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SNAREs pp 253-262 | Cite as

Assay of Lipid Mixing and Fusion Pore Formation in the Fusion of Yeast Vacuoles

  • Massimo D’Agostino
  • Andreas MayerEmail author
Protocol
Part of the Methods in Molecular Biology book series (MIMB, volume 1860)

Abstract

Fluorescence de-quenching can be used to analyze membrane lipid mixing during an in vitro fusion reaction. Here we describe a method to measure lipid mixing using vacuolar membranes purified from the yeast Saccharomyces cerevisiae. Labeling the isolated organelles with rhodamine-phosphatidylethanolamine allows to reveal ATP-dependent lipid mixing through fluorescence de-quenching in a spectrofluorometer. Combining this assay with content mixing indicators, such as the fusion-dependent maturation of a luminal vacuolar phosphatase, then permits the detection of hemifusion intermediates and the analysis of the requirements for fusion pore opening.

Key words

Membrane fusion Lipid mixing Yeast Vacuole 

1 Introduction

Membrane fusion is a fundamental process in cell biology that is at the heart of important physiological processes , such as transport between intracellular organelles, secretion of hormones and neurotransmitters, fertilization, or pathological processes such as virus invasion and metastasis [1, 2, 3, 4]. Before reaching completion, many fusion events transit through an intermediate hemifusion state, in which two engaged membranes mix lipids but leave their contents separated [5]. Although this situation has usually been considered only as a very transient moment during SNARE -driven membrane fusion, a growing number of in vitro and in vivo observations suggest that this intermediate could be more stable than expected [1, 6, 7, 8, 9, 10].

While hemifusion has been readily detectable in artificial membrane systems, its detection and analysis in physiological SNARE-dependent fusion events, both in vitro and in vivo, have remained much more challenging. This is a crucial limitation to the further elucidation of the fusion pathway because it is critical to study these intermediates not only in synthetic lipid systems but also in their physiological membrane environment. The composition of synthetic SNARE-dependent membrane systems can be chosen at will, which can grossly vary their fusion properties, including the occurrence of intermediate states, and allows to optimize them for the study of precisely defined, mechanistic questions. The advantage of studying a physiological membrane system is that here the parameters are much more constrained; that is, the lipid composition and the density of the fusion proteins are set by the cell. Since these parameters are critical determinants of the fusion pathway [11, 12, 13, 14, 15, 16, 17], the analysis of these complex systems remains necessary to judge the physiological lifetime and relevance of fusion intermediates, and their potential regulation by the cell.

A systematic characterization of the factors contributing to the formation of the hemifused state and its progression to full fusion requires experimental systems and assays that allow a robust detection of this intermediate and the characterization of protein interactions that accompany it. Both conditions are met by the lysosome-like vacuoles of yeast, which can be isolated in large quantities and good purity [18, 19]. They represent a physiological, SNARE-dependent membrane fusion system [20, 21]. Here, we describe a simple method to measure lipid mixing during the fusion of purified yeast vacuoles that can be combined with a content mixing assay in order to reveal a hemifused state [10, 22, 23, 24, 25].

Content mixing is measured through the activation of a luminal vacuolar enzyme, pro-alkaline phosphatase (contained, e.g., in strain BJ3505) by the vacuolar protease Pep4 (contained, e.g., in strain DKY6281), which is enclosed in the other fusion partner [26]. Assay of content mixing thus requires the separate preparation of two vacuole populations, followed by their mixing in vitro (Fig. 1a). To measure lipid mixing, one of the fusion partners is labeled with rhodamine-phosphatidylethanolamine (Rh-PE) at a self-quenching concentration [22]. Upon lipid mixing between the donor membranes, Rh-PE dilutes over the acceptor membrane, causing an increase in the fluorescence signal by de-quenching (Fig. 1c, d). Membrane proximity (Fig. 1b, “docking”) is not sufficient to achieve this de-quenching [10, 22, 27]. Upon fusion, the contents of both fusion partners are mixed and the protease Pep4 cleaves the pro-peptide from pro-alkaline phosphatase (p-pho8), maturing it into the form m- pho8 (Fig. 1e). The resulting activity of this enzyme provides a readout for content mixing and full fusion. It is measured by a simple colorimetric assay, the conversion of the colorless p-nitrophenyl phosphate into the yellow p-nitrophenol (Fig. 1f). Since vacuole fusion reactions proceed for up to 90 min in vitro, fusion samples can be split at numerous intermediate time points. Comparison of the lipid and content mixing signals then allows to identify bona fide hemifusion intermediates as a state in which Rh-PE de-quenching occurs but alkaline phosphatase remains immature. This approach has allowed to dissect the function of numerous fusion factors on the vacuolar membranes and assign their activities to distinct stages of the fusion reaction [10, 22, 23, 24, 25, 28, 29, 30].
Fig. 1

In vitro assay for content and lipid mixing. (a) Purified donor vacuoles expressing the protease Pep4 are labeled with the lipid probe rhodamine-phosphatidylethanolamine (Rh-PE) at self-quenching concentration and mixed in the fusion reaction with purified acceptor vacuoles expressing the precursor of alkaline phosphatase, p- Pho8 . (b) Membrane juxtaposition brings vacuoles in very close proximity (docking) without causing lipid mixing. (c) Hemifusion allows lipid exchange between the outer leaflets, causing a reduction of local Rh-PE concentration in the vacuolar membranes. (d) Rh-PE de-quenching results in a fluorescence signal enhancement in a time- and ATP -dependent manner. (e) Inner leaflet mixing allows content mixing and conversion of p-pho8 by Pep4 to produce the mature form m-pho8. (f) The ATP-dependent formation of m-pho8 is measured through its enzymatic activity, using a colorimetric assay

2 Materials

  1. 1.

    DTT buffer: 9 mL of 1 M Tris–HCl pH 8.9, 0.45 g of DTT, and H2O up to 300 mL.

     
  2. 2.

    YP medium: 400 g Yeast extract and 800 g of polypeptone are dissolved in 36 L of ddH2O, aliquoted in flasks and bottles, sterilized, and stored at room temperature.

     
  3. 3.

    Spheroblasting buffer: 15 mL of 4 M sorbitol, 10 mL of 500 mM KPi pH 7.5, and 75 mL of YP medium containing 0.2% glucose. Mix, remove 12–18 mL, and fill up with 12–18 mL of 0.1 mg/mL lyticase preparation [22]. Mix and leave at room temperature. Other, commercially available spheroblasting enzymes, such as zymolyase 100T, can also be used. Their suitable concentration for spheroblasting depends on the strain background used and must be empirically determined.

     
  4. 4.

    Rotor with 6 swing-out buckets, such as SW40Ti (Beckman).

     
  5. 5.

    3 mM Rhodamine-phosphatidylethanolamine (Rh-PE , Molecular Probes) is dissolved in DMSO. The solution is stored at −20 °C in aliquots of 60–80 μL.

     
  6. 6.

    Bradford solution is kept at 4 °C.

     
  7. 7.

    PS buffer: 10 mM PIPES-KOH pH 6.8, 200 mM sorbitol. This buffer is sterile filtered if longer storage is desired.

     
  8. 8.

    3 × 15 mL Falcon tubes with 0%, 4%, and 15% of Ficoll-400 in PS buffer is freshly prepared and kept at 4 °C.

     
  9. 9.

    10 mL 5% Milk powder (w/v) in ddH2O is kept at 20–25 °C.

     
  10. 10.

    3 M KCl, 10 mM MnCl2, 20% Triton TX-100 (w/v), 1 M glycine pH 11.5, and 1 M Tris-HCl pH 9.0 are kept at room temperature.

     
  11. 11.

    An ATP -regenerating system is prepared by mixing ATP, creatine phosphate (CP), and creatine kinase (CK) in the following ratio: 50 μL of 100 mM ATP, 25 μL of 50 mg/mL CK, 200 μL of 1 M CP, 275 μL Ficoll 0%.

     
  • 100 mM ATP: 60.5 mg ATP is dissolved in a solution containing 100 μL of 1 M MgCl2, 100 μL of 500 mM PIPES pH 6.8, 52.5 μL of 4 M KOH, and 747.5 μL of ddH2O. Aliquots were stored at −20 °C.

  • 50 mg/mL Creatine kinase: 25 mg of creatine kinase (from rabbit muscle, ~350 units/mg) is dissolved in a solution containing 0.5 mL of 10 mM PIPES-KOH pH 6.8 and 50% glycerol (v/v). Aliquots are stored at −20 °C. Note that creatine kinase slowly loses activity over several months at −20 °C. Once this activity drops below a critical threshold, the ATP regeneration system mixed from an aged stock suddenly stops working.

  • 1 M creatine phosphate: 1.31 g of creatine phosphate is dissolved in 4 mL ddH2O, aliquoted, and stored at −80°C.

  1. 12.

    100 mM p-nitrophenyl-phosphate (PNPP) is prepared by dissolving 461.4 mg in 10 mL ddH2O. The solution is aliquoted and stored at −20 °C.

     
  2. 13.

    Phosphate assay mix: 100 μL 1 M MgCl2, 100 μL 100 mM PNPP, 200 μL Triton TX-100 20% (w/v), 2.5 mL 1 M Tris–HCl pH 8.9, and 7.2 mL ddH2O. This solution is sufficient for 20 reactions and has to be freshly prepared before use.

     
  3. 14.

    Protein inhibitor cocktail (PIC), 1000× stock: 200 mM Pefabloc SC, 5 mg/mL leupeptin, 500 mM o-phenanthroline, and 5 mg/mL pepstatin A. Aliquots are stored at −20 °C.

     
  4. 15.

    Fusion buffer: 120 mM KCl and 0.33 mM MnCl2 in PS buffer. The mix is kept on ice.

     
  5. 16.

    Non-coated black 96-well plate.

     

3 Methods

Handle all vacuole-containing samples with pipette tips with a wide orifice (e.g., cut open with scissors) to avoid membrane rupture by shearing.

3.1 Vacuole Isolation

  1. 1.

    Incubate cells overnight in 1 L YPD, using baffled 2 L Erlenmeyer flasks at 30 °C while shaking at 150 rpm. Inoculate the cultures such that they are in logarithmic phase at the time of harvesting the next morning, with an OD600nm ranging between 1 and 1.5. Growth to higher densities should be avoided because it lowers fusion activity. Re-dilution of overgrown cultures for 1–2 h before harvesting does not cure the adverse effects of growth at higher cell density. Once the cultures have been taken out of the incubator, harvest the cells immediately and do not let the cultures stand for longer times without shaking.

     
  2. 2.

    Harvest 330 mL of culture at 2500 × g for 2–3 min at 4 °C. Preferably, a centrifuge with high acceleration and deceleration rates should be chosen.

     
  3. 3.

    Discard supernatant, resuspend cells in 50 mL of DTT buffer by vortexing, and incubate in the water bath at 30 °C for 5–6 min.

     
  4. 4.

    Centrifuge at 2500 × g for 2–3 min, discard supernatant, resuspend the cells in 15 mL of spheroblasting buffer by vortexing, and incubate in water bath at 30 °C for 25–30 min.

     
  5. 5.

    Transfer the suspension into 30 mL Corex tubes and harvest spheroblasts by centrifugation at 4 °C, 2500 × g, for 2 min. Discard supernatant, taking care that the pellet, which is quite loose, is not lost.

     
  6. 6.

    Resuspend spheroblasts in 2 mL of ice-cold 15% Ficoll-400 in PS buffer by gently vortexing or stirring with a rod.

     
  7. 7.

    Add 150–250 μL of ice-cold DEAE dextran solution, mix by gentle shaking, and leave tubes on ice for 2 min before incubating them in a water bath for other 2 min at 30 °C. The amount of DEAE dextran to be added must be optimized according to the strain background, growth conditions, and spheroblasting enzyme used.

     
  8. 8.

    Cool the suspension on ice and transfer it into an SW40 tube. Make discontinuous gradients by overlaying the suspension with steps of 8%, 4%, and 0% Ficoll-400 in PS buffer, such that the tube is filled up to the top.

     
  9. 9.

    Spin in an ultracentrifuge at 2 °C, 150,000 × g, for 90 min.

     
  10. 10.

    Remove lipids from the top of the tubes by using an aspiration pump and harvest vacuoles from the 0–4% Ficoll interface using cut pipette tips. Transfer the organelles into ice-cold reaction tubes. For optimal fusion results, the organelles should be used within an hour after harvesting. Longer storage leads to loss of activity.

     

3.2 Vacuole Membrane Labeling

  1. 1.

    A Rh-PE aliquot is thawed by incubating it at 37 °C for 20 min under strong agitation.

     
  2. 2.

    During this period of time, incubate a non-coated black 96-well plate with 200 μL of 5% milk powder per well at 20–25 °C. This coats the wells with protein and reduces the adhesion of vacuoles and proteins to the plastic.

     
  3. 3.

    Centrifuge Rh-PE for 15 min at 12,000 × g in a tabletop centrifuge to sediment non-dissolved material.

     
  4. 4.

    Meanwhile, collect vacuoles from the Ficoll gradient and determine their protein concentration using Bradford solution and BSA as a standard.

     
  5. 5.

    560 μg of DKY6281 vacuoles is mixed with 800 μL of PS buffer in a siliconized 2 mL reaction tube and equilibrated for 40 s at 32 °C under gentle agitation (500 rpm).

     
  6. 6.

    Rh-PE is slowly injected (3 × 17 μL) into the vacuole suspension under continuous vortexing at 500 rpm. After that, vacuoles are incubated in a water bath for 30 s at 27 °C (see Note 1 ).

     
  7. 7.

    Add 500 μL of pre-warmed 15% Ficoll buffer and gently mix by inverting the tubes four times. After a very short spin, put the sample on ice.

     
  8. 8.

    Prepare small discontinuous density gradients in 2 mL reaction tubes by overlaying vacuoles with 300 μL of pre-warmed 4% Ficoll buffer and 400 μL of pre-warmed PS buffer, taking care to create sharp interfaces.

     
  9. 9.

    Transfer the mini-gradients in a pre-cooled centrifuge equipped with a swing-out rotor. Spin for 7 min at 3 °C and 11,700 × g, using slow acceleration and deceleration.

     
  10. 10.

    Stained vacuoles can be harvested from the 4% Ficoll-PS interface by careful aspiration with a pipette. The organelles are kept on ice and their protein concentration is determined by Bradford assay.

     
  11. 11.

    Discard milk from the 96-well plate, wash the wells with ddH2O, and take care to remove all traces of water before proceeding to the next step.

     

3.3 Lipid Mixing Assay

  1. 1.

    After labeling with Rh-PE and determination of protein concentration, vacuoles from BJ3505 and DKY6281 are mixed at a ratio of 5 to 1.

     
  2. 2.
    A standard lipid mixing reaction contains:
    • 36 μg of vacuoles (30 μg non-labeled BJ3505 and 6 μg labeled DKY6281)

    • 112 mM KCl

    • 0.33 mM MnCl2

    • 60 μL Fusion buffer

    • 9.5 μL of ATP regeneration system

    • PS buffer up to 120 μL

     
  3. 3.

    Add 100 μL of each reaction mix into the corresponding wells of a 96-well plate pre-cooled on ice, reserving the remaining 90 μL for the parallel content mixing assay.

     
  4. 4.

    Put the plate into a microplate fluorescence reader with temperature control (e.g., SpectraMax Gemini XS) and let it to equilibrate at 27 °C for 2–5 min before starting the measurements.

     
  5. 5.

    Samples are excited at 544 nm and fluorescence changes are measured at 590 nm every 2 min for a total period of 32 min (from Ft = 0 min to Ft = 32 min). An emission cutoff filter (570 nm, long pass) is used.

     
  6. 6.

    After 32 min, add to every well 100 μL of 1% triton TX-100/Ficoll 0%, mix, and continue acquisition for the next 10 min taking measurements every 30 s. The corresponding average values will be F(TX100), the fluorescence expected upon maximal de-quenching of the vacuole-associated Rh-PE (see Note 2 ).

     

3.4 Content Mixing Assay

  1. 1.

    Use the remaining 90 μL of every reaction for the content mixing assay. Incubate the samples in the water bath at 27 °C for 90 min.

     
  2. 2.

    After 90 min, assay the generated activity of alkaline phosphatase by adding 0.5 mL of pre-warmed phosphate assay mix to every tube and continue the incubation at 27 °C for further 5 min.

     
  3. 3.

    Stop the reactions by adding 440 μL of 1 M glycine pH 11.5 and read the absorbance at 405 nm (Fig. 1f), using a vacuole-free sample as a reference (see Note 3 ).

     

3.5 Analysis of Lipid Mixing Data

All 17 fluorescence measurements (Ft) taken between t = 0 min and t = 32 min are divided by F(TX100) and the degree of de-quenching is calculated, (Ft – F0min)/F(TX100). For representing the values on a graph, the 0 min value is set to 0.01 and the values of all other time points are normalized to it. This operation facilitates comparisons between individual samples, even if their absolute starting values vary slightly (Fig. 1d). After the 30 min time point, the samples continue to show a slow increase in fluorescence, which typically is identical in slope for all samples, even for those incubated under conditions that do not support vacuole fusion. Therefore, we consider this as an unspecific background signal that is independent of vacuole fusion.

4 Notes

  1. 1.

    Membrane labeling with Rh-PE represents the critical point of the experiment. Rh-PE must be incorporated into the existing vacuole membrane at the right concentration to obtain strong fluorescence self-quenching, such that any dilution of the probe by fusion of these vacuoles with an unlabeled vacuole can dilute the probe enough to result in a significant decrease in self-quenching.

     
  2. 2.

    In order to directly compare lipid and content mixing signals, the two assays must be calibrated against each other. This is best achieved by titrating fusion inhibitors that inhibit very early reaction stages, such as the Rab-GTPase inhibitor Gdi1 or antibodies to vacuolar SNAREs or NSF /Sec18, which interfere with membrane docking [23]. This allows to optimize the Rh-PE concentrations for vacuole labeling such that the lipid mixing signals titrate in correspondence to the content mixing signals. This condition must be met in order to allow the identification of hemifusion states, which are defined as states in which lipid mixing occurs whereas content mixing is impaired.

     
  3. 3.

    Limitations of the assay: In this protocol, content mixing is detected by the transfer of an >30 kDa protease from one fusion partner into the other. This requires a fusion pore of sufficient size to let this protein pass, which should be >2.5 nm in diameter. The approach could not detect fusion pores that are narrower than this or remain open only for very short periods of time that would not suffice to transfer sufficient amounts of the protease to the other fusion partner. Thus, very small or flickering fusion pores, which can be detected by electrophysiological methods, may be missed by this approach.

     

References

  1. 1.
    Kweon D-H, Kong B, Shin Y-K (2017) Hemifusion in synaptic vesicle cycle. Front Mol Neurosci 10:65.  https://doi.org/10.3389/fnmol.2017.00065CrossRefPubMedPubMedCentralGoogle Scholar
  2. 2.
    Harrison SC (2015) Viral membrane fusion. Virology 479–480:498–507.  https://doi.org/10.1016/j.virol.2015.03.043CrossRefPubMedGoogle Scholar
  3. 3.
    Jahn R, Scheller RH (2006) SNAREs--engines for membrane fusion. Nat Rev Mol Cell Biol 7:631–643.  https://doi.org/10.1038/nrm2002CrossRefPubMedGoogle Scholar
  4. 4.
    Jackson MB, Chapman ER (2006) Fusion pores and fusion machines in Ca2+−triggered exocytosis. Annu Rev Biophys Biomol Struct 35:135–160.  https://doi.org/10.1146/annurev.biophys.35.040405.101958CrossRefPubMedGoogle Scholar
  5. 5.
    Chernomordik LV, Zimmerberg J, Kozlov MM (2006) Membranes of the world unite! J Cell Biol 175:201–207.  https://doi.org/10.1083/jcb.200607083CrossRefPubMedPubMedCentralGoogle Scholar
  6. 6.
    Spessott WA, Sanmillan ML, McCormick ME et al (2017) SM protein Munc18-2 facilitates transition of Syntaxin 11-mediated lipid mixing to complete fusion for T-lymphocyte cytotoxicity. Proc Natl Acad Sci U S A 114(11):E2176.  https://doi.org/10.1073/pnas.1617981114CrossRefPubMedPubMedCentralGoogle Scholar
  7. 7.
    Risselada HJ, Bubnis G, Grubmüller H (2014) Expansion of the fusion stalk and its implication for biological membrane fusion. Proc Natl Acad Sci U S A 111:11043–11048.  https://doi.org/10.1073/pnas.1323221111CrossRefPubMedPubMedCentralGoogle Scholar
  8. 8.
    Lai Y, Diao J, Liu Y et al (2013) Fusion pore formation and expansion induced by Ca2+ and synaptotagmin 1. Proc Natl Acad Sci U S A 110:1333–1338.  https://doi.org/10.1073/pnas.1218818110CrossRefPubMedPubMedCentralGoogle Scholar
  9. 9.
    Diao J, Grob P, Cipriano DJ et al (2012) Synaptic proteins promote calcium-triggered fast transition from point contact to full fusion. elife 1:e00109.  https://doi.org/10.7554/eLife.00109CrossRefPubMedPubMedCentralGoogle Scholar
  10. 10.
    Reese C, Mayer A (2005) Transition from hemifusion to pore opening is rate limiting for vacuole membrane fusion. J Cell Biol 171:981–990.  https://doi.org/10.1083/jcb.200510018CrossRefPubMedPubMedCentralGoogle Scholar
  11. 11.
    Mayer A (1999) Intracellular membrane fusion: SNAREs only? Curr Opin Cell Biol 11:447–452.  https://doi.org/10.1016/S0955-0674(99)80064-7CrossRefPubMedGoogle Scholar
  12. 12.
    Dennison SM, Bowen ME, Brunger AT, Lentz BR (2006) Neuronal SNAREs do not trigger fusion between synthetic membranes but do promote PEG-mediated membrane fusion. Biophys J 90:1661–1675.  https://doi.org/10.1529/biophysj.105.069617CrossRefPubMedGoogle Scholar
  13. 13.
    Chen X, Araç D, Wang T-M et al (2006) SNARE-mediated lipid mixing depends on the physical state of the vesicles. Biophys J 90:2062–2074.  https://doi.org/10.1529/biophysj.105.071415CrossRefPubMedGoogle Scholar
  14. 14.
    Zick M, Wickner W (2016) Improved reconstitution of yeast vacuole fusion with physiological SNARE concentrations reveals an asymmetric Rab(GTP) requirement. Mol Biol Cell 27:2590–2597.  https://doi.org/10.1091/mbc.E16-04-0230CrossRefPubMedPubMedCentralGoogle Scholar
  15. 15.
    Zick M, Orr A, Schwartz ML et al (2015) Sec17 can trigger fusion of trans-SNARE paired membranes without Sec18. PNAS 112:E2290–E2297.  https://doi.org/10.1073/pnas.1506409112CrossRefPubMedGoogle Scholar
  16. 16.
    Zick M, Stroupe C, Orr A et al (2014) Membranes linked by trans-SNARE complexes require lipids prone to non-bilayer structure for progression to fusion. elife 3:e01879CrossRefGoogle Scholar
  17. 17.
    Brunger AT, Cipriano DJ, Diao J (2015) Towards reconstitution of membrane fusion mediated by SNAREs and other synaptic proteins. Crit Rev Biochem Mol Biol 50:231–241.  https://doi.org/10.3109/10409238.2015.1023252CrossRefPubMedPubMedCentralGoogle Scholar
  18. 18.
    Dürr M, Boller T, Wiemken A (1975) Polybase induced lysis of yeast spheroplasts. Arch Microbiol 105:319–327.  https://doi.org/10.1007/BF00447152CrossRefPubMedGoogle Scholar
  19. 19.
    Boller T, Dürr M, Wiemken A (1975) Characterization of a specific transport system for arginine in isolated yeast vacuoles. Eur J Biochem 54:81–91CrossRefGoogle Scholar
  20. 20.
    Ostrowicz CW, Meiringer CTA, Ungermann C (2008) Yeast vacuole fusion: a model system for eukaryotic endomembrane dynamics. Autophagy 4:5–19CrossRefGoogle Scholar
  21. 21.
    Wickner W (2002) Yeast vacuoles and membrane fusion pathways. EMBO J 21:1241–1247.  https://doi.org/10.1093/emboj/21.6.1241CrossRefPubMedPubMedCentralGoogle Scholar
  22. 22.
    Reese C, Heise F, Mayer A (2005) Trans-SNARE pairing can precede a hemifusion intermediate in intracellular membrane fusion. Nature 436:410–414.  https://doi.org/10.1038/nature03722CrossRefPubMedGoogle Scholar
  23. 23.
    Pieren M, Schmidt A, Mayer A (2010) The SM protein Vps33 and the t-SNARE H(abc) domain promote fusion pore opening. Nat Struct Mol Biol 17:710–717.  https://doi.org/10.1038/nsmb.1809CrossRefPubMedGoogle Scholar
  24. 24.
    Pieren M, Desfougères Y, Michaillat L et al (2015) Vacuolar SNARE protein transmembrane domains serve as nonspecific membrane anchors with unequal roles in lipid mixing. J Biol Chem 290:12821–12832.  https://doi.org/10.1074/jbc.M115.647776CrossRefPubMedPubMedCentralGoogle Scholar
  25. 25.
    D’Agostino M, Risselada HJ, Mayer A (2016) Steric hindrance of SNARE transmembrane domain organization impairs the hemifusion-to-fusion transition. EMBO Rep 17:1590–1608.  https://doi.org/10.15252/embr.201642209CrossRefPubMedPubMedCentralGoogle Scholar
  26. 26.
    Haas A, Conradt B, Wickner W (1994) G-protein ligands inhibit in vitro reactions of vacuole inheritance. J Cell Biol 126:87–97CrossRefGoogle Scholar
  27. 27.
    Strasser B, Iwaszkiewicz J, Michielin O, Mayer A (2011) The V-ATPase proteolipid cylinder promotes the lipid-mixing stage of SNARE-dependent fusion of yeast vacuoles. EMBO J 30:4126–4141.  https://doi.org/10.1038/emboj.2011.335CrossRefPubMedPubMedCentralGoogle Scholar
  28. 28.
    Karunakaran S, Fratti RA (2013) The lipid composition and physical properties of the yeast vacuole affect the hemifusion-fusion transition. Traffic 14:650–662.  https://doi.org/10.1111/tra.12064CrossRefPubMedPubMedCentralGoogle Scholar
  29. 29.
    Desfougères Y, Neumann H, Mayer A (2016) Organelle size control - increasing vacuole content activates SNAREs to augment organelle volume through homotypic fusion. J Cell Sci 129:2817–2828.  https://doi.org/10.1242/jcs.184382CrossRefPubMedGoogle Scholar
  30. 30.
    Schwartz ML, Merz AJ (2009) Capture and release of partially zipped trans-SNARE complexes on intact organelles. J Cell Biol 185:535–549.  https://doi.org/10.1083/jcb.200811082CrossRefPubMedPubMedCentralGoogle Scholar

Copyright information

© Springer Science+Business Media, LLC, part of Springer Nature 2019

Authors and Affiliations

  1. 1.Département de BiochimieUniversité de LausanneEpalingesSwitzerland

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