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. 2020 May 22;9:e51358. doi: 10.7554/eLife.51358

A viral fusogen hijacks the actin cytoskeleton to drive cell-cell fusion

Ka Man Carmen Chan 1,2, Sungmin Son 2, Eva M Schmid 2, Daniel A Fletcher 1,2,3,4,✉
Editors: Michael M Kozlov5, Anna Akhmanova6
PMCID: PMC7244324  PMID: 32441254

Abstract

Cell-cell fusion, which is essential for tissue development and used by some viruses to form pathological syncytia, is typically driven by fusogenic membrane proteins with tall (>10 nm) ectodomains that undergo conformational changes to bring apposing membranes in close contact prior to fusion. Here we report that a viral fusogen with a short (<2 nm) ectodomain, the reptilian orthoreovirus p14, accomplishes the same task by hijacking the actin cytoskeleton. We show that phosphorylation of the cytoplasmic domain of p14 triggers N-WASP-mediated assembly of a branched actin network. Using p14 mutants, we demonstrate that fusion is abrogated when binding of an adaptor protein is prevented and that direct coupling of the fusogenic ectodomain to branched actin assembly is sufficient to drive cell-cell fusion. This work reveals how the actin cytoskeleton can be harnessed to overcome energetic barriers to cell-cell fusion.

Research organism: Human, Virus

Introduction

Cell-cell fusion plays a critical role in the development of multicellular organisms, beginning with fertilization and continuing with formation of muscles, osteoclasts, and the placenta in mammals. Viral pathogens, including some members of poxvirus, paramyxovirus, herpesvirus, retrovirus, aquareovirus and orthoreovirus, cause infected cells to fuse with their neighbors, creating syncytia that contribute to disease pathology (Compton and Schwartz, 2017; Domachowske and Rosenberg, 1999; Moss, 2006; Smith et al., 2009). While the basic steps of membrane fusion have been extensively investigated in the context of enveloped virus entry and SNARE-mediated intracellular vesicle fusion (Südhof and Rothman, 2009), the molecules and pathways responsible for cell-cell fusion are less well understood. The best studied cell-cell fusogens are those with similarities to enveloped viral fusogens, including syncytin-1 (placental syncytiotrophoblasts formation) (Gong et al., 2005; Renard et al., 2005), Hap2 (conserved in eukaryotic gamete fusion) (Fédry et al., 2017; Feng et al., 2018; Liu et al., 2008b; Mori et al., 2006; Valansi et al., 2017; von Besser et al., 2006), and Eff-1/Aff-1 (C. elegans epithelial fusion) (Mohler et al., 2002; Pérez-Vargas et al., 2014; Sapir et al., 2008; Shemer et al., 2004; Zeev-Ben-Mordehai et al., 2014).

A key feature of viral and cell-cell fusogens is their tall ectodomains, which in their metastable pre-fusion state typically extend more than 10 nm from the membrane. Since the plasma membrane of cells is densely decorated with glycoproteins and glycolipids that could sterically block membranes from getting close enough to fuse, the tall ectodomains of viral and cell-cell fusogens may allow them to reach across the membrane gap and anchor to the apposing membrane, involving insertion of a fusion peptide for Class I viral fusogens or a fusion loop for Class II (Harrison, 2015; Podbilewicz, 2014). Once the fusogen links the two membranes, conformational changes cause the fusogen to fold back, pulling the two membranes into close contact and forming a stable post-fusion structure that promotes membrane fusion (Bullough et al., 1994; Chen et al., 1999; Harrison, 2015; Ivanovic and Harrison, 2015; Pancera et al., 2014; Podbilewicz, 2014; Sapir et al., 2008; Wilson et al., 1981). This conformational change, together with fusogen oligomerization and local cooperativity, is believed to be sufficient to provide the energy required to overcome the repulsive hydration barrier, which prevents membranes from coming closer than ~2 nm (Chernomordik and Kozlov, 2003; Harrison, 2015; Ivanovic and Harrison, 2015; Pérez-Vargas et al., 2014; Rand and Parsegian, 1989).

However, in other instances of cell-cell fusion, transmembrane proteins required for fusion are short by comparison and do not appear to undergo conformational changes, raising the question of how they bring two plasma membranes into close contact (Figure 1a). One example is the reptilian orthoreovirus fusion protein p14, a non-structural, single-pass transmembrane protein, that is expressed after viral entry. One of seven members of the FAST family of reovirus fusion proteins discovered by Duncan and colleagues (Ciechonska and Duncan, 2014; Corcoran and Duncan, 2004; Dawe and Duncan, 2002; Duncan et al., 1995; Duncan et al., 2004; Guo et al., 2013; Kim et al., 2015; Racine et al., 2009; Shmulevitz et al., 2004; Shmulevitz and Duncan, 2000; Wilcox and Compans, 1982), p14 has a membrane-disruptive ectodomain that is necessary to drive cell-cell fusion but extends only 0.7–1.5 nm from the plasma membrane (Corcoran et al., 2004; Corcoran et al., 2006). This short ectodomain has minimal secondary structure but is sufficient to disrupt opposing membranes either when added as a soluble fragment or when bare liposomes containing p14 are brought into close apposition through divalent cations (Corcoran et al., 2004; Top et al., 2005). However, cell surfaces are crowded with proteins that sterically block close apposition, and it is unclear how p14 overcomes the energetic barrier of cell surface protein crowding, as well as the ~2 nm repulsive hydration barrier, to drive cell-cell fusion (Chernomordik and Kozlov, 2003; Harrison, 2015; Rand and Parsegian, 1989). Yet, expression of p14 alone in cultured cells is sufficient to drive fusion with neighboring naïve cells (Corcoran and Duncan, 2004; Duncan et al., 2004).

Figure 1. Expression of p14 drives cell-cell fusion and is quantified with a splitYFP fluorescent assay.

(a) Schematic of fusion-associated small transmembrane protein, p14, in proportion to post-fusion trimeric structure of cell-cell fusogens, Eff-1 (PDB:4OJC) and Hap2 (PDB: 5MF1), on the plasma membrane. (b) Expression of p14 in HEK293T cells drives cell-cell fusion forming large multinucleated cells that increases size and number of nuclei over time. (c) Average nuclei count of HEK293T cells expressing p14 at 12 hr and 24 hr. P-values are ks-test and error bars represent standard deviation from 3 independent transfections (See also Figure 1—source data 1). (d) Percent of p14 expressing nuclei in cells with 3 or more nuclei at 12 hr and 24 hr. P-values are two-tailed, two-sample Student’s t-test where ** = p<0.01, and error bars represent standard deviations from 3 independent transfections (e) Schematic of the splitYFP fluorescence assay to quantify cell-cell fusion. IYFP of sample is the YFP fluorescence intensity of the sample, IYFP of sample is the YFP fluorescence intensity of the non-transfected cells, and IYFP of sample is the YFP fluorescence intensity of cells transfected with p14 WT. (f) Average YFP fluorescence intensity of HEK293T cells expressing p14 at 12 hr and 24 hr with error bars representing standard deviations from 3 independent transfections of 3 wells each. P-values are two-tailed, two-sample Student’s t-test where *** = p<0.001 (See also Figure 1—figure supplement 1c, (d). (g) Average fusion index of p14 cytoplasmic truncation mutant with error bars representing standard deviations from 3 independent transfections of 3 wells each. P-values are two-tailed, two-sample Student’s t-test where *** = p<0.001 (See also Figure 1—figure supplement 1e, (f).

Figure 1—source data 1. Excel Spreadsheet of counts and distribution for p14-expressing HEK293T cells at 12 hr and 24 hr post transfection for Figure 1c and d.

Figure 1.

Figure 1—figure supplement 1. Characterization of p14 cytoplasmic tail cleavage and the splitYFP cell-cell fusion assay.

Figure 1—figure supplement 1.

(a) Schematic of hypothesized cleavage sites in p14 cytoplasmic tail based on molecular weights of cleaved fragments. (b) Membrane fractionation of p14 WT, p14 FVAI, Δcyto and GFP only, showing similar expression of each construct at the plasma membrane and molecular weights cleaved fragments. (c) Representative confocal image of HEK293T cells expressing mCherry (magenta) and nuclei stained with Hoechst 33342. (d) YFP intensity per cell vs number of nuclei in cell. Each point represents a single cell. (e) Representative confocal image of splitYFP cells expressing p14 WT mcherry (red). Nuclei are labeled with Hoechst 33342 (cyan), plasma membrane are labeled with CellMaskDeepRed (green). splitYFP has FOS and JUN coiled-coiled motif that directs splitYFP to the nucleus (yellow). (g) p14 WT and p14 Δcyto labeled with C-terminus mcherry tag are trafficked to the plasma membrane as visualized with TIRF microscopy. (g) Western blot of surface biotinylated GFP-tagged p14 WT, FVAI and Δcyto. (h) Average biotinylated p14 Δcyto normalized to that of p14 WT. Error bars represent standard deviations from three independent transfections and biotinylation. P-values are two-tailed two-sample Student’s t-test to p14 WT where n.s. = p>0.05. (i) Representative field of view of HEK293T cells expressing p14 Δcyto mCherry (magenta), nuclei stained with Hoechst 33342 (cyan).
Figure 1—figure supplement 1—source data 1. Excel Spreadsheet of surface biotinylation of p14-WT-GFP and p14-Δcyto-GFP for Figure 1—figure supplement 1h.
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To address the question of how p14 promotes close contact between cells and drives membrane fusion, we studied cell-cell fusion in HEK293T cells transiently expressing p14. We found that p14 drives cell-cell fusion by hijacking the host cell actin cytoskeleton. Using a phosphorylation-dependent motif in its cytoplasmic domain, p14 triggers N-WASP-mediated assembly of branched actin, directly coupling actin assembly with a short membrane-disruptive ectodomain. This work suggests that energetic barriers to cell-cell fusion can be overcome by harnessing force generated from local actin assembly, and it points to an alternate means of promoting cell-cell fusion in processes where tall fusogens have not been identified.

Results

Expression of p14 in HEK293T cells caused the cells to fuse with neighboring wild-type and p14-expressing cells, forming large multinucleated syncytia over the course of 24 hr (Figure 1b, Video 1), like previous reports for other cell types (Corcoran and Duncan, 2004). Partial cleavage of the p14 cytoplasmic tail, which also occurs during reptilian orthoreovirus infection (Top et al., 2009), liberates the C-terminus mCherry fluorescent tag from the transmembrane protein and serves as a convenient cytoplasmic marker of p14-expressing cells (Figure 1b, Figure 1—figure supplement 1a,b, and Video 2). At 12 hr post transfection, 5% of p14-expressing cells had 3 or more nuclei. At 24 hr post transfection, 34% of p14-expressing cells had 3 or nuclei (Figure 1c and d). In comparison, at 24 hr post transfection, no multinucleated cells were seen in cells transfected with mCherry alone (Figure 1—figure supplement 1c).

Video 1. Phase contrast timelapse of HEK293T cells expressing p14 WT showing extensive syncytium formation.

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Video 2. Confocal timelapse of a HEK293T expressing p14-mcherry (magenta) fusing with a WT HEK293T cell that appears dark.

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Plasma membrane is marked with gpi-anchored pHluorin (green). Scale bar is 5 μm.

To quantify cytoplasmic mixing during p14-mediated cell-cell fusion in a high-throughput manner, we expressed the two halves of splitYFP in two populations of HEK293T cells and mixed the cells together. When fusion between cells of the two populations occurred, the two halves of splitYFP self-associated in the mixed cytoplasm and fluoresced, allowing quantification by a plate reader (Figure 1e, and Figure 1—figure supplement 1d and e). Repeating the cell-cell fusion experiments above, the increase in YFP intensity in p14-expressing cells between 12 hr and 24 hr post transfection compared well with the increase in the percent of multinucleated cells, as quantified by counting nuclei (Figure 1f). Since YFP intensity is dependent on the percent of HEK293T cells expressing the two halves of splitYFP, the resultant YFP intensity was normalized to that of p14 WT and referred to as fusion index.

While the ectodomain of p14 is shorter than typical viral fusogens, its cytoplasmic domain is comparatively long (68 amino acids). To determine how the cytoplasmic domain of p14 might be involved in cell-cell fusion, we first truncated Q70-I125 (p14 Δcyto), retaining a polybasic motif needed for trafficking to the plasma membrane (Parmar et al., 2014). Although p14 Δcyto was properly trafficked to the plasma membrane (Figure 1—figure supplement 1f,g and h), cell-cell fusion was abrogated (Figure 1g, and Figure 1—figure supplement 1i). This is consistent with previous findings (Corcoran and Duncan, 2004), suggesting that p14 may be interacting with cellular components through its cytoplasmic tail to enable cell-cell fusion.

We next investigated whether post-translational modification of the cytoplasmic tail of p14 might play a role in cell-cell fusion. The p14 cytoplasmic tail is mostly disordered but has several tyrosines that could be phosphorylated (Figure 2a, and Figure 2—figure supplement 1a). To determine if these tyrosines are indeed phosphorylated, we immunoprecipitated p14 and probed with an anti-phosphotyrosine antibody, which confirmed p14 phosphorylation (Figure 2b). Next, we mutated each predicted tyrosine to phenylalanine (Y59F, Y77F, Y96F, Y100F, Y116F) and found that only one mutation (Y116F) decreased cell-cell fusion (Figure 2c and Figure 2—figure supplement 1b). We then used NetPhos3.1 and Scansite 4.0 to analyze the cytoplasmic tail of p14, and they predicted that Y116 is phosphorylated by c-src kinase (Figure 2d; Blom et al., 2004; Obenauer et al., 2003). To test this prediction, we mutated all other predicted phosphotyrosines of p14 to phenylalanine (YEY; Y59F, Y77F, Y96F, Y100F) and co-expressed it with constitutively active c-src mutant (CA c-src; Y527F). Y116 phosphorylation subsequently increased, indicating c-src phosphorylates p14 (Figure 2e). Consistent with this, Y116 phosphorylation was also increased when tyrosine phosphatases were broadly inhibited by addition of pervanadate (Figure 2e). Finally, to confirm that c-src is sufficient to phosphorylate Y116, we carried out a modified in vitro kinase assay using a peptide including Y116 (Y116-N121), along with CA c-src and kinase dead c-src (KD c-src; Y527F/K295R) mutants immunoprecipitated from HEK239T cells (Figure 2—figure supplement 1c and d). We found that CA c-src was sufficient to phosphorylate p14 cytoplasmic tail peptide, but KD c-src was not (Figure 2f), showing that c-src kinase is sufficient to phosphorylate Y116 in vitro. To verify this in live-cells, we expressed p14 WT in mouse embryonic fibroblasts lacking src, yes, and fyn kinase (SYF), as well as SYF cells supplemented with c-src (SYF + c-src). Despite lower transfection efficiency and reduced cell-cell fusion compared to HEK239T cells, SYF + c-src cells fused more than SYF cells (Figure 2g). Altogether, this demonstrates that c-src phosphorylates Y116 in the p14 cytoplasmic tail to drive cell-cell fusion.

Figure 2. p14 Y116 in the cytoplasmic tail is necessary for cell-cell fusion and is phosphorylated by c-src kinase.

(a) Schematic of predicted phosphotyrosines in p14 cytoplasmic tail (See also Figure 2—figure supplement 1a). (b) Western blot probed with α-phosphotyrosine confirming that p14 WT is phosphorylated. (c) Average fusion index of p14 phosphotyrosine mutants with error bars representing standard deviations from 3 independent transfections of 3 wells each. p-values are two-tailed, two-sample Student’s t-test where *** = p<0.001 (See also Figure 2—figure supplement 1b). (d) Schematic of c-src kinase and a tyrosine phosphatase activity on p14 Y116. (e) Western blot probed with α-phosphotyrosine confirming that p14 Y116 phosphorylation is increased with co-expression of constitutively active c-src kinase and with addition of pervanadate. (f) Dot blot of p14 cytoplasmic tail peptide phosphorylated in vitro with constitutively-active (Y527F) and kinase-dead c-src kinase (Y527F, K295R) (See also Figure 2—figure supplement 1c, (d). (g) Average percent of p14-expressing cells with 2 or more nuclei in SYF and SYF + c-src cells with error bars representing standard deviations from 3 independent transfections. p-values are two-tailed, two-sample Student’s t-test, where * = p<0.05 (See also Figure 2—figure supplement 1e).

Figure 2—source data 1. Excel Spreadsheet of counts for p14-expressing SYF and SYF+c-src cells for Figure 2g.

Figure 2.

Figure 2—figure supplement 1. Characterization of p14 cytoplasmic tail and Y116.

Figure 2—figure supplement 1.

(a) Disorder probability of p14 cytoplasmic tail as calculated with DisEMBL (http://dis.embl.de/). (b) Representative field of view of HEK293T cells expressing p14 WT and p14 mutants with p14 labeled with C-terminus mcherry (magenta), nuclei stained with Hoechst 33342 (cyan). (c) Coomassie stain of immunoprecipitation of FLAG-tagged constitutively active src kinase (CA src) and kinase dead src kinase (KD src) from HEK293T cells. (d) Western blot of immunoprecipitation of FLAG-tagged constitutively active src kinase (CA src) and kinase dead src kinase (KD src) probed with α-FLAG.
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To determine which cellular components could be interacting with p14 upon phosphorylation, we used the Eukaryotic Linear Motif (ELM) prediction tool to identify potential binding motifs (Dinkel et al., 2016). ELM predicted that phosphorylated Y116 binds to the SH2 domain of Grb2 as part of a Grb2 consensus-binding motif, YVNI (Figure 3a). To test this prediction, we carried out a co-immunoprecipitation assay and confirmed that p14 binds to Grb2 (Figure 3b). To determine if Grb2 binding is necessary for p14-mediated cell-cell fusion, we introduced two point mutations that disrupt the predicted Grb2 binding site both individually (Y116F, N118A) and together (FVAI; Y116F/N118A) (Figure 3b). While trafficking to the plasma membrane was intact, all three mutants severely attenuated cell-cell fusion (Figure 3c, Figure 3—figure supplement 1a,b and c), suggesting that Grb2 is important for p14-mediated fusion. To confirm that p14 is sufficient to recruit Grb2, we conjugated biotinylated p14 cytoplasmic tail peptide to streptavidin beads in vitro and incubated them with purified Grb2 fluorescently labeled with AF647 (Figure 3—figure supplement 1d). Consistent with our co-immunoprecipitation results, only phosphorylated Y116 bound to Grb2 (Figure 3d, and Figure 3—figure supplement 1e). When p14 Y116 phosphorylation is increased in cells with either co-expression of CA c-src or addition of the phosphatase inhibitor pervanadate, GFP-labeled Grb2 co-localized and enriched with p14 at the plasma membrane (Figure 3e and f). However, Grb2 did not co-localize or enrich with p14 at the plasma membrane when the Grb2 binding site was mutated to FVAI and co-expressed with CA src or treated with pervanadate (Figure 3e and f, and Figure 3—figure supplement 1f and g).

Figure 3. p14 Y116 in the cytoplasmic tail binds to Grb2.

(a) Schematic of p14 mutants that disrupt predicted Grb2 binding motif. (b) Western blot of co-immunoprecipitation of p14 with Grb2 (lane 7) and p14 mutants, Y116F, N118A, FVAI, that does not bind Grb2 (lane 8, 9, 10). (c) Average fusion index of p14 mutants with error bars representing standard deviations from 3 independent transfections of 3 wells each. P-values are two-tailed, two-sample Student’s t-test where *** = p<0.001 (See also Figure 3—figure supplement 1a, (b). (d) Streptavidin beads with biotinylated phosphorylated and non-phosphorylated Y116 p14 cytoplasmic tail peptide encoding (P113-N121) binds and did not bind to purified Grb2 respectively (See also Figure 3—figure supplement 1c, (d). (e) Confocal images of Grb2-GFP (green) enrichment to the plasma membrane of cells co-expressing p14 WT, p14 FVAI mCherry (magenta) and wildtype HEK293T with constitutively active c-src kinase (cyan). (f) Average normalized Grb2-GFP plasma membrane enrichment in cells co-expressing constitutively active c-src and either p14 WT (n = 33 cells) or p14 FVAI (n = 26 cells) or expressing constitively active c-src alone (n = 24 cells). Error bars represent standard deviations from three independent transfsections. P-values are two-tailed, two-sample Student’s t-test where *** = p<0.001 and n.s. = p>0.05.

Figure 3—source data 1. Excel Spreadsheet of plasma membrane enrichment of Grb2-GFP in cells expressing CA-src alone or with p14-WT-mcherry or with p14-FVAI mcherry.

Figure 3.

Figure 3—figure supplement 1. Characterization of Grb2 binding to p14.

Figure 3—figure supplement 1.

(a) Average biotinylated p14 FVAI to that of p14 WT. Error bars represent standard deviations from three independent transfections and biotinylation. P-values are two-tailed two-sample Student’s t-test to p14 WT n.s. = p>0.05. Representative western blot is in Figure 1—figure supplement 1g. (b) p14 mutants, Y116F, N118A, FVAI, are trafficked to the plasma membrane as visualized with TIRF microscopy. (c) Representative field of view of HEK293T cells expressing p14 WT and p14 mutants with p14 labeled with C-terminus mcherry (magenta), nuclei stained with Hoechst 33342 (cyan). (d) Coomaisse stain of purified human Grb2 (right) and labeled with AlexaFluor647 (left). (e) Biotin bead incubated with purified Grb2 has minimal binding. (f) p14-WT-mcherry (magenta) co-expressed with Grb2-GFP (green) in non-treated WT HEK293T cells. (g) Confocal images of Grb2 enrichment to the plasma membrane of cells upon treating with pervanadate with line scans of fluorescence intensity of each protein. Green line denotes the average fluorescence intensity from 20 p14 WT-expressing cells and 14 p14 FVAI expressing cells across 3 independent transfections. Grb2 does not re-localize to plasma membrane for p14 FVAI and HEK293T WT cells (bottom two rows).
Figure 3—figure supplement 1—source data 1. Excel Spreadsheet of surface biotinylation of p14-WT-GFP and p14-FVAI-GFP for Figure 3—figure supplement 1a.
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Having shown that Grb2 binds to the p14 cytoplasmic tail in a phosphorylation-dependent manner, we next sought to determine mechanistically how Grb2, an adaptor protein with two SH3 domains, plays a role in p14-mediated cell-cell fusion. The N-terminal SH3 domain of Grb2 binds to SOS, activating Ras, which in turn activates Raf kinase and the MAPK-ERK1/2 pathway, while the C-terminal SH3 domain of Grb2 binds to the actin nucleation promoting factor N-WASP, which binds to Arp2/3 and nucleates branched actin assembly (Figure 4a). To determine if one or both pathways are important for fusion, we first treated cells expressing p14 with sorafenib tosylate, an inhibitor of Raf kinase, but found no effect on cell-cell fusion at up to 100 times the IC50 (Figure 4b). We next considered whether branched actin networks could be directly involved in p14-mediated cell-cell fusion. Building on previous work showing that cytochalasin D disrupts fusion of p14-expressing cells (Salsman et al., 2008), we treated p14-expressing cells with wiskostatin, an inhibitor of N-WASP. We found that p14 trafficking to the plasma membrane fusion was unperturbed, but cell-cell fusion was significantly reduced (Figure 4b, Figure 4—figure supplement 1a and b). We then treated p14-expressing cells with CK-666 to inhibit the Arp2/3 complex, which nucleates branched actin filaments, and found that fusion was reduced in a dose-dependent manner despite proper trafficking of p14 to the plasma membrane (Figure 4b, Figure 4—figure supplement 1a and b). Interestingly, treating p14-expressing cells with smifH2, an inhibitor of formins, enhanced cell-cell fusion (Figure 4b), perhaps due to increased branched actin assembly that has been observed when formins are broadly inhibited (Burke et al., 2014).

Figure 4. N-WASP-dependent assembly of branched actin network is necessary for cell-cell fusion.

(a) Schematic of Grb2 binding to two potential downstream effectors, SOS and N-WASP (b) Extent of cell-cell fusion quantified with splitYFP fluorescence assay of p14 expressing cells treated sorafenib tosylate targeting Raf kinase, wiskostatin targeting N-WASP, CK-666 targeting Arp2/3 and smifH2 targeting formins, normalized to that of p14 WT treated with vehicle control, DMSO. Error bars indicate standard deviations from 3 independent transfections of 3 wells each. P-values are two-tailed, two-sample Student’s t-test to DMSO where ** = p<0.01, *** = p<0.001 and n.s. = p>0.05. (c) Average percent of p14-expressing cells with 3 or more nuclei in HEK293T WT cells and HEK293T cells overexpressing Grb2 SH2 domain, N-terminus SH2-SH3 mutant and C-terminus SH2-SH3 mutant. P-values are two-tailed, two-sample Student’s t-test where ** = p<0.01, *** = p<0.001 and n.s. = p>0.05. Error bars represent standard deviations from 3 independent transfections (See also Figure 4—figure supplement 1a,b,c,d, and Figure 4—source data 1). (d) Average percent of p14-expressing cells with 2 or more nuclei in N-WASP -/- and +/+ cells with error bars representing standard deviations from 3 independent transfections. P-values are two-tailed, two-sample Student’s t-test where ** = p<0.01 (See also Figure 4—figure supplement 1g). (e) In vitro actin bead motility of phosphorylated p14 cytoplasmic tail peptide conjugated to streptavidin beads in a purified actin motility mixture supplemented with Grb2. Polymerized actin is visualized with AlexaFluor488-labeled utrophin actin binding domain (See also Figure 4—figure supplement 1h).

Figure 4—source data 1. Excel Spreadsheet of counts and distribution for p14-expressing HEK293T cells over-expressing Grb2 mutants for Figure 4c.
Figure 4—source data 2. Excel Spreadsheet of counts and distribution for p14-expressing N-WASP -/- or +/+ mouse embryonic fibroblasts for Figure 4d.

Figure 4.

Figure 4—figure supplement 1. Characterization of over-expression of Grb2 mutants and purified components of actin motility assay.

Figure 4—figure supplement 1.

(a) Western blot of surface biotinylated GFP-tagged p14 WT treated with eitherDMSO, Wiskostatin or CK-666. (b) Average biotinylated p14 WT treated with Wiskostatin and CK-666 normalized to that of DMSO-treated p14 WT. Error bars represent standard deviations from three independent transfections and biotinylation. P-values are two-tailed, two-sample Student’s t-test to p14 WT where n.s. = p>0.05. (c) Representative confocal images of HEK293T cells over-expressing Grb2 mutants tagged with GFP before and after addition of pervanadate to increase p14 phosphorylation. Grb2 mutants are functional, and colocalizes with p14 at the plasma membrane. (d) Western blot of HEK293T cells over-expressing FLAG-tagged Grb2 mutants. (e) Representative field of view of HEK293T cells and HEK293T cells over-expressing Grb2 mutants transfected with p14 WT with p14 labeled with C-terminus mcherry (magenta), nuclei stained with Hoechst 33342 (cyan). (f) Average nuclei count of p14-expressing HEK293T cells and HEK293T overexpressing SH2 domain, N-terminus SH3 and SH2 and C-terminus SH3-SH2 domains of Grb2. P-values are ks-test and error bars represent standard deviation of 3 independent transfections. (g) Average nuclei count of p14-expressing N-WASP -/- and +/+ cells. P-values are ks-test and error bars represent standard deviation from 3 independent transfections. (h) Coomaisse stain of SDS-PAGE of each protein used in vitro actin motility assay.
Figure 4—figure supplement 1—source data 1. Excel Spreadsheet of surface biotinylation of p14-WT-GFP treated with wiskostatin and CK-666 for Figure 4—figure supplement 1b.
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To test whether the reduction in cell-cell fusion with wiskostatin and CK-666 was the result of a direct link between N-WASP and p14 or a more general inhibition of actin activity, we over-expressed Grb2 mutants that can only bind to either SOS or N-WASP by truncating either the N- or C-terminal SH3 domains, respectively. We found that both of these Grb2 mutants bound to p14 WT and co-localized with phosphorylated p14 in pervanadate-treated live cell images (Figure 4—figure supplement 1c), confirming that the mutations did not disrupt interactions with p14. However, when the Grb2 N-terminal mutant that can bind only to SOS (Nterm) was over-expressed, the extent of p14-mediated cell-cell fusion was reduced, similar to when endogenous Grb2 binding was reduced by overexpression of the Grb2 SH2 domain (Figure 4c, Figure 4—figure supplement 1d,e and f). When a Grb2 C-terminal mutant that can bind only to N-WASP (Cterm) was over-expressed, p14-mediated cell-cell fusion was restored to a level comparable to that of endogenous Grb2 in WT cells (Figure 4c, Figure 4—figure supplement 1d,e and f). To further confirm the specificity of N-WASP-nucleated branched actin assembly in p14-mediated cell-cell fusion, we expressed p14 WT in N-WASP null (N-WASP -/-) mouse embryonic fibroblasts. We found that the extent of p14-mediated cell-cell fusion was attenuated in N-WASP -/- cells compared to wild-type mouse embryonic fibroblasts (Figure 4d, Figure 4—figure supplement 1g).

To determine whether N-WASP binding to Grb2 and the p14 cytoplasmic tail is sufficient to nucleate actin assembly, we used an in vitro actin-based motility assay. We bound biotinylated p14 cytoplasmic tail peptides to streptavidin beads in a purified actin motility mixture containing N-WASP (lacking the EVH1 domain), Arp2/3, profilin, cofilin, capping protein and actin, supplemented with Grb2 (Figure 4—figure supplement 1h). When Y116 of the p14 cytoplasmic tail peptide was phosphorylated, actin tails were nucleated from the bead (Figure 4e), but when Y116 was not phosphorylated, actin tails were not observed (Figure 4e). This confirms that Grb2 is necessary and sufficient to recruit N-WASP to the p14 cytoplasmic tail and can nucleate localized branched actin networks when p14 is present.

We next investigated whether branched actin network assembly must be directly coupled to the fusogenic ectodomain, or whether the fusogenic ectodomain can simply be present in the same membrane as actin assembly by the cytoplasmic tail of p14. To test the necessity for direct coupling, we co-expressed the ectodomain deletion mutant (Δecto; ΔM1-T35), which traffics to the plasma membrane and binds Grb2 (Figure 5—figure supplement 1a and b) and the cytoplasmic tail deletion mutant (Δcyto) in the same cell. Interestingly, we found that cell-cell fusion was abolished (Figure 5a, and Figure 5—figure supplement 1c), despite the presence of both halves of p14. This indicates that actin assembly localized to the p14 ectodomain is necessary to drive fusion. This also suggests why native Arp2/3-generated branched actin networks at cell-cell contacts (Efimova and Svitkina, 2018) are not sufficient to cause spontaneous cell-cell fusion. We note that the mechanism described here is in addition to the role of the actin cytoskeleton at E-cadherin-mediated cell-cell adhesion sites (Salsman et al., 2008).

Figure 5. Branched actin assembly directly coupled to p14 cytoplasmic tail drives cell-cell fusion.

(a) Average fusion index of p14 truncation mutants normalized to that of p14 WT. P-values are two-tailed, two-sample Student’s t-test to p14 WT where *** = p<0.001. Error bars indicate standard deviations from 3 independent transfections of 3 wells each (See also Figure 5—figure supplement 1a, (b,c). (b) Schematic of fusion protein coupling actin assembly to p14 cytoplasmic tail consisting of Grb2 SH2 domain and 47 residues from EspFU. (c) Average percent of p14-expressing cell with 3 or more nuclei in HEK293T WT cells and HEK293T cells overexpressing Grb2 SH2 domain and SH2-R47. P-values are two-tailed, two-sample Student’s t-test where ** = p<0.01 and n.s. = p>0.05. Error bars represent standard deviations from 3 independent transfections (See also Figure 5—figure supplement 1d,e,f,g, and Figure 5—source data 1).

Figure 5—source data 1. Excel Spreadsheet of counts and distribution for p14-expressing HEK293T cells over-expressing R47 constructs for Figure 5c.

Figure 5.

Figure 5—figure supplement 1. Characterization of p14 Δectodomain and direct coupling of p14 to actin assembly.

Figure 5—figure supplement 1.

(a) Average biotinylated p14 Δecto to that of p14 WT. Error bars represent standard deviations from three independent transfections and biotinylation. P-values are two-tailed two-sample Student’s t-test to p14 WT where n.s. = p>0.05. Representative western blot is in Figure 1—figure supplement 1g. (b) Western blot of co-immunoprecipitation of p14 Δectodomain and p14 WT with Grb2. (c) Representative field of view of HEK293T cells expressing p14 WT and p14 truncation mutants with p14 labeled with C-terminus mcherry (magenta) and nuclei stained with Hoechst 33342 (cyan). (d) Representative confocal images of HEK293T cells over-expressing SH2-47 residues from EpsFu tagged with GFP and p14 WT mcherry (not shown) before and after addition of pervanadate to phosphorylate p14. (e) Western blot of HEK293T cells over-expressing FLAG-tagged SH2 and SH2-47 residues from EpsF(U). (f) Representative field of view of HEK293T cells and HEK293T cells over-expressing SH2 and SH2-47 residues from EpsF(U) transfected with p14 WT with p14 labeled with C-terminus mcherry (magenta) and nuclei stained with Hoechst 33342 (cyan). (g) Average nuclei count of p14-expressing in HEK293T cells and HEK293T over-expressing SH2 and SH2-R47. P-values are ks-test and error bars represent standard deviation from 3 independent transfections.
Figure 5—figure supplement 1—source data 1. Excel Spreadsheet of surface biotinylation of p14-WT-GFP and p14-Δecto-GFP for Figure 5—figure supplement 1a.

Figure 5—figure supplement 2. Characterization of p14 at time and site of fusion.

Figure 5—figure supplement 2.

(a) Representative fluorescence images of a fusion site 120 s prior to cytoplasmic mixing. Plasma membrane is marked with GPI-anchored pHluorin (green) and p14-WT mCherry in magenta. Dotted white lines mark the fusion site and a reference membrane site where p14-WT mCherry fluorescence intensity is normalized to. (b) Normalized p14-WT mCherry fluorescence intensity at the fusion site 200 s prior to cytoplasmic mixing for 6 fusion events. Bolded line is average normalized intensity and filled in area are the standard deviations. (c) Representative snapshots of a fusion site 90 s prior to cytoplasmic mixing. SH2-GFP (green) and p14-WT mCherry in magenta. Dotted white lines mark the fusion site and a reference membrane site where SH2-GFP fluorescence intensity is normalized to. (d) Normalized SH2-GFP fluorescence intensity at the fusion site 210 s prior to cytoplasmic mixing for 5 fusion events. Bolded line is average normalized intensity and filled in area are the standard deviations.
Figure 5—figure supplement 2—source data 1. Excel Spreadsheet of normalized intensity of p14-WT-mCherry and SH2-GFP at fusion site ~200 s prior to fusion for Figure 5—figure supplement 2b and d.
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If actin assembly directly coupled to p14 is necessary and sufficient for the fusogenic activity of p14, then it should be possible to drive cell-cell fusion by creating an alternate link between p14 and branched actin network assembly. To test this idea, we engineered a fusion protein that binds to p14 consisting of Grb2 SH2 domain and a 47-residue peptide from EspFU of enterohemorrhagic E. coli (EHEC) (Figure 5b). This 47-residue peptide binds to and relieves the auto-inhibition of endogenous N-WASP, nucleating branched actin network (Cheng et al., 2008; Sallee et al., 2008). Fusion of this peptide with the Grb2 SH2 domain, which we confirmed binds to phosphorylated p14 (Figure 5—figure supplement 1d), enables binding to WT p14. When we expressed this fusion protein (SH2-R47) together with p14 in HEK293T, cell-cell fusion was significantly higher than when the SH2 domain lacking R47 was expressed to a similar level (Figure 5c, Figure 5—figure supplement 1e,f and g). This result demonstrates that direct coupling of branched actin assembly to p14 is necessary and sufficient for fusion.

Discussion

This study reveals how a viral pathogen hijacks branched actin network assembly to drive cell-cell fusion, reminiscent of how the pathogens Listeria monocytogenes and vaccinia virus hijack branched actin network assembly to move within and between cells (Welch and Way, 2013). Here we show that the reptilian orthoreovirus fusogen p14 accomplishes this by presenting a c-src kinase substrate in its cytoplasmic tail, binding the host cell adaptor protein Grb2, and initiating branched actin assembly through the host nucleation promoting factor N-WASP. Since p14 ectodomain extends only 0.7–1.5 nm from the plasma membrane (Corcoran et al., 2006), it cannot interact with apposing membrane through the ~2 nm repulsive hydration barrier. If that barrier can be overcome, for example by using divalent cations to bring bare membranes containing negatively charged PS together in vitro, the p14 ectodomain is able to disrupt and mix membranes (Top et al., 2005). However, it is unclear how such close membrane-membrane interaction is achieved on the crowded plasma membrane where the lipid bilayer is sterically blocked by dense membrane proteins.

We propose that the primary role of branched actin network assembly by p14 is to physically push the fusogenic ectodomain into the opposing plasma membrane, a step normally carried out by conformational changes of tall fusogens. In this model, localized force generation by branched actin network assembly around p14’s cytoplasmic domain drives close membrane apposition by coupling to and pushing against the cortical actin cytoskeleton, much like other actin-based membrane protrusions. This close apposition in the presence of p14’s membrane-disruptive ectodomain, which contains hydrophobic residues and myristoylation that are known to be necessary for fusion (Corcoran et al., 2004; Corcoran and Duncan, 2004; Top et al., 2009), may be sufficient to drive cell-cell fusion. Several other roles of the actin cytoskeleton may also be important for p14-bsaed cell-cell fusion. The actin cytoskeleton could promote clustering of p14 ectodomains into sub-diffraction-limited domains (Köster and Mayor, 2016), although enrichment of p14 or phosphorylated p14 was not readily observed at sites of fusion (Figure 5—figure supplement 2a,b,c and d). The actin cytoskeleton could also change local membrane curvature and tension (Kozlov and Chernomordik, 2015), block fusion pore closure, and/or actively expand the fusion pore (Kozlov and Chernomordik, 2015), all of which would promote cell-cell fusion by p14.

Membrane fusion driven by other fusogens has long implicated the actin cytoskeleton (Kondo et al., 2015; Sanderson et al., 1998). The proposed roles of the actin cytoskeleton include regulating the activity, localization and enrichment of fusogens at fusion sites (Yang et al., 2017), regulating protein-protein interactions (Wakimoto et al., 2013), and enlarging the fusion pore (Wurth et al., 2010; Zheng and Chang, 1990). In cases where the putative fusogen remains unidentified or the mechanism of fusion remains unclear, the actin cytoskeleton has been suggested as a key player. Examples include formation of signaling scaffolds and protrusive structures during osteoclast fusion (Oikawa et al., 2012), macrophage fusion (Faust et al., 2019), and myoblast fusion in Drosophila (Kim et al., 2015; Sens et al., 2010). Protrusive membrane structures generated by branched actin assembly have also been observed at fusogenic synapses (Shilagardi et al., 2013) and can be formed in vitro (Liu et al., 2008a; Simon et al., 2019).

Our findings may be relevant to putative fusogens like myomixer, a short extracellular peptide that is required for myoblast fusion (Bi et al., 2018; Leikina et al., 2018; Millay et al., 2013; Quinn et al., 2017; Sampath et al., 2018; Zhang et al., 2017) and does not resemble conformational-change based fusogens. Like p14, myomixer, together with its partner myomaker, could harness the actin cytoskeleton to drive cell-cell fusion by physically forcing a short fusogenic ectodomain through dense cell surface proteins and into contact with an apposing membrane. In this model of cell-cell fusion, local forces generated by the cytoskeleton, rather than by conformational changes in a tall fusogen, provide a powerful tool for cells and pathogens to create multinucleated structures.

Materials and methods

Cloning

Reptilian reovirus membrane fusion protein, p14 (Accession number: Q80FJ1), was synthesized and inserted into mammalian expression vector pcDNA3.1 with C-terminus tags (mcherry, eGFP). Point mutations and truncations were introduced with primers.

splitYFPa and splitYFPb were amplified from pBiFC-bJun-YN155 and pBiFC-bFos-YC155 (a kind gift from Tom Kerppola) and inserted into lentiviral transfer plasmid, pHR, with Gibson assembly.

Constitutively active chick-src kinase was amplified from pLNCX chick src Y527F and inserted into pcDNA 3.1 with linker (GGGS) and C-terminus tags (FLAG and mTagBFP2). pLNCX chick src Y527F was a gift from Joan Brugge (Addgene plasmid # 13660). K295R was introduced to constitutive active chick-src kinase to render it kinase dead with primers. cDNA of Human Grb2 (GE Dharmacon, cloneID: 3345524) was amplified and inserted into pGEX4T2 with a N-terminus GST tag and TEV cleavage site for purification of Grb2. For overexpression of Grb2 mutants, IRES Puromycin was amplified from pLV-EF1a-IRES-Puro (a gift from Tobias Meyer, Addgene plasmid #85132) and inserted into lentiviral pHR backbone to create pHR-IRES-Puro. Grb2 N-terminus SH3 domain and SH2 domain (N-termSH3, 1–159), Grb2 C-terminus SH3 domain and SH2 domain (C-termSH3, 58–217), and Grb2 SH2 domain (58-159) were amplified from cDNA of Human Grb2 and inserted into pHR-IRES-Puro with C terminus FLAG tag.

47 residues from EspFU of enterohemorrhagic E. coli (268–314) was synthesized and inserted with GGGS linker downstream of Grb2 SH2 domain (58-159) and FLAG tag into pHR-IRES-Puro.

Cell culture, transfection and generation of mutant Grb2 overexpression cells

HEK293T cells were obtained from UCSF Cell Culture Facility. SYF (CRL-2459) and SYF + c-src (CRL-2498) cells were obtained from ATCC. N-WASP -/- and +/+ mouse embryonic fibroblasts were a kind gift from Scott Snapper. All cells were grown in DMEM (Life Technologies) supplemented with 10% heat-inactivated FBS (Life Technologies) and 1% Pen-Strep (Life Technologies), at 37°C, 5% CO2. Cells were negative for mycoplasma as verified with Mycoalert mycoplasma detection kit (Lonza).

Cells were transfected with TransIT-293 (Mirus Bio) according to manufacturer’s instructions.

To over-express Grb2 mutants to compete with endogenously expressed Grb2 and SH2 actin nucleators, pHR-Grb2NtermSH3-FLAG-IRES-Puro, pHR-Grb2CtermSH3-FLAG-IRES-Puro, pHR-Grb2SH2-FLAG-IRES-Puro, pHR-SH2-FLAG-R47 were co-transfected with second generation packaging plasmids, pMD2.G and p8.91 in HEK293T to generate lentivirus.

HEK293T cells were transduced with lentivirus, and 24 hr post transduction selected with 3 μg/ml puromycin (Clontech) to select for mutant Grb2 expression. Cultures were maintained in 3 μg/ml puromycin.

splitYFP cell-cell fusion assay pHR-splitYFPa and pHR-splitYFPb were co-transfected with second generation packaging plasmids, pMD2.G and p8.91 in HEK293T to generate lentivirus. WT HEK293T cells were transduced with splitYFPa and splitYFPb lentivirus. The cells were passaged for at least a week before use in cell-cell fusion assay.

To quantify cell-cell fusion, HEK293T cells stably expressing splitYFPa and splitYFPb were mixed at 50:50 ratios and 1.33 × 105 of cells were plated into each well of 48 well plate. The next day, the cells were transfected with TransitIT-293 (Mirus Bio). 18 hr post transfection, cells were moved to 30°C, 5% CO2 incubator to mature the splitYFP fluorophore. 24 hr post transfection, cells were lifted with 150 μl of 2 mM EDTA and placed into 96 well black bottom plate. splitYFP was excited at 510 nm and emission at 530 nm was quantified using a plate reader (Tecan).

Fusion index was quantified as (Isample – Icell) /(Ip14WT – Icell), where Icell is the YFP intensity of non-transfected HEK293T cells expressing splitYFPa and splitYFPb, Isample is the YFP intensity of HEK293T cells transfected with plasmid as specified, Ip14WT is the YFP intensity of HEK293T cells transfected with p14 WT and treated with DMSO as vehicle control. Average and standard deviation of fusion index is calculated from 3 independent transfections of 3 wells each. Statistical significance was determined using two-tailed, two-sample Student’s t-test.

Nuclei count

3.8 × 105 cells were plated into 24 well plates and transfected the next day with designated plasmid with TransIT-293 (Mirus Bio) according to manufacturer’s instructions. 2 hr post transfection, cells were lifted with 150 μl of 2 mM EDTA, re-suspended with 850 μl of media, and 300 μl of cell suspension was plated onto a fibronectin-coated glass bottom chamber (Cell-vis). At 6 hr and 18 hr post transfection, cells were transferred to 30°C, 5% CO2 incubator. After 6 hr incubation at 30°C, nuclei were labeled with 0.6% Hoechst 33342 (Life Technologies), and plasma membrane were labeled with 0.05% CellMask Deep Red (Thermo Fisher Scientific) for 20 min at 37°C. Cells were imaged using spinning disk confocal microscopy. About 80–100 random field of views are taken for each sample to image almost the entire imaging well, and the number of nuclei in p14 expressing cells are manually counted. Average and standard deviation of binned nuclei count is calculated from 3 independent transfections. Statistical significance was determined using two-tailed, two-sample Student’s t-test. Statistical significance between distributions of nuclei in p14-expressing cells were determined using Kolmogorov–Smirnov test (ks-test).

Drug treatment

To broadly inhibit the protein-tyrosine phosphatases, pervanadate is prepared by incubating 10 mM sodium orthovanadate with 0.15% hydrogen peroxide in 20 mM HEPES for 5 min at room temperature. Pervanadate is neutralized with catalase and added to cells immediately. For western blot, cells were lysed 10 mins after pervanadate addition, for live-imaging, cells were imaged immediately after pervanadate addition.

To perturb the actin cytoskeleton, 4 hr post transfection, the media was replaced with complete media supplemented with cytoskeletal drugs CK-666 (Sigma Aldrich), Wiskostatin (Krackeler Scientific) and smifH2 (EMD Millipore) at specified concentrations. DMSO was used as vehicle control. 18 hr post transfection, splitYFP was matured at 30°C. At 24 hr post transfection splitYFP fluorescence was quantified as described above.

To inhibit Raf kinase, 4 hr post transfection, the media was replaced with compete media supplemented with sorafenib tosylate (Selleckchem). DMSO was used as a vehicle control. 18 hr post transfection, splitYFP was matured at 30°C. At 24 hr post transfection splitYFP fluorescence was quantified as described above.

Co-immunoprecipitation

HEK293T were transfected with specified plasmids. 17–24 hr post transfection, HEK293T cells were washed with 1 mM CaCl2/PBS, lifted off the dish with 2 mM EDTA/PBS, pelleted and lysed by incubating in lysis buffer (150 mM NaCl, 25 mM HEPES, 1 mM EDTA, 0.5% NP-40, 1x PhosSTOP phosphatase inhibitor (Roche), 1x HALT protease inhibitor (Thermo Fisher Scientific) for 30 min, and bath sonicated in ice for 3 min. Cell debris was pelleted at 18,000 rcf for 10 min. Cell lysate were precleared with 15 μl of GFP-Trap (Chromotek) for 30 min at 4°C, and incubated with 15 μl of fresh GFP-Trap beads overnight at 4°C. The beads were washed with lysis buffer five times, before boiled in Laemmli sample buffer and separated on 4–20% acrylamide gradient gels by SDS-PAGE. Proteins were transferred onto nitrocellulose membrane and probed with primary antibodies, α-Grb2 (1:5000, Clone 81/Grb2, BD Biosciences), α-tubulin (1:5000, Clone YL1/2, Thermo), α-pTyr (1:2000, Phospho-Tyrosine (P-Tyr-1000) MultiMab Rabbit mAb mix #8954, Cell Signaling Technology), α-GFP (1:10000, Clone 3E6, Life Technologies or 1:5000, A21312, Life Technologies), and secondary antibodies, α-mouse HRP (1:10,000, Upstate Biotechnology or 1:5000, Jackson Labs), α-rabbit HRP (1:5000, 65–6120, Thermo Fisher), α-rat AlexaFluor 647 (1:5000, Life Technologies). Western blots were imaged on ChemiDoc (Bio-Rad).

Membrane fractionation

HEK293T were transfected with p14 WT, p14 Y116F/N118A, and p14 Δcyto. 18 hr post transfection, the cells were washed with 1 mM CaCl2/PBS, lifted off the dish with 2 mM EDTA/PBS. Cells were pelleted at 200 rcf for 5 min and re-suspended in fractionation buffer (20 mM HEPES, 10 mM KCl, MgCl2, 1 mM EDTA, 1 mM EGTA, 1 mM TCEP, 1x HALT protease inhibitor(Thermo Fisher Scientific)). The cell suspension lysed with five freeze/thaw cycles. Nuclei were pelleted via centrifugation (700 rcf, 5 min), and mitochondria were pelleted at 10,000 rcf, 5 min. The supernatant was then centrifuged at 100,000 rcf for an hour at 4°C to separate the membrane and cytoplasmic fraction. The membrane pellet was washed once in fractionation buffer and re-centrifuged at 100,000 rcf for an hour. The cell lysate, cytoplasmic fraction, and membrane pellet was boiled in Laemmli sample buffer, and separated on 4–20% acrylamide gradient gels by SDS-PAGE. Proteins were transferred onto nitrocellulose membrane and probed with primary antibodies, α-tubulin (1:5000, Clone YL1/2, Thermo Fisher Scientific), α-GFP (1:5000, A-21312, Life Technologies), and secondary antibodies, α-rabbit HRP (1:5000, 65–6120, Thermo Fisher) and α-rat AlexaFluor 647 (1:5000, Life Technologies). Western blots were imaged on a ChemiDoc (Bio-Rad).

Surface biotinylation

HEK293T cells were transfected with GFP tagged p14 WT, p14 FVAI, p14 Δcyto, p14 Δecto. P14 WT expressing cells were treated with 20 μM Wiskostatin or 100 μM CK-666 9 hr post transfection. 18–24 hr post transfection, cells were washed with 1 mM CaCl2/PBS on ice, lifted off the dish with ice-cold 2 mM EDTA/PBS. Cells were pelleted at 200 rcf for 5 min and washed with PBS (pH = 8) three times on ice and incubated with 1 mg NHS-Biotin in 0.5 mL PBS (pH = 8) for 1 hr on ice. The biotinylation reaction was quenched with 100 mM glycine in PBS for 10 min at room temperature. Cells were washed three times with 100 mM glycine in PBS and lysed with 1 mL lysis buffer (1x RIPA buffer supplemented with 1x HALT protease inhibitor (Thermo Fisher Scientific)) for 30 min on ice and sonicated for 3 min. Cell debris was pelleted for 5 min at 17,900 rcf at 4°C. 30 μl of streptavidin magnetic beads (ThermoFisher) were washed with RIPA buffer at 4°C, the supernatant was added and incubated overnight. Beads were washed 5 times with RIPA buffer, suspended in Laemmli sample buffer (30 ul), denatured at 95°C for 5 min and separated on 4–20% acrylamide gradient gels by SDS-PAGE. Proteins were transferred onto nitrocellulose membrane with iBlot (Thermo Fisher Scientific). The membrane was blocked with 5% milk powder in TBST for 1 hr and probed with primary antibodies α-GFP (1:5000, g1544, Sigma Aldrich), α-tubulin (1:5000, Clone YL1/2, Thermo) in 5% milk in TBST overnight at 4°C. The membrane was then probed with secondary antibodies, α-rabbit HRP (1:5000, 65–6120, Thermo Fisher) and, α-rat AlexaFluor 647 (1:5000, Life Technologies). Western blots were imaged on ChemiDoc (Bio-Rad).

Protein purification

GST-TEV-Grb2 (human) was expressed and purified from E. coli as previously described (Su et al., 2016).

N-WASP (ΔEVH1) was a kind gift from D. Wong and J. Taunton (University of California, San Francisco). Actin was purified from rabbit skeletal muscle as previously described (Spudich and Watt, 1971). Capping protein was a kind gift from S. Hansen and D. Mullins (University of California, San Francisco). Arp2/3 was purchased from Cytoskeleton, Inc Profilin, cofilin, and utrophin actin binding domain (1-261) were purified as previously described (Bieling et al., 2016; Harris et al., 2019).

Motility assay

Similar to a previously described motility assay (Okrut et al., 2015), 2 µl of 0.5% 3 µm streptavidin polystyrene beads (Bangs Laboratories) are incubated with 1 µM biotin-p14 cytoplasmic tail peptide in 10 mM HEPES (pH 7.5), 1 mg/ml BSA and 50 mM KCl for 10 min at room temperature. Peptide-coated beads are diluted eight-fold into motility buffer (10 mM HEPES, 2 mM MgCl2, 50 mM KCl, 50 mM NaCl, 1 mg/ml BSA, 2.5 mM ATP, 5 mM TCEP), containing 0.1 µM Grb2 (20% labeled), 0.2 µM N-WASP, 9 µM actin, 0.075 µM arp2/3, 0.05 µM capping protein, 2.6 µM profilin, 3.5 µM cofilin and incubated for 15 min at room temperature while rotating. 300 nM utrophin-AF488 is added to the mixture, and incubated for 5 min, before imaging.

In vitro kinase assay

As previously described (Dagliyan et al., 2016), HEK293T is transiently transfected with constitutively active chick src (Y527F) kinase and kinase dead (Y527F/K295R) src kinase with C-terminus FLAG tag with TransIT-293 (Mirus). 24 hr post transfection, cells were washed with 1 mM CaCl2/PBS, and lifted with 2 mM EDTA. Cells were pelleted, and lysed in 20 mM HEPES-KOH, 50 mM KCl, 100 mM NaCl, 1 mM EGTA, 1% NP-40, 1x PhosSTOP phosphatase inhibitor (Roche) and 1x HALT protease inhibitor (Thermo Scientific) for 30 min, 4°C while rotating. Cell debris was pelleted at 3000 g, 10 min, and FLAG-tagged kinase were immunoprecipitated with 3 µg of α-FLAG (M2 clone, Sigma) and 50 µl of Protein-G Dynabeads (Thermo Scientific) for 2 hr, 4°C, while rotating. Beads were washed twice with intracellular buffer (20 mM HEPES-KOH, 50 mM KCl, 100 mM NaCl, 1 mM EDTA, 1% NP-40), and twice with kinase buffer (25 mM HEPES, 5 mM MgCl2, 5 mM MnCl2, 0.5 mM EGTA). Beads were re-suspended in kinase buffer, supplemented with 0.2 mM ATP and 1.5 mM biotin-p14 cytoplasmic tail peptide, and incubated for 1 hr at room temperature. Protein-G dynabeads were removed, and the supernatant is incubated with 10 µl streptavidin magnetic beads (Pierce) for 30 min, room temperature. Streptavidin magnetic beads were washed twice with kinase buffer, and boiled in sample buffer. Sample are dotted onto nitrocellulose membrane (Bio-Rad), and blocked with 5% BSA, and probed with α-pTyr (1:5000, Phospho-Tyrosine (P-Tyr-1000) MultiMab Rabbit mAb mix #8954, Cell Signaling Technology) and α-biotin-AF647 (1:5000, BK-1/39, Santa Cruz Biotechnology) in 5% BSA overnight at 4°C. Blots were washed 3 times, 5 min each with TBST, and probed with secondary antibody, α-rabbit HRP (1:5000, 65–6120, Thermo Fisher) and washed 3 times, 15 min each. Blots were imaged on Chemi-Doc (Bio-rad).

Imaging

All live cells were maintained at 37°C, 5% CO2 with a stage top incubator (okolab) during imaging.

For confocal microscopy, cells were imaged with a spinning disk confocal microscope (Eclipse Ti, Nikon) with a spinning disk (Yokogawa CSU-X, Andor), CMOS camera (Zyla, Andor), and either a 4x objective (Plano Apo, 0.2NA, Nikon) or a 60x objective (Apo TIRF, 1.49NA, oil, Nikon). For total internal reflection fluorescence (TIRF) microscopy, cells were imaged with TIRF microscope (Eclipse Ti, Nikon), 60x objective (Apo TIRF, 1.49NA, oil, Nikon) and EMCCD camera (iXON Ultra, Andor). Both microscopes were controlled with Micro-Manager. Images were analyzed and prepared using ImageJ (National Institutes of Health).

Plasma membrane enrichment of Grb2

HEK293T cells were transfected with either constitutively active (CA) c-src-mTagBFP2 and Grb2-GFP alone, or together and mCherry tagged p14 WT or p14 FVAI. 18–24 hr post transfection, HEK293T cells were imaged with spinning disk confocal microscope with a 60x objective (Apo TIRF, 1.49NA, oil, Nikon). CA c-src BFP localizes to the plasma membrane and was used as a plasma membrane marker. Grb2-GFP fluorescence intensity at the plasma membrane was normalized to its fluorescence intensity at the cytoplasm in cells expressing either CA-src alone or together with p14 WT and p14 FVAI. The average normalized Grb2 enrichment at the plasma membrane from three independent transfections were compared.

Acknowledgements

The authors would like to thank R Duncan and E Chen for helpful discussion and the Fletcher Lab members, including MH Bakalar, BD Belardi, AR Harris, and MD Vahey, for useful feedback and technical consultation.

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Contributor Information

Daniel A Fletcher, Email: fletch@berkeley.edu.

Michael M Kozlov, Tel Aviv University, Israel.

Anna Akhmanova, Utrecht University, Netherlands.

Funding Information

This paper was supported by the following grants:

  • National Institute of General Medical Sciences R01GM114671 to Daniel A Fletcher.

  • Chan Zuckerberg Biohub Investigator Award to Daniel A Fletcher.

  • National Science Foundation DBI-1548297 to Daniel A Fletcher.

  • National Science Foundation GRFP to Ka Man Carmen Chan.

  • Life Sciences Research Foundation to Sungmin Son.

Additional information

Competing interests

No competing interests declared.

Author contributions

Conceptualization, Resources, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing.

Conceptualization, Funding acquisition, Investigation, Methodology, Writing - review and editing.

Conceptualization, Funding acquisition, Writing - review and editing.

Conceptualization, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing.

Additional files

Supplementary file 1. Key resources table.
elife-51358-supp1.docx (34.9KB, docx)
Transparent reporting form

Data availability

All data generated or analysed during this study are included in the manuscript and supporting files.

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Decision letter

Editor: Michael M Kozlov1
Reviewed by: Benjamin Podbilewicz2, Leonid Chernomordik3

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

The work explores mechanisms of cell fusion mediated by the intriguing protein fusogens of the FAST family of reovirus fusion proteins. The authors develop the hypothesis that these proteins promote close contact between cells and membrane fusion by controlling local rearrangements of actin cytoskeleton, and they present several well-conducted experiments in support of this hypothesis. However, additional future experiments, as suggested by one of the reviewers, will be needed to further clarify the mode by which this viral fusogen hijacks the host's cytoskeleton to drive cell-cell fusion.

Acceptance letter:

We include below the comments and reservations of one of the reviewers, which the authors might find useful for their future work and presentation of their results.

Some experiments showing Wb following surface biotinylation show that there is more wildtype p14 on the surface than in the mutants (for example cytotail delletion in Fig 1S1g & h) show lower expression and reduced fusion. If they had reduced the concentration of p14 by using lower DNA concentrations in the transfection or by inducing down regulation it would have been more convincing.

For example, Y116F and tailless mutant appear to have lower amounts of protein on the surface that correlates with reduced activity. Thus, it is not convincing that "trafficking to the plasma membrane was intact" for the different mutants and under the different treatments.

While they show that CK-666 treatments resulted in a dose-dependent reduction in fusion, they did not show what is the surface expression of p14 on the plasma membrane at different dosages. Fig 4S1a shows a reduction in surface expression compared with wt. In addition, it is not clear how they normalized for loading. Using cadherin, integrin or another surface protein could help here.

Response Figure 2B shows a reduction on the percentage of p14 expressing cells.

Response Figure 1A shows lower surface expression for some mutants that had lower fusion. The authors did not have loading controls and the surface expression was not normalized. In summary, using increasing amounts of DNA could have been used to try to correlate surface expression with fusion.

Knock down or knock out experiments: The authors were unable to show that N-WASP can be exchanged by a different actin nucleator. They did not show whether WASP(-/-) affects the surface expression of p14. They could have rescued N-WASP(-/-) instead of comparing with WT MEFs. Experiments on Grb2 shRNA and KO were not conclusive.

Do actin protrusions correlate with p14 localization and fusion?

The authors did try to look for p14 and actin filaments and networks at fusion sites. However, actin protrusions were not observed and the colocalization of p14 with actin fibers and branched actin was not shown to correlate with fusion. It appears that the proteins used in this study are highly overexpressed and the resolution of the colocalization studies is very low. Thus, it is not convincing that in cells p14 colocalizes with actin cytoskeleton and actin binding proteins. The comparison of the results of this study to the Listeria model is not convincing.

Decision letter after peer review:

Thank you for submitting your article "A viral fusogen hijacks the actin cytoskeleton to drive cell-cell fusion" for consideration by eLife. Your article has been reviewed by Anna Akhmanova as the Senior Editor, a Reviewing Editor, and four reviewers. The following individual involved in review of your submission has agreed to reveal his identity: Benjamin Podbilewicz (Reviewer #1).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This study by Ka Man Carmen Chan et al., focused on one of the most intriguing examples of membrane fusion, cell-cell fusion mediated by FAST proteins of non-enveloped reoviruses such as the reptilian orthoreovirus p14. The authors characterized the interactions between p14 and the actin cytoskeleton, showed that c-Src kinase can phosphorylate a specific tyrosine (Y116) on the cytoplasmic tail of p14, which can lead to interaction of this cytotail with the cellular adaptor protein Grb2. They further showed that in vitro this interaction can initiate the interaction with N-WASP nucleation factor that can drive localized assembly of the branched actin networks mediated by Arp2/3 complex.

The central message of the work, according to the current presentation of the results, is a proposal of a novel mechanism by which the polymerizing actin network is crucially involved in the process of membrane fusion. Since p14 is very short compared to the best characterized fusogens, and, therefore, could not reach the target membrane, the authors suggest that the primary role of the branched actin network assembly is to develop a force applied to the p14 ectodomain and push it into a close contact with a neighboring cell.

All the reviewers including the reviewing editor found the study to be elegant, the experiments well designed, and the article well written.

The reviewers agreed that the achievements of the work are:

a convincing demonstration of the p14-regulated assembly of branched actin, dissection of the way p14 regulates this assembly via p14/Grb2/N-WASP interactions, and demonstration of the importance of this assembly for p14-mediated fusion.

At the same time, the reviewers were not convinced that the proposed mechanism for the role of actin polymerization in p14-mediated fusion, which is presented by the authors as the major result of the work, was sufficiently supported by the presented data, and found it largely hypothetical. It is the opinion of the majority of the reviewers that substantiation of the mechanism is crucial for acceptance of the article for publication in eLife.

Essential revisions:

The first group of questions, which have to be addressed, is related to the uniqueness of the proposed mechanism. Specifically, can N-WASP mediated assembly of a branched actin network promote fusion by raising local concentration of p14 at the future fusion site? Can one exclude fusion dependence on branched actin dependent cell-cell adhesion (Efimova and Svitkina, 2018)? Is there any evidence (perhaps in the literature) that fusion depends on interactions between p14 ectodomain and the target membrane rather than on the p14 ectodomain interactions with the p14-expressing membrane? Can one decrease the dependence of p14-mediated fusion on the assembly of the branched actin networks by "shaving" the surface of the target cells with proteases?

The reviewers realize that some of these questions are most likely beyond the scope of this study so that they can be addressed by mentioning alternative interpretations and avoiding the definitive language such as "we demonstrate" while discussing the proposed mechanism.

At the same time, one of the alternative mechanisms has to be ruled out experimentally, namely, the mechanism in which the factor controlling fusion is the local concentration of p14, whereas the role of the polymerizing actin network is to modulate this concentration rather than to develop a pushing force. To address that point the reviewers suggest to:

1) Quantify the surface expression of p14 in wt and mutants under different experimental conditions and demonstrate that the differences in the fusion efficiency do not correlate with those in the p14 concentrations.

Addressing this point is especially important since differences in the number and density of fusogens on the surface can result in different levels of fusion. This has been shown for many fusogens including viral (e.g. Influenza's HA), exoplasmic (e.g. EFF-1) and endoplasmic (e.g. SNAREs). One of the reviewer suggested that the authors do surface biotinylation on ice for all the mutants under different pharmacological conditions (for example in Figure 1F, Figure 1—figure supplement 1B, Figure 2C, Figure 3B,C,E, Figure 4B, Figure 5A).

2) Perform knock down or knock out experiments using siRNAS or CRISPR-Cas9 in order to determine specifically whether Grb2, N-WASP, Arp2/3, Raf kinase and Ras are required for p14-mediated fusion.

3) Perform cell-cell imaging and colocalization of p14, actin and contact sites between membranes before during and after fusion can help to test the model.

The second group of questions is concerned with the physical background of the mechanism by which polymerizing actin could exert a force sufficient to press p14 ectodomains against the target membrane.

As an experimental substantiation of the mechanism the reviewers expect a test where, for example, another nucleator is used, so that the actin network (and, hence, the force) that pushes the protein is changed. In addition, p14 could be made longer, with repetitive motifs, to see whether the distance between the protein ends matters.

On the theoretical level, the way the force pushing p14 is generated is not clear. Indeed polymerizing actin network could push p14 only in case it is impeded from sliding back by some counter-force. For example, in the case of the actin-driven movement of leading edge of a lamellipodium, the counter-force is supposed to be a friction force between the actin gel and the external substrate mediated by cell adhesions. The origin of the counter-force in the proposed mechanism should be discussed.

Moreover, the force could have a direction opposite to that schematized in Figure 5D if actin acts through V shape pulling such that the edges of the V stick out and bind to another membrane (for pushing versus pulling by polymerizing actin see, e.g. (Simon et al., 2019)).

Finally, the possible values of the pushing force have to be estimated and compared to what one needs to push p14 through the membrane gap.

eLife. 2020 May 22;9:e51358. doi: 10.7554/eLife.51358.sa2

Author response

Essential revisions:

The first group of questions, which have to be addressed, is related to the uniqueness of the proposed mechanism. Specifically, can N-WASP mediated assembly of a branched actin network promote fusion by raising local concentration of p14 at the future fusion site?

We agree that the question of whether fusion is driven by raising the local concentration of p14 at the fusion site is a very important one. As described below, we have carried out additional experiments to address this question based on the reviewers’ suggestions. More findings and details of the results are provided below.

Can one exclude fusion dependence on branched actin dependent cell-cell adhesion (Efimova and Svitkina, 2018)?

This is an interesting topic. A prior study found that p14-mediated cell-cell fusion depends on E-cadherin and branched actin assembly associated with E-cadherin (Salsman et al., 2008). However, the ability of p14’s cytoplasmic tail to nucleate branched actin network assembly was not known at the time. Our work demonstrates that branched actin network assembly, mediated by Grb2 binding to the cytoplasmic tail of p14, is essential for robust cell-cell fusion. In particular, we find that p14 FVAI, which cannot bind to Grb2 or nucleate N-WASP-mediated actin assembly, causes HEK293T cells to fuse less readily even though branched actin dependent cell-cell adhesion is intact. We further find that co-expression of p14 truncation mutants, Δecto and Δcyto, is also insufficient to drive cell-cell fusion, indicating that branched actin assembly at the cytoplasmic tail must be directly coupled to the fusogenic ectodomain of p14. This suggests that branched actin assembly at cell-cell adhesion is unable to functionally replace actin assembly that is directly coupled to p14, and we have added this point to the revised manuscript’s discussion. The question of how p14 and cell-cell adhesions might cooperatively drive cell-cell fusion is a very interesting one that we would like to pursue in the future, especially considering our observation that cell-cell fusion is not completely abolished by the p14 FVAI mutant. However, since a putative role for E-cadherin has already been published (Salsman et al., 2008) and a mechanistic analysis of p14-adhesion interactions would take considerable time, we feel this topic would be best addressed in a separate study.

Is there any evidence (perhaps in the literature) that fusion depends on interactions between p14 ectodomain and the target membrane rather than on the p14 ectodomain interactions with the p14-expressing membrane?

Yes, prior in vitro work demonstrated that soluble myristoylated p14 ectodomain is able to disrupt pure liposomes without being anchored in a bilayer as a membrane protein (Corcoran et al., 2004). Full-length p14 embedded in liposomes can also fuse with liposomes lacking p14 if close contact is first established using DOPS and divalent cations (Top et al., 2005). Together, this suggests that p14 ectodomain could interact with either the target membrane or the p14-expressing membrane, however close apposition is required to fuse the two membranes together. We have clarified this point in our revised manuscript.

Can one decrease the dependence of p14-mediated fusion on the assembly of the branched actin networks by "shaving" the surface of the target cells with proteases?

This is a very interesting suggestion that we also investigated. If cell surface proteins represent a steric barrier to cell-cell fusion, shaving the surface of the target cells with proteases would decrease the barrier and possibly decrease p14’s dependence on branched actin network assembly. To test this, we mildly trypsinized HEK293T cells and mixed them with p14-expressing HEK293T cells, and compared the extent of fusion with untreated cells. Unfortunately, the trypsinized cells did not adhere to each other and were unable to form cell-cell contacts needed for p14-mediated cell-cell fusion. This raises the important point discussed above about the role of E-cadherin and suggests a possible role is to adhere cells so that p14-driven cell-cell fusion can occur.

The reviewers realize that some of these questions are most likely beyond the scope of this study so that they can be addressed by mentioning alternative interpretations and avoiding the definitive language such as "we demonstrate" while discussing the proposed mechanism.

Thank you for the comment. We agree and have changed the language in our revised manuscript to be less definitive regarding the proposed mechanism.

At the same time, one of the alternative mechanisms has to be ruled out experimentally, namely, the mechanism in which the factor controlling fusion is the local concentration of p14, whereas the role of the polymerizing actin network is to modulate this concentration rather than to develop a pushing force. To address that point the reviewers suggest to:

1) Quantify the surface expression of p14 in wt and mutants under different experimental conditions and demonstrate that the differences in the fusion efficiency do not correlate with those in the p14 concentrations.

Addressing this point is especially important since differences in the number and density of fusogens on the surface can result in different levels of fusion. This has been shown for many fusogens including viral (e.g. Influenza's HA), exoplasmic (e.g. EFF-1) and endoplasmic (e.g. SNAREs). One of the reviewer suggested that the authors do surface biotinylation on ice for all the mutants under different pharmacological conditions (for example in Figures 1F, Figure 1—figure supplement 1B, Figure 2C, Figure 3B,C,E, Figure 4B, Figure 5A).

Thank you for the helpful suggestions of experiments. To test whether the extent of fusion we measured were correlated with differences in surface expression of p14 WT, p14 mutants and in p14 WT-expressing cells treated with wiskostatin and CK-666, we biotinylated cells expressing the p14 mutants with a cell-impermeable reagent (NHS-Biotin) on ice to limit endocytosis. We then immunoprecipitated biotinylated surface molecules with streptavidin beads and probed for p14 with α-GFP antibody (Author response image 1A). We calculated the ratio of intensity of bands eluted from the streptavidin beads to the intensity of the lysate. We then normalized this ratio for either cells expressing p14 WT or non-treated cells. While there is some variation, we found no statistically significant differences between surface expression of p14 FVAI, p14 Δcyto, p14 Δecto with that of p14 WT (Author response image 1B). We similarly found no statistically significant differences in surface expression of p14 in cells treated with wiskostatin or CK-666, and no trend that correlates with fusion efficiency (Author response image 1B). We conclude that the changes in fusion efficiency that we report are not due to changes in surface concentration of p14. We have added this in the revised manuscript as part of Figure 1—figure supplement 1H, Figure 3 —figure supplement 1A, Figure 4 —figure supplement 1A and B, Figure 5 —figure supplement 1A.

Author response image 1. Surface biotinylation of p14, p14 mutants and cells treated with cytoskeletal drugs.

Author response image 1.

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(a) Representative western blot probed with α-GFP of lysate and streptavidin eluate of surface biotinylated cells expressing p14 WT, p14 FVAI, p14 Δcyto, p14 Δecto, and p14 WT expressing cells treated with Wiskostatin and CK-666. (b) Ratio of streptavidin eluate to lysate of p14 FVAI, p14 Δcyto, p14 Δecto, and p14 WT expressing cells treated with Wiskostatin and CK-666 normalizd to that of p14 WT. Average and standard deviations from three independent transfections and blots shown. P-values are two-tailed, two-sample Student’s t-test between condition and p14 WT n.s p>0.05.

2) Perform knock down or knock out experiments using siRNAS or CRISPR-Cas9 in order to determine specifically whether Grb2, N-WASP, Arp2/3, Raf kinase and Ras are required for p14-mediated fusion.

We thank the reviewers for the suggested experiments. The results of our new experiments are presented below.

Quantification of p14-mediated cell-cell fusion in Grb2 KO and KD cells

In this work, we over-express the SH2 domain (58-159) of Grb2 to compete with endogenous Grb2 binding to p14. The SH2 domain alone does not bind to either downstream effectors, N-WASP or SOS, and is used in a dominant negative manner. Prior work have demonstrated the specificity of the SH2 domain of Grb2 for ligands (Jadwin et al., 2016; Kessels et al., 2002; Songyang et al., 1993). Moreover, in our system, the SH2 domain is quite specific to the p14 phosphorylation-dependent motif YVNI. To investigate the specificity of SH2 domain for p14 YVNI motif, we used a tyrosine phosphatase inhibitor, pervanadate, to broadly increase tyrosine phosphorylation (Figure 3—figure supplement 1G). In cells expressing p14 WT and treated with pervanadate, Grb2 enriches readily at the plasma membrane. However in cells treated with pervanadate, Grb2 does not enrich at the plasma membrane of wildtype HEK293T cells or in p14 FVAI-expressing cells (Figure 3—figure supplement 1G). Hence, over-expression of the SH2 domain likely specifically competes with Grb2.

As suggested by reviewers, we knocked-down Grb2 in HEK293T cells using shRNA (Author response image 2A). Surprisingly, these knocked-down cells expressing p14 still readily fused (Author response image 2B). We hypothesize that while Grb2 expression is knocked-down in the population overall, a fraction of individual cells still express Grb2 at sufficient levels to drive fusion. As p14-expressing cells fuse with neighboring cells, the cytoplasm mixes, re-introducing Grb2.

Author response image 2. Extent of p14-mediated cell-cell fusion in Grb2 KD and KO cells.

Author response image 2.

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(a) Western blot probed with α-Grb2 of HEK293T with Grb2 knockdown. (b) Extent of p14-mediated cell-cell fusion in Grb2 KD and wild-type HEK293T cells. (c) Western blot probed with α-Grb2 of clone 1 and clone 2 of HEK293T with Grb2 knocked-out using Crispr-Cas9. (d) Western blot probed with α-Grb2 of clone 3 of HEK293T with Grb2 knocked-out using Crispr-Cas9. (e) Representative confocal images of wild-type HEK293T cells, KO clone 1, KO clone 2, KO clone 3 (magenta). Nuclei were stained with Hoechst 33342 (cyan).

To overcome this, we knocked-out Grb2 using CRISPR-Cas9, and characterized 3 clones (Author response image 2C and D). Expression of p14 in two of these clones (clone 1 and clone 2) had minimal cell-cell fusion (Author response image 2E). Surprisingly, expression of p14 in the other Grb2 knock-out clone (clone 3) fused robustly, independent of Grb2 (Author response image 2D and E). Clone 3 might fuse robustly due to various reasons, including (i) increased adhesion between cells resulting in closer apposition of the bilayers than WT cells, (ii) decreased fusion energy barrier resulting in a lowered dependence on actin assembly, or (iii) upregulation of a compensatory pathway. We are interested in investigating the biophysical and molecular basis that enable this clone to fuse in a Grb2-independent manner. However, we think this would be best pursued in a future study. For this revised manuscript, we have made it clear that there are likely additional mechanisms contributing to p14-mediated cell-cell fusion.

Quantification of p14-mediated cell-cell fusion in N-WASP null cells.

To determine if N-WASP is specifically involved in p14-mediated cell-cell fusion, we expressed p14-WT in N-WASP null mouse embryonic fibroblasts (a gift of Scott Snapper). Since this is a different cell type than our other experiments, we compared the extent of cell-cell fusion to that of p14 expressed in wild-type mouse embryonic fibroblasts (MEFs). The overall transfection efficiency in MEFs was much lower than that of HEK293T, and the extent of cell-cell fusion was also lower. We quantified the extent of fusion as a percent of p14-expressing cells with 2 or more nuclei. When transfected with p14-WT-mCherry, cell-cell fusion was significantly decreased in N-WASP null MEFs than in the control MEFs. We have included this data as Figure 4D of the revised manuscript.

Quantification of p14-mediated cell-cell fusion in SYF (src, yes, fyn kinase) null cells

Since inhibition of Raf kinase with sorafenib had no effect on p14-mediated fusion, and ourin vitro kinase results demonstrated that c-src kinase could phosphorylate Y116 in p14 cytoplasmic tail, we investigated the specificity of c-src kinase for Y116 in vivo. We compared the extent of cell-cell fusion in mouse embryonic fibroblasts lacking in c-src, Yes, Fyn kinase (SYF) and SYF + c-src cells expressing p14. Similar to the MEFs, the overall transfection efficiency in SYF cells were much lower than that of HEK293T, hence we quantified the extent of fusion as percent of p14-expressing nuclei with more than 2 nuclei. When transfected with p14-WT-mCherry, cell-cell fusion was more significantly decreased in SYF cells compared to SYF + c-src. We have included this data as Figure 2G of the revised manuscript.

3) Perform cell-cell imaging and colocalization of p14, actin and contact sites between membranes before during and after fusion can help to test the model.

We thank the reviewers for the suggested experiments. The results of our new experiments are presented below.

Live-cell imaging of p14 at fusion sites before, during and after fusion

We quantified the local concentration of p14 at sites of fusion using confocal microscopy in order to explore whether local concentration changes of p14 correlate with fusion. To visualize the plasma membrane we expressed a GPI-anchored pHluorin in p14-mcherry expressing cells and imaged every 20 seconds. Fusion sites were identified when a fusion pore expands and the plasma membrane peels back. We designated time of fusion as when the plasma membranes of two cells undergo full fusion and their cytoplasms mix. To identify the time when this diffraction-limited fusion pore opens, we used the cytoplasmic mCherry fluorophore from cleaved p14 as a marker of when the cytoplasm mixes between p14-mcherry expressing cell and a neighboring naive cell (Figure 5—figure supplement 5A). We then quantified the fluorescence intensity of p14-mcherry at the fusion site 200 seconds prior to cytoplasmic mixing and normalized it to the fluorescence intensity of p14-mcherry at another location on the plasma membrane. The normalized fluorescence intensity of p14-mcherry from 6 fusion events fluctuated around 1.0-1.1, indicating that p14-mcherry does not enrich at times and at sites of fusion, at least on the length scales resolvable by confocal microscopy (Figure 5—figure supplement 5B).

To test whether phosphorylated p14, in particular, might be locally enhanced during fusion, we expressed the GFP-tagged SH2 domain of Grb2 under a low-expressing promoter, the UBC promoter. We verified that this SH2 domain binds to phosphorylated p14 by treating p14-expressing cells with a broad tyrosine-phosphatase inhibitor, pervanadate. The GFP-tagged SH2 domain enriched at the plasma membrane, where p14 is localized to (Author response image 3). Using the cleaved mCherry fluorophore to again identify fusion pore opening, we quantified the fluorescence intensity of GFP-tagged SH2 domains at the fusion site 210 seconds prior to cytoplasmic mixing (Figure 5—figure supplement 5D). The normalized fluorescence intensity of SH2 domain from 5 fusion events also fluctuated around 1.0-1.1, indicating that phosphorylated p14 does not enrich at times and sites of fusion, at least on the length scales resolvable by confocal microscopy (Figure 5—figure supplement 5E). On average, we observed only a 10% increase in total and phosphorylated p14-mCherry, at times and sites of fusion. This is in contrast to previously reported plasma membrane enrichment, vesicle-docking and puncta-like structures of cell-cell fusogens, such as Eff-1 and Hap2, at fusion sites (Y. Liu et al., 2015; Neumann et al., 2015; Smurova and Podbilewicz, 2016; Y. Yang et al., 2017). This suggests that the p14-actin mechanism described in this manuscript does not require increased local concentration of p14 at fusion sites. We have included these results as Figure 5—figure supplement 2.

Author response image 3. Time-lapse imaging of p14 at sites of fusion using confocal microscopy.

Author response image 3.

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SH2-GFP (green) re-localizes to the plasma membrane when co-expressed with p14-WT mCherry (not shown) and treated with pervanadate to increase phosphorylation of p14.

Live-cell imaging of p14 and actin at fusion sites before, during and after fusion.

We carried out live imaging of cells expressing p14-WT-mCherry and actin with Lifeact-GFP to observe if there was co-localization of p14 with actin structures that would be expected from our proposed mechanism. We were able to capture 7 fusion events. To quantify p14 and actin colocalization, we measured the fluorescence intensity of Lifeact-GFP at the sites of fusion 210 seconds prior to cytoplasmic mixing and normalized. We were able to identify the site of fusion by determining where the fusion pore expands and the plasma membrane is pulled back.

For 6 of the fusion events, we did not observe actin structures at sites and times of fusion that can be readily identified as distinct from cortical actin structures of wild-type HEK293T cells (Author response image 4A and B). For 1 of the fusion events, we were able to capture increased local actin intensity at the site and time of fusion (Author response image 4B and C). However, we are unable to rule out that this was merely coincidental co-localization of fusion with endogenous actin assembly unrelated to fusion.

Author response image 4. Time-lapse imaging of p14 and actin at times and sites of fusion.

Author response image 4.

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(a) Representative snapshots of a fusion site 90 sec prior to cytoplasmic mixing. Actin is visualized with Lifeact-GFP(green) and p14-WT mCherry in magenta. Boxed region marks the fusion site, where the average lifeact-GFP fluorescence intensity is quantified and normalized to the fluorescence intensity at a reference site on the plasma membrane. Representative fluorescent images of a fusion site 90 sec prior to cytoplasmic mixing. Actin is visualized with Lifeact-GFP(green) and p14-WT mCherry in magenta. Boxed region marks the fusion site, where the average lifeact-GFP fluorescence intensity is quantified and normalized to the fluorescence intensity at a reference site on the plasma membrane. (b) Normalized lifeact-GFP fluorescence intensity at the fusion site 210 sec prior to cytoplasmic mixing for 7 fusion events. Bolded line is average normalized intensity and filled in areas are the standard deviations. (c) Fluorescent images of a fusion site 90 sec prior to cytoplasmic mixing where Lifeact-GFP fluorescence intensity is enriched at the fusion site.

Given ourin vitroevidence that the cytoplasmic tail of p14 can nucleate actin assembly, we were puzzled by the lack of co-localization of prominent actin structures and p14. However, we previously found that p14 Y116 was likely minimally phosphorylated (Figure 3—figure supplement F), suggesting that any nucleated actin structure was likely small, transient and difficult to distinguish from the endogenous cortical actin structures of the cells. Taken together, these experiments suggest that actin nucleation by p14, which must be phosphorylated at Y116, are likely sub-diffraction limited and transient, making them very difficult to identify separately from the endogenous actin structures.

The second group of questions is concerned with the physical background of the mechanism by which polymerizing actin could exert a force sufficient to press p14 ectodomains against the target membrane.

As an experimental substantiation of the mechanism the reviewers expect a test where, for example, another nucleator is used, so that the actin network (and, hence, the force) that pushes the protein is changed. In addition, p14 could be made longer, with repetitive motifs, to see whether the distance between the protein ends matters.

As suggested by reviewers, we experimentally changed the actin nucleator associated with p14. Branched actin assembly can exert up to 5 nN or up to 1250 nN/μm2 (Bieling et al., 2016; Mueller et al., 2017; Parekh et al., 2005; Prass et al., 2006), while formin-nucleated filopodia can exert up to 10 pN (Cojoc et al., 2007). We chose to test whether a constitutively active formin, made by truncating the wGBD domain of mDia2, could be used in place of N-WASP to drive p14-mediated fusion. We over-expressed ΔGBD-mDia2 as a fusion protein with the Grb2 SH2 domain to bind to phosphorylated p14. When transiently expressed in cells with p14 WT, we observed filopodia-like structures consistent with previously reported mDia2 nucleation activity (Author response image 5) (C. Yang et al., 2007). To compete with endogenous Grb2 across the population of cells involved in fusion, we stably expressed and selected cells expressing this construct. Unfortunately, cells that survived selection expressed a truncated, functionally null protein that did not exhibit the filopodia-rich phenotype seen in the transient transfection. Due to the competition with endogenous Grb2 and minimal expression of the full-length SH2-nucleator, we were unable to conclusively show that swapping N-WASP for a different actin nucleator preserves p14-mediated fusion.

Author response image 5. Confocal image of HEK293T cell over-expressing SH2-GFP-ΔGBD-mDia2 and p14-WT-mCherry.

Author response image 5.

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HEK293T cell expressing SH2-GFP- ΔGBD-mDia2 (green) and p14-WT-mCherry (magenta) with filopodia-like protrusions denoted with white arrows.

On the theoretical level, the way the force pushing p14 is generated is not clear. Indeed polymerizing actin network could push p14 only in case it is impeded from sliding back by some counter-force. For example, in the case of the actin-driven movement of leading edge of a lamellipodium, the counter-force is supposed to be a friction force between the actin gel and the external substrate mediated by cell adhesions. The origin of the counter-force in the proposed mechanism should be discussed.

Moreover, the force could have a direction opposite to that schematized in Figure 5D if actin acts through V shape pulling such that the edges of the V stick out and bind to another membrane (for pushing versus pulling by polymerizing actin see, e.g. (Simon et al., 2019)).

This is a good point, as simply the assembly of a branched network doesn’t guarantee formation of a protrusion. From the in vitro assay of Simon et al., 2019, based on actin polymerization from the surface of a GUV, the ability to deform the membrane is dependent on membrane tension. We have also seen similar spike-like membrane protrusions driven by branched actin networks assembled on the surface of GUVs in our own in vitro work (A. P. Liu et al., 2008). This model of membrane protrusions driven by local actin assembly that is coupled to cortical actin has been proposed for other WASP-dependent membrane protrusions in fusogenic synapses (Sens et al., 2010) and T-cell protrusions (Tamzalit et al., 2019). We hypothesize that p14-mediated actin polymerization will be coupled to and countered by the stiff cortical actin underneath the network, providing a counter-force that allows it to generate local membrane protrusions. However, it is possible that p14 will, on occasion, not form branched actin networks that are strongly coupled to the cortex, and we expect that these p14 will not be successful in driving cell-cell fusion. To clarify this, we have updated the revised manuscript to emphasize that the direction of force in our model is a hypothesis.

Finally, the possible values of the pushing force have to be estimated and compared to what one needs to push p14 through the membrane gap.

This is a good suggestion to see if the forces generated by p14 could be enough to overcome the membrane gap. First, to estimate the maximum force that p14 can generate, we use the measured stall force of branched actin assembly. We previously measured the force generated by Arp2/3 branched actin networks nucleated by WASP-family NPF in vitro and found the stall force to be approximately 1250 pN/µm2 (Bieling et al., 2016; Parekh et al., 2005). This is consistent with the stall force measured for a migrating keratocye (Mueller et al., 2017; Prass et al., 2006).

Next, we estimated the force that would be required to expose a patch of bare lipid bilayer at the site of fusion. Membrane proteins on a cell surface exert in-plane pressure due to protein-protein crowding, which we previously found to be sufficient to drive membrane tubulation (Stachowiak et al., 2012). In order for membranes to make close contact for fusion, these membrane proteins must be excluded. Assuming that proteins on a cell surface cover between 0.2 to 0.3 of the surface area and the projected area of average protein is 100 nm2, the in-plane pressure exerted by crowded proteins is between 3100 kT/µm2 to 6300 kT/µm2, based on a 2D Carnahan-Starling equation of state for hard discs (Stachowiak et al., 2012).

For an actin protrusion to successfully bring one membrane in contact with the other, the force generated by an actin-based protrusion must do enough work to overcome the in-plane pressure calculated above. Assuming an initial intermembrane distance of 20 nm (accounting for two cell surfaces with average cell surface protein heights of ~10 nm) and that the actin protrusion is pushing directly on the patch of membrane that must be cleared for fusion to take place at 500-1250 pN/ µm2, we estimate the work done by actin is between 2400 kT/µm2 to 6000 kT/µm2. This simple calculation suggests that branched actin network can provide a force that is sufficient to segregate proteins out of the membrane contact site.

Associated Data

    This section collects any data citations, data availability statements, or supplementary materials included in this article.

    Supplementary Materials

    Figure 1—source data 1. Excel Spreadsheet of counts and distribution for p14-expressing HEK293T cells at 12 hr and 24 hr post transfection for Figure 1c and d.
    elife-51358-fig1-data1.xlsx (24.2KB, xlsx)
    Figure 1—figure supplement 1—source data 1. Excel Spreadsheet of surface biotinylation of p14-WT-GFP and p14-Δcyto-GFP for Figure 1—figure supplement 1h.
    elife-51358-fig1-figsupp1-data1.xlsx (11.1KB, xlsx)
    Figure 2—source data 1. Excel Spreadsheet of counts for p14-expressing SYF and SYF+c-src cells for Figure 2g.
    elife-51358-fig2-data1.xlsx (12.1KB, xlsx)
    Figure 3—source data 1. Excel Spreadsheet of plasma membrane enrichment of Grb2-GFP in cells expressing CA-src alone or with p14-WT-mcherry or with p14-FVAI mcherry.
    elife-51358-fig3-data1.xlsx (13.2KB, xlsx)
    Figure 3—figure supplement 1—source data 1. Excel Spreadsheet of surface biotinylation of p14-WT-GFP and p14-FVAI-GFP for Figure 3—figure supplement 1a.
    elife-51358-fig3-figsupp1-data1.xlsx (9.5KB, xlsx)
    Figure 4—source data 1. Excel Spreadsheet of counts and distribution for p14-expressing HEK293T cells over-expressing Grb2 mutants for Figure 4c.
    elife-51358-fig4-data1.xlsx (79.3KB, xlsx)
    Figure 4—source data 2. Excel Spreadsheet of counts and distribution for p14-expressing N-WASP -/- or +/+ mouse embryonic fibroblasts for Figure 4d.
    elife-51358-fig4-data2.xlsx (342KB, xlsx)
    Figure 4—figure supplement 1—source data 1. Excel Spreadsheet of surface biotinylation of p14-WT-GFP treated with wiskostatin and CK-666 for Figure 4—figure supplement 1b.
    elife-51358-fig4-figsupp1-data1.xlsx (9.8KB, xlsx)
    Figure 5—source data 1. Excel Spreadsheet of counts and distribution for p14-expressing HEK293T cells over-expressing R47 constructs for Figure 5c.
    elife-51358-fig5-data1.xlsx (82KB, xlsx)
    Figure 5—figure supplement 1—source data 1. Excel Spreadsheet of surface biotinylation of p14-WT-GFP and p14-Δecto-GFP for Figure 5—figure supplement 1a.
    elife-51358-fig5-figsupp1-data1.xlsx (9.5KB, xlsx)
    Figure 5—figure supplement 2—source data 1. Excel Spreadsheet of normalized intensity of p14-WT-mCherry and SH2-GFP at fusion site ~200 s prior to fusion for Figure 5—figure supplement 2b and d.
    elife-51358-fig5-figsupp2-data1.xlsx (13.2KB, xlsx)
    Supplementary file 1. Key resources table.
    elife-51358-supp1.docx (34.9KB, docx)
    Transparent reporting form
    elife-51358-transrepform.docx (244.9KB, docx)

    Data Availability Statement

    All data generated or analysed during this study are included in the manuscript and supporting files.

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