©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
MOP2 (SLA2) Affects the Abundance of the Plasma Membrane H-ATPase of Saccharomyces cerevisiae(*)

(Received for publication, September 27, 1994; and in revised form, January 4, 1995)

Songqing Na(§)(¶) Marina Hincapie(¶)(**) John H. McCusker (§§) James E. Haber (¶¶)

From the Rosenstiel Basic Medical Sciences Research Center and the Department of Biology, Brandeis University, Waltham, Massachusetts 02254

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

The abundance of yeast plasma membrane H-ATPase on the cell surface is tightly regulated. Modifier of pma1 (mop) mutants were isolated as enhancers of the mutant phenotypes of pma1 mutants. mop2 mutations reduce the abundance and activity of Pma1 protein on the plasma membrane without affecting the abundance of other prominent plasma membrane proteins. The MOP2 gene encodes a 108-kDa protein that has previously been identified both as a gene affecting the yeast cytoskeleton (SLA2) (Holtzman, D.A., Yang, S., and Drubin, D. G.(1993) J. Cell Biol. 122, 635-644) and as a gene affecting endocytosis (END4) (Raths, S., Roher, J., Crausaz, F., and Riezman, H.(1993) J. Cell Biol. 120, 55-65). In some strains, MOP2 (SLA2) is essential for cell viability; in others, a deletion mutant is temperature sensitive for growth. mop2 mutations do not reduce the transcription of PMA1 nor do they lead to the accumulation of Pma1 protein in any intracellular compartment. An epitope-tagged MOP2 protein behaves as a plasma membrane-associated protein whose abundance is proportional to its level of gene expression. Over-expression of MOP2 relieved the toxicity caused by the over-expression of PMA1 from a high copy plasmid; conversely, the growth of mop2 strains was inhibited by the presence of a single extra copy of PMA1. We conclude that MOP2 (SLA2) encodes a plasma membrane-associated protein that is required for the accumulation and/or maintenance of plasma membrane H-ATPase on the cell surface.


INTRODUCTION

The plasma membrane proton-translocating ATPase (H-ATPase) of Saccharomyces cerevisiae is required for the maintenance of intracellular pH and the large electrochemical gradient that is necessary for the transport of many nutrients (Goffeau and Slayman, 1981; Bowman and Bowman, 1986; Slayman, 1986; Goffeau and Dufour, 1988; Serrano, 1988). Yeast H-ATPase is structurally and functionally analogous to the mammalian cation-translocating P-type ATPases including Na/K-, H/K-, and Ca-ATPase (Shull et al., 1985; MacLennan et al., 1986; Brandl et al., 1986). A comparison of the sequence of these enzymes has shown extensive homology that includes most functional domains (Serrano et al., 1986; Serrano, 1988). Like other P-type ATPases, the yeast plasma membrane H-ATPase has a catalytic subunit of 100 kDa, hydrolyzes ATP via a transient aspartylphosphate intermediate, and is inhibited by low concentrations of vanadate. The H-ATPase is a major cell protein, representing about 15% of the plasma membrane protein and 0.3% of total yeast protein (Serrano, 1991; Serrano, 1988; Bowman et al., 1981). Genetic studies have shown that the PMA1 gene encoding the H-ATPase is essential (Serrano et al., 1986).

The abundance of the yeast H-ATPase in the plasma membrane appears to be tightly regulated, as over-expression of the PMA1 gene on multicopy plasmids in yeast yields no significant increase in the amount of PMA1 protein (Pma1) in the plasma membrane (Eraso et al., 1987). This regulation does not appear to occur at the transcriptional level but must occur at any one of a number of steps during its proper entry into and progress through the entire secretory pathway or in its maintenance in the plasma membrane.

Many membrane proteins exist as part of homo-oligomeric or hetero-oligomeric structures. Many of the animal P-type ATPases form an active holoenzyme containing two subunits, a 100-kDa alpha subunit and a smaller 40-60-kDa glycoprotein beta subunit (Lingrel et al., 1988). The alpha subunit is considered the catalytic subunit while the beta subunit appears to be involved with the folding and maturation of the alpha subunit (Geering et al., 1987). The association of the alpha subunit with an appropriate beta protein is required for exit from endoplasmic reticulum (Jaunin et al., 1992) and subsequent transport to the plasma membrane (Noguchi et al., 1987; Takeyasu and Kawakami, 1989). For example, functional expression of Na/K-ATPase in various expression systems such as Xenopus oocytes or stably transfected cell lines requires the co-expression of a compatible endogenous or exogenous beta subunit (Noguchi et al., 1987; Takeyasu et al., 1988; Lemas et al., 1992). The requirement for the expression of both alpha and beta subunits is also seen when the mammalian Na/K-ATPase genes are expressed in yeast (Horowitz et al., 1990). In the case of the Na/K-ATPase, the beta subunit appears to play an important role in catalysis and in interactions with extracellular inhibitors (Geering, 1990). A beta subunit is not required for the catalytic activity of the Ca-ATPases (Wuytack and Racemackers, 1992). The yeast H-ATPase does not appear to require a beta-like protein for its maturation, transport, or functional expression on the plasma membrane. Although there is a 115-kDa glycoprotein tightly associated with the H-ATPase during the purification (Vai et al., 1988), the activity of the enzyme is normal in a mutant strain lacking the glycoprotein (Serrano et al., 1991).

Previous research has demonstrated that the yeast plasma membrane H-ATPase activity is regulated in response to growth conditions, increasing severalfold during glucose metabolism (Serrano, 1983) and in an acid medium (Eraso et al., 1987). This regulation is likely to occur by one of several mechanisms of post-translational modification. Protein phosphorylation, caused by several protein kinases, has been demonstrated both for the Ca-ATPase (Caroni and Carafoli, 1981) and for the yeast H-ATPase (Chang and Slayman, 1991). In addition, the C-terminal domain of the mammalian Ca-ATPase can interact with calmodulin to regulate its activity (Carafoli, 1992). Allosteric regulation of the plant H-ATPase enzyme by a synthetic C-terminal peptide has also been reported (Palmgren et al., 1991). Both mutation or deletion of the analogous C terminus of the yeast H-ATPase leads to the loss of glucose activation (Portillo et al., 1989). Whether such regulation in yeast depends on another interacting protein is unknown.

In this paper, we describe the identification and characterization of MOP2, a gene that is essential in some strains and nearly so in others. MOP2 protein behaves as a plasma membrane-associated protein. Mutations in MOP2 reduce dramatically the activity and abundance of Pma1 in the plasma membrane without affecting the abundance of other prominent plasma membrane proteins. Over-expression of MOP2 can rescue the inhibitory effect on cell growth caused by high expression of PMA1. Surprisingly, MOP2 has recently been identified in two very different searches of new yeast mutants. Holtzman et al. (1993) have identified mutations in the same gene as SLA2, synthetic lethal mutation in combination with a deletion of an actin binding protein. At the same time, Raths et al.(1993) have identified the same gene as END4, essential for endocytosis. (^1)These results suggest that MOP2 (SLA2) plays a central role in the maintenance and distribution of integral membrane proteins, including Pma1.


EXPERIMENTAL PROCEDURES

Yeast Strains and Cultivation

All yeast strains were isogenic derivatives of Y55 (HO gal3 MAL1 SUC1). The isolation and characterization of pma1-114 was described earlier (McCusker et al., 1987). Strain SN53 (MATaho::LEU2 ura3-1 arg4-1 mop2-2) was derived from a stable segregant of diploid SN46, which was created by crossing wild type SH89 (MATaho:LEU2 ade6-1 arg4-1 trp5-1 ura3-1 leu2-1) with MATalpha spores of diploid Y55-300 (HO ade6-1 mop2-2). The diploid strain Y55-296 (HO ura3-1 leu2-1) was used to study the over-expression of MOP2, PMA1, or both. Strain YPS3-2A (ho MATaleu2-Delta1, his3-Delta200, trp1-Delta63, pma1-Delta::HIS3, pma2-Delta::TRP1) was provided by A. Goffeau. Yeast strains were transformed using the lithium acetate procedure of Ito et al.(1983) as modified by Schiestl and Gietz(1989).

A haploid-viable, temperature-sensitive derivative of a mop2::URA3 deletion was obtained by transforming diploid DDY288 (Holtzman et al., 1993) with the URA3-marked disruption of MOP2 (SLA2) in plasmid pSN101, described below.

The media used in these experiments have been previously described (Perlin et al., 1989). Cells were grown in YEPD (1% (w/v) yeast extract, 2% (w/v) Bacto-peptone, 2% (w/v) dextrose, pH 5.5) or standard defined media lacking uracil or leucine. The ability of cells to grow under conditions of low external pH was tested in YEPD medium adjusted to pH 3.0 with HCl. Hygromycin B resistance was scored using YEPD containing 300 or 500 µg/ml hygromycin B. Cycloheximide hypersensitivity was scored using YEPD containing 0.5 µg/ml cycloheximide.

Plasmid Construction

The MOP2 gene was originally obtained as a genomic DNA insert into the multicopy plasmid YEp24. The MOP2 containing clone recovered from the screening of a genomic DNA library was designated pSN12. A restriction map is shown in Fig. 1. Plasmids with different deletions of MOP2 were derived from pSN12 as follows. Plasmid pSN33 was constructed by cutting pSN12 with BglII and ligated so that a 3.6-kb (^2)BglII-BglII fragment was deleted from the insert DNA. Plasmid pSN35 was constructed by cutting pSN12 with PvuII and ligated so that a 2.9-kb fragment was deleted. Plasmid pSN42 was constructed by inserting a 4.6-kb BamHI-XbaI fragment from pSN12 into YEp24 digested with BamHI and NheI. Plasmid pSN47 was constructed by inserting a 3.4-kb BglII-BglII fragment from pSN12 into BamHI-digested YEp24. pSN48 was constructed by inserting the 4.6-kb BamHI-XbaI fragment from pSN12 into BamHI and XbaI-digested pBluscript KS(-) (Stratagene). A MOP2 null mutation was created by cutting pSN48 with SphI, end-filling with Klenow fragment, and then ligating with a blunt-ended 1.1-kb BamHI-BamHI fragment of the URA3 gene, creating plasmid pSN101 (Fig. 1B).


Figure 1: Cloning and analysis of MOP2. A, deletion analysis of the MOP2-containing DNA fragment from a genomic library. Both the 4.6-kb BamHI-XbaI and the 3.3-kb BamHI-PflMI fragment can complement all mop2 phenotypes. DNA sequence reveals that only the BamHI-XbaI fragment contains the intact MOP2 open reading frame. Restriction endonuclease sites are labeled as follows: E, BstEII; B, BamHI; Bg, BglII; P, PvuII; Pf, PflMI; X, XbalI. B, replacement of MOP2 open reading frame with URA3 gene in diploid SN189. Deletion of MOP2 caused cell lethality, as evidenced by the recovery of only 2 viable (Ura) spores per tetrad. C, location of a putative membrane-spanning domain and point of insertion of the c-Myc epitope.



Epitope Tagging

Plasmid pSN117 is identical to pSN42 except for the addition of the c-Myc epitope at the C terminus of the MOP2 open reading frame. Epitope tagging of Mop2 was performed by inserting an 11-amino acid c-Myc epitope at the C terminus of Mop2 between amino acid residues 904 and 905 (Mop2::c-Myc). Two complementary oligomer DNAs, with 30 nucleotides encoding the c-Myc epitope flanked on both ends by six nucleotides that define the unique PflMI restriction site, were synthesized. The DNA sequence of the coding strand is 5`-GAGCAAAAATTGATTTCTGAGGAGGACTTGAATGTC-3`. pSN48 was digested with PflMI, gel purified, and then ligated with the annealed c-Myc epitope oligomers, creating plasmid pSN110. Insertion of the c-Myc epitope was checked by both restriction enzyme digestion and DNA sequencing. To test if Mop2::c-Myc was fully functional, plasmid pSN117 was transformed into the mop2-2 mutant strain SN53, to test the ability of Mop2::c-Myc to complement all mop2 phenotypes.

Plasmid pSN145 is a LEU2-marked high copy plasmid in which the BamHI-XbaI fragment of plasmid pSN117 carrying the epitope-tagged MOP2 gene was inserted in place of the BamHI-NheI fragment of plasmid YEp21.

Plasmid pNS146 is a derivative of pNS110 containing a 1.1-kb URA3 fragment inserted at an NruI site in the 3`-non-coding region of the epitope-tagged MOP2 gene.

DNA Sequence Analysis

The nucleotide sequence of the MOP2 gene was determined by DNA sequence analysis of the 4.6-kb genomic BamHI-XbaI fragment, which fully complements all mop2 phenotypes. Nested deletions of this insert in pSN48 were generated with the Erase-a-Base system according to the protocol provided by the manufacturer (Promega). Single-stranded DNA was made following infection with helper phage M13K07. DNA was sequenced by the dideoxy chain termination method (Sanger et al., 1977) modified for single-stranded and double-stranded template with sequenase as described by the enzyme supplier (U. S. Biochemical). Protein homology searches were performed at the National Center for Biotechnology Information using the Blast network service (Altschul et al., 1990). The accession number L12352 has been assigned to the MOP2 sequence by GenBank.

Plasma Membrane Isolation and Measurements of Abundance and ATP Hydrolysis of H-ATPase

For the preparation of plasma membranes, cells were grown overnight in 5-ml cultures in either YEPD or selective media for plasmid retention. 1 ml of the saturated culture was transferred to 100 ml of the same media and grown to a cell density of 5.0 times 10^6. 50 ml of these cells were then subcultured into 1 liter of YEPD. Cells were harvested when they reached an A = 4-5 absorbance units. Yeast plasma membranes were purified by differential centrifugation and sucrose gradient centrifugation (Perlin et al., 1989).

ATPase activity was assayed as described (Perlin et al., 1989). Total protein concentration was determined by the Lowry method. Abundance of H-ATPase in purified plasma membrane preparations was examined by SDS-PAGE electrophoresis. Samples were solubilized in SDS-PAGE loading buffer and incubated at 37 °C for 10 min before loading on a 10% SDS-PAGE gel. Proteins were visualized with Coomassie Blue or by Western blotting. For Western blot analysis, proteins were transferred to nitrocellulose membranes. Transfer buffer was 25 mM Tris, 192 mM glycine, 10% methanol, 0.03% SDS, pH 8.4. Transfer was carried out for 1.5 h at a constant current of 275 mA. Protein on immunoblots was detected using a chemiluminescence detection system according to the protocol provided by the manufacturer (Amersham Corp.).

The effect of mop2 on the abundance of Pma1 was determined by comparing a wild type (SN197) and a mop2-2 mutant (SN53) strain. Cells were grown at 25 °C in YEPD media as described above. When cells reached an A = 1, the cultures were divided into three flasks. Growth was continued in one flask at 25 °C for 4 h and for either 2 or 4 h at 37 °C in the other two flasks. Cells were harvested, and plasma membranes were prepared.

Solubilization of Mop2 from plasma membranes was carried out by treating the purified plasma membrane with chemical reagents. 100 µg of purified plasma membranes from SN330 (transformant of Y55-296 with pSN117) were resuspended in 200 µl of one of the following solubilizing reagents: 1% Triton X-100, 1% SDS, 1% Zwittergent-14 (Boehringer Mannheim), 2 M urea, or 0.1 M Na(2)CO(3), pH 11. Samples were incubated on ice for 1 h. As a control, plasma membrane was incubated with membrane buffer (10 mM Tris-Cl, pH 7.5, 1 mM EDTA, 10% glycerol) under the same conditions. Each mixture was subjected to sedimentation for 1 h at 100,000 rpm in a table top ultracentrifuge. Each pellet was resuspended in 200 µl of 10 mM Tris buffer, pH 7.5. Equivalent amounts of the high speed supernatant and the solubilized pellet were then analyzed by Western blot, probed with the 9E10 anti c-Myc monoclonal antibody.

Fluorescence Microscopy

Staining of fixed yeast cells by indirect immunofluorescence was carried out as described by Davis and Fink(1990).

Immunoprecipitation

Purified plasma membranes were diluted to 1.0 mg/ml with immunoprecipitation buffer (20 mM Tris, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, pH 7.5) (Ip buffer). 100 µl (100 µg) were immunoprecipitated overnight with 10 µg of purified anti-Pma1 rabbit polyclonal antibody and 20 µl of protein A-Sepharose. Immunoprecipitates were washed four times with 500 µl of Ip buffer, and the beads were resuspended in 20 µl of SDS-PAGE buffer. Samples were heated at 37 °C for 10 min and subjected to electrophoresis in a 10% SDS-PAGE gel. Transfer of proteins to nitrocellulose was carried out as described above.


RESULTS

Isolation of mop2 Mutants

Mutations in the PMA1 gene encoding the yeast plasma membrane H-ATPase cause a sensitivity to lowered pH and a reduction in cellular membrane potential, with a consequent resistance to hygromycin B. Modifier of pma1 (mop) mutants were isolated as enhancers of the mutant phenotypes of several pma1 mutations. Three spontaneous mutations were initially discovered that increased the hygromycin B resistance of strains carrying a pma1 mutation from 300 to 500 µg/ml. All three of these mutations proved to confer hypersensitivity to cycloheximide, preventing growth at 0.5 µg/ml, a concentration at which both wild type and pma1 mutants can grow. All three mop mutants (mop1, -2, and -3) were shown to be unlinked to pma1 by crossing the mop pma1 strains to a wild type strain and following the segregation of hygromycin B resistance, sensitivity to acid-loading conditions (0.2 M acetate, pH 5.0 or 4.8), and cycloheximide hypersensitivity. The cycloheximide hypersensitivity of the mop2 mutants segregated as a Mendelian trait (2:2) and segregated independently of the sensitivity to acid-loading conditions that marked pma1 (McCusker, 1986). While pma1 by itself is resistant to hygromycin B at 300 µg/ml, resistance on plates containing 500 µg/ml was only seen for segregants carrying both mutations. mop mutants are not, by themselves, hygromycin B resistant (Fig. 2). Complementation tests showed that the three mop mutations defined separate complementation groups. Further analysis of mop1 and mop3 will be described elsewhere.


Figure 2: Phenotypes of pma1-114, mop2-1, mop2-2, SH89, and the wild type strain double mutant pma1-114 mop2-1. pma1-114 (Perlin et al., 1989) is resistant to hygromycin B at a concentration of 300 µg/ml but sensitive to hygromycin B at 500 µg/ml and sensitive to YEPD + 0.2 M acetate, pH 4.8. mop2-1 is hypersensitive to cycloheximide (YEPD + 0.5 µg/ml) and sensitive to YEPD + 0.2 M acetate, pH 5.0. mop2-2 has the same phenotype as mop2-1 but is also temperature sensitive (no growth at 37 °C). The phenotypes of higher concentration hygromycin B resistance, low pH sensitivity, and high osmotic stress sensitivity are only seen for the double mutant pma1-114 mop2-1.



Three additional mop2 alleles (mop2-2, mop2-3, and mop2-4) were subsequently isolated by their hyper-hygromycin B resistance of pma1-114. Both mop2-2 and mop2-4 were also found to be temperature sensitive for growth at 37 °C.

Strains carrying mutations in both MOP2 and PMA1 have several interesting phenotypes that are absent in the mop2 and pma1 mutants. Although some severe pma1 alleles are unable to grow at pH 3.0 and are sensitive to osmotic stress (McCusker et al., 1987), pma1-114 is resistant to both of these conditions, although it is hygromycin B resistant and sensitive to acid loading (YEPD + 0.2 M acetate, pH 4.8). mop2 pma1-114 strains are sensitive to low external pH (YEPD, pH 3.0) and high osmotic medium (YEPD plus 2 M glycerol) in addition to showing higher resistance to hygromycin B (Fig. 2). This proved to be true of both mop2-1 and mop2-3 and for the temperature-sensitive mop2-2 and mop2-4 alleles grown at their permissive temperature of 25 °C.

mop2 Reduces Pma1 Abundance in the Plasma Membrane

Plasma membrane H-ATPase is a very abundant membrane protein representing about 15% of total plasma membrane proteins (Bowman et al., 1981). All four mop2 alleles caused a remarkable reduction in the activity of Pma1. At the permissive temperature, the abundance of Pma1 in a mop2-2 mutant was estimated to be 60% of wild type. At the non-permissive temperature, a decrease in the abundance of both wild type and mop2-2 was observed. The decrease in abundance of wild type Pma1 has been previously reported in heat-stressed cells (40 °C) (Panaretou and Piper, 1992). They showed that the reduction of Pma1 is accompanied by the induction of a 30-kDa heat shock protein. The enrichment of a 30-kDa protein is also seen in our strains at 37 °C (Fig. 3A). However, the induction of this protein is more pronounced in the mop2-2 mutant. There are other low molecular weight proteins that are also enriched in the mop2-2, but not wild type. Western blots probed with the monoclonal antibody 9E10 (Fig. 3B), as well as a panel of anti-Pma1 polyclonal antibodies (data not shown), indicated that there was no reactivity to any of the low molecular weight bands. We conclude that these low molecular weight proteins are not degradation products of Pma1.


Figure 3: Abundance of H-ATPase in wild type (SN197) and mop2-2 (SN53) strains. A, plasma membranes derived from SN197 and SN53 were analyzed by SDS-gel electrophoresis on a 10% gel (40 µg/lane). Membranes were prepared from both SN197 and SN53 cells grown at 25 °C (the permissive temperature for mop2-2) and 37 °C (non-permissive temperature) as described under ``Experimental Procedures.'' Lane1, wild type (25 °C); lane2, wild type (37 °C, 2.0 h); lane3, wild type (37 °C, 4.0 h); lane4, mop2-2 (25 °C); lane5, mop2-2 (37 °C, 2.0 h); lane6, mop2-2 (37 °C, 4.0 h). B, immunoblot of the same plasma membranes as in A. 5 µg/lane were loaded on a 10% mini SDS-PAGE gel and transferred to nitrocellulose membrane. Blot was probed with anti-Pma1 monoclonal antibody.



PMA1 Transcription Is Not Affected by the mop2 Mutation

The decreased abundance of Pma1 in the plasma membrane could be due to alterations in the transcription, translation, or post-translational processing of the protein. To investigate the effect of mop2 on transcription, we carried out a primer extension analysis of the abundance of PMA1 mRNA in wild type and temperature-sensitive mop2-2 cells. There was no difference in PMA1 mRNA abundance between wild type and the mop2-2 mutant at either the permissive or non-permissive temperature (data not shown). This indicates that Pma1 reduction by mop2 was not caused by reducing PMA1 transcription. Similarly, the effect of mop2 mutants could not be suppressed by introducing additional copies of the PMA1 gene; in fact, extra copies of PMA1 were highly toxic to mop2 strains (see below).

A further experiment was done by replacing the PMA1 promoter with the galactose-inducible GAL1 promoter (Cid et al., 1987) to test whether the effect of mop2 on the abundance of the Pma1 could be suppressed by placing the PMA1 gene under the control of another promoter. Strain SN225 was constructed by crossing the mop2-2 strain, SN234 (MATalpha ho::LEU2 ura3-1 leu2-1 arg4-1 mop2-2 GAL3), with a MATalpha spore of strain SH132, which contains PMA1 under galactose-inducible promoter. The correct haploid meiotic segregants, confirmed by both Southern blot and scoring for the mop2-2 spore colonies that grow on galactose, were used for measurement of Pma1 abundance in plasma membranes. SDS protein gels of the purified plasma membrane showed that abundance of Pma1 was still reduced by mop2 to the same extent as the cells with the normal PMA1 promoter (data not shown). These experiments demonstrated that MOP2 does not act at the level of PMA1 transcription.

Cloning and Mapping of MOP2

The MOP2 gene was cloned by complementation of the mop2-2 allele for its temperature sensitivity (YEPD at 37 °C) and cycloheximide hypersensitivity (YEPD plus 0.5 µg/ml cycloheximide) at 25 °C. The mop2 mutant strain, SN53, was transformed with genomic DNA libraries made in both the centromere vector YCp50 and the multicopy vector YEp24. Ura transformants were replica plated onto YEPD plus 0.5 µg/ml cycloheximide grown at 25 °C and onto YEPD at 37 °C. 2 out of 900 Ura transformants (one each from the single copy and multicopy library) were found to complement the mop2 phenotypes. Restriction mapping reveals that both clones had overlapping genomic DNA insertions. All subsequent work was carried out with the YEp24 clone, designated pSN12 (Fig. 1A).

MOP2 was mapped to chromosome XIV by probing a Southern blot of a yeast chromosome separating gel (data not shown) using P-labeled cloned MOP2-containing DNA fragment. Tetrad analysis was then carried out to establish that MOP2 lies approximately 7 centimorgans proximal to RAD50 on the left arm of chromosome XIV (data not shown). There are no other known genes in this vicinity (Mortimer et al., 1992).

In the clone from the YEp24 genomic bank, MOP2 was contained in a 10-kb insert. To narrow down the fragment that encodes MOP2, several deletions of the MOP2 clone were made and tested for their functional complementation (Fig. 1A). The smallest fragment that complemented a mop2 mutant was the 3.47-kb BamHI-PflMI fragment. However, DNA sequencing of MOP2 shows that the Mop2 open reading frame extends 63 amino acids beyond the PflMI site (see below). This suggests that at least the last 63 amino acids of C terminus are not essential for Mop2 function. Moreover, an insertion of c-Myc epitope at the PflMI site of MOP2 did not affect the function of Mop2 (see below).

DNA Sequence of MOP2

The nucleotide sequence of the 4.6-kb BamHI-XbaI fragment, which complemented the mop2 phenotypes showed that MOP2 encodes an open reading frame of 968 amino acids. Hydropathy analysis of the deduced Mop2 polypeptide predicts a single putative membrane-spanning segment (Fig. 1C). A search of the data base revealed that the same gene has recently been cloned and sequenced under the name SLA2 (Holtzman et al., 1993). The two different versions of the same sequence are virtually identical. There are no amino acid differences, but at positions 910 and 911, our sequence reads GC while that of Holtzman et al.(1993) is CG. The protein has several motifs that are characteristic of DNA binding proteins, including stretches of polyglutamic acids and a putative leucine zipper, although Mop2 is not involved in transcription. The most striking homology revealed by searching the data base was found for the last 190 amino acids of the C terminus of Mop2 (amino acid residues 689-968), which are highly homologous (33% identity) to the C terminus of the integral membrane-associated mammalian protein talin (Reed et al., 1990); however, since a deletion of the terminal 63 amino acids of Mop2 is fully functional (see above), it is not clear if this homologous region is important for Mop2 function.

A Deletion of MOP2 Is Lethal in Some Strains and Temperature Sensitive in Others

To demonstrate that we had indeed cloned the MOP2 gene rather than a suppressor of mop2-2, we integrated a linear fragment carrying the MOP2 gene with a URA3 gene inserted in the 3`-non-coding region. Approximately half of the Ura transformants of temperature-sensitive mop2-2 strain SN53 became Ts, while the rest remained Ts. Southern blot analysis (not shown) demonstrated that all of the insertions were in the same location. Thus, approximately half of the time, the targeted insertion of URA3 co-converted the adjacent mop2-2 allele to wild type, as expected if the cloned sequence integrated at mop2-2.

A null mutation of MOP2 was created by removal of almost the entire protein coding sequence and replacement of this segment with the selectable marker URA3 (see ``Experimental Procedures''). This construct, mop2Delta::URA3, was transformed into a derivative of diploid strain, Y55 (HO ura3-1). Stable Ura transformants were selected, and the deletion of MOP2 was confirmed by Southern blot analysis (data not shown). A Ura diploid was sporulated and dissected. Only two viable spores were obtained in each tetrad, all of which were Ura, indicating that MOP2 encodes an essential protein for cell growth. This result is in contrast to that of Holtzman et al.(1993), who found that a virtually identical deletion of the same gene in a different strain yielded viable haploids that were temperature sensitive for growth. It was likely that our strains, based on strain Y55 (McCusker and Haber, 1988), harbor a genetic difference from those used by Holtzman et al.(1993). To confirm this genetic difference, we transformed the same mop2::URA3 deletion construct we had introduced into our strain Y55 into strain DDY228 used by Holtzman et al.(1993). Tetrads of Ura derivatives of DDY228 were sporulated and dissected at 25 °C. After 3 days, there were only two visible (Ura) segregants. However, in contrast with strain Y55, by 5 days two Ura colonies become visible. These very slow growing colonies proved to be unable to grow at 37 °C, in agreement with Holtzman et al.(1993). Thus, there appears to be a genetic difference between the two strains so that deletions of MOP2 (SLA2) are lethal in one strain but viable (though very debilitated) in another.

MOP2 Encodes a Membrane-associated Plasma Membrane Protein

To establish the intracellular localization of Mop2 and to gain some insight into its in vivo function, we marked the C terminus of Mop2 with the c-Myc epitope (see ``Experimental Procedures'' and Fig. 1C). Yeast strain Y55-296 (HO leu2-1 ura3-1) was transformed with pSN117 containing the epitope-tagged MOP2::Myc construct. Cells transformed with pSN42 containing untagged MOP2 were prepared in parallel as a control.

To determine the nature of the membrane association of Mop2, the purified plasma membranes from strain SN330, containing a high copy plasmid with MOP2::c-Myc, were treated with a variety of reagents and resedimented at 100,000 rpm. Equivalent portions of the resulting supernatant were analyzed by immunoblotting (see ``Experimental Procedures''). Mop2 protein was found exclusively in the membrane fraction. No immunoreactivity was found in the high speed supernatant fraction (Fig. 4B, lane2). The results showed that Mop2 is partially extracted by the reagents known to extract peripherally associated membrane proteins and is efficiently extracted by SDS and Zwittergent-14 known to solubilize integral membrane proteins (Fig. 4B). Triton X-100 is less efficient compared with the other two detergents. Taken together, these data indicate that Mop2 itself is a plasma membrane-associated protein.


Figure 4: Mop2 is a membrane-associated protein. A, immunoreactivity of untagged Mop2 (lane1) and tagged Mop2::c-Myc (lane2) Western blot was probed with anti-c-Myc mAb 9E10. B, solubilization of Mop2 from the purified plasma membrane. Purified plasma membrane was isolated from SN330 carrying pSN117 (see ``Experimental Procedures''). 10 µl of high speed pellet (P) and supernatant (S) fractions were analyzed by Western blot. The nitrocellulose membrane was probed with the 9E10 anti-c-Myc mAb.



In some preparations we find two bands, approximately 92 and 108 kDa, that were highly stained with anti c-Myc monoclonal antibody 9E10 (Fig. 4A). The size of 108 kDa is consistent with the predicted molecular size from the MOP2 sequence. The 92-kDa band is most likely a proteolytic digestion product of Mop2. Another protein of approximately 100 kDa is found both in cells expressing MOP2::c-Myc and those expressing an untagged Mop2 protein. This second protein is most likely Pma1, as shown by immunoprecipitation with anti-Pma1 antibodies (data not shown). Immunoreactivity of anti-c-Myc antibody with this protein has been seen previously (Kuchler et al., 1993). Immunoprecipitation of Pma1 does not precipitate the 108-kDa Mop2::c-Myc protein (data not shown). The strength of anti-c-Myc immunoreactivity with Pma1 is strain dependent, as it is less evident in strains provided by A. Goffeau (Supply et al., 1993) (see below).

Further evidence of the plasma membrane location of Mop2 came from immunofluorescence staining by monoclonal anti c-Myc antibody of the cells carrying the plasmid containing epitope-tagged Mop2. The cells showed bright staining of the cell periphery (Fig. 5), as do antibodies against Pma1 (Harris, et al., 1994) and other plasma membrane proteins (Ljungdahl et al., 1992). Despite the fact that anti-c-Myc antibody reacts with Pma1 on Western blots, it does not exhibit any significant indirect immunofluorescence in cells not expressing Mop2::c-Myc (Fig. 5). We conclude that Mop2 is located at the plasma membrane.


Figure 5: Localization of Mop2 by indirect immunofluorescence. Strain YPS3-2A carrying pSN145 was grown in standard defined complete medium lacking leucine for selection of plasmid. Cells carrying the MOP2::c-Myc gene on a high copy plasmid pSN145 (A) and the same strain carrying a similar plasmid but lacking the MOP2 gene, YEp21 (B), were examined by indirect immunofluorescence using an anti-c Myc mAb antibody (toppanels). Lowerpanels, cells viewed by phase contrast and stained with 4,6-diamidino-2-phenylindole.



Over-expression of MOP2 Rescues the Toxicity of Over-expressing PMA1

It has been previously shown that the amount of yeast H-ATPase is only slightly increased by introducing extra copies of PMA1 gene in both single copy and high copy plasmids (Eraso et al., 1987). Moreover, the over-expression of PMA1 is detrimental and slows cell growth (Eraso et al., 1987). One interesting question is whether over-expression of MOP2 can rescue the inhibitory effect caused by over-expression of PMA1. The growth rate of cells containing PMA1 on either centromere or 2-µm plasmids were measured with or without a second high copy plasmid carrying MOP2. While a single extra copy of PMA1 has no effect on growth, high copy over-expression of PMA1 markedly inhibited cell growth (Fig. 6A). This inhibition was suppressed when cells carrying the high copy PMA1 plasmid also contained a high copy MOP2 plasmid (Fig. 6A).


Figure 6: Growth curve of both the wild type (Y55-296) and mop2-2 mutant (SN53) strains carrying one extra copy or high copy plasmid with PMA1, MOP2, or both PMA1 and MOP2. All of the strains were grown at 25 °C with continuous shaking. Measurement of cell growth was done by measuring absorbance at 600 nm. A, growth curve of wild type cells with high copy PMA1 or MOP2 or both (bullet, + YEp24; circle, + high copy plasmid containing PMA1; box, + high copy plasmid containing MOP2 (pSN117); [tirf], + both high copy plasmids containing MOP2 and PMA1). B, growth curve of wild type cells and mop2-2 mutant cells with different copies of PMA1 (bullet, wild type + YEp24; , mop2-2 + YEp24; circle, mop2-2 + high copy plasmid containing PMA1; times, mop2-2 + single copy (centromeric) plasmid containing PMA1 (pSN141).



We also measured the abundance of Pma1 protein in these cells. The SDS-PAGE gel and Western blot depicted in Fig. 7show that the level of Pma1 in the plasma membrane does not change in cells containing different numbers of PMA1 genes. Moreover, the addition of a high copy MOP2 plasmid has no effect on the amount of Pma1 observed. We conclude that the relief of toxicity of over-expressing PMA1 by over-expressing MOP2 may involve a change in the way excess Pma1 protein is transported or turned over and that this requires the participation of the Mop2 protein.


Figure 7: Effect of over-expressing PMA1 and MOP2 on the abundance of Pma1 protein. A, 30 µg of purified plasma membranes were analyzed by SDS-PAGE on a 10% mini gel. Lane1, wild type; lane2, single copy PMA1 (centromeric plasmid); lane3, high copy PMA1 plasmid; lane4, both high copy PMA1 and high copy MOP2 plasmids; lane5, high copy MOP2 plasmid; lane6, molecular weight markers. B, immunoblot of the same plasma membrane samples as in A. Nitrocellulose membrane was probed with anti-Pma1 mAb. In some lanes, a band consistent with being a Pma1 dimer is seen.



Interestingly, wild type and mop2 mutant cells exhibited different sensitivity to the over-expression of PMA1. In mop2-2 cells grown at permissive temperature, a single extra copy of the PMA1 on a centromere plasmid was as deleterious to the rate of cell growth as with the high copy PMA1 plasmid, while growth is inhibited only by high copy PMA1 in wild type cells (Fig. 6B). These results suggest that MOP2 interacts with PMA1 protein and apparently regulates the transport accumulation or turnover of Pma1 protein.

Mop2 Abundance Increases with Increasing Gene Copy Number

One possible role for Mop2 is that it acts like the beta subunits of some mammalian cation ATPases to form a heterodimer with the catalytic subunit to aid in the transport of the alpha subunit to the plasma membrane. This would suggest that the level of Mop2 should regulate the level of Pma1 in the plasma membrane and vice versa. Over-production of Pma1 without over-production of Mop2 does not result in an increase in Pma1 in the plasma membrane. In contrast, the level of Mop2::c-Myc protein in the plasma membrane is much higher in a cell carrying a multicopy plasmid expressing Mop2::c-Myc than in a cell carrying a single copy (centromere-containing) plasmid (data not shown). This suggests that while Mop2 is required for the accumulation of Pma1, the converse is not true; Mop2 can accumulate under conditions where Pma1 does not increase.


DISCUSSION

MOP2 was identified by mutations that enhance the phenotypes of pma1 mutants. This enhancement appears to be caused by markedly reducing the abundance of the mutant Pma1 protein in the plasma membrane. All of the evidence suggests that MOP2 encodes an essential (in our strains) plasma membrane-associated protein that is required for the proper accumulation of the H-ATPase in the plasma membrane. This effect on Pma1 protein is not simply a general change in membrane protein abundance of large (>30 kDa) plasma membrane proteins. Further evidence of a direct relationship between the abundance of functional Mop2 and the abundance of Pma1 protein comes from our finding that the detrimental effects on cell growth by over-expression of PMA1 can also be rescued by over-expression of MOP2. Conversely, a mop2 mutant strain becomes hypersensitive to an increase in PMA1 copy number. In this discussion, we try to understand what the primary role of Mop2 might be, especially in view of the finding that MOP2 (SLA2) has recently been identified by two other mutant screens based on very different aspects of cell biology (Holtzman et al., 1993; Raths et al., 1993). (^3)

Reduction of Pma1 on the plasma membrane could, in principle, be due to any defect in the transport or subsequent degradation pathway caused by the mop2 mutation. Based on both biochemical characterization and immuno-electron microscopic study of the transport vesicles (Holcomb et al., 1988; Brada and Schekman, 1988), it has been shown that Pma1 is transported in the same secretory vesicles as other secretory proteins, including invertase and acid phosphatase. Defects in several SEC genes block the transport of Pma1 protein to the plasma membrane and result in the accumulation of secretory vesicles (Nakamoto et al., 1991; Chang et al., 1993). It does not appear that mop2 blocks Pma1 protein transport in the same way, since we do not see any accumulation of Pma1 protein in the endoplasmic reticulum or any other intracellular locations when we examine mop2-2 strains by immunofluorescence staining at either their permissive (but still mutant) or non-permissive temperature (data not shown). We know such accumulation can be seen by our immunofluorescence methods, as we find very significant accumulation of Pma1 in cells expressing dominant lethal pma1 mutations (Harris et al., 1994). Also, no change in the abundance of Pma1 was observed in cells that overexpress PMA1. This suggests that the regulation of Pma1 abundance might occur at the level of the plasma membrane.

The fact that MOP2 is a plasma membrane-associated protein is compatible with several suggestions about the way it affects the abundance of Pma1. One possibility is that Mop2 functions as a necessary and specific transport protein in the secretion of H-ATPase, similar to the SHR3 protein identified for amino acid permeases (Ljungdahl et al., 1992). Mop2 differs from classic chaperone proteins in that it does not remain in the endoplasmic reticulum or Golgi apparatus but instead is itself found at the plasma membrane.

Alternatively, Mop2 might act similarly to the beta subunit of some mammalian P-type ATPases to form a heterodimer with the alpha subunit that is required for its transport to the plasma membrane. There has been no evidence for such a transport subunit for H-ATPase in yeast, but several recent findings are consistent with the idea that such a beta-like protein exists in yeast and is required for H-ATPase transport to the plasma membrane. For example, when the Arabidopsis plasma membrane H-ATPase is expressed in S. cerevisiae, the plant enzyme is abundant and is fully functional in terms of ATP hydrolysis, formation of a phosphorylated intermediate, and ability to pump protons in reconstituted vesicles (Villalba et al., 1992). However, the plant plasma membrane H-ATPase is not transported to the yeast plasma membrane but remains trapped in the endoplasmic reticulum. Since the accumulated plant H-ATPase is fully functional in vitro, the presence of plant H-ATPase in the endoplasmic reticulum seems unlikely to be due to the improper folding of the enzyme. The plant and yeast H-ATPase share extensive homology in their membrane structure and function; nevertheless, the plant homologue is unable to traverse the secretory apparatus, possibly because it cannot interact properly with a Pma1-specific transport protein. One argument against Mop2 being a beta subunit is that its abundance in the plasma membrane can be significantly increased in the plasma membrane by over-expressing MOP2, but at the same time Pma1 does not hyperaccumulate.

Another possibility is that Mop2 might be needed to combine with and stabilize H-ATPase once it has reached the plasma membrane. It is conceivable that mop2 mutations might accelerate the endocytosis and turnover of Pma1 protein on the plasma membrane. Pma1 is normally a long-lived protein with a half-life of 30 h (Benito et al., 1991). Its half-life in mop2 mutants has not been measured.

Accommodating the Many Phenotypes of mop2, end4, and sla2 Mutants

Three different sets of mutations in the same gene have been identified by very different criteria. mop2 mutants appear to reduce the abundance of a normally very abundant plasma membrane protein. end4 mutants were found because they prevent receptor-mediated and fluid-phase endocytosis (Raths et al., 1993). More recently, end4 mutants have been shown to prevent the normally rapid turnover of the normally low abundance uracil permease protein (Volland et al., 1994). One possible interpretation of these results would be that MOP2 (END4) is required for normal endocytosis but, further, that the apparent stability of Pma1 in the plasma membrane depends on a continual recycling of Pma1 into early endosomes and then back to the plasma membrane. A prediction of this idea would be that Pma1 would have a much reduced half-life in mop2 strains. On the other hand, one might imagine that the earliest steps in endocytosis would require the establishment of the normally high membrane potential generated by Pma1. Consequently, reducing the abundance of Pma1 might prevent normal endocytosis. One argument against this proposition is that the effect of temperaturesensitive mutations of end4 on endocytosis is very rapid, whereas an effect on the accumulation of Pma1 per se would be expected to be relatively slow.

At the same time, sla2 mutations of the same gene were found as synthetic lethal mutations in a strain already deleted for the actin binding protein, ABF1 (Holtzman et al., 1993). sla2 mutations appear to profoundly affect the actin-based cytoskeleton of the cell, as evidenced by a disruption of the polarized distribution of actin cortical granules. Moreover, temperature-sensitive alleles of sla2 frequently have multinucleate cells with altered morphology (a phenotype we do not see in our cells). The fact that MOP2 (SLA2) shares apparently significant homology with the cytoskeleton-associated protein, talin, in mammalian cells (Reed et al., 1990) provides a satisfying analogy, although we have shown that a deletion of the last 63 amino acids including about of this homology has no effect on the activity of MOP2 (SLA2).

Part of the difficulty in understanding how mop2 mutant phenotypes relate to sla2 mutant phenotypes comes from the fact that the strains used to analyze the two sets of mutations are quite different, and, in fact, a deletion of the MOP2 (SLA2) gene in our strains is lethal at any temperature, while an identical deletion in the strains used to isolate sla2 leads only to very delayed germination, slow growth, and temperature-sensitive conditional lethality. The genetic difference in our strains has not been elucidated.

One way to understand the effect of sla2 mutants on Pma1 abundance comes from the observation in mammalian cells of another member of the cation ATPase superfamily. The mammalian Na/K-ATPase is found asymmetrically distributed in polarized epithelia and that this distribution involves associations with a number of membrane cytoskeleton proteins including ankyrin and fodrin (Marrs et al., 1993; Hammerton et al., 1991; Nelson and Veshnock, 1987). In this view, an alteration in the distribution and disruption of the yeast cytoskeleton would alter sites where Pma1 would normally be located in the cell membrane. Given previous evidence that new secretory vesicle insertions into the plasma membrane occur predominantly in the growing yeast cell bud (Holcomb et al., 1988; Brada and Schekman, 1988), it is understandable that a mutation that altered this normal polarity would reduce the abundance of Pma1 in the plasma membrane. Similarly, defects in the cytoskeleton might prevent normal endocytosis.

Although we can demonstrate an interaction between Mop2 and Pma1 in several ways, it is not yet clear that all of the properties of mop2 mutants can be explained by its role in regulating the abundance of Pma1. mop2 also exhibits hypersensitivity to cycloheximide. This might imply that mop2 might also affect other membrane properties. Cycloheximide hypersensitivity cannot simply be explained as the lowering of membrane potential by affecting Pma1 abundance because pma1 mutants, even those such as pma1-155 that decrease the abundance of wild type protein, do not share this phenotype (Perlin et al., 1989).

How Direct Is the Interaction between Pma1 and Mop2?

Although Mop2 and Pma1 apparently do not form a 1:1 stoichiometric interaction, our genetic and physiological studies have indicated a very intimate relationship between Mop2 and Pma1. If Mop2 is required for H-ATPase transport to plasma membrane, an interesting question is why does Pma1 need a unique factor for its functional expression on plasma membrane? There are many examples where membrane enzyme assembly and transport need another particular protein. Both the assembly of yeast vacuolar H-ATPase subunits into the vacuolar membrane (Hirata et al., 1993) and the expression of mitochondrial ATPase require the participation of proteins that are not part of the functional enzyme (Ackerman et al., 1990a, 1990b; Bowman et al., 1991; Ackerman et al., 1992).

The yeast Pma1 is a major plasma membrane protein accounting for 15% of total plasma membrane protein. The abundance of Pma1 is quite constant during the changes in growth conditions or after increasing PMA1 gene dosage; in fact, over-production of the PMA1 slows yeast cell growth (Eraso et al., 1987). There may be structural constraints against increasing the amount of H-ATPase without affecting the integrity of the membrane. Another possibility is that H-ATPase is a major ATP consumer, and its over-production may compromise the energy charge of the cell (Serrano, 1980). Therefore, it is possible the yeast cells need to regulate the abundance of H-ATPase in the plasma membrane. Mop2 appears to be one of the components that controls H-ATPase abundance.


FOOTNOTES

*
This work was supported by National Institutes of Health Grant GM39739 (to J. E. H.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Current address: Pfeizer Corp., Groton, CT 06340.

A major contributor to this paper.

**
A National Institutes of Health Macromolecular Structure and Mechanism pre-doctoral trainee (Training Grant GM07596).

§§
Current address: Dept. of Biochemistry, Stanford University, Stanford, CA 94305.

¶¶
To whom correspondence should be addressed. Tel.: 617-736-2462; Fax: 617-736-2405; haber{at}hydra.rose.brandeis.edu.

(^1)
H. Riezman, personal communication.

(^2)
The abbreviations used are: kb, kilobase(s); PAGE, polyacrylamide gel electrophoresis.

(^3)
H. Riezman, personal communication.


ACKNOWLEDGEMENTS

We thank Sandra Harris for many yeast strains, providing c-Myc monoclonal antibody 9E10, and many suggestions for the experiments, John Teem for providing a monoclonal anti-H-ATPase antibody, and Ed Louis for providing a chromosome separating blot. Xiaochun Zhu carried out the measurements of the effect of mop2 on Pma1 activity. We also thank David Elliott for instructions and suggestions for the immunofluorescence experiments. Discussions with both Howard Reizman and David Drubin are gratefully acknowledged.

Note Added in Proof-Recently, we have found that under some physiological conditions, overexpression of PMA1 can suppress the temperature sensitivity of mop2-2 (M. Hincapie and J. E. Haber, unpublished results).


REFERENCES

  1. Ackerman, S. H., Gatti, L., Gellefors, P., Douglas M. G., and Tzagoloff, A. (1990a) Proc. Natl. Acad. Sci. U. S. A. 87, 4986-4990 [Abstract]
  2. Ackerman, S. H., and Tzagoloff, A. (1990b) J. Biol. Chem. 265, 9952-9959 [Abstract/Free Full Text]
  3. Ackerman, S. H., Martin, J., and Tzagoloff, A. (1992) J. Biol. Chem. 267, 7386-7394 [Abstract/Free Full Text]
  4. Altschul, S. F., Gish, W., Miller, W., Myers, E. W., and Lipman, D. J. (1990) J. Mol. Biol. 215, 403-410 [CrossRef][Medline] [Order article via Infotrieve]
  5. Benito, B., Moreno, E., and Lagumas, R. (1991) Biochim. Biophys. Acta 1063, 265-268 [Medline] [Order article via Infotrieve]
  6. Bowman, B. J., and Bowman, E. J. (1986) J. Membr. Biol. 94, 83-97 [Medline] [Order article via Infotrieve]
  7. Bowman, B. J., Blasco, F., and Slayman, C. W. (1981) J. Biol. Chem. 256, 12343-12349 [Abstract/Free Full Text]
  8. Bowman, S., Ackerman, S. H., Griffiths, D. E., and Tzagoloff, A. (1991) J. Biol. Chem. 266, 7517-7523 [Abstract/Free Full Text]
  9. Brada, D., and Schekman, R. (1988) J. Bacteriol. 170, 2775-2783 [Medline] [Order article via Infotrieve]
  10. Brandl, C. J., Green, N. M., Korczak, B., and MacLennan, D. H. (1986) Cell 44, 597-607 [Medline] [Order article via Infotrieve]
  11. Carafoli, E. (1992) J. Biol. Chem. 267, 2115-2118 [Free Full Text]
  12. Caroni, P., and Carafoli, E. (1981) J. Biol. Chem. 256, 9371-9373 [Abstract/Free Full Text]
  13. Chang, A., and Slayman, C. W. (1991) J. Cell Biol. 115, 289-295 [Abstract]
  14. Chang, A., Rose, M. D., and Slayman, C. W. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 5808-5812 [Abstract]
  15. Cid, A., Perona, R., and Serrano, R. (1987) Curr. Genet. 12, 105-110 [Medline] [Order article via Infotrieve]
  16. Davis, L. I., and Fink, G. R. (1990) Cell 61, 965-978 [Medline] [Order article via Infotrieve]
  17. Eraso, P., Cid, A., and Serrano, R. (1987) FEBS Lett. 224, 193-197 [CrossRef][Medline] [Order article via Infotrieve]
  18. Geering, K. (1990) J. Membr. Biol. 115, 109-121 [Medline] [Order article via Infotrieve]
  19. Geering, K., Kraehenbuhl, J. P., and Rossier, B. C. (1987) J. Cell Biol. 105, 2613-2619 [Abstract]
  20. Goffeau, A., and Slayman, C. W. (1981) Biochim. Biophys. Acta 639, 197-223 [Medline] [Order article via Infotrieve]
  21. Goffeau, A., and Dufour, J. P. (1988) Methods Enzymol. 157, 528-533 [Medline] [Order article via Infotrieve]
  22. Hammerton, R. W., Krzeminski, K. A., Mays, R. W., Ryan, T. A., Wollner, D. A., and Nelson, W. J. (1991) Science 254, 847-850 [Medline] [Order article via Infotrieve]
  23. Harris, L. S., Na, S., Zhu, X., Seto-Young, D., Perlin, D. S., Teem, J. L., and Haber, J. E. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 10531-10535 [Abstract/Free Full Text]
  24. Hirata, R., Umemoto, N., Ho, M. N., Ohya, Y., Stevens, T. H., and Anraku, Y. (1993) J. Biol. Chem. 268, 961-967 [Abstract/Free Full Text]
  25. Holcomb, C. L., Hansen, W. J., Etcheverry, T., and Sheckman, R. (1988) J. Cell Biol. 106, 641-648 [Abstract]
  26. Holtzman, D. A., Yang, S. and Drubin, D. G. (1993) J. Cell Biol. 122, 635-644 [Abstract]
  27. Horowitz, B., Eakle, K. A., Scheiner-Bobis, G., Randolph, G. R., Chen, C. Y., Hitzeman, R. A., and Farley, R. A. (1990) J. Biol. Chem. 265, 4189-4192 [Abstract/Free Full Text]
  28. Ito, H., Fufuda, Y., Murata, K., and Kimura, A. (1983) J. Bacteriol. 153, 163-168 [Medline] [Order article via Infotrieve]
  29. Jaunin, P., Horisberger, J. D., Richter, K., Good, P. J., Rossier, B. C., and Geering, K. (1992) J. Biol. Chem. 267, 577-585 [Abstract/Free Full Text]
  30. Kuchler, K., Dohlman, H. G., and Thoner, J. (1993.) J. Cell Biol. 120, 1203-1215
  31. Lemas, M. V., Takeyasu, K., and Fambrough, D. M. (1992) J. Biol. Chem. 267, 20987-20991 [Abstract/Free Full Text]
  32. Lingrel, J. B, Orlowski, J. Shull, M. M., and Price, E. M. (1988) Prog. Nucleic Acid Res. Mol. Biol. 38, 37-89
  33. Ljungdahl, P. O., Gimeno, C. J., Styles, C. A., and Fink, G. R. (1992) Cell 71, 463-478 [Medline] [Order article via Infotrieve]
  34. MacLennan, D. H., Brandl, C. J., Korczak, B., and Green, N. M. (1987) Soc. Gen. Physiol. Ser. 41, 287-300 [Medline] [Order article via Infotrieve]
  35. Marrs, J. A., Napolitano, E. W., Murphey-Erdosh, C., Mays, R. W., Reichard, L. F., and Nelson, W. J. (1993) J. Cell Biol. 123, 149-161 [Abstract]
  36. McCusker, J. H. (1986) Ph.D. thesis, Brandeis University
  37. McCusker, J. H., and Haber, J. E. (1988) Genetics 119, 317-327 [Abstract/Free Full Text]
  38. McCusker, J. H., Perlin, D. S., and Haber, J. E. (1987) Mol. Cell. Biol. 7, 4082-4088 [Medline] [Order article via Infotrieve]
  39. Mortimer, R. K., Contopoulou, C. R., and King, J. S. (1992) Yeast 8, 817-902 [Medline] [Order article via Infotrieve]
  40. Nakamoto, R. K., Rao, R., and Slayman, C. W. (1991) J. Biol. Chem. 266, 7940-7949 [Abstract/Free Full Text]
  41. Nelson, W. J., and Veshnock, P. J. (1987) Nature 328, 533-536 [CrossRef][Medline] [Order article via Infotrieve]
  42. Noguchi, S., Mishina, M., Kawamura, M. M., and Numa, S. (1987) FEBS Lett. 225, 27-32 [CrossRef][Medline] [Order article via Infotrieve]
  43. Palmgren, M. G., Sommarin, M., Serrano, R., and Larsson, C. (1991) J. Biol. Chem. 266, 20470-20475 [Abstract/Free Full Text]
  44. Panaretou, B., and Piper, P. W. (1992) Eur. J. Biochem. 206, 635-640 [Abstract]
  45. Perlin, D. S., Harris, S. L., Seto-Young, D., and Haber, J. E. (1989) J. Biol. Chem. 264, 21857-21864 [Abstract/Free Full Text]
  46. Portillo, F., DeLarrinoa, I. F., and Serrano, R. (1989) FEBS Lett. 247, 381-386 [CrossRef][Medline] [Order article via Infotrieve]
  47. Raths, S., Roher, J., Crausaz, F., and Riezman, H. (1993) J. Cell Biol. 120, 55-65 [Abstract]
  48. Reed, D. J. G., Ades, S. E., Singer, S. J., and Hynes, R. O. (1990) Nature 347, 685-689 [CrossRef][Medline] [Order article via Infotrieve]
  49. Sanger, F., Nicklen, S., and Coulson, A. R. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 5463-5467 [Abstract]
  50. Schiestl, R. H., and Gietz, R. D. (1989) Curr. Genet. 16, 339-346 [Medline] [Order article via Infotrieve]
  51. Serrano, R. (1980) Eur. J. Biochem. 105, 419-424 [Abstract]
  52. Serrano, R. (1983) FEBS Lett. 156, 11-14 [CrossRef][Medline] [Order article via Infotrieve]
  53. Serrano, R. (1988) Biochim. Biophys. Acta 947, 1-28 [Medline] [Order article via Infotrieve]
  54. Serrano, R. (1991) Molecular Biology of the Yeast Saccharomyces (Broach, J. R., Pringle, J., and Jones, E. W., eds) pp. 523-586, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
  55. Serrano, R., Kielland-Brandt, M. C., and Fink, G. R. (1986) Nature 319, 689-693 [Medline] [Order article via Infotrieve]
  56. Serrano, R., Montesinod, C. C., Roldan, M., Garrido, G., Ferguson, C., Leonard, K., Monk, B. C., Perlin, D. S., and Weiler, E. W. (1991) Biochim. Biophys. Acta 1062, 157-164 [Medline] [Order article via Infotrieve]
  57. Shull, G. E., Schwartz, A., and Lingrel, J. B (1985) Nature 316, 691-695 [Medline] [Order article via Infotrieve]
  58. Slayman, C. L. (1986) J. Bioenerg. Biomembr. 19, 1-20
  59. Supply, P., Wach, A., Thines-Sempoux, D., and Goffeau, A. (1993) J. Biol. Chem. 268, 19744-19752 [Abstract/Free Full Text]
  60. Takeyasu, K., and Kawakami, K. (1989) Seikagaku 61, 394-401 [Medline] [Order article via Infotrieve]
  61. Takeyasu, K., Tamkun, M. M., Renaud, K. J., and Fambrough, D. M. (1988) J. Biol. Chem. 263, 4347-4354 [Abstract/Free Full Text]
  62. Vai, M., Poplo, L., and Alberghina, L. (1986) FEBS Lett. 206, 135-141 [CrossRef][Medline] [Order article via Infotrieve]
  63. Villalba, J. M., Palmgren, M. G., Berberian, G. E., Ferguson, C., and Serrano, R. (1992) J. Biol. Chem. 267, 12341-12349 [Abstract/Free Full Text]
  64. Volland, C., Urban-Grimal, D., Geraud, G., and Haguenauer-Tsapis, R. (1994) J. Biol. Chem. 269, 9833-9841 [Abstract/Free Full Text]
  65. Wuytack, F., and Racemackers, L. (1992) J. Bioenerg. Biomembr. 24, 285-300 [Medline] [Order article via Infotrieve]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.