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SNARE ‐mediated membrane fusion arrests at pore expansion to regulate the volume of an organelle

2018; Springer Nature; Volume: 37; Issue: 19 Linguagem: Inglês

10.15252/embj.201899193

1460-2075

M. D’Agostino, Herre Jelger Risselada, Laura Josefine Endter, Véronique Comte‐Miserez, Andreas Mayer,

Advanced Fluorescence Microscopy Techniques

Article17 August 2018free access Source DataTransparent process SNARE-mediated membrane fusion arrests at pore expansion to regulate the volume of an organelle Massimo D'Agostino orcid.org/0000-0002-4756-6330 Département de Biochimie, Université de Lausanne, Epalinges, Switzerland Search for more papers by this author Herre Jelger Risselada Department of Theoretical Physics, Georg-August University, Göttingen, Germany Leiden Institute of Chemistry, Leiden University, Leiden, The Netherlands Search for more papers by this author Laura J Endter Leiden Institute of Chemistry, Leiden University, Leiden, The Netherlands Search for more papers by this author Véronique Comte-Miserez Département de Biochimie, Université de Lausanne, Epalinges, Switzerland Search for more papers by this author Andreas Mayer Corresponding Author [email protected] orcid.org/0000-0001-6131-313X Département de Biochimie, Université de Lausanne, Epalinges, Switzerland Search for more papers by this author Massimo D'Agostino orcid.org/0000-0002-4756-6330 Département de Biochimie, Université de Lausanne, Epalinges, Switzerland Search for more papers by this author Herre Jelger Risselada Department of Theoretical Physics, Georg-August University, Göttingen, Germany Leiden Institute of Chemistry, Leiden University, Leiden, The Netherlands Search for more papers by this author Laura J Endter Leiden Institute of Chemistry, Leiden University, Leiden, The Netherlands Search for more papers by this author Véronique Comte-Miserez Département de Biochimie, Université de Lausanne, Epalinges, Switzerland Search for more papers by this author Andreas Mayer Corresponding Author [email protected] orcid.org/0000-0001-6131-313X Département de Biochimie, Université de Lausanne, Epalinges, Switzerland Search for more papers by this author Author Information Massimo D'Agostino1,†, Herre Jelger Risselada2,3, Laura J Endter3, Véronique Comte-Miserez1 and Andreas Mayer *,1 1Département de Biochimie, Université de Lausanne, Epalinges, Switzerland 2Department of Theoretical Physics, Georg-August University, Göttingen, Germany 3Leiden Institute of Chemistry, Leiden University, Leiden, The Netherlands †Present address: Department of Molecular Medicine and Medical Biotechnology, University of Naples Federico II, Naples, Italy *Corresponding author. Tel: +41 21 6925704; E-mail: [email protected] EMBO J (2018)37:e99193https://doi.org/10.15252/embj.201899193 See also: TH Söllner & J Malsam (October 2018) PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Constitutive membrane fusion within eukaryotic cells is thought to be controlled at its initial steps, membrane tethering and SNARE complex assembly, and to rapidly proceed from there to full fusion. Although theory predicts that fusion pore expansion faces a major energy barrier and might hence be a rate-limiting and regulated step, corresponding states with non-expanding pores are difficult to assay and have remained elusive. Here, we show that vacuoles in living yeast are connected by a metastable, non-expanding, nanoscopic fusion pore. This is their default state, from which full fusion is regulated. Molecular dynamics simulations suggest that SNAREs and the SM protein-containing HOPS complex stabilize this pore against re-closure. Expansion of the nanoscopic pore to full fusion can thus be triggered by osmotic pressure gradients, providing a simple mechanism to rapidly adapt organelle volume to increases in its content. Metastable, nanoscopic fusion pores are then not only a transient intermediate but can be a long-lived, physiologically relevant and regulated state of SNARE-dependent membrane fusion. Synopsis Membrane tethering and docking have been assumed to control fusion reactions. New data show that non-expanding, nanoscopic pores connect yeast vacuoles in vivo and regulate fusion between two organelles according to the physiological status of the cell. Non-expanding, nanoscopic fusion pores connect vacuoles in vivo. Fusion pores are long-lived structures. Metastable fusion pores allow organelles to rapidly adapt to osmotic pressure changes. SNARE proteins and the HOPS complex stabilize pores against re-closure. Metastable nanoscopic pores serve as a control point for the fusion reaction. Introduction Membrane fusion reactions in eukaryotic cells traverse a series of intermediate steps. Docking is followed by outer membrane leaflet fusion, establishing a hemifusion stalk, which can expand into a larger hemifused zone, the hemifusion diaphragm. Fusion of the inner leaflets leads to the formation and subsequent expansion of the fusion pore, which allows content mixing. Trans-SNARE complexes transmit force to the membranes, which drives fusion reactions through these steps (Gao et al, 2012; Hernandez et al, 2012, 2014; Shi et al, 2012). Theory indicates that the expansion of an existing fusion pore faces a major energy barrier, which may be overcome through membrane tension (Chizmadzhev et al, 2000; Kozlov et al, 2010; Long et al, 2012; Kozlov & Chernomordik, 2015; Ryham et al, 2016). This suggests that expansion of fusion pores might be rate-limiting, reversible, and give rise to a potentially long-lived intermediate. In line with this, in vitro data show that the number of trans-SNARE complexes around a fusion pore and SNARE-associated proteins influences the diameter and dynamics of that pore (Shi et al, 2012; Lai et al, 2013; Bao et al, 2018). Also in vivo, exocytic fusion pores are highly dynamic and their properties depend on the expression level of exocytic SNAREs and membrane tension in vivo (Bao et al, 2018; Shin et al, 2018). Despite this dynamics, exocytic fusion pores usually remain relatively short-lived (sub-second range) (Bao et al, 2018; Shin et al, 2018). Pore flickering for seconds could be observed during exocytosis of very compact or colloidal content of secretory vesicles, which can counteract the completion of fusion. In this special case, pore opening requires actin-dependent force generation for active extrusion (Tse et al, 1993; Zimmerberg et al, 1994; Chen et al, 2008; Tran et al, 2015; Rousso et al, 2016; Zhao et al, 2016), suggesting that pore opening might be hindered through interactions between the membrane and its contents. Stable, long-lived intermediates arrested in hemifusion or at the state of non-expanding fusion pores have remained elusive in living cells. Particularly for all non-exocytic fusion events, in vivo fusion intermediates are hence usually presumed to be short-lived. Visualization of such intermediates in living cells is challenging, but it might be favored in a situation in which pore expansion is governed by limited membrane tension (Kozlov & Chernomordik, 2015). We used yeast vacuoles to probe for such fusion intermediates in vivo. The structure of the vacuolar compartment is dynamically controlled by two antagonistic processes. On the one hand, membrane fission reduces the total volume of the vacuolar compartment by fragmenting it into multiple smaller vacuoles. On the other hand, fusion can re-assemble these small vacuoles into a single bigger structure. The equilibrium between both processes determines vacuole structure (Weisman, 2003; Peters et al, 2004; Baars et al, 2007; Brett & Merz, 2008; Michaillat et al, 2012; Alpadi et al, 2013). Fusion is favored when vacuolar content increases, either by accumulation of macromolecules, e.g., during autophagy, or through osmotic water influx in hypotonic media. Fragmentation is triggered by lack of luminal content or hypertonic media (Bonangelino et al, 2002; Zieger & Mayer, 2012; Desfougères et al, 2016a). Homotypic fusion of yeast vacuoles in vitro could be dissected into a series of well-characterized steps: Priming by Sec18/NSF promotes SNARE activation through dissociation of cis-SNARE complexes (Mayer et al, 1996; Ungermann et al, 1998). Tethering, governed by coordinated action of the Rab-GTPase Ypt7 and the HOPS complex, permits the formation of trans-SNARE complexes (Mayer & Wickner, 1997; Price et al, 2000; Zick & Wickner, 2014; Orr et al, 2015; Lürick et al, 2017). These, together with V0 proteolipids, induce lipid mixing (Peters et al, 2001; Reese et al, 2005; Strasser et al, 2011; Desfougères et al, 2016b; Mattie et al, 2017). Content mixing requires several trans-SNARE complexes (D'Agostino et al, 2016) and their anchoring in the membrane through peptidic transmembrane domains in helical continuity with the SNARE domain (Pieren et al, 2015). Content mixing also requires the association of trans-SNARE complexes with HOPS and its SM protein subunit Vps33 in order to deform the hemifusion zone sufficiently to facilitate fusion pore formation (Pieren et al, 2010; Zick & Wickner, 2014; D'Agostino et al, 2017). The transition from hemifusion to content mixing is rate-limiting for vacuole fusion in vitro (Reese & Mayer, 2005), but it could not yet be resolved whether the limiting step is fusion pore formation or pore expansion. It has also remained unclear whether the transition from hemifusion to pore formation or expansion is kinetically relevant in vivo and whether the respective intermediates might be of physiological importance. Since, in living cells, yeast vacuoles always appear as a cluster of tethered vesicles, we explored whether these organelles are only docked, or whether they are connected through hemifusion, non-expanding or expanded fusion pores. Results When multiple vacuolar vesicles are present, as is the case in many wild-type strains under conditions of logarithmic growth on standard rich media, these vacuoles are always tethered to each other. Given that vacuoles are tethered over a large surface in living yeast cells, we tested whether expanded fusion pores might be detectable. We stained the vacuoles in living cells with FM4-64, a vital dye that inserts into the outer leaflet of the plasma membrane. FM4-64 enters the cells by endocytosis and finally accumulates in the vacuolar membrane (Vida & Emr, 1995). Stained cells were analyzed by serial confocal sectioning with a spinning disk microscope and 3D reconstruction. Only ~1% of the cells showed a pore connecting two adjacent vacuoles (Fig 1A), and we never observed more than a single pore in a contact zone. Figure 1. In vivo visualization of an expanding fusion pore z-stacks were acquired from FM4-64-labeled vacuoles in wild-type cells (BJ3505), using a spinning disk microscope (step size 200 nm). 3D reconstructions from these stacks reveal a peripherally located fusion pore. Scale bar: 2 μm. Wild-type (BJ3505) and isogenic pmc1Δ/vcx1Δ mutant cells were labeled with FM4-64. The number of vacuoles per cell was determined in three independent experiments evaluating 200 cells each. Scale bar: 2 μm. Frequency of visible fusion pores. Cells were grown as in (B). The percentage of cells showing visible fusion pore was determined from z-stacks. Means and SD are shown for 100 stained vacuoles from three independent experiments. Schematic view of Vph1-GFP and FM4-64 distribution over the vacuolar membranes in the presence of a nanoscopic fusion pore or of an extended hemifusion diaphragm. Distribution of Vph1-GFP and FM4-64 in vacuole–vacuole contact sites. Vph1-GFP-expressing cells, labeled with FM4-64, were analyzed under a spinning disk confocal microscope equipped with two cameras for the simultaneous acquisition of GFP and FM4-64 fluorescence signals. Separate channels are shown on the right. The entire contact surface, but not the pore, is accessible to both probes. Scale bar: 2 μm. Expansion of a vacuolar fusion pore in real time. pmc1Δ/vcx1Δ cells were labeled with FM4-64 and immobilized in a 50-μl flow chamber (Ibidi). The osmotic value of the medium was changed by perfusion with water, using a pump with a flow of 30 μl/s. A frame was acquired every 2 s for a total period of the 30 s, using the laser at minimal intensity. Source data are available online for this figure. Source Data Figure 1 [embj201899193-sup-0003-SDataFig1.xlsx] Download figure Download PowerPoint Osmotic pressure controls vacuolar fusion pores Membrane tension and osmotic pressure determine the kinetics and the degree of fusion pore expansion (Cohen et al, 1980; Chizmadzhev et al, 1999, 2000; Bretou et al, 2014; Tran et al, 2015; Rousso et al, 2016). Based on this notion, we searched for mutants that show pores in the contact zone with higher frequency. Cells lacking the two main vacuolar Ca2+ importers, Pmc1 and Vcx1, had this property. pmc1Δ vcx1Δ mutants show increased cytosolic Ca2+ concentration, which activates calcineurin, an important controller of osmo-homeostasis that induces the expression of pumps extruding monovalent cations from the cells (Cyert & Philpott, 2013). This can result in an osmotic imbalance between cytosol and vacuoles. Only 35% of pmc1Δ vcx1Δ cells contain two or more vacuoles (Fig 1B) and could be analyzed for pores in the contact zones. Strikingly, 85% of these contact zones showed pores (Fig 1B and C), which were identified by simultaneous exclusion of FM4-64 and of a GFP-tagged transmembrane protein, the V-ATPase a subunit Vph1 (Fig 1D and E). We always found only a single pore, and it was adjacent to the vertex of the contact zone, which accumulates many fusion-relevant proteins and lipids (Wang et al, 2002, 2003; Fratti et al, 2004; Karunakaran et al, 2012). Simulation approaches suggested that fusion pores can open through direct disruption of a fusion stalk, or at the rim of a hemifusion diaphragm (Risselada et al, 2014). Since in vitro studies had provided evidence for extended hemifusion diaphragms between giant unilamellar vesicles (Nikolaus et al, 2010) as well as between tethered vacuoles (Mattie et al, 2017), which were large enough to be detectable by light microscopy, we tested the existence of these structures in living yeast cells. Transmembrane proteins and proteins bound to outer leaflet lipids should be excluded from a hemifusion diaphragm, whereas the lipidic probe FM4-64 might label the diaphragm (Fig 1D). We found Vph1-GFP and FM4-64 to be distributed over the entire contact zone between two vacuoles (Fig 1E). Similar observations were made with all the GFP-tagged transmembrane proteins (not shown) as well as with proteins binding the outer vacuolar leaflet that we used below (see, e.g., PX-GFP in Movie EV1). The only non-labeled areas that we found (mostly in pmc1Δ vcx1Δ mutants) were also devoid of FM4-64 and we hence consider those as expanded fusion pores. This suggests that, in living cells, vacuoles do not form extended hemifusion diaphragms and favors the notion that vacuolar fusion pores might originate directly from a stalk, or from small hemifusion diaphragms that would escape detection by the technique that we applied. When the cells were challenged by hypotonic media, their vacuoles fused within 30 s. Time-lapse microscopy under minimal illumination showed that fusion occurred through gradual expansion of the single existing pore over the entire contact surface (Fig 1F). Formation of lumenal vesicles or the excision of membrane disks, which had been reported previously (Wang et al, 2002), was not observed. We noticed, however, that this phenomenon was triggered upon continuous exposure of the cells to intense illumination from the microscope for 30 s, or upon heat-stressing the cells at 40°C. Both treatments strongly increased the frequency of lumenal, FM4-64-stained membranes (Fig EV1; McNally et al, 2017), which were mobile. When fusion was triggered by hypotonic media for 30 s at the normal growth temperature of 30°C, and only a single snapshot was taken, lumenal fragments were not observed (Fig EV1). The excision of membrane disks or lumenal vesicles thus appears to be thermally nucleated. It may require a transition to more inverted membrane phases (Risselada et al, 2014). Click here to expand this figure. Figure EV1. Effect of osmotic and temperature shocks on vacuole structure in vivo Effect of excessive excitation light. Vacuole fusion was stimulated in BY4741 cells, labeled with FM4-64, by adding water. Pictures were acquired either under continuous laser excitation for 30 s, or with only a single laser flash before and 30 s after water addition. Effect of elevated temperature. BY4741 cells were labeled with FM4-64, and vacuole morphology was analyzed by spinning disk confocal microscopy before and after exposure to 40°C for 2 min. Arrows indicate vacuoles with lumenal vesicles. Data information: All scale bars: 5 μm. Download figure Download PowerPoint Tethered yeast vacuoles in living cells exchange lipid but not content We explored the possibility that also the tethered vacuoles of wild-type cells, which only very rarely show microscopically visible pores in their contact sites, might be much more advanced in the fusion pathway than we commonly presume and exist in a partially fused state, such as a hemifusion intermediate or a nanoscopic, non-expanding fusion pore. We labeled vacuoles with FM4-64, which served as a lipidic tracer to evaluate continuity of vacuolar membranes by fluorescence recovery after photobleaching (FRAP). We chose cells showing a vacuole that was in contact with neighboring vacuoles, yet in a sufficiently peripheral position in the cluster to allow selective photobleaching of the FM4-64 in its membranes by a laser pulse. The FM4-64 signal could be completely bleached but recovered within a few seconds after the laser pulse (Fig 2A and B). Cells deleted for the vacuolar SNAREs Nyv1 or Vam3, or for the vacuolar Rab-GTPase Ypt7, which are deficient for lipid and content mixing in vitro and carry numerous small vacuoles in vivo, did not show any recovery. We also analyzed a strain carrying a single amino acid substitution in a subunit of the proteolipid cylinder of the V-ATPase (vma16F190Y). In vitro, vma16F190Y vacuoles dock and form trans-SNARE complexes, but they are impaired in lipid mixing (Strasser et al, 2011). Consistent with the in vitro data, vacuoles in vma16F190Y cells did not show fluorescence recovery after photobleaching in vivo (Fig 2A). Transfer of FM4-64 between tethered wild-type vacuoles did not depend on the chemical nature of this compound, because similar transfer was observed with other vacuolar lipid probes, such as the lipophilic dye MDY-64 or a GFP bound to the cytoplasmic leaflet of vacuoles through the PI3P-binding PX or FYVE domains (Burd & Emr, 1998; Gillooly et al, 2000; Cheever et al, 2001; Fig EV2). Also, the transfer of these lipid probes between vacuoles was suppressed in fusion-deficient mutants (Fig EV2E for PX-GFP and see below for FYVE2-GFP). Together, these observations suggest that transfer of lipid probes between vacuoles does not occur through the cytosol and requires the vacuolar fusion machinery. Figure 2. In vivo assay for lipid and content mixing of vacuolar membranes A. Vacuolar membranes of wild-type (BY4741), nyv1Δ vam3Δ, ypt7Δ, or vma16F190Y cells were labeled with the vital dye FM4-64. A vacuole that was in a sufficiently peripheral location to be selectively photobleached was exposed to a laser pulse (white arrows indicate the bleaching area). Recovery of fluorescence in this area was assayed 10 s later by spinning disk confocal microscopy. Scale bar: 2 μm. B. Kinetics of FM4-64 recovery in (A). Means and SD are shown for 20 vacuole clusters from three independent experiments. C. In vivo lipid and content mixing. Vacuoles in living yeast were labeled with the indicated lumenal (CDCFDA) and membrane (FM4-64 or PX-GFP) probes. Note that CRY1 cells lack the ADE2 gene and hence naturally accumulate 5-amino-1-(5-phospho-D-ribosyl)imidazole as fluorescent fluid phase marker in the vacuolar lumen. FRAP experiments were performed as in (A) (arrows indicate the bleaching area). (i) Non-bleached lumenal area. (ii) Bleached lumenal area. (iii) Bleached membrane area. Scale bar: 2 μm. D, E. Kinetics of fluorescence recovery after photobleaching of the probes in (C). Numbers denote the areas shown in (C): (i) Non-bleached lumenal area. (ii) Bleached lumenal area. (iii) Bleached membrane area. Means and SD are shown for 20 vacuole clusters from three independent experiments. Source data are available online for this figure. Source Data Figure 2 [embj201899193-sup-0004-SDataFig2.xlsx] Download figure Download PowerPoint Click here to expand this figure. Figure EV2. FRAP-based assay of in vivo lipid mixing with different fluorescent probes A–E. Wild-type cells from two different backgrounds (BJ3505 and BY4741) were labeled with (A) FM4-64 or (B) MDY-64, or transformed with expression plasmids for (C) PX-GFP or (D) FYVE2-GFP in order to decorate the outer leaflet of the vacuolar membranes. The cells were subjected to FRAP assays as described in Fig 2. White arrows indicate the area of bleaching. (E) Same experiment as in (C), but using BJ ypt7Δ to illustrate that the transfer of PX-GFP is impaired in fusion-deficient mutants. Scale bars: 2 μm. Download figure Download PowerPoint Next, we measured the exchange of two soluble lumenal markers in combination with lipid probes. We used FM4-64 in combination with an ade2 mutant strain, which lacks the activity of phosphoribosylaminoimidazole carboxylase, an enzyme necessary for de novo synthesis of purines. ade2 mutant cells (in this case in the CRY1 strain) grow well on rich media, but they accumulate brightly fluorescent adenine precursors (5-amino-1-(5-phospho-D-ribosyl)imidazole), which concentrate in the lumen of their vacuoles and provide a convenient vacuolar fluid phase marker (Smirnov et al, 1967; Weisman et al, 1987). Alternatively, we used cells (BJ3505) expressing the lipid probe PX-GFP, which binds to phosphatidylinositol-3-phosphate (PI3P) in the vacuolar membrane (Cheever et al, 2001), and stained them with CDCFDA (5(6)-carboxy-2′,7′-dichlorofluorescein diacetate), which is hydrolyzed by esterases in the vacuolar lumen and hence accumulates there. Both approaches lead to vacuoles with stained membranes and diffuse labeling of the aqueous lumen (Fig 2C). The dye combinations were chosen such that the lumenal and membrane probes could be simultaneously bleached by the same laser. After a laser pulse had bleached both the lumenal and membrane markers, FM4-64 and PX-GFP signals in the membrane recovered within 3 s, whereas the lumenal probes showed barely any recovery (Fig 2C–E). The small signal recovery that is visible in the lumenal area most likely represents out-of-focus fluorescence stemming from the recovering membrane probe that fills the membrane above and below the imaging plane (this background is visible, for example, in Fig 2A, particularly for smaller vacuoles). Recovery of the lumenal signal is also not limited by restricted diffusion because, when we bleached only a smaller area within a single, very large vacuole, recovery is instantaneous, i.e., too fast to be imaged by our microscopy setup, which has a switch-time between bleaching and acquisition of around 50 ms. Thus, vacuoles in living yeast cells efficiently exchange lipid markers but not content markers. Tethered vacuoles are connected by non-expanding fusion pores This behavior could indicate either a state of hemifusion, in which the outer leaflets mix but the inner leaflets remain separated, or the presence of a nanoscopic fusion pore, through which both leaflets are connected but which is small enough to restrain the passage of fluorescent lumenal tracers. We tried to distinguish between these possibilities by establishing lipid tracers that allow to selectively probe the outer or inner membrane leaflet. FM4-64 inserts into the outer leaflet of the plasma membrane and reaches vacuoles by endocytosis (Vida & Emr, 1995), which suggests that it might decorate the inner leaflets of vacuolar membranes. We tested this by placing GFP domains either at the inner or outer surface of the vacuolar membrane. This allows to assay fluorescence energy transfer (FRET) from GFP to FM4-64 because FM4-64 has its excitation maximum exactly at the emission wavelength of GFP (Fig 3A). Since FRET diminishes with the inverse 6th power of the distance between donor and acceptor, much more efficient quenching of GFP is expected if the GFP domain resides on the same leaflet as FM4-64. In order to place GFP at the surface of either leaflet, we fused it to the luminal C-terminus of the vacuolar transmembrane protein Nyv1 or to C-terminus of the vacuolar V-ATPase subunit Vph1, which is placed at the cytosolic face of the membrane (Mazhab-Jafari et al, 2016; Fig 3B). We compared the fluorescence intensity of these two markers before and after FM4-64 labeling by FACS. Labeling intensity with FM4-64 was similar for both strains (Fig 3C–E). However, the fluorescence emission of the lumenal GFP (Nyv1-GFP) was significantly diminished by the presence of FM4-64, whereas GFP emission at the cytosolic leaflet (Vph1-GFP) was not reduced (Fig 3E). This suggests that FM4-64 is concentrated in the lumenal leaflet of vacuoles. Figure 3. FRET-based localization of FM4-64 at the inner membrane leaflet FM4-64 and GFP excitation/emission spectra. FRET-based in vivo strategy to distinguish localization of FM4-64 in the inner or outer leaflets of vacuolar membranes. EGFP tags were added to the cytosolic C-terminus of Vph1 or to the lumenal C-terminus of Nyv1. Assay by microscopy. Half of two cultures of cells expressing Vph1-EGFP or Nyv1-EGFP were labeled with FM4-64, washed, and mixed with the other, non-labeled half of the same culture. Cells were analyzed by spinning disk confocal microscopy. The intensities of GFP fluorescence were tracked along the dashed lines and compared between neighboring FM4-64-labeled and unlabeled cells. FACS analysis. The strains from (C) were labeled as described, washed, and analyzed by FACS. Shown are side (SSC-A) and forward scatter (FSC-A), as well as the fluorescence intensity distributions for FM4-64 and for GFP in the absence or presence of FM4-64. Comparison of the GFP mean fluorescence from FM4-64-stained cells and unstained cells (20,000 each). Possibilities for FM4-64 transfer between the inner leaflets of two tethered vacuoles. Flipping between the trans-leaflets of a hemifusion structure; reversible flipping into the outer leaflet, establishing a minor pool that then transfers through the buffer or through the outer leaflet; or passage through a fusion pore. Source data are available online for this figure. Source Data Figure 3 [embj201899193-sup-0005-SDataFig3.xlsx] Download figure Download PowerPoint The transfer of FM4-64 between two vacuoles might then occur through continuity of the inner leaflets, or through the outer leaflets if the probe can flip between the two leaflets (Fig 3F). We developed a reporter that allows to stringently test the continuity of the inner leaflets (Fig 4), based on the following prediction: If the organelles are connected by a nanoscopic fusion pore in which both leaflets are fused, transmembrane proteins might be able to pass between these vacuoles, whereas hemifused membranes would not allow this passage. Passage through the pore should only be possible provided that the transmembrane domains (TMDs) carry no bulky hydrophilic parts that are exposed to the lumen and might hinder their diffusion through the small pore. As a basis for this reporter, we used the TMD of the vacuolar protein Nyv1, because it does not contain hydrophilic sequences at its lumenal, C-terminal end (Fig 4A and B). A yeGFP domain was added to the cytosolic N-terminal end of this TMD. FRAP experiments showed that this reporter (yeGFP-LT) passed freely between tethered vacuoles and behaved similarly as FM4-64 (Fig 4C and G). An equivalent construct with the TMD of an unrelated vacuolar protein, alkaline phosphatase (GFP-TMDALP), showed the same behavior (Fig EV3), suggesting that the ability to pass between adjacent vacuoles was independent of the chemical nature of the TMD. Figure 4. Protein-based assay for a nanoscopic fusion pore A. Synthetic reporter proteins carrying EGFP either at the lumenal or cytosolic end of the single-spanning transmembrane domain of Nyv1, which carries no hydrophilic extensions at its lumenal C-terminus. B. The topology of these constructs in the vacuolar membrane is indicated. C, D. FRAP assays for fusion pores. The indicated cells expressing (C) yeGFP-LT or (D) LT-yeGFP were labeled with FM4-64 and subjected to FRAP experiments as in Fig 2. Cells were imaged before and 0 (bleached) or 30 s after photobleaching (recovery). The bleached areas are indicated by an arrow. E, F. FRAP assays for fusion pores were performed as in (C, D), but with nyv1Δ cells. Scale bar: 2 μm. G. The histogram reports the fraction of cells showing FRAP. Means and SD are shown from three independent experiments, with 100 cells analyzed in each. Source data are available online for this figure. Source Data Figure 4 [embj201899193-sup-0006-SDataFig4.xlsx] Download figure Download PowerPoint Click here to expand this figure. Figure EV3. Passage of a GFP-fusion with the TMD of the vacuolar alkaline phosphatase (ALP) between vacuoles A–C. The synthetic reporter protein GFP-TMDALP was created (A), which carries GFP at the cytosolic, N-terminal en