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Role of the V-ATPase in Regulation of the Vacuolar Fission–Fusion Equilibrium

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Like numerous other eukaryotic organelles, the vacuole of the yeast Saccharomyces cerevisiae undergoes coordinated cycles of membrane fission and fusion in the course of the cell cycle and in adaptation to environmental conditions. Organelle fission and fusion processes must be balanced to ensure organelle integrity. Coordination of vacuole fission and fusion depends on the interactions of vacuolar SNARE proteins and the dynamin-like GTPase Vps1p. Here, we identify a novel factor that impinges on the fusion–fission equilibrium: the vacuolar H+-ATPase (V-ATPase) performs two distinct roles in vacuole fission and fusion. Fusion requires the physical presence of the membrane sector of the vacuolar H+-ATPase sector, but not its pump activity. Vacuole fission, in contrast, depends on proton translocation by the V-ATPase. Eliminating proton pumping by the V-ATPase either pharmacologically or by conditional or constitutive V-ATPase mutations blocked salt-induced vacuole fragmentation in vivo. In living cells, fission defects are epistatic to fusion defects. Therefore, mutants lacking the V-ATPase display large single vacuoles instead of multiple smaller vacuoles, the phenotype that is generally seen in mutants having defects only in vacuolar fusion. Its dual involvement in vacuole fission and fusion suggests the V-ATPase as a potential regulator of vacuolar morphology and membrane dynamics.


Cellular compartments are very dynamic structures that adapt their shape, size, and number to cellular metabolism, differentiation state, and environmental conditions. Changes in mitochondrial shape during animal development (Chan, 2006), proliferation and degradation of peroxisomes in response to nutrient availability (Yan et al., 2005), and extensive fragmentation of the mammalian Golgi during mitosis (Shorter and Warren, 2002) are only a few examples highlighting organellar dynamics. Organelle homeostasis in the face of permanent membrane remodeling requires an equilibration and coordination of the fundamental processes of membrane fission and fusion.

Important insights into membrane dynamics have come from studies of the vacuole of the budding yeast Saccharomyces cerevisiae. Vacuoles are highly dynamic structures that undergo regulated cycles of membrane fission and fusion in the course of the cell cycle and in adaptation to changing environmental conditions. When a yeast cell buds and initiates growth of a daughter cell, the vacuole pinches off vesicles that migrate into the growing daughter cell where they fuse to form the new vacuolar compartment (Weisman, 2003). Vacuolar rearrangements also occur when yeast cells are faced with nutrient limitation or osmotic stress. Exposure of yeast cells to hypertonic medium induces fragmentation of the vacuole into numerous small vacuolar vesicles, a process involving Fab1p-mediated phosphatidylinositol-3,5-bisphosphate synthesis (Efe et al., 2005). Conversely, in fast adaptation to hypotonic shock, vacuoles fuse to give rise to a more voluminous vacuole. These rapid changes of the surface-to-volume ratio of the vacuole via fission or fusion allow uptake or release of water to restore the osmotic balance of the cell.

Vacuolar membrane dynamics can be readily analyzed in living yeast cells by fluorescence microscopy (Vida and Emr, 1995). Moreover, homotypic vacuole fusion can be assayed cell free on isolated organelles (Conradt et al., 1992; Haas et al., 1994). Detailed studies of vacuolar fusion have allowed to dissect the process into different stages and to categorize the molecular players accordingly (Wickner, 2002). In the first stage of vacuole fusion, priming, the ATPase Sec18p/NSF and its cofactor Sec17p/α-SNAP disassemble cis-soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) complexes, thereby activating them for fusion. Vacuoles are then tethered by the action of the Rab-GTPase Ypt7p, the homotypic fusion and vacuole protein sorting (HOPS) complex, and the Ccz1p–Mon1p complex (Wang et al., 2003). This allows trans-SNARE complexes to form and fusion partners to associate more tightly (docking). Addition of recombinant Vam7p to the in vitro fusion reaction promotes docking of vacuoles while bypassing the requirement for priming and reducing the level of Ypt7p needed to drive fusion (Thorngren et al., 2004). The final membrane fusion step involves further factors, such as the armadillo repeat protein Vac8p and the membrane sector of the vacuolar H+-ATPase (V0) (Peters et al., 2001; Veit et al., 2001; Wang et al., 2001; Bayer et al., 2003; Subramanian et al., 2006).

Vacuolar-type ATPases (V-ATPases) are multisubunit enzymes mediating ATP-driven translocation of protons from the cytosol into intracellular compartments or extracellular space (Nishi and Forgac, 2002; Graham et al., 2003; Kane, 2006). The V-ATPase complex is composed of the peripheral V1 domain (subunits A–H) and the membrane integral V0 domain (subunits a, c, c′, c″, d, and e). ATP hydrolysis by the V1 domain drives proton translocation through the V0 sector. In budding yeast, two isoforms of subunit a, Stv1 and Vph1, are expressed. Stv1-containing complexes are targeted to the late Golgi, whereas Vph1-containing complexes are found on vacuoles (Manolson et al., 1994). Both isoforms can partially substitute for each other. Besides their crucial role in intracellular pH regulation, V-ATPases have been implied in vacuole fusion in yeasts (Peters et al., 2001; Bayer et al., 2003), in exocytosis of multivesicular bodies in worms (Liegeois et al., 2006), in synaptic exocytosis in flies (Hiesinger et al., 2005), and in insulin secretion in mammalian cells (Sun-Wada et al., 2006). In all these systems, fusion requires physical presence of the V-ATPase complex, but not its proton translocation activity.


General Methods

Yeasts were cultured in YPD. For growth of V-ATPase mutants, YPD was buffered to pH 5.5 with 50 mM 2-(N-morpholino)ethanesulfonic acid.


Concanamycin A was purchased from MP Biomedicals (Irvine, CA), N- (3-triethylammoniumpropyl)-4-(p-diethylaminophenylhexatrienyl)-pyridinium dibromide (FM4-64; SynaptoRed C2) was from Biotium (Richmond, CA), and G-418 sulfate was from Calbiochem (San Diego, CA). Vam7p was recombinantly expressed and purified from Escherichia coli (Merz and Wickner, 2004).

Strains and Genetic Modifications

Yeast strains used are listed in Table 1. BJ3505 and DKY6281 are standard fusion strains (Haas et al., 1994). OMY1 is a pep4- and prb1-deficient strain (Muller et al., 2002). Deletion mutants for vam3 and nyv1 (Nichols et al., 1997), for ypt7 (Haas et al., 1995), and for vph1 (Bayer et al., 2003) have been described previously. BJ3505 vph1Δ GFP-Pho8p was generated by transformation of BJ3505 vph1Δ with pRS316 GFP-PHO8 expressing green fluorescent protein (GFP)-Pho8p under its endogenous promotor. The plasmid was a kind gift from Robert Piper (Department of Molecular Physiology and Biophysics, University of Iowa, Iowa City, IA). vma4-1 (JWY1) and the corresponding wild-type (WT) strain (SF838-5A) were generously provided by Patricia Kane (Department of Biochemistry and Molecular Biology, SUNY Upstate Medical University, Syracuse, NY) (Stevens et al., 1986; Zhang et al., 1998). CUY369a (vac8 cys4,5,7ala) was a kind gift from Christian Ungermann (Department of Biochemistry, University of Osnabrück, Osnabrück, Germany) (Subramanian et al., 2006). BY4741 and its vma1::kanMX4, vma2::kanMX4, vma5::kanMX4, and vph1::kanMX4 derivates as well as FY1679-13A and its mon1::kanMX4 derivative were purchased from Euroscarf (Frankfurt, Germany). BY4732 was obtained from the American Type Culture Collection (Manassas, VA). pep4::URA3 derivatives of FY1679, FY1679 mon1Δ, and BY4732 were constructed by one-step gene disruption with pTS15 (from Tom Stevens, Institute of Molecular Biology, University of Oregon, Eugene, OR). VAM2 was deleted in BY4732 pep4Δ by disruption with the HIS3 cassette by using pYVQ213 (from Yoh Wada, Division of Biological Sciences, Institute of Scientific and Industrial Research, Osaka University, Osaka, Japan) as described previously (Nakamura et al., 1997).

Table 1. Yeast strains used in this study

StrainGenotypeSource or reference
BJ3505MATa pep4::HIS3 prb1-Δ1.6R lys2-208 trp1-Δ101 ura3-52 gal2 canJones et al. (1982)
BJ3505 ccz1ΔBJ3505; ccz1::kanMXThis study
BJ3505 nyv1ΔBJ3505; nyv1::TRP1Nichols et al. (1997)
BJ3505 nyv1Δvph1ΔBJ3505 nyv1Δ; vph1::kanMXThis study
BJ3505 stv1ΔBJ3505; stv1::TRP1This study
BJ3505 stv1Δvph1ΔBJ3505 vph1Δ; stv1::TRP1This study
CUY369a (vac8 cys4,5,7ala)BJ3505; vac8::TRP1, VAC8pr::pRS406-VAC8pr-vac8cys4,5,7alaSubramanian et al. (2006)
BJ3505 vam3ΔBJ3505; vam3::TRP1Nichols et al. (1997)
BJ3505 vam3Δvma3ΔBJ3505 vma3Δ;vam3::kanMXThis study
BJ3505 vam6ΔBJ3505; vam6::kanMXThis study
BJ3505 vma3ΔBJ3505; vma3::TRP1This study
BJ3505 vma6ΔBJ3505; vma6::TRP1This study
BJ3505 vph1ΔBJ3505; vph1::kanMXBayer et al. (2003)
BJ3505 vph1Δ GFP-Pho8pBJ3505; vph1::kanMX; pRS316-GFP-PHO8This study
BJ3505 ypt7ΔBJ3505; ypt7::URA3Haas et al. (1995)
BY4732MATα his3Δ200 met15Δ0 trp1Δ63 ura3Δ0Brachmann et al. (1998)
BY4732 pep4ΔBY4732; pep4::URA3This study
BY4732 pep4Δvam2ΔBY4732 pep4Δ; vam2::HIS3This study
BY4741MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0Brachmann et al. (1998)
BY4741 vma1ΔBY4741; YDL185w::kanMX4Euroscarf
BY4741 vma2ΔBY4741; YBR127c::kanMX4Euroscarf
BY4741 vma3ΔBY4741; YEL027w::kanMX4Euroscarf
BY4741 vma5ΔBY4741; YKL080w::kanMX4Euroscarf
BY4741 vma6ΔBY4741; YLR447c::kanMX4Euroscarf
BY4741 vph1ΔBY4741; YOR270c::kanMX4Euroscarf
DKY6281MATα leu2-3 leu2-112 ura3-52 his3-Δ200 trpl-Δ901 lys2-801 suc2-Δ9 pho8::TRP1Haas et al. (1994)
DKY6281 vma6ΔDKY6281; vma6::kanMXThis study
DKY6281 vph1ΔDKY6281; vph1::kanMXBayer et al. (2003)
FCRN044-04B (AL)FY; MATα ura3-52 his3Δ200 leu2Δ1 LYS2 TRP1 YGL124c (23, 1893)::kanMX4Euroscarf
FY1679 pep4Δmon1ΔFCRN044-04B (AL); pep4::URA3This study
FY1679-2BMATα ura3-52 leu2Δ1 TRP1 his3Δ200 GAL2Winston et al. (1995)
FY1679-2B pep4ΔFY1679-2B; pep4::URA3This study
JWY1 (vma4-1ts)SF838-5A; vma4-1Zhang et al. (1998)
OMY1BY4732; pep4::URA3 prb1::TRP1Muller et al. (2002)
OMY1 vma6ΔOMY1; vma6::HIS3This study
SF838-5AMATα ade6 leu2-3 leu2-112 ura3-52 gal2Stevens et al. (1986)
W303aMATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100John York (Duke University, Durham, NC)
W303a vph1ΔW303a; vph1::kanMXThis study

All other deletion mutants were generated by polymerase chain reaction (PCR)-mediated gene disruption by using either the loxP-kanMX-loxP module from pUG6 (Guldener et al., 1996) or the auxotrophic marker genes HIS3 orTRP1 from pRS403 and pRS404, respectively (Brachmann et al., 1998). To generate the double knockout of nyv1 and vph1, the VPH1 open reading frame (ORF) was deleted in BJ3505 nyv1Δ by one-step gene replacement with the kanMX cassette from plasmid pUG6 (Guldener et al., 1996). Primers used have been described previously (Bayer et al., 2003). For generation of W303a vph1Δ, the VPH1 ORF was replaced by the kanMX module analogously. Parental strain W303a was kindly provided by John York (Department of Pharmacology and Cancer Biology, Duke University Medical Center, Durham, NC). CCZ1 was disrupted in BJ3505 with the same strategy by using primers Ccz1_Kan_fwd: 5′-CTG ATT CAA TAC CCT CTT TAA ACG AAA AAC TGT CCA TCA TAG CAG CTG AAG CTT CGT ACG C-3′ and Ccz1_Kan_rev: 5′-ATT TCC CTG GAT TCC CAC CAG TCA GTC ACG TCA CGA CCT AAA CGC ATA GGC CAC TAG TGG ATC TG-3′. For the generation of deletion mutants of VMA3, VMA6, or STV1 in BJ3505 or in BJ3505vph1Δ, the respective ORF was replaced by the TRP1 cassette from pRS404 (Brachmann et al., 1998) using the following primers: Vma3_Del_pRS_fwd: 5′-GAC TGA CCC TTG ATA GTT TTG TAC AAT TAT ACA CTC GTT CTG AGA TTG TAC TGA GAG TGC AC-3′, Vma3_Del_pRS_rev: 5′-CTC TAT TCC TGC TTT AGT GAT TCA GAA GCT GCC TTA ACA GAC CTG TGC GGT ATT TCA CAC CG-3′, Vma6_Del_pRS_fwd: 5′-CGC TAA GAG CTA GCA AAA GAG TAT ACC ATT CGG ATC GTG TTG CAG ACG CAG ATT GTA CTG AGA GTG CAC-3′, Vma6_Del_pRS_rev: 5′-CAC TAA CAC CAC ACG CTT GTA ACA AAC TAA GGA TGT CCT GAC CCT GTG CGG TAT TTC ACA CCG-3′, Stv1_Del_pRS_fwd: 5′-CGA GGC TTA CAT TAA GGT GCA TAT CAT AAT TCG CAA CAG ACG CAG ATT GTA CTG AGA GTG CAC-3′ and Stv1_Del_pRS_rev: 5′-CCA CTT GAC CGA CCC AGA CAT ATA TAT ACC GTA AAC TAT GTG CTG TGC GGT ATT TCA CAC CG-3′. VAM6 was replaced in BJ3505 by the kanMX module from pUG6 by using primers VAM6 Kan fw: 5′-GTC TTA TAT TGA TCA GCA AAA ACC CTT CAA AAT ATC AAT TCA GCT GAA GCT TCG TAC GC-3′ and VAM6 Kan rev: 5′-GAA ATA CTA ACA ACA ATA ACA GCA GCT GTT AAG GGA TCT TGC ATA GGC CAC TAG TGG ATC TG-3′.

FM4-64 Staining

To visualize vacuolar membranes in vivo, cells were stained with the vital, lipophilic dye FM4-64 as described previously (Vida and Emr, 1995) with the following modifications: Cells were grown overnight at 25°C to logarithmic phase (OD600 < 1) in YPD, YPD, pH 5.5, or selective medium. Cultures were adjusted to OD600 = 0.4, and 10 μM FM4-64 was added from a 10 mM stock solution in H2O. Cells were incubated for 1 h at 25°C, harvested (1 min; 3000 × g), washed twice in fresh medium, resuspended in YPD at OD600 = 0.4, and shaken for 2–3 h at 25°C. For microscopy, cells were pelleted (15 s; 8000 × g) and resuspended in YPD at OD600 = 10, and then they were immediately analyzed by spinning disk confocal microscopy by using an excitation laser at 488 nm and a 100× objective. To quantify vacuole morphology, photos of at least 10 random fields were taken. The number of visible vacuolar vesicles in 100 cells per experiment and condition was determined, and cells were accordingly grouped into one of four categories: one to two, three to four, five to eight, or more than eight vacuoles per cell. Data from three or more independent experiments were averaged, and the corresponding standard deviations were calculated.

Concanamycin A Treatment

Cells were grown and stained with FM4-64 as described above. Concanamycin A at 1 μM (MP Biomedicals) was added to the cultures from a 250× stock in dimethyl sulfoxide (DMSO) or ethanol. For kinetic analysis, cells were incubated for 10, 20, or 40 min at 25°C with shaking. For all other experiments, incubation was 2–3 h at 25°C with shaking. Control samples were treated with the corresponding solvent.

In Vivo Fragmentation Test

Yeast cells were grown to logarithmic phase, stained with FM4-64, and treated with concanamycin A if applicable. The culture medium was supplemented with 0.4 M NaCl from a 5 M stock solution in H2O. After 10-min incubation at 25°C, 200 μl of cells was centrifuged (15 s; 8000 × g), resuspended in 10 μl of their supernatant, and then immediately analyzed by fluorescence microscopy.

Heat Inactivation of the Conditional vma4 Allele

Cells carrying the conditional vma4-1 allele were grown logarithmically overnight at 25°C. Vacuolar membranes were labeled with FM4-64 as described above. For inactivation of vma4-1, an aliquot of the culture was shifted to 37°C for 40 min. Isogenic wild-type cells were treated in the same way as the mutant cells.

Vacuole Isolation and Fusion

Yeast was precultured in YPD medium (6–8 h; 30°C). Overnight cultures were inoculated (30°C; 14–16 h; 225 rpm) in baffled 2-l Erlenmeyer flasks with 1 l of YPD medium. At an OD600 of 1–1.5, cells were centrifuged (3 min; 4000 × g; 23°C; JA10 rotor), resuspended in 50 ml of 0.1 M Tris-Cl, pH 8.9, with 10 mM dithiothreitol, incubated (5 min; 30°C), centrifuged (3 min; 4000 × g; 2°C; JA10 rotor), resuspended in 15 ml of SB (50 mM K-phosphate, pH 7.5, 600 mM sorbitol in YPD with 0.2% dextrose and 3600 U ml−1 lyticase for BJ3505 or OMY1 and 1800 U ml−1 for DKY6281), and transferred into 30-ml Corex tubes. Cells were incubated (20 min; 30°C), reisolated (1 min, 800 × g; then 1 min, 1500 × g, 2°C; JA20 rotor), and resuspended in 2.5 ml of 15% Ficoll 400 in PS buffer [10 mM piperazine-N,N′-bis(2-ethanesulfonic acid) PIPES/KOH, pH 6.8, and 200 mM sorbitol] by gentle stirring with a glass rod. DEAE-dextran (200 μl for BJ3505 or OMY1 and 100 μl for DKY6281) was added from a frozen stock (0.4 g l−1) in 15% Ficoll in PS buffer. The cells were incubated (2 min at 0°C; 75 s at 30°C), chilled again, transferred to a SW41 tube, and overlaid with 3 ml of 8% Ficoll, 3 ml of 4% Ficoll, and 1.5 ml of 0% Ficoll (all in PS buffer). After centrifugation (90 min; 150,000 × g), the vacuoles were harvested from the 0%/4% interphase. A standard fusion reaction contained 3 μg of each vacuole type (isolated from strains BJ3505 or OMY1 and DKY6281) in a total volume of 30–35 μl of reaction buffer (20 mM PIPES/KOH, pH 6.8, 200 mM sorbitol, 150 mM KCl, 0.5 mM MgCl2, 0.5 mM MnCl2, 0.5 mM ATP, 7.5 μM Pefablock SC (Roche, Basel, Switzerland), 7.5 μg l−1 leupeptin, 3.75 μM o-phenanthroline, 37.5 μg l−1 pepstatin A, 20 mM creatine phosphate, and 35 U ml−1 creatine kinase). The ATP regenerating system, containing MgCl2, ATP, creatine phosphate, and creatine kinase, was added from a frozen 20× stock solution in PS buffer with 150 mM KCl. After 60 min at 27°C, alkaline phosphatase activity was determined as described previously (Wickner and Haas, 2000). One unit of fusion activity is defined as 1 μmol of p-nitrophenol developed per minute and micrograms of BJ3505 vacuoles at 27°C.


In previous reports, we have addressed the involvement of the vacuolar H+-ATPase in homotypic fusion of the yeast vacuole (Peters et al., 2001; Bayer et al., 2003; Muller et al., 2003). Using a cell-free reaction of vacuole fusion, we have shown that the V0 sector of the V-ATPase plays a direct role in vacuole fusion that is distinct from its function in proton translocation. Mutation of fusion-relevant components often results in fragmentation of vacuoles into multiple small vesicles in vivo (Raymond et al., 1992; Wada et al., 1992). However, most deletion mutants of the V-ATPase do not display fragmented vacuoles in vivo (Seeley et al., 2002; Graham et al., 2003). This poses the question whether V-ATPase is either not necessary for fusion of vacuoles in vivo or whether the reverse process, vacuole fragmentation, could be affected by it. Vacuolar integrity in a given strain should be determined by the differential rates of vacuole fusion and fission. If fusion prevails over fission, only a few larger vacuoles should be observed. If, in contrast, fusion is outweighed by fission, vacuoles should be more numerous and smaller. We sought to understand the seemingly contradictory phenotype of the V-ATPase mutants; hence, we began to characterize the role of V-ATPase in the reaction reversing vacuole fusion, i.e., the fission of vacuoles into smaller fragments.

Vacuole Structure of V-ATPase Mutants Correlates to Their Ability to Acidify the Vacuole

Loss of V-ATPase function is associated with the vacuolar membrane H+-ATPase (Vma) phenotype. This phenotype is characterized by complete loss of vacuolar acidification and ATPase activity, inability to grow on neutral media, and sensitivity to high Ca2+ concentrations (Kane, 2006). Deletion of any of the core V-ATPase subunits, except for subunit a, results in the Vma phenotype. Subunit a is the only V-ATPase subunit in yeast that is expressed in two isoforms, the vacuolar isoform Vph1p and the Golgi/endosomal isoform Stv1p. They can substitute for one another in proton translocation (Manolson et al., 1994). Deletion of Vph1p does not result in the full Vma phenotype (Manolson et al., 1994) because the Stv1p-containing V-ATPase isoform remains. Vacuoles of vph1Δ cells still show basal vacuole acidification (Perzov et al., 2002).

We assayed the vacuolar morphology of V-ATPase mutants by microscopy after labeling the vacuolar membranes with the red fluorescent vital dye FM4-64 (Vida and Emr, 1995). vma mutants deleted for the V1-subunit Vma1p or the V0 subunits Vma3p or Vma6p typically showed one enlarged vacuole per cell (Figure 1A). In contrast to partial deletions of the V0 subunit Vph1p in W303 background, for which no fragmentation was observed (Perzov et al., 2002), complete knockouts of Vph1p displayed numerous small vesicle-like structures, independently of the strain background (BY4741, BJ3505, or W303a; Figure 1, A and B). Visualization of vacuolar membranes in vph1Δ by fluorescence microscopy of the vacuolar marker proteins GFP-Pho8p (Figure 1A) and Vtc1p-GFP (data not shown) identified these structures as vacuolar fragments. This link between vacuolar acidification and morphology suggested that proton translocation by the V-ATPase might have an impact on vacuole morphology, possibly by interfering with the balance between fusion and fission events at vacuolar membranes.

Figure 1.

Figure 1. Vacuolar morphology of V-ATPase mutants. (A) vma1Δ, vma3Δ, vph1Δ, and WT (BY4741) yeast cells were grown in YPD, pH 5.5, to logarithmic phase, stained with FM4-64, and analyzed by spinning disk confocal microscopy by using an excitation laser at 488 nm and a 100× objective. As an alternative means to visualize vacuolar membranes, GFP-Pho8p was expressed in vph1Δ cells and revealed by fluorescence microscopy as described above. (B) Vacuolar membranes of wild-type and vph1Δcells (in BJ3505 and W303a background) were labeled with FM4-64 and visualized by fluorescence microscopy.

The V-ATPase Is Required for Both Vacuole Fusion and Vacuole Fragmentation

Vacuole fusion can be studied on isolated organelles in a cell-free assay. Vacuoles are purified from a strain that expresses the enzymatically inactive proform of the alkaline phosphatase (pro-Pho8p) and from a strain that lacks alkaline phosphatase but provides the vacuolar proteases (Pep4p and Prb1p) required for maturation of pro-Pho8p. Vacuoles from both strains are mixed and incubated in the presence of ATP and salt. When vacuoles fuse, their contents mix; hence, Pep4p and Prb1p gain access to proalkaline phosphatase, resulting in activation of Pho8p. Alkaline phosphatase activity serves as readout for fusion (Haas et al., 1994).

We analyzed vacuole fusion activity of mutants deleted for the V0 subunits Vph1p or Vma6p by using this system. Vacuoles from vph1Δ incubated either alone or in combination with wild-type vacuoles did not fuse efficiently (Figure 2). Addition of the recombinant target membrane–associated- SNARE subunit Vam7p to the fusion reaction of vph1Δ vacuoles enhanced fusion only slightly (from 13 to 18% of wild type). Similarly, vacuoles from vma6Δ showed only marginal fusion activity (18%), when incubated alone. Recombinant Vam7p increased fusion of vma6Δ to 34% of wild-type. If vma6Δ vacuoles were fused in combination with wild-type vacuoles; however, 64–68% of wild-type fusion activity was reached. In the presence of recombinant Vam7p, fusion was even stimulated to wild-type levels. For the moment, we cannot explain the differential fusion defects of vacuoles from vph1Δ and vma6Δ cells with certainty. We speculate that the severe Vma growth phenotype associated with the complete loss of the V-ATPase in vma6Δ cells (Graham et al., 2003) might trigger adaptive changes in cellular traffic that could alleviate the vacuolar fusion defects resulting from disruption of VMA6. Because vph1Δ cells do not show the severe Vma phenotype, they might not develop similar compensation. Further studies will be necessary to analyze the differences in the vacuolar fusion machineries between the two strains.

Figure 2.

Figure 2. Fusion activity of V0 mutants. Vacuoles were prepared from WT and mutant cells (vma6Δ and vph1Δ). Their fusion activity was assayed in standard reactions run in the presence or absence of recombinant 0.1 g l−1 Vam7p. After 60 min on ice or at 27°C, alkaline phosphatase (ALP) activities were determined. ALP activity of the wild-type control at 27°C was 3–5 U. Ice values varied from 0.2 to 0.35 U, and they were subtracted from the respective 27°C values.

We tested a potential role of the V-ATPase in vacuole fission by using mutations resulting in conditional and constitutive deficiency of V-ATPase function. Fragmentation of yeast vacuoles in response to high salt can be readily analyzed in living yeast cells (Bonangelino et al., 2002). Vacuolar membranes were stained with the vital dye FM4-64. Subsequently, cells were subjected to a 10-min incubation with 0.4 M NaCl, and they were immediately analyzed by fluorescence microscopy. Wild-type cells typically displayed one to four large vacuoles that were readily split into several vacuolar fragments upon salt treatment (Figures 3, A and B, and 4; note that different wild-type strains may differ in steady-state vacuole size and number). In contrast to the wild-type, deletion mutants of the V-ATPase catalytic headgroup subunits Vma1p and Vma2p, of the V1-stalk-subunit Vma5p, of the proteolipid-ring constituent Vma3p, and of the V0 rotor-subunit Vma6p maintained their single enlarged vacuole in high salt (Figure 4A). Although these mutations abolish all cellular V-ATPase activity, knockout of Stv1p removes only the Golgi/endosomal isoform of the V-ATPase, but leaves the vacuolar isoform intact. Deletion of Stv1 did not reduce fragmentation activity (Figure 4B). The observed fragmentation defects thus suggest that both the V1 and the V0 sectors are necessary for vacuolar fragmentation and that only the vacuole-localized isoform of the V-ATPase (containing Vph1p) is necessary for fission.

Figure 3.

Figure 3. In vivo test for vacuole fragmentation. (A) Vacuole fragmentation in response to hypertonic stress. Wild-type cells (BJ3505) were grown in YPD at 25°C to logarithmic phase. Cells were stained with FM4-64, incubated for 10 min in the presence or absence of 0.4 M NaCl, and analyzed by fluorescence microscopy as described in Figure 1. (B) Quantification of vacuole morphology. The number of vacuolar vesicles per cell was determined, and cells were accordingly grouped into the indicated categories. For each experiment, 100 cells per strain and condition were analyzed. Three independent experiments were averaged, and standard deviations were calculated.

Figure 4.

Figure 4. Fragmentation defects of V-ATPase mutants. Salt-induced vacuole fragmentation in V0 and V1 mutants. Yeasts were logarithmically grown in YPD, pH 5.5, at 25°C, stained with FM4-64, and subjected to a 10-min salt shock. Vacuolar morphology was quantified as described in Figure 1. (A) Wild-type, vma1Δ, vma2Δ, vma5Δ, vma3Δ, vma6Δ (in BY4741). (B) Wild-type and stv1Δ (in BJ3505). (C) vma4-1ts and its isogenic wild-type SF838-5A were treated and analyzed as described above, but the cells had been transferred to 37°C, or they were kept at 25°C for 40 min before the salt treatment.

To ensure that the observed fragmentation defects were due to a direct role of the V-ATPase in fission rather than to potential secondary effects resulting from the Vma phenotype, we analyzed the fragmentation activity of a temperature-conditional V-ATPase mutant. The vma4-1ts allele provides normal V-ATPase function at 25°C, but it results in loss of vacuolar acidification upon shift to the restrictive temperature of 37°C (Zhang et al., 1998). vma4-1ts mutants grown at 25°C displayed wild-type–like vacuole morphology and fragmentation activity (Figure 4C). If cells were shifted to the restrictive temperature of 37°C 40 min before salt treatment, vacuole fragmentation was strongly impaired. A partial inhibition of fragmentation was apparent also in the wild-type at 37°C, however, to a significantly lesser degree than in the vma4-1ts cells.

Proton Translocation by the V-ATPase Is Necessary for Fission

To further analyze the function of the V-ATPase in fission, we tested the effects of the V-ATPase inhibitor concanamycin A on salt-induced fragmentation of wild-type cells. Concanamycin A is a macrolide antibiotic that blocks V-ATPase–dependent proton translocation at nanomolar concentrations (Drose and Altendorf, 1997). We treated yeast cultures for 10, 20, or 40 min in the presence of 1 μM concanamycin A before testing fragmentation activity. Wild-type cells treated only with the solvent of concanamycin A fragmented their vacuoles in response to high salt. However, cells that had received concanamycin A showed significantly reduced fragmentation activity (Figure 5) already after 10 min of concanamycin A treatment. A small fraction of cells (5%) displayed a single large vacuole surrounded by multiple smaller vacuolar fragments. This phenotype is reminiscent of vacuole structures seen in the deletion mutant of the yeast dynamin-like GTPase Vps1p, which is defective in vacuole fission (Peters et al., 2004).

Figure 5.

Figure 5. Proton translocation by the V-ATPase is indispensable for vacuole fragmentation. Effect of concanamycin A on in vivo vacuole fragmentation of wild-type cells (BJ3505). To block proton pumping by the V-ATPase, cells were incubated with 1 μM concanamycin A for 10, 20, or 40 min before hyperosmotic shock in 0.4 M NaCl.

The apparent suppression of fragmentation by concanamycin A likely results from reduced fission. An alternative explanation would be enhanced fusion activity that immediately reverses and thus hides the fragmentation event. If enhancement of fusion underlies the fragmentation defect, inactivation of a fusion component by mutation should render vacuole fragmentation insensitive toward concanamycin A treatment. Hence, we tested the effect of concanamycin A on salt-induced fragmentation of a mutant lacking the fusion-relevant vacuolar vesicle (v)-SNARE Nyv1p (Nichols et al., 1997). In this mutant, fragmentation activity was blocked by concanamycin A to the same extent as in wild-type cells (Figure 6, A and B). Hence, the abolishment of fragmentation by concanamycin A is unlikely to be due to enhanced fusion activity. We conclude that vacuole fragmentation requires proton translocation by the V-ATPase.

Figure 6.

Figure 6. Suppression of fragmentation by concanamycin A results from reduced fission and not from enhanced fusion. Effect of concanamycin A on in vivo vacuole fragmentation of wild-type (A) and fusion-defective nyv1Δ cells (B; both in BJ3505). To block proton pumping by the V-ATPase, cells were incubated with 1 μM concanamycin A for 2 h before hyperosmotic shock in 0.4 M NaCl.

Fission Defects Are Epistatic to Fusion Defects

Because the V-ATPase fulfills a dual role in vacuolar fusion and fission, we sought to define whether one role might predominate over the other in vivo. Vacuole fusion requires the physical presence of the V-ATPase, but, in contrast to fission, it does not depend on the proton pump function of the V-ATPase (Bayer et al., 2003; Muller et al., 2003). The V-ATPase mutant vph1Δ shows strongly reduced fusion activity (Figure 2), but it retains basal vacuole acidification and it shows vacuolar fragmentation in vivo. Hence, it offered a possibility to directly test the possibility of epistasis of fission over fusion. If the fission defect caused by elimination of vacuolar acidification was epistatic to a fusion defect caused by the physical absence of the V0 complex from the vacuole, it should be possible to cure vacuolar fragmentation in a vph1Δ mutant by abolishing V-ATPase pump activity. To test this hypothesis, we prevented vacuole acidification in vph1Δ both genetically and pharmacologically. We deleted STV1, the gene encoding for the Golgi/endosomal isoform of the yeast a subunit, in the vph1Δ background. The resulting vph1Δ stv1Δ double mutants show a complete loss of vacuole acidification (Manolson et al., 1994). The additional deletion of Stv1p restored morphology of the fragmented vph1Δ vacuoles and yielded cells with only few enlarged vacuoles (Figure 7A). As an independent means to block proton translocation by the V-ATPase, we incubated wild-type and vph1Δ cells with 1 μM of the pump inhibitor concanamycin A before microscopic analysis. Concanamycin A treatment drastically reduced the fraction of vph1Δ cells displaying highly fragmented vacuoles (>8 vacuoles/cell) from 55 to 11% while increasing the number of cells showing one to two vacuoles from 14 to 62%. Concanamycin A only had small effects on vacuole structure of wild-type cells (Figure 7, B and C). The fraction of cells having one to two vacuoles per cell increased from 81 to 90%, whereas the percentage of cells with three to four vacuoles dropped from 18 to 8%. Cells generally displayed large spherical vacuoles. These results indicate that the defects of fission caused by the lack of vacuolar proton pump activity dominate over the defects in vacuole fusion caused by physical absence of the V0 sector of the V-ATPase.

Figure 7.

Figure 7. Epistasis of fragmentation and fusion defects. (A) Vacuole morphology of subunit a deletion mutants. Vacuolar membranes of wild-type, vph1Δ, stv1Δ, and stv1Δvph1Δ cells (all in BJ3505 background) were labeled with FM4-64 and visualized by fluorescence microscopy. (B) Effect of concanamycin A on vacuolar morphology of wild-type and vph1Δ cells. Cells were stained with FM4-64, and subsequently they were incubated for 2 h with 1 μM concanamycin A or control buffer (DMSO). Microscopy was as described in A. (C) Quantification of the experiment in B. (D) Vacuole morphology of nyv1Δvph1Δ cells after 2-h concanamycin A treatment as described in B.

We tested whether the reversal of fragmentation occurred along the authentic pathway of SNARE-dependent fusion by measuring the effect of concanamycin A on the vacuole structure of a vph1Δ strain deleted for the vacuolar v-SNARE Nyv1p. Similar to vph1Δ, the double-knockout %vph1Δnyv1Δ displayed highly fragmented vacuoles. Unlike the single knockout, however, %vph1Δnyv1Δ cells maintained fragmented vacuoles in the presence of concanamycin A (Figure 7D). Quantification revealed only a moderate decrease in the fraction of cells with highly fragmented vacuoles (>8 vacuoles/cell) from 69 to 40%. The percentage of cells with five to eight vacuoles rose from 13 to 22%, of cells showing three to four vacuoles from 10 to 17% and of cells having one to two vacuoles from 8 to 20%. Thus, deletion of Nyv1p counteracted the reversion of vacuole structure in vph1Δ. This observation indicates that the phenotypic conversion is SNARE dependent and that it follows the normal fusion pathway.

Deletion of early-acting fusion factors that eliminate docking of the membranes completely prevents fusion. Deletion of late-acting factors can be slightly less severe, because spontaneous fusion of docked membranes can ensue if they are simply held in contact long enough. Consequently, we compared with which degree the vacuolar morphology of early-acting fusion mutants could be reverted by blocking proton translocation. Deletion of the V0 subunit Vma3p in the SNARE knockout vam3Δ had no effect on vacuolar structure: vacuoles seemed highly fragmented in both strains (Figure 8A). Equally, blocking V-ATPase activity with concanamycin A in vam3Δ and in deletion mutants of other early-acting fusion factors such as the SNARE Vam7p, the Rab-GTPase Ypt7p, and the HOPS complex components Vam2p and Vam6p as well as Ccz1p and Mon1p did not influence vacuole morphology (Figure 8, B–E; data not shown). However, in a mutant of Vac8p, a factor that is required in a late stage of the fusion reaction (Wang et al., 2001), a reduction in the fraction of cells having five and more vacuoles from 18 to 4% (>8 vacuoles/cell) and from 19 to 10% (5–8 vacuoles/cell) was observed after concanamycin A treatment (Figure 8F). Concurrently, the percentage of cells with one to two and three to four vacuoles increased from 42 to 60% and from 20 to 25%, respectively. These experiments argue for an essential role of early-acting fusion factors in phenotypic conversion, and they suggest that only the effect of late-acting factors, such as Vac8p and Vph1p, can be suppressed.

Figure 8.

Figure 8. Rescue of fusion-defective mutants by blocking proton pumping. (A) Vacuole structure in wild-type, vam3Δ, vma3Δ, and %vam3Δvma3Δ cells (all in BJ3505). Labeling and microscopy are as described in Figure 7A. (B–F) Effect of concanamycin A on the morphology of the indicated fusion-defective mutants (all in BJ3505 background). Labeling, concanamycin A treatment, and microscopy are as described in Figure 7B.


Here, we have identified a requirement of V-ATPase proton pump activity for vacuole fission. This requirement of V-ATPase activity for vacuole fragmentation may relate to previous findings on this process: Vacuole fragmentation in response to hyperosmotic shock is initiated by Fab1p-dependent synthesis of phosphatidylinositol-3,5-bisphosphate [PtdIns(3,5)P2]. PtdIns(3,5)P2 may promote fission through recruitment of effectors such as Svp1p/Atg18p to the vacuole (Dove et al., 2004). However, in addition, fab1 mutants show a reduction of vacuolar acidification, which led to the suggestion that PtdIns(3,5)P2 could modulate V-ATPase activity (Weisman, 2003). Because we have now found that vacuolar acidification itself is crucial for fission, future studies should address the possibility that PtdIns(3,5)P2 supports vacuolar fission by regulating V-ATPase activity.

Because the V-ATPase has a dual role in vacuolar fission (as a pump) and in fusion (by physical presence of the V0 sector and its interaction with SNAREs) (Peters et al., 2001; Bayer et al., 2003), disruption of the V-ATPase will affect both fission and fusion. Our data are consistent with the hypothesis that the apparent vacuole morphology results from a true equilibrium of the competing reactions of fusion and fission. The final outcome depends on the ratio of these reaction rates rather than on their absolute magnitude. Interfering with either one or both reactions by genetic manipulation or drug administration results in a shift of the fission–fusion equilibrium (Figure 9), which can explain the phenotype of V-ATPase mutants: The V-ATPase mutant vph1Δ retains basal vacuolar acidification (which may still support fission, even though perhaps at lower rate), and it has, in addition, the most pronounced fusion defect. Therefore, fission still prevails and leads to a cell with fragmented vacuoles (Figure 9, type 3). Residual vacuolar acidification in vph1Δ could be eliminated either by additional deletion of Stv1p or by concanamycin A treatment. These manipulations may have reduced fission activity enough to allow it to be outweighed by fusion. Thus, the equilibrium of these two reduced rates now favors the restoration of a single large vacuole in vph1Δ cells (Figure 9, type 2). This produces the same vacuolar phenotype as seen in all other V-ATPase deletion mutants that completely lack vacuolar proton pump activity. The epistasis of fission over fusion provides a satisfactory explanation for the fact that, in contrast to numerous other vacuolar fusion mutants, V-ATPase mutants do not show a vacuole fragmentation phenotype.

Figure 9.

Figure 9. Schematic representation of the vacuolar fission-fusion equilibrium. Vacuolar morphology is determined by the equilibrium of the antagonistic processes of fission and fusion that constantly takes place with the specific rates kfis and kfus. Manipulations interfering with fission and/or fusion result in modified reaction rates. The relative changes in fission and fusion rates determine whether and to what extent the equilibrium point is shifted and hence the phenotypic outcome. Three cases can be distinguished: 1) Fission and fusion rates are balanced. Vacuole morphology is normal. 2) Fission is more strongly affected than fusion. As a result fusion prevails, leading to a cell with a single large vacuole. and 3) Fission is less strongly affected than fusion. As a consequence, fission outweighs fusion, resulting in a cell with fragmented vacuoles.

Vacuole fusion proceeds through various steps. The Rab-GTPase Ypt7p, the HOPS complex of tethering factors, and a set of SNARE proteins are required for docking the membranes and for keeping them in close contact. Subsequently, further protein factors such as the V0 sector and Vac8p are necessary to efficiently induce fusion of the apposed membranes. Disruption of proton pumping by the V-ATPase restored vacuolar morphology in both vph1Δ and in vac8 mutant cells. In contrast, mutants of early fusion factors such as SNAREs, the Rab-GTPase Ypt7p, the HOPS complex, Mon1p, and Ccz1p were not rescued to a similar extent. This can be explained by the observation that the early factors are needed to keep the membranes in tight apposition, which is an irreplaceable prerequisite for fusion. Bilayer fusion itself requires catalysis by late-acting factors, but it may also occur spontaneously, although at modest rates, if only the vacuoles are tethered and docked. Due to spontaneous fusion disruption of late-acting factors may lead to less profound blocks of fusion than defects in early-acting factors. Hence, fusion defects of early-acting factors may be less readily compensated by reductions of fission activity that can be produced by eliminating vacuolar proton pump activity (Figure 9, types 2 and 3).

Differential reduction of fusion rates may also account for the in vivo phenotype of other mutants in vacuolar fusion factors that do not show fragmented vacuoles, such as the Vtc proteins Vtc1p and Vtc4p, which are involved in SNARE priming (Muller et al., 2002, 2003). The in vitro fusion activity of vacuoles from these strains is less severely impaired than that of vam3Δ vacuoles. This becomes especially evident if mutant vacuoles are fused in combination with wild-type vacuoles. A vam3Δ/WT combination is virtually nonfusogenic (<10%), whereas a vtc1Δ/WT or a vac8Δ/WT combination fuses well, i.e., with 50–70% of the activity of a WT/WT combination (Nichols et al., 1997; Veit et al., 2001) (Müller and Mayer, unpublished data). Consequently, the equilibrium of activities in vtc mutants might still favor the maintenance of large vacuoles.

The electrochemical gradient produced by proton translocation could influence fission in several ways. Association of peripheral membrane proteins with lipid bilayers often depends on the membrane potential. It is conceivable that cytosolic components of the fission machinery might interact with vacuolar membranes, depending on the pH at the membrane surface. Precedence for such a mode of interaction exists, as exemplified by the recruitment of the cytosolic small GTPase Arf6p and its cognate GDP/GTP exchange factor ADP-ribosylation factor nucleotide site opener (ARNO) to endosomal membranes (Hurtado-Lorenzo et al., 2006). Here, increasing acidification of endosomes along the degradative pathway presumably triggers conformational rearrangements of the V-ATPase that permit subsequent association of Arf6p and ARNO with the V0 sector. The electrochemical gradient could also influence transport processes across vacuolar membranes, e.g., the pH-dependent activity of vacuolar amino acid or ion transporters and exchangers. In this way, V-ATPase activity might modulate the flux of solutes and water across the vacuolar membrane. Such flux may be connected to vacuole fission, and it may even be essential to it because the fragmentation of a large vacuole into multiple small vesicles will alter the surface to volume ratio. If the available membrane surface stays constant multiple vacuolar fragments will provide much less volume than a single large vacuole. Fragmentation of a large vacuole will hence require an extrusion of water and perhaps also solutes from the organelle to adapt the surface-to-volume ratio.

Finally, the vacuolar proton gradient could also influence membrane structure directly. The transbilayer distribution of phospholipids in large unilamellar vesicles is highly pH sensitive (Hope et al., 1989). And a redistribution of only a small fraction of phospholipids suffices to induce shape changes in giant liposomes (Farge and Devaux, 1992). At the yeast vacuole, elimination of the vacuolar pH gradient might similarly modulate the transmembrane distribution of phospholipids, and in this way it may change the fission characteristics of the membrane. Moreover, the pH at the membrane surface could affect membrane curvature through modification of the charge of lipids and membrane proteins, which can change their conformation, shape, spontaneous curvature, and assembly state. In line with this, the pH-dependent self-assembly of lysobisphosphatidic acid is essential for the formation of multivesicular liposomes in vitro (Matsuo et al., 2004). Also, other vacuolar processes that involve significant shape changes require a pH gradient, e.g., formation of autophagic tubes (large vacuolar invaginations forming during microautophagy) and scission of vesicles from their tip into the vacuole lumen (Kunz et al., 2004).


This article was published online ahead of print in MBC in Press ( on July 25, 2007.

Abbreviations used:

homotypic fusion and vacuole protein sorting




membrane sector of the vacuolar H+-ATPase


peripheral sector of the vacuolar H+-ATPase


vacuolar-type ATPase


vacuolar membrane H+-ATPase.


We thank Monique Reinhardt, Andrea Schmidt, Véronique Comte, and Susanne Bühler for technical assistance and members of the Mayer laboratory for helpful discussion. We are grateful to Patricia Kane, Robert Piper, Tom Stevens, Christian Ungermann, Yoh Wada, and John York for sharing strains and plasmids. This work was supported by grants from Fonds National Suisse, Roche, and Human Frontier Science Program to A.M. and C.P.


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