL, Corver J. 2006. Important role for the transmembrane domain of severe acute respiratory syndrome coronavirus spike protein during entry. J. Virol. 80:1302–1310. http://dx .doi.org/10.1128/JVI.80.3.1302-1310.2006. Chen BJ, Takeda M, Lamb RA. 2005. Influenza virus hemagglutinin (H3 subtype) requires palmitoylation of its cytoplasmic tail for assembly: M1

Annexin A1 and Syncytium Formation

June 2014 Volume 88 Number 11

63. Ma H, Kien F, Maniere M, Zhang Y, Lagarde N, Tse KS, Poon LL, Nal B. 2012. Human annexin A6 interacts with influenza a virus protein M2 and negatively modulates infection. J. Virol. 86:1789 –1801. http://dx.doi .org/10.1128/JVI.06003-11. 64. Bizzarro V, Belvedere R, Dal Piaz F, Parente L, Petrella A. 2012. Annexin A1 induces skeletal muscle cell migration acting through formyl peptide receptors. PLoS One 7:e48246. http://dx.doi.org/10.1371/journal .pone.0048246. 65. Cristante E, McArthur S, Mauro C, Maggioli E, Romero IA, Wylezinska-Arridge M, Couraud PO, Lopez-Tremoleda J, Christian HC, Weksler BB, Malaspina A, Solito E. 2013. Identification of an essential endogenous regulator of blood-brain barrier integrity, and its pathological and therapeutic implications. Proc. Natl. Acad. Sci. U. S. A. 110:832– 841. http://dx.doi.org/10.1073/pnas.1209362110. 66. Chen A, Leikina E, Melikov K, Podbilewicz B, Kozlov MM, Chernomordik LV. 2008. Fusion-pore expansion during syncytium formation is restricted by an actin network. J. Cell Sci. 121:3619 –3628. http://dx.doi .org/10.1242/jcs.032169. 67. Richard JP, Leikina E, Chernomordik LV. 2009. Cytoskeleton reorganization in influenza hemagglutinin-initiated syncytium formation. Biochim. Biophys. Acta 1788:450 – 457. http://dx.doi.org/10.1016/j.bbamem.2008.09 .014. 68. Richard JP, Leikina E, Langen R, Henne WM, Popova M, Balla T, McMahon HT, Kozlov MM, Chernomordik LV. 2011. Intracellular curvature-generating proteins in cell-to-cell fusion. Biochem. J. 440:185– 193. http://dx.doi.org/10.1042/BJ20111243. 69. Wurth MA, Schowalter RM, Smith EC, Moncman CL, Dutch RE, McCann RO. 2010. The actin cytoskeleton inhibits pore expansion during PIV5 fusion protein-promoted cell-cell fusion. Virology 404:117–126. http://dx.doi.org/10.1016/j.virol.2010.04.024. 70. Hayes MJ, Rescher U, Gerke V, Moss SE. 2004. Annexin-actin interactions. Traffic 5:571–576. http://dx.doi.org/10.1111/j.1600-0854 .2004.00210.x. 71. Cornely R, Rentero C, Enrich C, Grewal T, Gaus K. 2011. Annexin A6 is an organizer of membrane microdomains to regulate receptor localization and signalling. IUBMB Life 63:1009 –1017. http://dx.doi.org/10.1002 /iub.540. 72. Hayes MJ, Merrifield CJ, Shao D, Ayala-Sanmartin J, Schorey CD, Levine TP, Proust J, Curran J, Bailly M, Moss SE. 2004. Annexin 2 binding to phosphatidylinositol 4,5-bisphosphate on endocytic vesicles is regulated by the stress response pathway. J. Biol. Chem. 279:14157–14164. http://dx.doi.org/10.1074/jbc.M313025200. 73. Babiychuk EB, Monastyrskaya K, Draeger A. 2008. Fluorescent annexin A1 reveals dynamics of ceramide platforms in living cells. Traffic 9:1757– 1775. http://dx.doi.org/10.1111/j.1600-0854.2008.00800.x. 74. Corcoran JA, Salsman J, de Antueno R, Touhami A, Jericho MH, Clancy EK, Duncan R. 2006. The p14 fusion-associated small transmembrane (FAST) protein effects membrane fusion from a subset of membrane microdomains. J. Biol. Chem. 281:31778 –31789. http://dx.doi.org /10.1074/jbc.M602566200. 75. Darbellay B, Arnaudeau S, Konig S, Jousset H, Bader C, Demaurex N, Bernheim L. 2009. STIM1- and Orai1-dependent store-operated calcium entry regulates human myoblast differentiation. J. Biol. Chem. 284:5370 – 5380. http://dx.doi.org/10.1074/jbc.M806726200. 76. Hindi SM, Tajrishi MM, Kumar A. 2013. Signaling mechanisms in mammalian myoblast fusion. Sci. Signal. 6:re2. http://dx.doi.org/10.1126 /scisignal.2003832. 77. Draeger A, Monastyrskaya K, Babiychuk EB. 2011. Plasma membrane repair and cellular damage control: the annexin survival kit. Biochem. Pharmacol. 81:703–712. http://dx.doi.org/10.1016/j.bcp.2010.12.027. 78. Hyser JM, Utama B, Crawford SE, Broughman JR, Estes MK. 2013. Activation of the endoplasmic reticulum calcium sensor STIM1 and storeoperated calcium entry by rotavirus requires NSP4 viroporin activity. J. Virol. 87:13579 –13588. http://dx.doi.org/10.1128/JVI.02629-13.

jvi.asm.org 6147

Downloaded from http://jvi.asm.org/ on November 3, 2015 by UNIVERSITY OF NEW SOUTH WALES

46. Tahara M, Takeda M, Yanagi Y. 2007. Altered interaction of the matrix protein with the cytoplasmic tail of hemagglutinin modulates measles virus growth by affecting virus assembly and cell-cell fusion. J. Virol. 81: 6827– 6836. http://dx.doi.org/10.1128/JVI.00248-07. 47. Cathomen T, Mrkic B, Spehner D, Drillien R, Naef R, Pavlovic J, Aguzzi A, Billeter MA, Cattaneo R. 1998. A matrix-less measles virus is infectious and elicits extensive cell fusion: consequences for propagation in the brain. EMBO J. 17:3899 –3908. http://dx.doi.org/10.1093/emboj/17.14 .3899. 48. Dutch RE, Lamb RA. 2001. Deletion of the cytoplasmic tail of the fusion protein of the paramyxovirus simian virus 5 affects fusion pore enlargement. J. Virol. 75:5363–5369. http://dx.doi.org/10.1128/JVI.75.11.5363 -5369.2001. 49. Gerke V, Creutz CE, Moss SE. 2005. Annexins: linking Ca2⫹ signalling to membrane dynamics. Nat. Rev. Mol. Cell Biol. 6:449 – 461. http://dx.doi .org/10.1038/nrm1661. 50. McNeil AK, Rescher U, Gerke V, McNeil PL. 2006. Requirement for annexin A1 in plasma membrane repair. J. Biol. Chem. 281:35202–35207. http://dx.doi.org/10.1074/jbc.M606406200. 51. Bouter A, Gounou C, Berat R, Tan S, Gallois B, Granier T, d’Estaintot BL, Poschl E, Brachvogel B, Brisson AR. 2011. Annexin-A5 assembled into two-dimensional arrays promotes cell membrane repair. Nat. Commun. 2:270. http://dx.doi.org/10.1038/ncomms1270. 52. Potez S, Luginbuhl M, Monastyrskaya K, Hostettler A, Draeger A, Babiychuk EB. 2011. Tailored protection against plasmalemmal injury by annexins with different Ca2⫹ sensitivities. J. Biol. Chem. 286:17982– 17991. http://dx.doi.org/10.1074/jbc.M110.187625. 53. Patel DM, Ahmad SF, Weiss DG, Gerke V, Kuznetsov SA. 2011. Annexin A1 is a new functional linker between actin filaments and phagosomes during phagocytosis. J. Cell Sci. 124:578 –588. http://dx.doi.org/10 .1242/jcs.076208. 54. Gavins FN, Hickey MJ. 2012. Annexin A1 and the regulation of innate and adaptive immunity. Front. Immunol. 3:354. http://dx.doi.org/10 .3389/fimmu.2012.00354. 55. Perretti M, D’Acquisto F. 2009. Annexin A1 and glucocorticoids as effectors of the resolution of inflammation. Nat. Rev. Immunol. 9:62–70. http://dx.doi.org/10.1038/nri2470. 56. Woodham AW, Da Silva DM, Skeate JG, Raff AB, Ambroso MR, Brand HE, Isas JM, Langen R, Kast WM. 2012. The S100A10 subunit of the annexin A2 heterotetramer facilitates L2-mediated human papillomavirus infection. PLoS One 7:e43519. http://dx.doi.org/10.1371/journal.pone .0043519. 57. Raynor CM, Wright JF, Waisman DM, Pryzdial EL. 1999. Annexin II enhances cytomegalovirus binding and fusion to phospholipid membranes. Biochemistry 38:5089–5095. http://dx.doi.org/10.1021/bi982095b. 58. Yang SL, Chou YT, Wu CN, Ho MS. 2011. Annexin II binds to capsid protein VP1 of enterovirus 71 and enhances viral infectivity. J. Virol. 85: 11809 –11820. http://dx.doi.org/10.1128/JVI.00297-11. 59. De Meyer S, Gong Z, Depla E, Maertens G, Yap SH. 1999. Involvement of phosphatidylserine and non-phospholipid components of the hepatitis B virus envelope in human Annexin V binding and in HBV infection in vitro. J. Hepatol. 31:783–790. http://dx.doi.org/10.1016/S0168-8278(99) 80278-5. 60. Rai T, Mosoian A, Resh MD. 2010. Annexin 2 is not required for human immunodeficiency virus type 1 particle production but plays a cell typedependent role in regulating infectivity. J. Virol. 84:9783–9792. http://dx .doi.org/10.1128/JVI.01584-09. 61. Saxena V, Lai CK, Chao TC, Jeng KS, Lai MM. 2012. Annexin A2 is involved in the formation of hepatitis C virus replication complex on the lipid raft. J. Virol. 86:4139 – 4150. http://dx.doi.org/10.1128/JVI.06327-11. 62. Ryzhova EV, Vos RM, Albright AV, Harrist AV, Harvey T, GonzalezScarano F. 2006. Annexin 2: a novel human immunodeficiency virus type 1 Gag binding protein involved in replication in monocyte-derived macrophages. J. Virol. 80:2694 –2704. http://dx.doi.org/10.1128/JVI.80.6 .2694-2704.2006.