JBC Biosymposia, Inc.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M309445200 on September 5, 2003

J. Biol. Chem., Vol. 278, Issue 47, 47299-47306, November 21, 2003
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
278/47/47299    most recent
M309445200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Zhang, Z.
Right arrow Articles by Wilkens, S.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Zhang, Z.
Right arrow Articles by Wilkens, S.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Yeast V1-ATPase

AFFINITY PURIFICATION AND STRUCTURAL FEATURES BY ELECTRON MICROSCOPY*

Zhenyu Zhang{ddagger}, Colleen Charsky§, Patricia M. Kane§, and Stephan Wilkens{ddagger}

From the {ddagger}Department of Biochemistry, University of California, Riverside, Riverside, California 92521 and §Department of Biochemistry and Molecular Biology, State University of New York Upstate Medical University, Syracuse, New York 13210

Received for publication, August 26, 2003


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
V1-ATPase from the yeast Saccharomyces cerevisiae was purified via a FLAG affinity tag introduced into the N terminus of the G subunit. The preparation migrated as a single band in native gel electrophoresis and contained subunits ABCDEFGH (with subunit C present at substoichiometric amounts) as determined by matrix-assisted laser desorption ionization time-of-flight mass spectrometry. The initial specific Ca-ATPase activity was ~6 µmol/min/mg. The structure of the yeast V1-ATPase was studied by electron microscopy of negatively stained and frozen hydrated samples. A 25-Å resolution three-dimensional model of the complex was calculated from two-dimensional projections by the angular reconstitution technique. The model shows six elongated densities arranged in pseudo-3-fold symmetry around a large central cavity. At the top of the molecule, various protrusions can be seen. At the bottom of the complex, two large masses are visible that are connected to the main body of the molecule. Comparison of the yeast V1 structure with the structure of the intact V1V0-ATPase from bovine brain clathrin-coated vesicles (Wilkens, S., Vasilyeva, E., and Forgac, M. (1999) J. Biol. Chem. 274, 31804–31810) indicates that the structure of the isolated V1 from yeast is very similar to the structure of the V1 domain in the intact V-ATPase complex.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Vacuolar ATPases (V-ATPases)1 are large membrane-bound, multi-subunit complexes that utilize the free energy change generated during ATP hydrolysis to translocate protons or sodium ions across biological membranes (1–4). In eukaryotic cells, the vacuolar ATPase is found in the endomembrane system, where it functions to acidify subcellular compartments such as endosomes, lysosomes, Golgi-derived vesicles, clathrin-coated vesicles, synaptic vesicles, the plant vacuole, and chromaffin granules. Acidification of these organelles plays a vital role in processes like protein trafficking, receptor-mediated endocytosis, neurotransmitter release, intracellular pH maintenance, and waste management. A vacuolar type ATPase is also found in the plasma membrane of certain specialized cells such as renal intercalated cells or osteoclasts, where acidification of an enclosed extracellular space is required. Despite their widespread nature, V-ATPases are remarkably similar with respect to their functionality and overall architecture (5). Early studies using electron microscopy have shown that the complex is organized in two major domains, a membrane extrinsic V1 and a membrane-embedded V0 (6). ATP is hydrolyzed on the membrane extrinsic V1, and the energy released during that process is transmitted to the membrane-bound V0 to drive ion translocation. This energy coupling occurs via the so-called "stalk" structure, an assembly of several polypeptides that forms the functional and structural interface between the two active domains of the complex. The eukaryotic V-ATPase is composed of at least 13 different polypeptides with molecular masses ranging from 12 to 100 kDa. The water-soluble V1 contains subunits ABCDEFGH, and the membrane-embedded V0 is made of subunits a(c,c')c''d. The subunit stoichiometry has been determined in the bovine enzyme to 3:3:1:1:1:1:2:2 for the V1 and 1:4–5:1:1 for the V0 domain (7, 8). As in the related F-ATPase (9), energy coupling involves rotation of a central stalk domain with respect to the static remainder of the complex. In the V-ATPase, the rotor has been shown to contain subunits D and F (10) and the ring of proteolipids (11) whereas subunit G has been found to be part of the stator (11). Electron microscopy and image analysis has provided a detailed picture of the overall structure of the V-ATPase from eubacteria (12), bovine (13), and plant (14). A remarkable feature of the V-ATPase is its nutrient-dependent reversible dissociation in vivo (15, 16). The reversible dissociation of V1 from V0 allows isolation of cytoplasmic V1-ATPase as has been demonstrated for the insect and yeast systems (17, 18). The yeast V-ATPase is well characterized, both by biochemical and genetic means, and its polypeptides can be manipulated by site-directed mutagenesis, making the yeast V-ATPase a powerful model system for structural and functional studies.

Here we describe isolation of highly purified yeast V1-ATPase via an affinity tag introduced into the N terminus of the G subunit. The preparation has a high specific Ca-ATPase activity that is inhibited in the presence of even low concentrations of Mg2+ ions. Furthermore, we present structural features of the V1-ATPase from electron microscopy and image reconstruction. A comparison of the electron microscopic images of the yeast V1 with the images of the bovine V1V0 indicates that the structure of the V1 when detached from the V0 is very similar to the structure of the V1 domain in the intact V-ATPase complex.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—All chemicals were from Sigma unless noted otherwise. Premixed YEPD (1% yeast extract, 2% peptone, 2% glucose) and leucine dropout medium was from Difco and Invitrogen, respectively. M2 anti-FLAG affinity-agarose, anti-FLAG IgG, and FLAG peptide were from Sigma. Anti-subunit A and B monoclonal antibodies were from Molecular Probes, Inc. Protein concentrations were determined with the BCA assay system (Pierce).

Construction of a FLAG-tagged Vma10p—A single FLAG epitope was placed immediately after the N-terminal methionine of VMA10 by fusion PCR as follows. Two fragments of VMA10 containing extensions corresponding to overlapping portions of the FLAG epitope were amplified from yeast genomic DNA using oligonucleotide pairs VMA10-P1 and VMA10-P9 FLAG and VMA10-P2 and VMA10-P8 FLAG and a combination of LA-Taq (Panvera) and Native Pfu (Stratagene) polymerases. The sequences of the oligonucleotides (purchased from MWG Biotech) were as follows: P1, 5'-AGCGTTGTAATGCCTATAG-3'; P2, 5'-GATAGTTGTAGTCCCTCGG-3'; P8 FLAG, 5'-CTTGTCATCGTCG-TCCTTGTAGTCCATTCTGCTTTGTATACCTTGCA-3'; P9 FLAG, 5'-GACTACAAGGACGACGATGACAAGTCCCAAAAAAACGGAATTGC-CA-3'.

PCR fragments from the initial amplification were purified and combined. The fusion product was PCR-amplified by adding oligonucleotides P1 and P2 to the fragments, along with the polymerase mixture described above. The fusion PCR product was treated with Taq polymerase to add a non-templated dATP and then recovered in plasmid pGEM-T Easy (Promega) according to the manufacturer's instructions. The fusion placed a single FLAG epitope immediately after the ATG and also deleted the intron present in VMA10; the sequence was confirmed by DNA sequencing (SUNY Upstate Medical University Sequencing Facility). The tagged VMA10 gene was excised from pGEM-T Easy with BamHI and XhoI and cloned into the low copy (CEN) vector pRS315 (19) using the same enzymes. The plasmid was transformed into yeast strain SF838–5Aa vma10{Delta}::URA3 (20) using an overnight lithium acetate procedure (21), and transformants were selected by growth on supplemented minimal medium lacking leucine. Vacuoles isolated from this strain as described (15) had levels of concanamycin A-sensitive ATPase activity comparable with wild type vacuoles.2

Yeast V1-ATPase Purification—Yeast V1-ATPase was purified via a FLAG tag introduced into the N terminus of the VMA10 gene product (Vma10p, subunit G). Briefly, yeast was grown in small scale over night to an A600 of around 3 in leucine dropout medium, and 25 ml of the resulting inocculum was added to 1 liter of unbuffered YEPD (1% yeast extract, 2% peptone, 2% glucose) in a 2.8-liter Fernbach flask. The flasks were incubated at 30 °C under vigorous shaking until an A600 of around 3 (early log phase). Yeast was harvested by centrifugation and resuspended in 50 mM Tris, pH 7.5, 1.2 M sorbitol, 2% glucose. Yeast was converted to spheroplasts by incubation with zymolase (100 units per gram of yeast) for 20 min at 30 °C. Spheroplasts were collected by centrifugation, and V1 was dissociated from vacuolar membranes by incubation for 5 min at 30 °C with gentle shaking in YP medium without glucose. Spheroplasts were collected by centrifugation and resuspended in the same volume of TBSE (20 mM Tris/Cl, pH 7.2, 150 mM NaCl, 1 mM EDTA) and the protease inhibitors leupeptin, aprotinin, pepstatin, and phenylmethylsulfonyl fluoride (at concentrations of 1 µg/ml, 5 µg/ml, 1 µg/ml, and 1 mM, respectively) were added shortly before cells were lysed by two to three passages through a French pressure cell at 20,000 p.s.i. The cell lysate was centrifuged at 275,000 x g for 1.5 h at 4 °C. The resulting supernatant was then applied to a small column packed with 1 ml of anti-FLAG M2 agarose. The anti-FLAG column was washed with 10 column volumes of TBSE, and pure V1-ATPase was eluted with a solution containing 100 µg/ml FLAG peptide in TBSE. V1-ATPase containing fractions were pooled and precipitated by addition of 60% ammonium sulfate, and the redissolved protein was further purified by gel filtration over a 1 x 30-cm Superdex200 column attached to an Äkta fast protein liquid chromatography system (Amersham Biosciences). Gel filtration was carried out in TBSE (+1 mM DTT). V1-ATPase-containing fractions were pooled and precipitated by addition of saturated ammonium sulfate to 60% saturation, and the redissolved protein was desalted by passage over two consecutive Sephadex G25 spin columns in TBSE (+1 mM DTT).

ATPase Activity—ATP hydrolysis activity of V1-ATPase was determined in the presence of 5 mM ATP, 1.6 mM CaCl2, 30 units/ml each of lactate dehydrogenase and pyruvate kinase, 0.5 mM NADH, 2 mM phosphoenol pyruvate in 50 mM HEPES, pH 7.5, at 37 °C. The ATP hydrolysis reaction was started by adding between 0.5 and 5 µg of purified V1-ATPase to 1 ml of the assay mix. After a given time (between 0.5 and 40 min), the amount of ADP produced by the V1-ATPase was determined by adding 4 mM MgCl2 and monitoring the absorbance decrease at 340 nm. No significant ATP hydrolysis activity could be measured in this assay system when both Mg2+ and Ca2+ were present in the assay before adding the V1-ATPase.

In-gel Trypsin Digestion of V-ATPase Subunits—Protein bands were excised from SDS gels, and samples for mass spectrometry were obtained from the gel slices essentially as described (22). Briefly, gel slices were dehydrated with acetonitrile and dried. Buffer containing sequencing grade trypsin (Promega) at a concentration of 12 µg/ml was used to rehydrate the gel pieces, which were incubated with the protease overnight at 37 °C. The resulting peptide fragments were extracted twice with acetonitrile plus 5% trifluoroacetic acid. The extractions were pooled and completely dried in a speed vac. The eluted peptides were resuspended in a 1:1 mixture of acetonitrile/water containing 0.1% trifluoroacetic acid and mixed with the appropriate matrix in the manner described below.

MALDI-TOF Mass Spectrometry and Protein Identification—All mass spectra were acquired on a PerSeptive Biosystems (Framingham, MA) Voyager DE-STR equipped with an N2 laser (337 nm, 3-ns pulse width, 3-Hz repetition rate). The mass spectra were acquired in the positive reflector mode with delayed extraction. The matrix used was {alpha}-cyano-4-hydroxycinnaminic acid, 10 mg/ml in a 1:1 solution of acetonitrile and 0.1% trifluoroacetic acid. Typically, the extracted peptides were diluted in the matrix 1:10 prior to applying 1 µl to the sample plate. Data searches were carried out against the National Center for Biotechnology Information, nonredundant data base using Protein Prospector (prospector.ucsf.edu) to identify the protein bands. Internal mass calibration was obtained by using trypsin fragments at 842.51 and 2211.10 Da allowing a mass error tolerance of less than 30 ppm in most samples. Without mass calibration, the error tolerance was increased to 200 ppm during the data base search.

Electron Microscopy—For negative staining, protein at a concentration of between 20 and 50 µg/ml in TBS (+1 mM DTT) was applied to glow discharged carbon-coated copper grids. Grids were washed once with water, stained with 1% uranyl acetate, and air-dried. Grids were examined in Philips CM300 and Tecnai12 transmission electron microscopes operating at 100 kV. Images were recorded in low dose mode with a 1024 x 1024 (CM300) and 2048 x 2048 (Tecnai12) pixel charge-coupled device (Gatan Inc.) in single frame or montage mode or on Eastman Kodak Co. SO163 film, which was developed in D19 developer for 12 min. For cryoelectron microscopy, V1-ATPase at a concentration of 0.1–0.2 mg/ml in TBS (+1 mM DTT) was applied to holey carbon-coated copper grids (Structure Probe Inc. or in-house production) and plunged into liquid ethane cooled by liquid nitrogen. Frozen samples were mounted in a Gatan cryo stage and examined under low dose conditions. Images of stained samples were recorded with an underfocus of between 382 and 576 nm and an electron optical magnification of x47,000, placing the 1st zero of the contrast transfer function at around 1/20 Å-1. Images of frozen samples were recorded at an underfocus of -960 nm.

Analysis of Stained Images—Images were analyzed with the EMAN (23) and IMAGIC 5 (24) software packages running on SGI workstations (O2 and octane) essentially as described (13, 25). Briefly, two data sets from different preparations (5000 and 7600 images, respectively) were selected from 20 3 x 3 charge-coupled device frames (CM300) and 49 2048 x 2048 charge-coupled device frames (Tecnai12), respectively. The images were normalized and band pass filtered to remove low (<0.015 Å-1) and high (>0.1 Å-1) spatial frequencies. The data sets were then analyzed by the alignment by classification (26) procedure leading to sets of initial references that were used in a first multireference alignment step. The multireference alignment was iterated until no further improvement in the class sums was obtained. A visual inspection of the results revealed that the analysis of the two different data sets produced very similar class sums. At this point, the two data sets were merged for further analysis. To start up the three-dimensional reconstruction, 100 of the best class sums were selected, normalized, and surrounded by a soft mask. A set of initial Euler angles was then assigned to the projections with the startAny program within EMAN, assuming 3-fold symmetry of the complex. The resulting 3-fold symmetric model was then used to produce an anchor set for refinement (all subsequent refinement was done within IMAGIC 5) with the original input projections, this time assuming C1 symmetry (no symmetry at all). After seven rounds of refinement, the resulting model was filtered to remove noise (density not connected to the main volume of the model) and forward-projected along 49 directions uniformly distributed over the Euler sphere. The 49 projections were normalized and used as references in a new multi-reference alignment step. Fifty of the images aligned to each reference were summed and used as input projections for a new three-dimensional model. This process was iterated increasing the number of forward projections, and images were summed for each reference until no further improvement was observed. At this point, classification of the aligned data set produced very similar class sums compared with the ones used to start up the three-dimensional reconstruction. The resolution of the final reconstruction was estimated by calculating the Fourier shell correlation between two 3D models each calculated from half the input projections (27, 28). The final model was filtered to 0.05 Å-1, corresponding to the first zero in the contrast transfer function.

Analysis of Cryoimages—Electron micrographs recorded at x47,000 were digitized on an Optronics drum scanner with a sampling rate of 18.75 µm corresponding to 4 Å/pixel on the specimen level. A total of 40,000 individual images were selected from 18 micrographs using the automatic particle picking routine within EMAN. The images were treated as above and sorted into 400 classes by the alignment by classification procedure. A visual inspection of the alignment by classification classums revealed that virtually all the averages showed the pseudo-trigonal top (or bottom) view of the molecule with few classes showing a slightly tilted top or bottom view and essentially no classes showing a clearly identifiable side view, thus far preventing the calculation of a reliable 3D model of the V1-ATPase from the cryoimages with the angular reconstitution procedure.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Yeast V1-ATPase Purification—Introduction of a FLAG epitope tag into the N terminus of the yeast vacuolar ATPase G subunit (Vma10p) allows efficient and rapid isolation of highly purified V1-ATPase complex. Fig. 1A shows SDS-PAGE of V1-ATPase purified this way. The left gel in Fig. 1A shows elution of the anti-FLAG-agarose with FLAG peptide. On the gel, eight protein bands (excluding the weak band running at the top of the gel) can be clearly identified (close inspection of the 2nd major band from the top showed that this band is composed of two bands running close together). To verify the identity of the bands, peptide mass fingerprinting by MALDI-TOF mass spectrometry was performed. Fig. 1B shows the results of the peptide mass fingerprinting from the protein bands excised from the SDS-polyacrylamide gel. The data show that the eight bands in the order from top to bottom could be unambiguously identified as yeast V1-ATPase subunits A, B, H, C, D, E, F, and G. Mass spectra for the contaminating band running above the A subunit (slice 1) did not allow unambiguous identification of the protein(s) contributing to this band. To remove any potential excess of G subunit that might be present in the cytosol, the V1-containing fractions were pooled, concentrated by ammonium sulfate precipitation, and treated by size exclusion chromatography. The middle gel shows analysis of the fractions eluted from the gel filtration column. As can be seen, pure V1-ATPase is eluted in a single peak including fractions 19–22, well resolved from the following peaks containing subunits G, E, C, and H (fraction 26) and a peak containing essentially only subunit G (fraction 31). V1-containing fractions from the gel filtration column were pooled, precipitated by addition of 60% ammonium sulfate, dissolved in TBSE (+DTT), and desalted over two consecutive Sephadex G25 centrifuge columns in the same buffer. The preparation eluted from the second spin column is shown in the right gel of Fig. 1A. As can be seen, all the subunits of the V1-ATPase are present in the preparation. The low intensity of the subunit C band suggests that this subunit is present in sub-stoichiometric amounts. To analyze the oligomeric state of the complex, native gel electrophoresis was performed. Fig. 1C shows a 1% agarose gel of FLAG tag-purified V1-ATPase. The complex runs as a single band on the native gel, which, when analyzed by a second-dimension polyacrylamide gel in the presence of SDS, contains all the polypeptides of the V-ATPase identified above.



View larger version (54K):
[in this window]
[in a new window]
  		FIG. 1.
Purification and subunit analysis of yeast V1-ATPase. A, SDS-PAGE of V1-ATPase purification. Left gel, right lane, V1-ATPase eluted from anti-FLAG affinity column. Left gel, left lane, molecular weight standards. Middle gel, size exclusion chromatography of pooled V1-ATPase eluted from anti-FLAG column. Right gel, V1-ATPase after size exclusion chromatography, ammonium sulfate precipitation, and two consecutive centrifuge columns. B, identification of V1-ATPase subunits by MALDI-TOF mass spectrometry. The protein bands in the left gel (1-9 from top to bottom) were excised and digested with trypsin in-gel, and the resulting peptides were analyzed by MALDI-TOF mass spectrometry. C, native gel electrophoresis of 20 µg of FLAG tag-purified yeast V1-ATPase in 1% agarose, 20 mM Tris acetate, pH 7. After staining with Coomassie Blue, the band (see arrow) was excised from the agarose gel, soaked in SDS/DTT containing gel loading buffer, and loaded onto a denaturing polyacrylamide gel in the presence of SDS (right gel).

 
Ca2+-ATPase Activity—The ATPase activity of the affinity purified V1 was determined in presence of 5 mM ATP, 1.6 mM Ca2+. The amount of ADP released after given amounts of time was determined with an ATP regenerating system containing phosphoenol pyruvate, pyruvate kinase, NADH, and lactate dehydrogenase. Because V1-ATPase is inhibited by even low concentrations of magnesium (see below), the ATP hydrolysis reaction and the following ADP determination could be performed in the same assay mix. The ATP hydrolysis reaction was started by adding V1-ATPase to an assay mix containing CaATP and the components necessary for ADP rephosphorylation. After given amounts of time, ATP hydrolysis by the V1-ATPase was stopped by addition of 4 mM Mg2+. At the same time, addition of magnesium activated the magnesium-dependent pyruvate kinase, which could then rephosphorylate the ADP coupled to the oxidation of NADH. Under these conditions, an initial ATP hydrolysis rate of ~6 µmol/min/mg was observed (±1 µmol/min/mg; two preparations). Over time, the rate of ATP hydrolysis by the yeast V1-ATPase was decreasing from the initial rate of 6 µmol/min/mg to below 0.5 µmol/min/mg after 30 min. Fig. 2 shows a representative measurement of the time dependence of the Ca-ATPase activity of FLAG tag-purified V1-ATPase. A short (<1 min) activity lag after addition of V1-ATPase to the assay mix was seen in some experiments but not others. When V1-ATPase was added to the assay mix in the presence of 1.6 mM Ca2+ and even low concentrations of Mg2+, only a short burst of ATPase activity could be observed with a specific activity of less than 0.5 units/mg when measured over the first 30 s after addition of V1 to the assay mix. This suggests that MgATP competes with CaATP and that MgATP binds more tightly to the catalytic sites than CaATP but is not turned over. No ATPase activity could be measured in the absence of Ca2+.



View larger version (18K):
[in this window]
[in a new window]
  		FIG. 2.
ATPase activity of V1-ATPase. ATPase activity was measured in the presence of 1.6 mM Ca2+,5mM ATP in 50 mM Hepes, pH 7.5, in the presence of pyruvate kinase, lactate dehydrogenase, phosphoenol pyruvate, and NADH. The ATP hydrolysis reaction was started by addition of 1 µg of V1-ATPase to the assay mix. After given times, 4 mM Mg2+ was added, and the decrease in the absorbance at 340 nm was measured. No significant ATP hydrolysis could be measured when adding V1 to the assay mix, which contained both Ca2+ and Mg2+. The time dependence of the ATP hydrolysis rate (•) and the total accumulated amount of ATP hydrolyzed ({blacksquare}) is shown.

 
Electron Microscopy of Yeast V1-ATPase—The data summarized above indicate that the FLAG tag-isolated V1-ATPase is highly purified, enzymatically active, stable, and monodisperse, properties that are prerequisites for structural and functional studies by biochemical and biophysical methods. For negative stain electron microscopy, yeast V1-ATPase was applied to glow discharged carbon-coated copper grids and washed on the grid with 1% uranyl acetate. Fig. 3A shows a section of a typical electron micrograph of negatively stained yeast V1-ATPase. As can be seen, the preparation is highly monodisperse, consistent with the native gel electrophoresis results (see above). Only few smaller protein molecules are visible on the electron microscopy images, suggesting that no significant dissociation of the V1-ATPase complex occurs under the conditions used to prepare the specimen (1% uranyl acetate, pH 5). Two data sets of V1 molecules from different preparations were analyzed by alignment and classification procedures. Fig. 3B shows a selection of the most characteristic projections obtained after several rounds of multireference alignment. Most distinct are the top and side view projections in which the molecule is seen perpendicular or parallel to the membrane surface, respectively. The top view projection shows two sets of each three large densities arranged alternatingly around a central cavity (see black and white arrows in Fig. 3B, image 3). The densities can be interpreted as the alternating A and B subunits of the V-ATPase. A clear handedness is present in these projections making it possible to distinguish whether the molecule is seen from the top or the bottom. Other views represent projections of the molecule from the side. Fig. 3C shows the side view projection of the bovine V-ATPase (taken from Ref. 13). Comparison of the yeast V1-ATPase side view projections with the bovine holo enzyme projections allows orientation of the V1-side views with respect to the membrane surface. The comparison also shows that the projection structure of the isolated yeast V1 is very similar to the projected structure of the V1-domain in the bovine holo enzyme. Several features visible in the bovine V1 domain are also present in the yeast V1-ATPase: the elongated and knob-like densities at the top of the molecule (see arrows in Fig. 3B, images 5 and 10, respectively) and the stalk domain consisting of two to three densities in the yeast V1 (see arrowheads in images 1 and 7). The dimensions of the yeast V1-ATPase are ~14 x 20 nm, including the stalk and the protruding densities at the top of the molecule.



View larger version (202K):
[in this window]
[in a new window]
  		FIG. 3.
Electron microscopy and image analysis of negatively stained V1-ATPase. A, area from a typical electron micrograph of V1-ATPase stained with 1% uranyl acetate. Some of the molecules are indicated by black circles. B, ten of the best class sums after several rounds of multireference alignment/classification. Most characteristic are top (images 2, 3, and 6) and side view projections (images 1, 4, 5, 7, 8, 9, and 10). C, side view projection of the bovine V-ATPase (taken from Ref. 13). The averages shown in B have been calculated from between 60 and 90 aligned images.

 
Three-dimensional Reconstruction of the Yeast V1-ATPase—To start up the three-dimensional reconstruction, 100 of the best projections from the merged data set including the ones shown in Fig. 3B were used. A 3-fold symmetry was assumed in this initial step of the three-dimensional reconstruction based on the composition of the catalytic core of the complex, A3B3, which accounts for ~85% of the total mass of the V1 excluding the stalk region. In the following refinement steps, C1 (no symmetry at all) was assumed. After seven rounds of refinement, the 3D model was projected along 49 directions uniformly distributed on the Euler sphere, and the projections were used as references in a multireference alignment step. This procedure was iterated with increasing numbers of forward projections until no further improvement in the final model was observed. Fig. 4A shows surface representations of the final 3D model of the V1-ATPase. A selection of final input projections (images 1, 3, 5, 7, 9, and 11) and the corresponding re-projections (images 2, 4, 6, 8, 10, and 12) is shown in Fig. 4B. Fig. 4C shows contoured cross-sections of the 3D model from bottom to top as indicated by the white lines on the left side of image 1 in Fig. 4A.



View larger version (71K):
[in this window]
[in a new window]
  		FIG. 4.
Three-dimensional reconstruction of the yeast V1-ATPase. A, surface representation of the final 3D model of the yeast V1-ATPase. The resolution in the final model was 25 Å as determined with the Fourier shell correlation method using a cut off of 0.5. The threshold has been set so that the volume of the model corresponds to the expected molecular mass of the complex with a stoichiometry of A3B3DEFG2H (530 kDa). The first four views are views parallel to the membrane surface, rotated with respect to image 1 by 60, 120, and 180°, respectively. Images 5 and 6 are views perpendicular to the membrane surface from the cytoplasm and membrane surface, respectively. The pseudo-3-fold symmetry is indicated by asterisks (image 5). B, final input (images 1, 3, 5, 7, 9, and 11) and corresponding re-projections (images 2, 4, 6, 8, 10, and 12) from the final 3D model. C, contoured cross-sections at the positions indicated on the left side of image 1 in part A.

 
Overall Features of the Three-dimensional Model—The yeast V1-ATPase is composed of subunits AB(C)DEFGH in the presumed ratio of 3:3:(1):1:1:1:2:1. The ratios of the A and B subunits is well established, but the stoichiometry of the stalk subunits has not been rigorously determined in the yeast system. In the bovine V-ATPase, the ratio of the CDEFGH subunits has been measured to 1:1:1:1:2:2 (7, 8), but the presence of as many as three copies each of subunits E (29) and G (30, 31) has been suggested. Subunit C is generally not considered a subunit of the membrane-detached V1-ATPase, but its presence in the affinity-purified yeast V1 suggests that subunit C does nevertheless have a significant affinity for the rest of the complex. The total molecular mass of the complex (excluding subunit C) is ~530 kDa, assuming a subunit stoichiometry as determined for the bovine enzyme (except for subunit H, which might be present in only one copy in the yeast enzyme). As mentioned earlier, more than 85% of the mass of the V1-ATPase without the stalk domain comes from the A and B subunits, which are arranged in the pseudo-trigonal arrangement seen in the top view projection shown in image 3 of Fig. 3B (see also the cryoelectron microscopy results in the following section). A high degree of pseudo-3-fold symmetry is present over much of the complex as can be seen in the contoured cross-sections shown in Fig. 4C. Strong trigonal symmetry is also seen in the related F1-ATPase structure, where it is broken most by the presence of the single copy stalk subunits at the bottom the complex and the differential nucleotide binding to the catalytic/non-catalytic sites on the {beta} and {alpha} subunits, respectively (32, 33). This is also seen in the bottom cross-section of the three-dimensional model of the yeast V1-ATPase (image 2 in Fig. 4C). The highest symmetry in the F1-ATPase ({alpha}3{beta}3{gamma}{delta}{epsilon}, subunit nomenclature of the mitochondrial enzyme) is seen in the top part of the complex where the N-terminal domains of both the {alpha} and {beta} subunits form a ring of interacting {beta} barrels. This interaction is probably required for stability of the complex. The eukaryotic V1-ATPase is different in the top portion of the complex in that it shows additional features such as the pronounced elongated and knob-like densities (13). Based on above symmetry considerations, we proposed earlier (13) that the knob-shaped and elongated densities seen in the projections of the V1 domain in the bovine V-ATPase might be present in two or three copies, and it was furthermore suggested that the knob-like features are formed at least in part by the ~100-amino acid inserts near the N termini of the A subunits. From Fig. 4A it becomes clear that whereas the knob-like features are present in three copies (see arrows in images 1-3 in part A of Fig. 4), there seem to be only two of the elongated densities (see arrowheads in Fig. 4A, images 1 and 2). The two large subunits to which the elongated densities are attached extend slightly higher than the third large subunit (see arrowheads in image 6 of part C of Fig. 4).

Features of the Stalk Region—At the bottom of the complex two large densities can be seen, one of which is right beneath the cavity formed by the A and B subunits, and the other one is off center by about 3 nm. Candidates for these protein densities are subunits C, D, E, F, G, and H. The weak staining intensity of the subunit C band in SDS-polyacrylamide gels (see Fig. 1) suggests that this subunit is only present in substoichiometric amounts (20% or less), and it is unlikely that its presence will produce significant density in the 3D model of the complex. Subunits F and G, both about 12 kDa, are most likely too small to be seen as individual densities at the current resolution leaving subunits D (32 kDa), E (27 kDa), possibly together with F and G, and subunit H (54 kDa). Recently, cross-linking data have been presented that imply that subunits E and G bind along the outside of the complex, with subunit E spanning the entire length of the V1 and part of G binding at the very top of the complex (34, 35). This arrangement would leave subunits D (most likely together with F) and H for the densities seen at the very bottom of the complex.

Analysis of the V1-ATPase Structure by Cryoelectron Microscopy—To complement the negative stain data for the V1-ATPase, we decided to analyze the structure of the complex embedded in amorphously frozen buffer by cryoelectron microscopy (Fig. 5A). A data set of ~40,000 individual molecular images of the complex was sorted into 400 classes by the alignment by classification procedure. A visual inspection of the 400 averaged classes showed that essentially all of the molecules were oriented to produce the pseudo-trigonal top or bottom view of the molecule with only very few classes showing different views (data not shown). No clear side views were present, preventing the calculation of a reliable three-dimensional model from the two-dimensional projections of the ice-embedded molecule thus far. Fig. 5B shows one of the top view classes of the ice-embedded V1-ATPase after several rounds of multireference alignment and classification. For comparison the top view as obtained in negative staining is shown in Fig. 5C. As can be seen, both averages show very similar features including the pseudo-trigonal symmetry, the clear handedness, the cavity in the center of the molecule, and the small, wing-like domains extending from one set of the large subunits (see arrows in part B) of Fig. 5). In the projections shown in B and C of Fig. 5, the molecule is seen from the cytoplasmic side (top view) based on an Euler angle assignment with respect to the final three-dimensional model (not shown). By aligning the averages to the top view projection of the final three-dimensional model, they have been brought into the same orientation as the cross-sections shown in Fig. 4C. This indicates that the subunits showing the wing-like structures (see arrows in part B) correspond to the subunits that bind the second stalks (presumably the B subunits; see "Discussion"). This assignment would be consistent with the knob-like structures (that are present in three copies; see arrows in images 1, 2, and 3 in part A of Fig. 4) being part of the A subunits of the complex.



View larger version (130K):
[in this window]
[in a new window]
  		FIG. 5.
Cryoelectron microscopy of the yeast V1-ATPase. A, area from a typical cryoelectron micrograph of yeast V1-ATPase. Some of the molecules are indicated by black circles. B, top view projection of the V1-ATPase. The average was obtained from a data set of ~40,000 molecules by alignment and classification procedures. An average of 150 molecules is shown. C, for comparison, the top view average of the negatively stained data set is shown (80 images). Arrows in B indicate the wing-like densities giving the molecule its characteristic handedness in projection. The clear similarity in the projections of stained and ice-embedded V1 indicates that the stain embedding allows a reasonably good representation of the yeast V1-ATPase structure at its current resolution of 2.5 nm.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Purification and Ca-ATPase Activity of the Yeast V1-ATPase—In the current study we show that by introducing a FLAG affinity tag into the VMA10 gene product (subunit G) N terminus, highly purified and active yeast V1-ATPase can be isolated with a few steps. We have shown that the preparation forms a stable complex containing subunits ABCDEFGH. Consistent with earlier reports, subunit C is present in substoichiometric amounts (18). The preparation has an initial specific Ca-ATPase activity of ~6 µmol/min/mg when measured at 37 °C in presence of 5 mM ATP, 1.6 mM Ca2+. This value is somewhat higher compared with the initial CaATP hydrolysis rate of 1.7 µmol/min/mg obtained for wild type V1-ATPase purified without the affinity tag (18). The higher activity of the FLAG tag-isolated V1-ATPase might be because of the different assay or purification conditions used, but it cannot be ruled out that the FLAG tag in the G subunit N terminus has an influence on the Ca2+-ATPase activity. No significant ATPase activity could be measured in the presence of even low concentrations of magnesium ions, regardless of whether Ca2+ was present. This suggests that MgATP competes for the catalytic sites with high affinity but is not hydrolyzed by the yeast V1-ATPase. A similar inhibitory effect of magnesium on the Ca-ATPase activity had been reported for the V1-ATPase from Manduca sexta (17), indicating that the inhibitory effect of Mg2+ is a common feature of the eukaryotic V1-ATPase. Consistent with previous activity measurements conducted for V1-ATPase isolated from wild type yeast without affinity tag, Ca2+-ATPase activity of the FLAG tag-purified enzyme decreases essentially to zero over a time period of 20–30 min (18). Whether this decrease in activity is because of product (ADP) inhibition or dissociation of the complex or both is not clear at this time. Although it could be shown that a double mutation in subunit A of the bacterial V1-ATPase essentially eliminated the inhibition caused by MgADP (10), cryo and negative stain electron microscopy of the yeast V1-ATPase pre-incubated with CaADP and inorganic phosphate or Ca2+ and AMPPNP showed some dissociation of the complex (data not shown), suggesting that a slow dissociation of the complex in the assay mix might be involved in the deactivation of the eukaryotic enzyme.

Structure of the V1-ATPase—Fig. 6 shows our current working model for the subunit arrangement in the yeast V1-ATPase. The primary structures of the A (excluding the non-homologous region; see below) and B subunits of the V1-ATPase are ~20–25% identical to the {beta} and {alpha} subunits of the related F-ATPase, respectively, suggesting that the catalytic A3B3 domain of the V1-ATPase has a similar tertiary structure as the F1-ATPase {alpha}3{beta}3 catalytic core. The x-ray structure of the bovine mitochondrial F1-ATPase shows the {alpha} and {beta} subunits arranged pairwise in a pseudo-trigonal fashion around a central cavity (32, 33) whereas perfect 3-fold symmetry is seen in the x-ray model for the enzyme from rat liver (36). In the case of the F1-ATPase, the central cavity is filled with the {alpha} helical N and C terminus of the {gamma} subunit, which extends about 45 Å below the {alpha}3{beta}3 domain to form the central stalk, which also includes the {delta} and {epsilon} subunits (subunit nomenclature of the mitochondrial F-ATPase (32, 33)). The peripheral stalk of the F-ATPase, not seen so far in any of the x-ray crystallographic models, has been most thoroughly analyzed in the bacterial enzyme, and it has been shown by means of chemical cross-linking and electron microscopy that it is formed by the {delta} and b subunits (subunit nomenclature of the bacterial enzyme). There is no convincing sequence homology between the single copy F- and V-ATPase subunits, and the structural organization of the V-ATPase central and peripheral stalks remains controversial. Genetic (37) and mechanistic studies (10, 11) provide strong evidence that the D and F subunits of the V-ATPase (Vma8p and Vma7p of the yeast enzyme, respectively) function as rotating central stalk, together with the membrane-bound ring of proteolipids, whereas the E, G, and H subunits contribute to the stator domain (11, 34, 35). Consistent with this picture, there are two large protein densities visible at the bottom of the complex, candidates for which are subunits D (together with F) and H. Subunit H has been shown to interact both with the large A subunit of the V1 and the cytoplasmic domain of the V0 {alpha} subunit (38), as well as the N terminus of subunit E (39). The crystal structure of the isolated yeast H subunit shows an elongated three-domain structure for the subunit (40), measuring ~100 x 40 Å, suggesting that in the current 3D model of the yeast V1, subunit H is only partly resolved. Fig. 3B shows that there is an additional weak density (see white arrowhead in image 7) connected to the density right next to the central stalk density right underneath the A3B3 domain. This fuzzy density is seen in other averages (images 4 and 10 in Fig. 3B; see also Ref. 13), but its weak appearance might be the explanation why it is not resolved in the final 3D model of the complex. The eukaryotic V-ATPase has been shown to interact with other proteins in the cell, and this interaction can be via subunit H of the V1 domain as in the case of the yeast ectoapyrase (41) or human immunodeficiency virus Nef (42). The absence of these binding partners in the isolated yeast V1 might be a reason why this subunit is poorly defined in the three-dimensional model of the V1 complex. No comparable density is seen in the electron microscopy-derived three-dimensional model of the V1 from M. sexta (43, 44) consistent with subunit H being flexible in the V1 domain when detached from the V0.



View larger version (64K):
[in this window]
[in a new window]
  		FIG. 6.
Model of the subunit arrangement in the yeast V1-ATPase. The model is based on the presented three-dimensional structure of the yeast V1-ATPase and the available biochemical and genetic subunit interaction data discussed in the text.

 
The presence of two (and possibly three) stators has been proposed for the V-ATPase based on two-dimensional projections of the enzyme from eubacteria (12) and bovine brain (13). In the here presented three-dimensional model of the yeast V1-ATPase, two of the large subunits show the elongated density bound at the periphery whereas the third one seems to lack this additional density at the top of the complex. This would be consistent with the presence of (at least) two stator domains in the isolated yeast V1-domain. Candidates for the stator domains are subunits E and G, which have been shown to interact in vivo (45) and which can be cross-linked to the periphery of subunit B (34, 35). The stoichiometry for these subunits has not been measured in the yeast V-ATPase, and although a ratio of two Gs and one E has been reported for the enzyme from bovine brain (7, 8), the presence up to three copies of these subunits has been proposed based on staining intensities of the bands in SDS-PAGE gels (29–31).

In the related F-ATPase, it has been shown that the stator is interacting with the F1 via an {alpha} subunit (46, 47). Based on the subunit homology between F- and V-type ATPases, this would suggest that the stators in the V-ATPase interact with the V1 via the B subunit, consistent with the cross-linking results involving subunits B, E, and G reported for the yeast V-ATPase complex (34, 35). This assignment would also be consistent with the knob-like densities corresponding to the non-homologous inserts in the V-ATPase A subunits. The 100-amino acid non-homologous insertion occurs at the connection between the N-terminal {beta} barrel and middle domains of the F-ATPase {beta} subunit, right at the position of the knob-like densities seen in the V1-ATPase structure.

The yeast V1 structure is remarkably similar to the projection structures of the coated vesicle enzyme (13). All the features present in the bovine V-ATPase V1 domain seem to be present in the yeast V1-ATPase, including the knob-shaped and elongated densities at the very top of the molecule. This suggests that there might be no major involvement of V0 subunits in the formation of the peripheral stalk(s) at the top of the V1 as has been suggested for the plant V-ATPase (14). Recently, we have presented a three-dimensional structural model of the V0 domain from the coated vesicle enzyme determined at a resolution of 21 Å (25). The model shows that the N terminus of the 100-kDa {alpha} subunit is folded as a rather globular protein domain, close to the membrane surface and in contact with a protein density of about the same size, possibly subunit d, which is sitting in the middle of the cytoplasmic opening of the proteolipid ring. As for the structure of the isolated V1, projections of three-dimensional model of the isolated V0 are very similar to the projection structure of the V0 domain in the intact bovine V-ATPase (25). Taken together, these data indicate that, although there are significant changes in enzymatic characteristics when V1 is released from the V0 sector (Fig. 2), the subunit rearrangements involved in the binding or release process of V1 from V0 might be more subtle than what can be resolved in the current three-dimensional models of the protein domains. Further experiments will be required to define the exact locations of the stalk subunits in the V1 and V0 domains and the intact V-ATPase, experiments which are ongoing in our laboratories.


   FOOTNOTES
 
* This work was supported in part by National Institutes of Health Grants GM58600 (to S. W.) and GM50322 (to P. M. K.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed. Tel.: 909-787-3131; Fax: 909-787-4434; E-mail: stephan.wilkens{at}ucr.edu.

1 The abbreviations used are: V-ATPase, vacuolar ATPase; 3D, three-dimensional; MALDI-TOF, matrix-assisted laser desorption ionization time-of-flight; DTT, dithiothreitol; F-ATPase, F1F0-ATP synthase. Back

2 M. Tarsio and P. M. Kane, manuscript in preparation. Back


   ACKNOWLEDGMENTS
 
We thank Norton Kitagawa for help with the mass spectrometry analysis.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Nishi, T., and Forgac, M. (2002) Nat. Rev. Mol. Cell Biol. 3, 94-103[CrossRef][Medline] [Order article via Infotrieve]
  2. Stevens, T. H., and Forgac, M. (1997) Annu. Rev. Cell Dev. Biol. 13, 779-808[CrossRef][Medline] [Order article via Infotrieve]
  3. Finbow, M. E., and Harrison, M. A. (1997) Biochem. J. 324, 697-712[Medline] [Order article via Infotrieve]
  4. Nelson, N., and Harvey, W. R. (1999) Physiol. Rev. 79, 361-385[Abstract/Free Full Text]
  5. Wilkens, S. (2001) Cell Biochem. Biophys. 34, 191-208[CrossRef][Medline] [Order article via Infotrieve]
  6. Dschida, W. J., and Bowman, B. J. (1992) J. Biol. Chem. 267, 18783-18789[Abstract/Free Full Text]
  7. Arai, H., Terres, G., Pink, S., and Forgac, M. (1988) J. Biol. Chem. 263, 8796-8802[Abstract/Free Full Text]
  8. Xu, T., Vasilyeva, E., and Forgac, M. (1999) J. Biol. Chem. 274, 28909-28915[Abstract/Free Full Text]
  9. Cross, R. L. (2000) Biochim. Biophys. Acta 1458, 270-275[Medline] [Order article via Infotrieve]
  10. Imamura, H., Nakano, M., Noji, H., Muneyuki, E., Ohkuma, S., Yoshida, M., and Yokoyama, K. (2003) Proc. Natl. Acad. Sci. U. S. A. 100, 2313-2315
  11. Hirata, T., Iwamotot-Kihara, A., Sun-Wada, G.-H., Okajima, T., Wada, Y., and Futai, M. (2003) J. Biol. Chem. 278, 23714-23719[Abstract/Free Full Text]
  12. Boekema, E. J., Ubbink-Kok, T., Lolkema, J. S., Brisson, A., and Konings, W. N. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 14291-14293[Abstract/Free Full Text]
  13. Wilkens, S., Vasilyeva, E., and Forgac, M. (1999) J. Biol. Chem. 274, 31804-31810[Abstract/Free Full Text]
  14. Domgall, I., Venzke, D., Lüttge, U., Ratajczak, R., and Böttcher, B. (2002) J. Biol. Chem. 277, 13115-13121[Abstract/Free Full Text]
  15. Kane, P. M. (1995) J. Biol. Chem. 270, 17025-17032[Abstract/Free Full Text]
  16. Sumner, J. P., Dow, J. A., Earley, F. G., Klein, U., Jäger, D., and Wieczorek, H. (1995) J. Biol. Chem. 270, 5649-5653[Abstract/Free Full Text]
  17. Gräf, R., Harvey, W. R., and Wieczorek, H. (1996) J. Biol. Chem. 271, 20908-20913[Abstract/Free Full Text]
  18. Parra, K. J., Keenan, K. L., and Kane, P. M. (2000) J. Biol. Chem. 275, 21761-21767[Abstract/Free Full Text]
  19. Sikorsky, R. S., and Hieter, P. (1989) Genetics 122, 19-27[Abstract/Free Full Text]
  20. Charsky, C. M., Schumann, N. J., and Kane, P. M. (2000) J. Biol. Chem. 275, 37232-37239[Abstract/Free Full Text]
  21. Elble, R. (1992) BioTechniques 13, 18-20[Medline] [Order article via Infotrieve]
  22. Gharahdahgi, F., Weinberg, C., Meagher, D., Imai, B., and Mische, S. (1999) Elctrophoresis 20, 601-605[CrossRef]
  23. Lüdtke, S. J., Baldwin, P. R., and Chiu, W. (1999) J. Struct. Biol. 128, 82-97[CrossRef][Medline] [Order article via Infotrieve]
  24. van Heel, M., Harauz, G., Orlova, E. V., Schmidt, R., and Schatz, M. (1996) J. Struct. Biol. 116, 17-24[CrossRef][Medline] [Order article via Infotrieve]
  25. Wilkens, S., and Forgac, M. (2001) J. Biol. Chem. 276, 44064-44068[Abstract/Free Full Text]
  26. Dube, P., Tavares, P., Lurz, R., and van Heel, M. (1993) EMBO J. 12, 1303-1309[Medline] [Order article via Infotrieve]
  27. Schatz, M., Orlova, E. V., Dube, P., Jäger, J., and van Heel, M. (1995) J. Struct. Biol. 114, 28-40[CrossRef][Medline] [Order article via Infotrieve]
  28. Böttcher, B., Wynne, S. A., and Crowther, R. A. (1997) Nature 386, 88-91[CrossRef][Medline] [Order article via Infotrieve]
  29. Xie, X. S. (1996) J. Biol. Chem. 271, 30980-30985[Abstract/Free Full Text]
  30. Supekova, L., Supek, F., Ma, Y., and Nelson, N. (1995) J. Biol. Chem. 270, 13726-13732[Abstract/Free Full Text]
  31. Wieczorek, H., Grüber, G., Harvey, W. R., Huss, M., Merzendorfer, H., and Zeiske, W. (2000) J. Exp. Biol. 203, 127-135[Abstract]
  32. Abrahams, J. P., Leslie, A. G., Lutter, R., and Walker, J. E. (1994) Nature 370, 621-628[CrossRef][Medline] [Order article via Infotrieve]
  33. Gibbons, C., Montgomery, M. G., Leslie, A. G. W., and Walker, J. E. (2000) Nat. Struct. Biol. 7, 1055-1061[CrossRef][Medline] [Order article via Infotrieve]
  34. Arata, Y., Baleja, J. D., and Forgac, M. (2002) J. Biol. Chem. 277, 3357-3363[Abstract/Free Full Text]
  35. Arata, Y., Baleja, J. D., and Forgac, M. (2002) Biochemistry 41, 11301-11307[CrossRef][Medline] [Order article via Infotrieve]
  36. Bianchet, M. A., Hullihen, J., Pedersen, P. L., and Amzel, L. M. (1998) Proc. Natl Acad. Sci. U. S. A. 95, 11065-11070[Abstract/Free Full Text]
  37. Xu, T., and Forgac, M. (2000) J. Biol. Chem. 275, 22075-22081[Abstract/Free Full Text]
  38. Landolt-Marticorena, C., Williams, K. M., Correa, J., Chen, W., and Manolson, M. F. (2000) J. Biol. Chem. 275, 15449-15457[Abstract/Free Full Text]
  39. Lu, M., Vergara, S., Zhang, L., Holliday, L. S., Aris, J., and Gluck, S. L. (2002) J. Biol. Chem. 277, 38409-38415[Abstract/Free Full Text]
  40. Sagermann, M., Stevens, T. H., and Matthews, B. W. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 7134-7139[Abstract/Free Full Text]
  41. Zhang, X., Malhotra, R., and Guidotti, G. (2000) J. Biol. Chem. 275, 35592-35599[Abstract/Free Full Text]
  42. Lu, X., Yu, H., Liu, S. H., Brodsky, F. M., and Peterlin, B. M. (1998) Immunity 8, 647-656[CrossRef][Medline] [Order article via Infotrieve]
  43. Grüber, G., Radermacher, M., Ruiz, T., Godovac-Zimmermann, J., Canas, B., Kleine-Kohlbrecher, D., Huss, M., Harvey, W. R., and Wieczorek, H. (2000) Biochemistry 39, 8609-8616[CrossRef][Medline] [Order article via Infotrieve]
  44. Radermacher, M., Ruiz, T., Wieczorek, H., and Grüber, G. (2001) J. Struct. Biol. 135, 26-37[CrossRef][Medline] [Order article via Infotrieve]
  45. Tomashek, J. J., Graham, L. A., Hutchins, M. U., Stevens, T. H., and Klionsky, D. J. (1997) J. Biol. Chem. 272, 26787-26793[Abstract/Free Full Text]
  46. Ogilvie, I., Aggeler, R., and Capaldi, R. A. (1997) J. Biol. Chem. 272, 16652-16656[Abstract/Free Full Text]
  47. Weber, J., Muharemagic, A., Wilke-Mounts, S., and Senior, A. E. (2003) J. Biol. Chem. 278, 13623-13626[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Biol. Chem.Home page
N. Kitagawa, H. Mazon, A. J. R. Heck, and S. Wilkens
Stoichiometry of the Peripheral Stalk Subunits E and G of Yeast V1-ATPase Determined by Mass Spectrometry
J. Biol. Chem., February 8, 2008; 283(6): 3329 - 3337.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
J. M. Rizzo, M. Tarsio, G. A. Martinez-Munoz, and P. M. Kane
Diploids Heterozygous for a vma13{Delta} Mutation in Saccharomyces cerevisiae Highlight the Importance of V-ATPase Subunit Balance in Supporting Vacuolar Acidification and Silencing Cytosolic V1-ATPase Activity
J. Biol. Chem., March 16, 2007; 282(11): 8521 - 8532.
[Abstract] [Full Text] [PDF]

Home page