![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
Vol. 18, Issue 10, 3873-3882, October 2007
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
,
*Département de Biochimie, Université de Lausanne, 1066 Epalinges, Switzerland; and Friedrich-Miescher-Laboratorium der Max-Planck-Gesellschaft, 72076 Tübingen, Germany
Submitted March 5, 2007;
Revised July 9, 2007;
Accepted July 16, 2007
Monitoring Editor: Akihiko Nakano
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
), 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/
-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.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Reagents
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 vph1
GFP-Pho8p was generated by transformation of BJ3505 vph1 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 mon1, 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 pep4 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).
|
) 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 nyv1 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 vph1, 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 BJ3505vph1, 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 x 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 x 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 100x 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 250x 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 x 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 x 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 x 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 x g; then 1 min, 1500 x 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 x 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 20x 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.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
; 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 vph1 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 vph1 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.
|
).
We analyzed vacuole fusion activity of mutants deleted for the V0 subunits Vph1p or Vma6p by using this system. Vacuoles from vph1 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 vph1 vacuoles enhanced fusion only slightly (from 13 to 18% of wild type). Similarly, vacuoles from vma6 showed only marginal fusion activity (18%), when incubated alone. Recombinant Vam7p increased fusion of vma6 to 34% of wild-type. If vma6 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 vph1 and vma6 cells with certainty. We speculate that the severe Vma– growth phenotype associated with the complete loss of the V-ATPase in vma6 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 vph1 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.
|
). 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.
|
|
). 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).
|
). 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.
|
; Muller et al., 2003). The V-ATPase mutant vph1 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 vph1 mutant by abolishing V-ATPase pump activity. To test this hypothesis, we prevented vacuole acidification in vph1 both genetically and pharmacologically. We deleted STV1, the gene encoding for the Golgi/endosomal isoform of the yeast a subunit, in the vph1 background. The resulting vph1 stv1 double mutants show a complete loss of vacuole acidification (Manolson et al., 1994). The additional deletion of Stv1p restored morphology of the fragmented vph1 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 vph1 cells with 1 µM of the pump inhibitor concanamycin A before microscopic analysis. Concanamycin A treatment drastically reduced the fraction of vph1 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.
|
strain deleted for the vacuolar v-SNARE Nyv1p. Similar to vph1, the double-knockout %vph1nyv1 displayed highly fragmented vacuoles. Unlike the single knockout, however, %vph1nyv1 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 vph1. 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 vam3 had no effect on vacuolar structure: vacuoles seemed highly fragmented in both strains (Figure 8A). Equally, blocking V-ATPase activity with concanamycin A in vam3 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.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). 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 vph1 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 vph1 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 vph1 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.
|
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 vam3 vacuoles. This becomes especially evident if mutant vacuoles are fused in combination with wild-type vacuoles. A vam3/WT combination is virtually nonfusogenic (<10%), whereas a vtc1/WT or a vac8/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).
|
ACKNOWLEDGMENTS |
|---|
|
Footnotes |
|---|
Present address: Zentrum für Molekulare Biologie der Universität Heidelberg, Im Neuenheimer Feld 282, 69120 Heidelberg, Germany. 
Address correspondence to: Andreas Mayer (andreas.mayer{at}nil.ch)
Abbreviations used: HOPS, homotypic fusion and vacuole protein sorting; PtdIns(3,5)P2, phosphatidylinositol-3,5-bisphosphate; V0, membrane sector of the vacuolar H+-ATPase; V1, peripheral sector of the vacuolar H+-ATPase; V-ATPase, vacuolar-type ATPase; Vma, vacuolar membrane H+-ATPase.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Bonangelino, C. J., Nau, J. J., Duex, J. E., Brinkman, M., Wurmser, A. E., Gary, J. D., Emr, S. D., and Weisman, L. S. (2002). Osmotic stress-induced increase of phosphatidylinositol 3,5-bisphosphate requires Vac14p, an activator of the lipid kinase Fab1p. J. Cell Biol 156, 1015–1028.
Brachmann, C. B., Davies, A., Cost, G. J., Caputo, E., Li, J., Hieter, P., and Boeke, J. D. (1998). Designer deletion strains derived from Saccharomyces cerevisiae S288C: a useful set of strains and plasmids for PCR-mediated gene disruption and other applications. Yeast 14, 115–132.[CrossRef][Medline]
Chan, D. C. (2006). Mitochondria: dynamic organelles in disease, aging, and development. Cell 125, 1241–1252.[CrossRef][Medline]
Conradt, B., Shaw, J., Vida, T., Emr, S., and Wickner, W. (1992). In vitro reactions of vacuole inheritance in Saccharomyces cerevisiae. J. Cell Biol 119, 1469–1479.
Dove, S. K. et al. (2004). Svp1p defines a family of phosphatidylinositol 3,5-bisphosphate effectors. EMBO J 23, 1922–1933.[CrossRef][Medline]
Drose, S., and Altendorf, K. (1997). Bafilomycins and concanamycins as inhibitors of V-ATPases and P-ATPases. J. Exp. Biol 200, 1–8.[Abstract]
Efe, J. A., Botelho, R. J., and Emr, S. D. (2005). The Fab1 phosphatidylinositol kinase pathway in the regulation of vacuole morphology. Curr. Opin. Cell Biol 17, 402–408.[CrossRef][Medline]
Farge, E., and Devaux, P. F. (1992). Shape changes of giant liposomes induced by an asymmetric transmembrane distribution of phospholipids. Biophys. J 61, 347–357.[Medline]
Graham, L. A., Flannery, A. R., and Stevens, T. H. (2003). Structure and assembly of the yeast V-ATPase. J. Bioenerg. Biomembr 35, 301–312.[CrossRef][Medline]
Guldener, U., Heck, S., Fielder, T., Beinhauer, J., and Hegemann, J. H. (1996). A new efficient gene disruption cassette for repeated use in budding yeast. Nucleic Acids Res 24, 2519–2524.
Haas, A., Conradt, B., and Wickner, W. (1994). G-protein ligands inhibit in vitro reactions of vacuole inheritance. J. Cell Biol 126, 87–97.
Haas, A., Scheglmann, D., Lazar, T., Gallwitz, D., and Wickner, W. (1995). The GTPase Ypt7p of Saccharomyces cerevisiae is required on both partner vacuoles for the homotypic fusion step of vacuole inheritance. EMBO J 14, 5258–5270.[Medline]
Hiesinger, P. R. et al. (2005). The v-ATPase V0 subunit a1 is required for a late step in synaptic vesicle exocytosis in Drosophila. Cell 121, 607–620.[CrossRef][Medline]
Hope, M. J., Redelmeier, T. E., Wong, K. F., Rodrigueza, W., and Cullis, P. R. (1989). Phospholipid asymmetry in large unilamellar vesicles induced by transmembrane pH gradients. Biochemistry 28, 4181–4187.[CrossRef][Medline]
Hurtado-Lorenzo, A. et al. (2006). V-ATPase interacts with ARNO and Arf6 in early endosomes and regulates the protein degradative pathway. Nat. Cell Biol 8, 124–136.[CrossRef][Medline]
Jones, E. W., Zubenko, G. S., and Parker, R. R. (1982). PEP4 gene function is required for expression of several vacuolar hydrolases in Saccharomyces cerevisiae. Genetics 102, 665–677.
Kane, P. M. (2006). The where, when, and how of organelle acidification by the yeast vacuolar H+-ATPase. Microbiol. Mol. Biol. Rev 70, 177–191.
Kunz, J. B., Schwarz, H., and Mayer, A. (2004). Determination of four sequential stages during microautophagy in vitro. J. Biol. Chem 279, 9987–9996.
Liegeois, S., Benedetto, A., Garnier, J. M., Schwab, Y., and Labouesse, M. (2006). The V0-ATPase mediates apical secretion of exosomes containing Hedgehog-related proteins in Caenorhabditis elegans. J. Cell Biol 173, 949–961.
Manolson, M. F., Wu, B., Proteau, D., Taillon, B. E., Roberts, B. T., Hoyt, M. A., and Jones, E. W. (1994). STV1 gene encodes functional homologue of 95-kDa yeast vacuolar H(+)-ATPase subunit Vph1p. J. Biol. Chem 269, 14064–14074.
Matsuo, H. et al. (2004). Role of LBPA and Alix in multivesicular liposome formation and endosome organization. Science 303, 531–534.
Merz, A. J., and Wickner, W. T. (2004). Trans-SNARE interactions elicit Ca2+ efflux from the yeast vacuole lumen. J. Cell Biol 164, 195–206.
Muller, O., Bayer, M. J., Peters, C., Andersen, J. S., Mann, M., and Mayer, A. (2002). The Vtc proteins in vacuole fusion: coupling NSF activity to V(0) trans-complex formation. EMBO J 21, 259–269.[CrossRef][Medline]
Muller, O., Neumann, H., Bayer, M. J., and Mayer, A. (2003). Role of the Vtc proteins in V-ATPase stability and membrane trafficking. J. Cell Sci 116, 1107–1115.
Nakamura, N., Hirata, A., Ohsumi, Y., and Wada, Y. (1997). Vam2/Vps41p and Vam6/Vps39p are components of a protein complex on the vacuolar membranes and involved in the vacuolar assembly in the yeast Saccharomyces cerevisiae. J. Biol. Chem 272, 11344–11349.
Nichols, B. J., Ungermann, C., Pelham, H. R., Wickner, W. T., and Haas, A. (1997). Homotypic vacuolar fusion mediated by t- and v-SNAREs. Nature 387, 199–202.[CrossRef][Medline]
Nishi, T., and Forgac, M. (2002). The vacuolar (H+)-ATPases—nature's most versatile proton pumps. Nat. Rev. Mol. Cell Biol 3, 94–103.[CrossRef][Medline]
Perzov, N., Padler-Karavani, V., Nelson, H., and Nelson, N. (2002). Characterization of yeast V-ATPase mutants lacking Vph1p or Stv1p and the effect on endocytosis. J. Exp. Biol 205, 1209–1219.
Peters, C., Baars, T. L., Buhler, S., and Mayer, A. (2004). Mutual control of membrane fission and fusion proteins. Cell 119, 667–678.[CrossRef][Medline]
Peters, C., Bayer, M. J., Buhler, S., Andersen, J. S., Mann, M., and Mayer, A. (2001). Trans-complex formation by proteolipid channels in the terminal phase of membrane fusion. Nature 409, 581–588.[CrossRef][Medline]
Raymond, C. K., Howald-Stevenson, I., Vater, C. A., and Stevens, T. H. (1992). Morphological classification of the yeast vacuolar protein sorting mutants: evidence for a prevacuolar compartment in class E vps mutants. Mol. Biol. Cell 3, 1389–1402.[Abstract]
Seeley, E. S., Kato, M., Margolis, N., Wickner, W., and Eitzen, G. (2002). Genomic analysis of homotypic vacuole fusion. Mol. Biol. Cell 13, 782–794.
Shorter, J., and Warren, G. (2002). Golgi architecture and inheritance. Annu. Rev. Cell Dev. Biol 18, 379–420.[CrossRef][Medline]
Stevens, T. H., Rothman, J. H., Payne, G. S., and Schekman, R. (1986). Gene dosage-dependent secretion of yeast vacuolar carboxypeptidase Y. J. Cell Biol 102, 1551–1557.
Subramanian, K., Dietrich, L. E., Hou, H., LaGrassa, T. J., Meiringer, C. T., and Ungermann, C. (2006). Palmitoylation determines the function of Vac8 at the yeast vacuole. J. Cell Sci 119, 2477–2485.
Sun-Wada, G. H., Toyomura, T., Murata, Y., Yamamoto, A., Futai, M., and Wada, Y. (2006). The a3 isoform of V–ATPase regulates insulin secretion from pancreatic {beta}-cells. J. Cell Sci 119, 4531–4540.
Thorngren, N., Collins, K. M., Fratti, R. A., Wickner, W., and Merz, A. J. (2004). A soluble SNARE drives rapid docking, bypassing ATP and Sec17/18p for vacuole fusion. EMBO J 23, 2765–2776.[CrossRef][Medline]
Veit, M., Laage, R., Dietrich, L., Wang, L., and Ungermann, C. (2001). Vac8p release from the SNARE complex and its palmitoylation are coupled and essential for vacuole fusion. EMBO J 20, 3145–3155.[CrossRef][Medline]
Vida, T. A., and Emr, S. D. (1995). A new vital stain for visualizing vacuolar membrane dynamics and endocytosis in yeast. J. Cell Biol 128, 779–792.
Wada, Y., Ohsumi, Y., and Anraku, Y. (1992). Genes for directing vacuolar morphogenesis in Saccharomyces cerevisiae. I. Isolation and characterization of two classes of vam mutants. J. Biol. Chem 267, 18665–18670.
Wang, C. W., Stromhaug, P. E., Kauffman, E. J., Weisman, L. S., and Klionsky, D. J. (2003). Yeast homotypic vacuole fusion requires the Ccz1-Mon1 complex during the tethering/docking stage. J. Cell Biol 163, 973–985.
Wang, Y. X., Kauffman, E. J., Duex, J. E., and Weisman, L. S. (2001). Fusion of docked membranes requires the armadillo repeat protein Vac8p. J. Biol. Chem 276, 35133–35140.
Weisman, L. S. (2003). Yeast vacuole inheritance and dynamics. Annu. Rev. Genet 37, 435–460.[CrossRef][Medline]
Wickner, W. (2002). Yeast vacuoles and membrane fusion pathways. EMBO J 21, 1241–1247.[CrossRef][Medline]
Wickner, W., and Haas, A. (2000). Yeast homotypic vacuole fusion: a window on organelle trafficking mechanisms. Annu. Rev. Biochem 69, 247–275.[CrossRef][Medline]
Winston, F., Dollard, C., and Ricupero-Hovasse, S. L. (1995). Construction of a set of convenient Saccharomyces cerevisiae strains that are isogenic to S288C. Yeast 11, 53–55.[CrossRef][Medline]
Yan, M., Rayapuram, N., and Subramani, S. (2005). The control of peroxisome number and size during division and proliferation. Curr. Opin. Cell Biol 17, 376–383.[CrossRef][Medline]
Zhang, J. W., Parra, K. J., Liu, J., and Kane, P. M. (1998). Characterization of a temperature-sensitive yeast vacuolar ATPase mutant with defects in actin distribution and bud morphology. J. Biol. Chem 273, 18470–18480.
This article has been cited by other articles:
|
|
 |
|
  Y. Lee, E.-S. Kim, Y. Choi, I. Hwang, C. J. Staiger, Y.-Y. Chung, and Y. Lee The Arabidopsis Phosphatidylinositol 3-Kinase Is Important for Pollen Development Plant Physiology, August 1, 2008; 147(4): 1886 - 1897. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||
