Liposome reconstitution of a minimal protein-mediated membrane fusion machine

Article4 August 2005free access Liposome reconstitution of a minimal protein-mediated membrane fusion machine Deniz Top Deniz Top Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Roberto de Antueno Roberto de Antueno Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Jayme Salsman Jayme Salsman Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Jennifer Corcoran Jennifer Corcoran Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, CanadaPresent address: Department of Medical Microbiology and Immunology, Heritage Medical Research Building, University of Alberta, Edmonton, AB, Canada T6G 2S2 Search for more papers by this author Jamie Mader Jamie Mader Department of Pathology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author David Hoskin David Hoskin Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Department of Pathology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Ahmed Touhami Ahmed Touhami Department of Physics, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Manfred H Jericho Manfred H Jericho Department of Physics, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Roy Duncan Corresponding Author Roy Duncan Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Deniz Top Deniz Top Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Roberto de Antueno Roberto de Antueno Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Jayme Salsman Jayme Salsman Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Jennifer Corcoran Jennifer Corcoran Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, CanadaPresent address: Department of Medical Microbiology and Immunology, Heritage Medical Research Building, University of Alberta, Edmonton, AB, Canada T6G 2S2 Search for more papers by this author Jamie Mader Jamie Mader Department of Pathology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author David Hoskin David Hoskin Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Department of Pathology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Ahmed Touhami Ahmed Touhami Department of Physics, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Manfred H Jericho Manfred H Jericho Department of Physics, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Roy Duncan Corresponding Author Roy Duncan Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada Search for more papers by this author Author Information Deniz Top1, Roberto de Antueno1, Jayme Salsman1, Jennifer Corcoran1, Jamie Mader2, David Hoskin1,2, Ahmed Touhami3, Manfred H Jericho3 and Roy Duncan 1 1Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada 2Department of Pathology, Dalhousie University, Halifax, Nova Scotia, Canada 3Department of Physics, Dalhousie University, Halifax, Nova Scotia, Canada *Corresponding author. Department of Microbiology and Immunology, Faculty of Medicine, Dalhousie University, Halifax, Nova Scotia, Canada B3H 1X5. Tel.: +1 902 494 6770; Fax: +1 902 494 5125; E-mail: [email protected] The EMBO Journal (2005)24:2980-2988https://doi.org/10.1038/sj.emboj.7600767 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Biological membrane fusion is dependent on protein catalysts to mediate localized restructuring of lipid bilayers. A central theme in current models of protein-mediated membrane fusion involves the sequential refolding of complex homomeric or heteromeric protein fusion machines. The structural features of a new family of fusion-associated small transmembrane (FAST) proteins appear incompatible with existing models of membrane fusion protein function. While the FAST proteins function to induce efficient cell–cell fusion when expressed in transfected cells, it was unclear whether they function on their own to mediate membrane fusion or are dependent on cellular protein cofactors. Using proteoliposomes containing the purified p14 FAST protein of reptilian reovirus, we now show via liposome–cell and liposome–liposome fusion assays that p14 is both necessary and sufficient for membrane fusion. Stoichiometric and kinetic analyses suggest that the relative efficiency of p14-mediated membrane fusion rivals that of the more complex cellular and viral fusion proteins, making the FAST proteins the simplest known membrane fusion machines. Introduction Biological membrane fusion requires protein catalysts to overcome the thermodynamic barriers that prevent spontaneous merger of membranes. Studies of viral and cellular fusion proteins have provided detailed structural and functional insights into the protein complexes that mediate membrane merger (Skehel and Wiley, 2000; Ungar and Hughson, 2003). In the case of enveloped viruses, the fusion catalyst is provided by multimeric glycoproteins that exist as metastable structures embedded in the virus envelope (Earp et al, 2005). Following structural rearrangements, a previously sequestered hydrophobic fusion peptide motif is extended toward, and inserts into, the target cell membrane, anchoring the viral fusion protein in both membranes (Tamm et al, 2002). In contrast, intracellular fusion of transport vesicles is dependent on the pairing of a preformed vesicle (v)-SNARE (soluble N-ethylmaleimide-sensitive fusion protein attachment protein receptor) complex with a cognate target membrane (t)-SNARE to form a trans-acting heteromeric complex (Ungar and Hughson, 2003). In spite of extensive structural diversity in these fusion protein complexes, there are striking similarities in the structural rearrangements that accompany the fusion reaction (Skehel and Wiley, 1998; Weber et al, 1998). However, a clear understanding of the energetics and lipid rearrangements involved in protein-mediated membrane fusion remains incomplete. A new family of viral proteins involved in cell–cell membrane fusion has recently been described, whose features are incongruent with existing models of protein-mediated membrane fusion. The fusion-associated small transmembrane (FAST) proteins are encoded by the fusogenic reoviruses, an unusual group of nonenveloped viruses that induce syncytium formation (Duncan et al, 2004). The FAST protein family is comprised of the p10 proteins of avian reovirus and Nelson Bay reovirus, the p14 protein of reptilian reovirus, and the p15 protein of baboon reovirus (Shmulevitz and Duncan, 2000; Dawe and Duncan, 2002; Corcoran and Duncan, 2004). Unlike enveloped virus fusion proteins, the FAST proteins are nonstructural viral proteins and are not involved in virus entry. The sole defined function of the FAST proteins is the induction of cell–cell fusion following their expression in reovirus-infected cells and trafficking through the ER–Golgi pathway to the plasma membrane (Shmulevitz et al, 2004a). Structural and functional properties of the FAST proteins distinguish them from both enveloped virus fusion proteins and SNARE proteins, although they share certain features with each of these distinct groups of membrane fusion proteins. The FAST proteins assume a bitopic Nexoplasmic/Ccytoplasmic (Nexo/Ccyt) membrane topology with small, approximately equal-sized ecto- and endodomains (Shmulevitz and Duncan, 2000; Corcoran and Duncan, 2004; Dawe et al, 2005). At 95–140 residues in size, the FAST proteins are more similar in size to SNARE proteins than enveloped virus fusion proteins. They also function, in effect, as 'cellular' fusion proteins mediating cell–cell rather than virus–cell membrane fusion. However, the FAST proteins lack the heptad repeat sequences responsible for formation of the helical bundles characteristic of the SNARE proteins and of the class I enveloped virus fusion proteins (Skehel and Wiley, 1998). Similarly, the p10 and p14 FAST proteins contain short regions (18–20 residues) of moderate hydrophobicity in their ectodomains that may function in an analogous manner as the enveloped virus fusion peptide or fusion-loop motifs, serving to destabilize lipid bilayers (Tamm et al, 2003; Gibbons et al, 2004; Modis et al, 2004). However, the fusion peptide motifs of the p10 and p14 FAST proteins are considerably less hydrophobic than typical fusion peptides, they comprise approximately half of the ectodomains of these proteins, and the p14 fusion peptide motif has an obligate requirement for myristoylation (Corcoran et al, 2004; Shmulevitz et al, 2004b). The unusual structural features of the FAST proteins are difficult to reconcile with existing models of protein-mediated membrane fusion, all of which invoke significant energy input derived from extensive rearrangement of homo- or heteromeric complexes (Kozlov and Chernomordik, 1998; Bentz, 2000; Bentz and Mittal, 2003). Expression of individual FAST proteins in transfected cells induces efficient syncytium formation in different cell types. In addition, numerous point mutations eliminate cell–cell fusion activity without altering FAST protein topology or trafficking, clearly indicating that the FAST proteins are intimately involved in the membrane fusion reaction and are the only reovirus proteins required for syncytium formation (Shmulevitz and Duncan, 2000; Shmulevitz et al, 2003; Corcoran et al, 2004). The remarkable physical attributes of the FAST protein family, however, questioned whether the FAST proteins are membrane fusion proteins per se and can function by themselves to induce membrane fusion, or whether they function through unidentified cellular fusion factors that serve as the actual membrane fusion complex. Using proteoliposomes containing the purified p14 FAST protein, we now show that p14 is both necessary and sufficient for membrane fusion, and that the activity of this minimal protein-mediated membrane fusion machine rivals that of the more complex viral and cellular fusion complexes. Results p14 does not form homomeric trans-acting complexes The p14 FAST protein of reptilian reovirus (Figure 1) has an ∼36-residue ectodomain containing an N-terminal myristate moiety and a 20-residue moderately hydrophobic patch that includes a loop structure (residues 5–13). Both the myristate moiety and the hydrophobic patch are essential for p14 membrane fusion activity (Corcoran et al, 2004). The 68-residue endodomain contains a membrane-proximal polybasic region (nine residues of 18 in this region are basic) of no known function and a nonessential C-proximal polyproline motif. To determine whether p14 must be present in both membranes undergoing fusion, p14-transfected cells were monitored for fusion to nontransfected target cells using a fluorescent heterotypic cell–cell fusion assay. As indicated by the presence of numerous large syncytia, all of which contained both donor (red) and target (blue) cell nuclei (Supplementary Figure 1), p14 does not need to form trans-acting homomeric complexes to effect membrane fusion. Figure 1.Structural motifs in the p14 FAST protein. The 125-residue p14 protein is depicted in its functional Nexo/Ccyt membrane topology. Structural motifs include an N-terminal myristate (open triangle) that may interact with the external leaflet of the bilayer, a moderately hydrophobic patch (shaded rectangle) that shares certain features with fusion peptide motifs, a transmembrane domain (black rectangle), a membrane-proximal polybasic region (hatched rectangle), and a C-proximal polyproline motif (open rectangle). Download figure Download PowerPoint Reconstitution of p14-proteoliposomes Baculovirus-expressed p14 was purified to near homogeneity in the presence of n-octyl β-D-glucopyranoside (OG) (Supplementary Figure 2). The purified protein was reconstituted into large (400 nm diameter) unilamellar vesicles (LUVs) by mixing detergent-suspended p14 with LUVs presaturated with detergent, followed by removal of the detergent. Sucrose gradient fractionation and SDS–PAGE analyses of the resulting protein–lipid mixtures revealed inefficient incorporation of p14 into liposomes when the detergent was removed by dialysis at 4°C or room temperature (Figure 2B, lanes 1–6), as evidenced by the presence of p14 in the protein–lipid aggregate (20–30% sucrose) and free protein (30% sucrose) fractions of the gradient. However, stepwise dilutions at 4°C, but not at room temperature, to gradually lower the OG concentration in 0.1% increments resulted in efficient incorporation of p14 into liposomes (Figure 2B, lanes 7–9 versus 10–12). Apparently, the rate of detergent depletion and temperature influences on membrane fluidity, protein–detergent, and/or lipid–detergent interactions influence the efficiency of p14 incorporation into proteoliposomes. Figure 2.Reconstitution of p14 into liposomes. (A) Suspensions of p14-liposomes were fractionated by centrifugation through a discontinuous sucrose gradient. The liposomes migrated primarily to the 0–20% sucrose interface (arrowhead). (B) p14-liposomes were prepared using the dilution or dialysis approaches at room temperature or 4°C. Fractions obtained from the indicated sucrose gradient interfaces were analyzed by SDS–PAGE and silver staining to detect p14 (P) and lipid (L). (C) Analysis of p14-proteoliposomes by electron microscopy indicated the presence of LUVs. Scale bar=0.5 μm. (D) Flow cytometric analysis of p14-liposomes immunostained using normal rabbit serum (shaded histogram), antibody against the C-terminal enterokinase tag (black line tracing), or anti-p14 ectodomain antibody (gray line tracing) and fluorescent secondary antibody. Download figure Download PowerPoint Electron microscopy confirmed that the p14-liposomes consisted of LUVs (Figure 2C). Immunofluorescent staining of the p14-liposomes, using antisera specific for the N-terminal ectodomain or for the C-terminal enterokinase tag, revealed that a significant proportion of p14 resides in the correct Nout/Cin topology (Figure 2D). The low level of staining observed with the anti-enterokinase antibody was consistently above background levels, suggesting that at least a proportion of p14 may reside in the inverse Nin/Cout topology, as previously reported for other membrane proteins inserted into detergent-saturated liposomes (Rigaud and Levy, 2003). Calculations based on quantitative analysis of phospholipid and protein concentrations in the isolated proteoliposomes estimated an average protein:lipid molar ratio of ∼1:300 (i.e. 3–3.5 p14 molecules per thousand lipid molecules). Reconstituted p14-liposomes mediate liposome–cell lipid mixing at the plasma membrane Lipid mixing between p14-liposomes and cell membranes was assessed using a fluorescence resonance energy transfer assay (Struck et al, 1981). Flow cytometry, rather than fluorimetry, was used to quantify increased cell-associated NBD fluorescence, indicative of transfer of the fluorescent lipids from liposomes to cell membranes. Flow cytometry provided a clearer indication of the range of fusion events (e.g. cells that have fused to multiple liposomes fluoresce more intensely and can be distinguished from cells that have fused to fewer liposomes), and allowed a clear distinction between fluorescent cells and background fluorescence due to residual liposomes adhered to cells. The fluorescence intensity of cells incubated with p14-liposomes at 37°C was compared to control samples representing p14-liposomes incubated at 4°C (to prevent fusion) or liposomes lacking p14 incubated at 37°C. Time-course analysis revealed a progressive increase in cellular fluorescence when p14-liposomes were incubated with target cells at 37°C (Figure 3). Within 5–10 min after the shift to 37°C, fluorescence intensity increased, as indicated by the rightward shift of the histogram, and continued to increase to maximal levels by 30–40 min. As indicated by the distribution of the histogram (Figure 3A), cell fluorescence intensity varied over two orders of magnitude, suggesting a wide variation in the number of fusion events per target cell. A graphical representation of the time points (Figure 3B), plotting the percent of cells in the target population with fluorescence intensities above background (t=0 intensity), indicated that lipid mixing reached maximal levels of ∼35–40% of the cells fluorescing above background. Using quantitative video microscopy, a similar situation was observed with fusion of HA-expressing cells to target red blood cells, where 25–30% of cells undergo full lipid mixing, with 70–80% of cells showing any level of lipid mixing (Mittal et al, 2003). Increased cell fluorescence over time was not observed when LUVs lacking p14 (i.e. standard liposomes) were incubated with cells at 37°C or when cells treated with p14-liposomes were incubated at 4°C (Figure 3B), suggesting that purified p14 mediates lipid mixing and, by inference, the hemifusion phase of membrane fusion. Figure 3.p14 induces liposome–cell lipid mixing. (A) Time-course analysis by flow cytometry of lipid mixing at 37°C between fluorescent p14-proteoliposomes and target QM5 cells. The line tracings indicate that fluorescence intensity increases over the indicated durations, in triplicate. The shaded histogram represents cell autofluorescence. (B) Overton subtraction analysis of the flow cytometry results obtained with p14-liposomes incubated with target cells at 37°C (filled diamonds) or 4°C (open diamonds), and standard liposomes lacking p14 incubated with target cells at 37°C (filled squares). Results are presented as the percent of cells fluorescing above background autofluorescence over time, and are the mean±standard error of three separate experiments in duplicate. Download figure Download PowerPoint The increase in fluorescence over time observed with p14-liposomes could have resulted from increased p14-liposome adherence to target cells, followed by 'spontaneous' lipid transfer or endocytic disruption of the adhered liposomes. Analysis by atomic force microscopy revealed that p14-liposomes do exhibit increased adhesive properties compared to standard liposomes (Supplementary Figure 3). Cell-binding analysis also indicated that p14-liposomes adhere more strongly to target cells (Figure 4A). To exclude that this increased adherence was responsible for the observed difference in lipid mixing between p14-liposomes and standard liposomes, the dose of input liposomes was adjusted to yield equivalent numbers of liposomes adhered to cells. Under such conditions, standard liposomes exhibited no increase in fluorescence while p14-liposomes underwent a time-dependent increase in lipid mixing (Figure 4B), suggesting that the fluorescence increase observed in the lipid-mixing assay was specifically due to p14-mediated lipid mixing. Additional support for this conclusion was obtained by demonstrating that lysophosphatidylcholine (LPC), a monoacylated fatty acid known to inhibit membrane fusion (Chernomordik and Kozlov, 2003), effectively inhibited lipid mixing between p14-liposomes and target cells in a dose-dependent manner (Supplementary Figure 4A). Furthermore, endocytosis inhibitors reduced p14-induced lipid mixing by only 20% (Supplementary Figure 4B), suggesting that endocytic entry pathways make only a minor contribution to the observed lipid-mixing. We conclude that p14 induces liposome–cell lipid mixing, indicative of the hemifusion phase of membrane fusion, and that membrane fusion occurs primarily at the plasma membrane, independent of cellular endocytic pathways. Figure 4.Adherence properties of p14-liposomes. (A) Fluorescently labeled p14-liposomes (diamonds) or standard liposomes (squares) at the indicated doses were bound to target cell monolayers at 4°C, and the relative binding efficiency (in arbitrary fluorescence units) was determined by fluorimetry. Results are the mean±s.d. from a representative experiment conducted in triplicate. (B) Target QM5 cell monolayers were incubated with 50 μl of p14-liposomes or 300 μl of standard liposomes (liposome doses that gave equivalent moles of liposomes bound to cells as shown in panel A) and the time course of liposome–cell lipid mixing was followed by flow cytometry as described in Figure 3. Results are presented as the percent of cells treated with p14-liposomes that fluoresced above levels observed in cells treated with standard liposomes, as determined by Overton subtraction. Results are the mean±s.d. from a representative experiment conducted in triplicate. Download figure Download PowerPoint Reconstituted p14-liposomes mediate content mixing The ability of p14 to mediate pore formation was assessed using the aqueous fluor calcein to monitor delivery of liposome contents to the cytosol of target cells. As with the lipid-mixing assay, cells treated with calcein-containing p14-liposomes at 37°C showed increased fluorescence over time, reaching maximal levels by 20 min that corresponded to ∼25% of the target cell population (Figure 5A). There was no increase in cellular fluorescence when cells were incubated at 37°C with standard liposomes or with p14-liposomes incubated at 4°C. Addition of similar levels of soluble calcein to cell monolayers resulted in no increased cell fluorescence (data not shown), implying that the cell fluorescence observed with calcein-containing p14-liposomes was not due to leakage of calcein out of the liposomes and into cells. Unlike the rapid onset of lipid mixing in <5 min, the intracellular delivery of calcein was not detected until after 10 min (Figure 5A). Whether this apparent lag simply reflects a decreased sensitivity of the content-mixing assay (i.e. the extent of calcein that must be delivered to cells before increased cell fluorescence is detectable) or something more interesting, such as the uncoupling of the lipid mixing and pore formation phases of the p14 fusion reaction, is unclear. Figure 5.p14 induces liposome–cell content mixing. (A) Target QM5 cells were incubated with calcein-loaded p14-liposomes at 37°C (filled diamonds) or 4°C (open diamonds), or with standard liposomes lacking p14 (open squares). At the indicated durations, cell fluorescence was quantified by flow cytometry and the percent cells fluorescing above background cell autofluorescence was determined by Overton subtraction. Results are the mean±standard error of three separate experiments. (B) Lipid mixing and content mixing assays were conducted as described in Figure 3 and above, using Vero cells (white bars) or QM5 cells (black bars) incubated for 20 min with p14-liposomes at 37 or 4°C, or with standard liposomes at 37°C. Cell fluorescence was quantified by flow cytometry and the percent cells fluorescing above background cell autofluorescence was determined by Overton subtraction. Results are the mean±s.d. from a representative experiment conducted in triplicate. Download figure Download PowerPoint Repeating these assays with Vero epithelial cells indicated levels of lipid mixing and content mixing comparable to the results obtained with QM5 fibroblasts (Figure 5B). Using fluorescence microscopy, cellular fluorescence was observed for both the aqueous and lipidic fluors when Vero cells were incubated with p14-liposomes at 37°C, but not when cells were incubated at 4°C or with liposomes that lacked p14 at 37°C (Figure 6). Examination of the z-angles revealed an asymmetric distribution of the two fluors, with the red lipidic fluor associated preferentially with the extremity of cells and the green aqueous fluor with the cell interior, consistent with lipid mixing at the plasma membrane and cytosolic delivery of the aqueous liposome contents. Therefore, p14 mediates liposome–cell membrane fusion and intracellular content delivery to different cell types. Figure 6.Fluorescence microscopy analysis of lipid mixing and content mixing. Vero cells were treated with p14-liposomes at 37°C (top row) or 4°C (middle row), or with liposomes lacking p14 (bottom row). Liposomes were labeled with calcein (green, left columns) to detect content mixing and with a lipidic fluor (red, middle columns) to monitor lipid mixing. The right-hand columns are an overlay of the DIC image and the lipid and content mixing of a single cell. The flanking panels are z-angles showing the asymmetric distribution of the lipidic fluor near the apical surface of the cell (top and right) and the aqueous calcein fluor in the interior of the cell (bottom and left). Arrows indicate residual liposomes adhered to cells. Size bar=10 μm. Download figure Download PowerPoint p14 is both necessary and sufficient for membrane fusion Liposome–liposome fusion assays were conducted to exclude the possibility that p14 might de dependent on formation of a trans-acting complex with a protein cofactor present in the target cell membrane. Initial experiments using our neutral liposomes did not detect p14-induced lipid mixing (data not shown). While p14-liposomes adhered to target cells (Figure 4), we suspected that the same might not be true for p14-liposome adherence to artificial phospholipid bilayers, and that the lack of liposome–liposome fusion was due to the absence of sufficiently close liposome–liposome interactions. Therefore, p14 was reconstituted into the anionic liposome formulation (POPC (1-palmitoyl-2-oleoyl-phosphatidylcholine):DOPS (dioleoyl-phosphatidylserine) at 85:15 mol%) widely used to assess SNARE-mediated liposome fusion (Weber et al, 1998; Parlati et al, 1999; McNew et al, 2000; Chen et al, 2004), and liposome–liposome interactions were promoted using divalent cations. In the absence of divalent cations, p14-induced lipid mixing barely exceeded the low level of spontaneous lipid mixing (Figure 7A and B, curve c). The addition of divalent cations resulted in a dose-dependent increase in lipid mixing for p14-liposomes, but had little to no effect on lipid mixing in the absence of p14 (Figure 7A and B, curves a and b). This situation applied to both Ca2+ and Mg2+, with Ca2+ exerting a more pronounced effect (Figure 7C). Figure 7.p14 mediates liposome–liposome fusion. (A) Unlabeled anionic (POPC:DOPS at 85:15 mol%) p14-liposomes were prepared using the standard detergent depletion protocol, and were mixed with fluorescently labeled target anionic liposomes in the presence of 10, 1, or 0 mM Ca2+ (curves a, b, and c, respectively). Increased NBD fluorescence due to lipid mixing was monitored by fluorimetry over time at 37°C. Results are reported as the percent maximal fluorescence, as determined by detergent solubilization of the liposomes. Inset: The 0–20% (lane 1), 20–30% (lane 2), and 30% (lane 3) fractions from a sucrose gradient were analyzed by SDS–PAGE and silver staining (see Figure 2B legend) to reveal efficient incorporation of p14 into the POPC–DOPS liposomes by the detergent dilution approach. (B) The same experimental conditions as described in panel A were performed using POPC-DOPS liposomes lacking p14. (C) Unlabeled p14-liposomes (curves a and b) or unlabeled liposomes lacking p14 (curves c and d) were mixed with fluorescently labeled target liposomes in the presence of 10 mM Ca2+ (curves a and c) or 10 mM Mg2+ (curves b and d) and lipid mixing was quantified as described in panel A. (D) Liposome–liposome fusion assays were conducted as described in panel A in the presence of 10 mM Ca2+, under conditions where p14 was present in both donor and target liposomes (curve a), donor liposomes only (curve b), or absent from both liposomes (curve c). Download figure Download PowerPoint Under optimal conditions for p14-induced lipid mixing, there was a sharp increase in NBD fluorescence in the first few minutes, which leveled off by 5–7 min at ∼15–20% of the theoretical maximum lipid mixing followed by a slow progressive increase in fluorescence from 10 to 30 min that paralleled the low rate of spontaneous lipid mixing observed between liposomes lacking p14 (Figure 7A). As with liposome–cell fusion, additional studies indicated that p14 need only be present in the donor membrane to effect lipid mixing between liposomes (Figure 7D). A modest increase in fusion activity was noted when p14 was present in both membranes; however, this effect was most likely the result of the 25% increase in the total amount of p14 in the system (i.e. a 4:1 ratio of donor:target membranes) rather than the need for p14 in both membranes, as suggested by the ability of p14-transfected cells to fuse to nontransfected cells (Supplementary Figure 1). The ability of p14 to induce liposome–liposome fusion suggests that the p14 FAST protein is both necessary and sufficient to induce the lipid rearrangements required for merger of two lipid bilayer