JBC PeproTech; Our Business is Cytokines!

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


     

Originally published In Press as doi:10.1074/jbc.M601818200 on June 2, 2006

J. Biol. Chem., Vol. 281, Issue 33, 23887-23898, August 18, 2006
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
281/33/23887    most recent
M601818200v1
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 Google Scholar
Google Scholar
Right arrow Articles by Mason, A. B.
Right arrow Articles by Slayman, C. W.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Mason, A. B.
Right arrow Articles by Slayman, C. W.
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?

Effects of C-terminal Truncations on Trafficking of the Yeast Plasma Membrane H+-ATPase*

A. Brett Mason{ddagger}, Kenneth E. Allen{ddagger}, and Carolyn W. Slayman{ddagger}§1

From the Departments of {ddagger}Genetics and §Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, Connecticut 06510

Received for publication, February 24, 2006 , and in revised form, April 19, 2006.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Within the large family of P-type cation-transporting ATPases, members differ in the number of C-terminal transmembrane helices, ranging from two in Cu2+-ATPases to six in H+-, Na+,K+-, Mg2+-, and Ca2+-ATPases. In this study, yeast Pma1 H+-ATPase has served as a model to examine the role of the C-terminal membrane domain in ATPase stability and targeting to the plasma membrane. Successive truncations were constructed from the middle of the major cytoplasmic loop to the middle of the extended cytoplasmic tail, adding back the C-terminal membrane-spanning helices one at a time. When the resulting constructs were expressed transiently in yeast, there was a steady increase in half-life from 70 min in Pma1{Delta}452 to 348 min in Pma1{Delta}901, but even the longest construct was considerably less stable than wild-type ATPase (t1/2 = 11 h). Confocal immunofluorescence microscopy showed that 11 of 12 constructs were arrested in the endoplasmic reticulum and degraded in the proteasome. The only truncated ATPase that escaped the ER, Pma1{Delta}901, traveled slowly to the plasma membrane, where it hydrolyzed ATP and supported growth. Limited trypsinolysis showed Pma1{Delta}901 to be misfolded, however, resulting in premature delivery to the vacuole for degradation. As model substrates, this series of truncations affirms the importance of the entire C-terminal domain to yeast H+-ATPase biogenesis and defines a sequence element of 20 amino acids in the carboxyl tail that is critical to ER escape and trafficking to the plasma membrane.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The single most abundant protein in the yeast plasma membrane is a 100-kDa H+-ATPase that accounts for at least 10% of total plasma membrane protein (1). Encoded by the PMA1 gene (2) and belonging to the widely distributed family of P-type cation-transporting ATPases (3-5), the yeast H+-ATPase acts physiologically to pump protons out of the cell, creating a transmembrane potential that drives the uptake of amino acids, sugars, and other nutrients (1). Because of its abundance, the H+-ATPase has attracted considerable attention over the past two decades, both for mechanistic studies of ATP-coupled proton transport and, more recently, as a model system to explore the biogenesis of polytopic membrane proteins.

A particularly fruitful approach to biogenesis studies has come from the use of site-directed mutagenesis to probe structure-function relationships in key regions of the H+-ATPase polypeptide (1). Of the mutations that have been made to date, a significant subset cause partial or complete blocks in ATPase biogenesis, giving direct evidence for the critical nature of proper protein folding. Some of the mutant forms are severely defective and become arrested in the endoplasmic reticulum, from where they are shipped to the proteasome for degradation (e.g. see Refs. 6-8). Others, with more subtle abnormalities in folding, are transported further along the secretory pathway to reach the Golgi (pma1-7, Ref. 9) or even the plasma membrane (pma1-10, Ref. 10 and pma1G381A, Ref. 11), before being recognized as abnormal and committed to the endosomal/vacuolar pathway for degradation. In addition to their use in mapping amino acid residues within the Pma1 polypeptide that are important for folding and trafficking, both kinds of mutant have been used to identify novel components of the yeast secretory pathway, such as Eps1p, a protein-disulfide isomerase implicated in ER quality control (7), and Ast1p, a protein involved in transport from the Golgi to the plasma membrane (9).

The work to be described in this paper has taken a systematic look at features of the yeast H+-ATPase structure and topology that are required for successful delivery to the plasma membrane. We have focused in particular on the C-terminal portion of the molecule, given the variability of this region during the evolution of P-type ATPases (12). On the one hand, members of the P2 subfamily, including mammalian Na+,K+- and Ca2+-ATPases as well as the fungal and plant H+-ATPases, have six C-terminal transmembrane helices, designated M5 through M10 (3). At the other extreme, heavy metal ATPases of the P1 family have only two C-terminal helices, whereas the P3-type Kdp ATPase of Escherichia coli is intermediate between the first two groups, with four such helices.

To dissect the role of the C-terminal portion of yeast Pma1 ATPase in biogenesis and trafficking, we have constructed a set of mutants with truncations after each of the last six transmembrane segments, from M5 through M10. Whereas the results reveal a clear correlation between protein length and intracellular stability, all but one of the mutant ATPases fails to exit the endoplasmic reticulum. Only Pma1{Delta}901, truncated midway along the cytoplasmic C-terminal tail, is successfully targeted to the plasma membrane, where it is capable of splitting ATP and supporting growth. Thus, all six C-terminal membrane helices and at least a portion of the C terminus are essential for proper folding and trafficking of the H+-ATPase through the secretory pathway.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Yeast Strains and Growth Conditions—Saccharomyces cerevisiae strains used in this study are listed in Table 1. For transient expression experiments, the mutant pma1 allele to be studied was introduced on either a centromeric or integrating plasmid into yeast strains with the appropriate mutant background (BMY40, W303-1B pep4{Delta}, WG4a 11/22a) and in which the chromosomal copy of PMA1 had been placed under control of the GAL1 promoter (see Table 1). Transformants were selected on 2% galactose in complete synthetic medium lacking uracil and the relevant auxotrophic amino acid(s) (Bio 101 Inc., Carlsbad, CA). For the transient expression of hemagglutinin HA2-tagged pma1 alleles, chromosomal GAL1p-PMA1 expression was first turned off by incubating the cells for 3 h in medium containing 2% glucose instead of galactose. Expression of the HA-tagged Pma1 mutant was then induced by incubating the cells at 39 °C for the appropriate time (15).


View this table:
[in this window]
[in a new window]
  		TABLE 1
Yeast strains used in this study

 

For the stable expression of chromosomally integrated pma1 alleles, yeast strain NY13 was used to make strain BMY58, a uracil auxotroph in which PMA2 is disrupted with the kanamycin resistance cassette (pma2::KanMX4).

Standard yeast media and genetic manipulations were as described by Sherman et al. (16). Yeast transformations were performed as described (17) using the Alkali-Cation Yeast Transformation kit (Bio 101).

Plasmids—Truncated pma1{Delta} alleles were made in plasmid YCplac22-2HSEpr-HA-PMA1-MluI, a derivative of YCplac22-2HSEpr-HA-PMA1 (11) containing an MluI site at the TAA stop codon of the PMA1 gene. Another MluI site was built into mutagenic oligonucleotides used to introduce a second TAA stop codon at the desired location within the PMA1 coding sequence in the plasmid template. Intervening DNA sequences were then removed by digestion with MluI, and the construct encoding a truncated ATPase product, with 3'-flanking sequence intact, was generated by circularization of the purified vector DNA fragment. The same strategy was used to create 2HSEpr-HA-pma1 constructs in the integrative vector pRS303-2HSE (11) for targeting to the chromosomal HIS3 locus. Plasmid-encoded wild-type and mutant alleles of PMA1 used in this study were tagged with a single copy of the 9-amino acid HA epitope (YPYDVPDYA), inserted in frame between codons 2 and 3 of the PMA1 coding sequence (6). The QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA) was used for in vitro mutagenesis, and all mutant constructs were verified by DNA sequencing.

For the chromosomal integration of mutant pma1 alleles by homologous recombination at the PMA1 locus, plasmid pGW201 (18) was modified by adding two unique restriction sites: an NcoI site at the PMA1 ATG start codon and an MluI site at the PMA1 TAA stop codon. These two new restriction sites were used to introduce an HA-tagged copy of the PMA1 gene, producing plasmid pBM432. The pma1{Delta}901 allele was constructed in pBM432 by replacement of the 1.1-kb BamHI-MluI fragment with the corresponding fragment from YCplac22-2HSE-HA-pma1{Delta}901. A 6-kb HindIII fragment from this construct was used to transform yeast strain BMY58 (Table 1). Transformants selected on complete synthetic medium lacking uracil were screened by immunoblotting for expression of truncated ATPase Pma1{Delta}901.

Immunofluorescence—Intracellular structures containing HA-tagged ATPase were visualized by confocal microscopy as described previously (6). Cells were fixed, treated with Zymolyase T-100, permeabilized with 0.1% Triton X-100, and stained for immunofluorescence (11). Primary antibodies included rabbit anti-HA polyclonal (Medical and Biological Laboratories Co., Ltd., Nagoya, Japan), diluted 1:100; anti-HA.11 monoclonal 16B12 (Babco, Richmond, CA), diluted 1:150; anti-vacuolar H+-ATPase 60-kDa subunit monoclonal 13D11-B2 from raw ascites fluid (Molecular Probes, Inc., Eugene, OR), diluted 1:150; and rabbit anti-Kar2p polyclonal antibody (provided by M. Rose), diluted 1:2500. Goat anti-rabbit and goat anti-mouse IgGs conjugated, respectively, to FITC and Texas Red (Jackson Immunoresearch, West Grove, PA), served as fluorescent secondary antibodies and were used at 1:100 dilutions. For double labeling experiments, both primary antibodies were present during the initial incubation, and both secondary antibodies were present during the subsequent incubation. To control for spurious antibody cross-reactivity, non-induced cells were simultaneously fixed and stained with each set of antibodies. Samples were then mounted in Citifluor (Ted Pella, Redding, PA).

Cells were observed under a Zeiss L510 scanning confocal microscope using dual channel filters for simultaneous visualization of FITC and Texas Red fluorochromes. All images were taken with a 63 x 1.4 numerical aperture Plan-Apochromat III differential interference contrast objective (Zeiss). In time course experiments, microscope settings were unchanged throughout the experiment in order to obtain a semi-quantitative signal. Cross-talk between FITC and Texas Red was avoided through the use of the Zeiss L510 digital signal processor. The absence of bleed-through was confirmed by checking that the signal disappeared when viewed with single-wave-length filter blocks. Images were collected with LSM5 software (Zeiss) and modified by contrast stretching and merging, using Adobe Photoshop 7.0 (Adobe Systems Inc., San Jose, CA).

Total Protein Extracts and Immunoblotting—Total protein extracts were prepared as described by Volland et al. (19). Log phase cells (A600 nm = 1.5) were suspended in 0.5 ml of water and then lysed by adding 50 µl of 1.85 M NaOH, 5% (v/v) beta-mercaptoethanol. Proteins were precipitated by adding 50 µl of 50% (m/v) trichloroacetic acid. The resulting pellets were resuspended in 15 µl of 1 M Trizma Base and 30 µl of 3x Laemmli SDS-PAGE loading buffer (20), resolved on 8% polyacrylamide gels (SDS-PAGE), and transferred to a polyvinylidene difluoride membrane (Immobilon, Millipore Corp., Bedford, MA). HA-tagged ATPases were quantified by immunoblotting with rabbit anti-HA polyclonal antibody diluted 1:4000 (Medical and Biological Laboratories, Nagoya, Japan) and Kar2p by immunoblotting with rabbit anti-Kar2p polyclonal antibody diluted 1:20,000 (provided by M. Rose), followed by incubation with 125I-protein A, autoradiography, and phosphorimaging analysis (Storm 840; Amersham Biosciences). Alternatively, blots were developed using horseradish peroxidase-conjugated secondary antibodies and the ECL chemiluminescent detection system (Amersham Biosciences). For quantitative immunoblotting of the HA-tagged ATPase in sucrose gradient fractions (see below), proteins were trichloroacetic acid-precipitated from 100-µl samples. Dolichol phosphate mannose synthase (Dpm1p) was detected on immunoblots using monoclonal antibody 5C5 (Molecular Probes, Inc., Eugene, OR).

Subcellular Fractionation—Sixty-ml cultures were grown to an A600 nm of ~1.0, shifted to glucose for 3 h, and then induced at 39 °C for 15 min as detailed above (see "Yeast Strains and Growth Conditions"). Samples equivalent to 25 A600 nm units were taken immediately before the temperature shift (t = 0) and at specified time points after induction. Sodium azide was added immediately to a final concentration of 10 mM, and cells were then centrifuged at 2000 x g for 5 min prior to resuspension in STE10 buffer (10% sucrose in 10 mM Tris, pH 7.5, 10 mM EDTA) supplemented with a protease inhibitor mixture (leupeptin, pepstatin, aprotinin, and chymostatin, at 2 µg/ml each, plus phenylmethylsulfonyl fluoride to 1 mM final concentration). Cells were broken by vortexing with 0.45-mm glass beads (4 x 1-min pulses). Lysates were recovered, and the remaining beads were washed with 400 µl of STE10 plus protease inhibitors. Washes were combined with primary lysates, and these suspensions were centrifuged at 3000 x g for 5 min. Supernatants were applied to 5-ml 20-60% linear sucrose gradients, ultracentrifuged in an SW55Ti rotor for 16 h at 100,000 x g, and the gradients fractionated into 300-µl volumes.

Limited Trypsinolysis of Gradient-purified Plasma Membranes—Plasma membranes were prepared as described (21). Trypsinolysis was performed according to Nakamoto et al. (15) using trypsin/protein ratios as stated.

ATP Hydrolysis and Protein Determination—Activity of the plasma membrane H+-ATPase was assayed at 30 °C using a standard (21) or microscale ATPase assay (22).

Protein concentrations were measured as described (23) or as modified by Ambesi et al. (24), using bovine serum albumin as a standard.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Design and Expression of Truncated ATPases—Fig. 1 illustrates the topology of yeast Pma1 H+-ATPase, which is embedded in the lipid bilayer by four transmembrane helices (M1-M4) toward the amino terminus of the protein and six (M5-M10) toward the C terminus (25). The starting point for this study was a truncation at Glu901, midway along the C-terminal tail. This modest deletion of 18 amino acids was studied previously and was shown to allow both measurable ATPase activity and growth (26). To judge the effects of longer C-terminal deletions on ATPase biogenesis and stability, stepwise terminations were introduced at eight additional points upstream of Glu901 (Fig. 1). These were as follows: Ala881 (immediately following M10); Tyr873 (in the middle of M10); Asp851, Ile820, Ile785, Pro753, and Asp718 (in the small loops following M9, M8, M7, M6, and M5, respectively); and His686 (at the C-terminal end of the large cytoplasmic loop). The final and most amino-terminal truncation was at residue Asp452, near the middle of the large cytoplasmic loop and within the N (nucleotide-binding) domain of the ATPase (25). This mutation occurred spontaneously and was included in the series after preliminary studies suggested that it behaved distinctively when expressed in yeast.

Mutant pma1{Delta} constructs tagged with a single HA epitope at the amino terminus were introduced into the yeast centromeric expression plasmid YCplac22-2HSE-HA-PMA1-MluI, placing them under control of two heat-activated promoter elements (2HSE). They were then transformed into yeast strain BMY40, in which the chromosomal copy of the PMA1 gene is under control of the GAL1 promoter (11). For expression studies, cells were grown in galactose medium at 23 °C (wild-type allele ON; mutant allele OFF) and then shifted to glucose medium at 23 °C for 3 h to turn off expression of the chromosomal PMA1 gene and empty the secretory pathway of wild-type ATPase. Cells were then incubated for 15 min at 39 °C to induce a short pulse of synthesis of mutant ATPase before being returned to 23 °C, with the HA tag providing a convenient way to track the synthesis, stability, and intracellular location of the mutant protein (11).


Figure 1
View larger version (33K):
[in this window]
[in a new window]
  		FIGURE 1.
Truncations of Pma1 H+-ATPase. A topological model shows the transmembrane disposition of the ATPase and the residues at which truncations were made by introducing TAA stop codons (crossed circles). Nucleotide binding (ATP) and phosphorylation (P) sites are indicated along with the position of a single HA sequence tag at the NH2 terminus.

 


Figure 2
View larger version (18K):
[in this window]
[in a new window]
  		FIGURE 2.
Stability of transiently expressed wild-type and truncated ATPases. Cultures were shifted to 39 °C at t = 0 min to induce expression of plasmid-encoded, HA-tagged ATPases and then returned to 23 °C at t = 15 min. Total protein extracts (see "Experimental Procedures") were separated by SDS-PAGE, immunoblotted using polyclonal anti-HA, and detected by 125I-protein A and autoradiography. Each point represents the average of at least three determinations. Error bars have been omitted for clarity, but S.E. values were typically less than 20%. Solid squares, wild type; open inverted triangles, Pma1{Delta}452; open circles, Pma1{Delta}686; open triangles, Pma1{Delta}753; open squares, Pma1{Delta}820; solid triangles, Pma1{Delta}881; solid circles, Pma1{Delta}901.

 
Stability of Truncated ATPase as a Function of Protein Length—The first step was to examine the stability of the truncated ATPases. For this purpose, total protein extracts were made from cells taken immediately before (t = 0 min) and at various times after expression had been induced at 39 °C. Quantitative immunoblotting with anti-HA antibody revealed a striking relationship between the length of the mutant ATPase and its half-life (Fig. 2 and Table 2). The shortest polypeptide (Pma1{Delta}452) was by far the least stable, with a t1/2 of only ~70 min. Adding back the remainder of the large cytoplasmic loop (in Pma1{Delta}686) increased the t1/2 to 116 min, and the successive addition of transmembrane helices M5-M10 led to further stepwise increases from 152 to 288 min. Pma1{Delta}901, which retained part of the C-terminal tail, had the longest half-life (~348 min), although still only about one-half that of wild-type Pma1 ATPase (~11 h) (27). Thus, virtually the entire 100-kDa polypeptide chain is required for maximal stability, with critical contributions coming from all six transmembrane helices downstream of the major cytoplasmic domain and even from the C-terminal tail.


View this table:
[in this window]
[in a new window]
  		TABLE 2
Stability of truncated H+-ATPase mutants

 
Trafficking of Truncated ATPases—To learn more about the processing of the truncated ATPases, their intracellular trafficking was monitored by confocal immunofluorescence microscopy (Fig. 3). In keeping with the results of earlier studies (11), HA-tagged wild-type ATPase could be detected as early as 15 min after induction of expression, and it was then exported efficiently through the secretory pathway to reach the plasma membrane by 60 min post-induction. The shortest truncation product, Pma1{Delta}452, was trapped in a predominantly perinuclear location and degraded rapidly, becoming undetectable after 60 min. By contrast, the next shortest truncation product (Pma1{Delta}686) gave a punctate staining pattern that persisted throughout the 180-min time course. Similar patterns were seen for Pma1{Delta}718 through Pma1{Delta}881, which also failed to reach the cell surface but remained visible for at least 180 min; confocal images for three of these mutants are included in Fig. 3. Of the mutants that were studied, only Pma1{Delta}901 was transported to the plasma membrane, with a trafficking behavior very similar to that of the wild-type ATPase.


Figure 3
View larger version (67K):
[in this window]
[in a new window]
  		FIGURE 3.
Expression and trafficking of wild-type (WT) and Pma1{Delta} ATPases. Cell samples (total A600 nm ~5.0 units) were taken from the cultures described in the legend to Fig. 2. Cells were fixed and processed for immunofluorescent confocal microscopy using monoclonal anti-HA and Texas Red-conjugated goat anti-mouse IgG to identify HA-tagged ATPase as described (see "Experimental Procedures"). Confocal microscope settings were unchanged between time points in order to obtain a semiquantitative signal. Bar,5 µm.

 
The punctate staining patterns seen in Fig. 3 resemble ER-derived tubulo-vesicular structures that have been reported previously for misfolded full-length Pma1p mutants (7, 11, 28) as well as for trafficking-defective mutants of other plasma membrane proteins (29, 30). It therefore seemed likely that mutants Pma1{Delta}452 through Pma1{Delta}881 were arrested in the ER. Indeed, as illustrated in Fig. 4, these HA-tagged proteins co-localized strongly with the molecular chaperone Kar2p, a classical marker of the ER lumen (31). By comparison, wild-type and {Delta}901 ATPases could be seen at the cell surface, clearly distinct from the Kar2p marker.

Collectively, the results from confocal microscopy made it possible to divide the truncated ATPases into three distinct groups based on their trafficking patterns. The single mutant in Group I (Pma1{Delta}452) was unique in its perinuclear ER distribution with peripheral extensions. Eight mutants in Group II (Pma1{Delta}686 through Pma1{Delta}881) were sequestered in discrete, punctate, ER-associated bodies; and Pma1{Delta}901, the only example of Group III, was able to exit the ER and travel to the plasma membrane.

Truncated ATPases in Groups I and II Are Degraded in the Proteasome—Immunoblotting and immunofluorescence results indicated that the aggregates formed by the ER-arrested ATPases (Groups I and II) were gradually cleared from the cell, most likely by ER-associated degradation in the proteasome. To test this idea directly, the truncated ATPases were expressed in a pre1-1 pre2-2 yeast strain deficient in chymotrypsin-like activity of the proteasome (WG4a-11/22) (Table 1) (32). In the experiment of Fig. 5, total protein samples were taken both before induction at 39 °C and at intervals after induction, and assayed for HA-tagged ATPase by quantitative immunoblotting. Results are compared with control measurements carried out in the proteasomally active parent strain (WG4a) (Table 1).


Figure 4
View larger version (40K):
[in this window]
[in a new window]
  		FIGURE 4.
Arrest of group I and group II ATPases in the endoplasmic reticulum. Expression of HA-tagged wild-type (WT) and truncated ATPases was induced, and cell samples were prepared for immunofluorescent labeling as described in the legend to Fig. 3. A combination of monoclonal anti-HA and Texas Red-conjugated goat anti-mouse IgG was used to detect HA-tagged ATPase, whereas polyclonal rabbit anti-Kar2p and FITC-conjugated goat anti-rabbit IgG detected the ER lumen marker Kar2p. Co-localization of ATPase and Kar2p can be seen in the merge panels as a yellow-orange hue. Images of cells expressing wild type, Pma1{Delta}820, and Pma1{Delta}901 were taken 120 min after induction, whereas images of cells expressing ATPases with shorter half-lives (Pma1{Delta}452, Pma1{Delta}686, and Pma1{Delta}753) were taken at 30, 60, and 60 min, respectively. Bar,5 µm.

 
In several cases (wild type, Pma1{Delta}452, and Pma1{Delta}686), ATPase levels in the pre1-1 pre2-2 mutant background were elevated even before induction (Fig. 5). This was probably due to some leakiness of the 2HSE promoter (15), resulting in accumulation of newly synthesized ATPase in the proteasome-defective strain. More important was the finding that all nine truncated ATPases in Groups I and II (Pma1{Delta}452 through Pma1{Delta}881) were stabilized when expressed in the pre1-1 pre2-2 mutant background, consistent with the idea that these mutant polypeptides undergo preferential degradation in the proteasome. This effect was most pronounced for early truncations, such as Pma1{Delta}452, Pma1{Delta}686, and Pma1{Delta}753, which were almost completely degraded by the 180-min time point without pre1-1 pre2-2 protection (Fig. 5); however, some stabilization was also evident for later truncations, such as Pma1{Delta}820 and Pma1{Delta}881.

Two strategies were tested without success in an attempt to override the ER quality control system and enable shipment of truncated ATPases to the plasma membrane. First, the eps1{Delta} mutation was introduced into strain BMY40 to see whether elimination of Eps1p, a member of the protein-disulfide isomerase family, might promote export of ER-arrested Pma1{Delta}p mutants, as has been shown for a severely misfolded full-length Pma1 ATPase mutant (D378N) (7). Our experiments confirmed the published observations for HA-tagged Pma1-D378N but showed no effect of eps1{Delta} on the stability or trafficking of the truncated ATPases. The second, more general strategy, involved the addition of glycerol at concentrations as high as 10% to the galactose growth medium and/or the glucose medium used during induction of mutant ATPase expression. Other researchers have reported that glycerol stabilizes and even promotes the export of certain endogenous integral membrane proteins (33). However, no change was observed in either the ER arrest or instability of the truncated Pma1 ATPases (not shown).

Overall, the results up to this point show that truncations from the middle of the large cytoplasmic loop (Pma1{Delta}452) to the end of M10 (Pma1{Delta}881) create a mutant protein that is recognized as abnormal by the quality control machinery in the ER and degraded, with no discernible ability to be exported further along the secretory pathway.

Pma1{Delta}901 ATPase Is Partially Functional—By contrast with the behavior of truncations Pma1{Delta}452 through Pma1{Delta}881, Pma1{Delta}901 (the longest polypeptide of the present series) was able to reach the plasma membrane (Fig. 3), making it of interest to study under steady-state growth conditions. For this purpose, the mutant allele was inserted in place of the chromosomal copy of the PMA1 gene in a strain (BMY58) carrying a disruption of the closely related PMA2 gene (pma2::KanMX4), creating strain BMY62 (Table 1). PMA2 is not expressed under standard laboratory conditions (34), but it has been shown to undergo gene conversion with certain mutant pma1 alleles under selective growth conditions (28) and could thus have interfered with studies of Pma1{Delta}901 expression.


Figure 5
View larger version (29K):
[in this window]
[in a new window]
  		FIGURE 5.
Group I and Group II ATPases are degraded by the proteasome. HA-tagged wild-type (WT) and mutant ATPases were expressed transiently in either the WG4a parental background (open squares) or the WG4a-11/22a pre1-1 pre2-2 strain (solid squares) using constructs based on plasmid YCplac22-2HSE-HA-Pma1-MluI. Mid-logarithmic phase cells growing at 23 °C in medium containing 2% galactose were shifted to medium containing 2% glucose for 90 min before the amino acid analog, canavanine, was added to a final concentration of 20 µg/ml to further stress the proteasome. After another 90-min incubation at 23 °C, HA-tagged ATPase expression was induced by shifting cultures to 39 °C for 15 min. Total protein extracts made from cell samples obtained immediately before (t = 0 min) and at intervals after induction were analyzed as described in the legend to Fig. 2.

 
In the presence of glucose (glucose-metabolizing conditions), the integrated pma1{Delta}901 allele supported growth and was expressed in the plasma membrane at 67% of the wild-type level (Table 3). After correction for expression, the {Delta}901 polypeptide could be seen to hydrolyze ATP at almost one-half the rate measured for the wild-type control (3.83 versus 8.09 µmol/min/mg). Carbon starvation revealed contrasting responses between wild-type and Pma1{Delta}901 ATPases, consistent with results reported from studies in a different yeast background (26). Under carbon-starved (CS) conditions, wild-type ATPase displayed a rapid, 4-fold down-regulation of activity from 8.09 to 2.02 µmol/min/mg (Table 3). By contrast, Pma1{Delta}901 was constitutively active, with equal ATPase activities under GM and CS conditions (3.83 and 3.81 µmol/mg/min). The difference between the mutant and wild-type strains provides added support for the widely accepted view that the C-terminal tail of Pma1 H+-ATPase acts as an autoinhibitory domain to down-regulate activity during carbon starvation (35).


View this table:
[in this window]
[in a new window]
  		TABLE 3
H+-ATPase activity in plasma membranes from carbon-starved and glucose-metabolizing cells

 


Figure 6
View larger version (84K):
[in this window]
[in a new window]
  		FIGURE 6.
Pma1{Delta}901 is misfolded. Plasma membranes (5 µg of protein) from strains BMY58 (Pma1p) or BMY62 (Pma1{Delta}901) were digested using a trypsin/protein ratio of 1:5. Aliquots of each digestion mixture were removed at the times indicated, separated by SDS-PAGE, and immunoblotted against a polyclonal rabbit anti-Neurospora crassa Pma1p antibody. WT, wild type.

 
Pma1{Delta}901 ATPase Is Folded Poorly, and Its Passage through the Secretory Pathway Is Slowed—To ask whether structural abnormalities could be detected that would account for the abnormal behavior of Pma1{Delta}901 ATPase, isolated plasma membranes from BMY58 (wild-type Pma1) and BMY62 (Pma1{Delta}901) were analyzed by limited trypsinolysis, which provides a sensitive way to examine the folded structure of yeast H+-ATPase (e.g. see Ref. 36). Pma1{Delta}901 proved to be significantly more sensitive to trypsin than its wild-type counterpart, with the mutant protein being almost completely degraded in 15 min at a trypsin/protein ratio of 1:5 (Fig. 6). Apart from the absence of a small amount of an ~71-kDa digestion product, the pattern and size of trypsin degradation products for Pma1{Delta}901 were almost identical to those of wild-type Pma1p, suggesting that the increased susceptibility of the mutant form to trypsin was due to the exposure of cleavage sites normally masked by the C terminus.

Not surprisingly, further investigation revealed that this structural abnormality was capable of slowing the rate at which Pma1{Delta}901 ATPase travels to the plasma membrane. For these experiments, lysates from cells expressing either wild-type or Pma1{Delta}901 ATPase were prepared 15, 60, or 120 min after induction at 39 °C and analyzed by ultracentrifugation on linear 20-60% sucrose gradients. Fractionation profiles were then analyzed by immunoblotting for the presence of HA-tagged wild-type or Pma1{Delta}901 ATPase (Fig. 7). At the earliest time point (15 min after induction), a substantial difference could be seen between the distributions of the two ATPases. Unlike the wild-type protein, which was spread uniformly throughout the gradient, most of the 15-min Pma1{Delta}901 ATPase was confined to the top half, where it co-fractionated with the ER membrane marker Dpm1p (dolichol phosphate mannose synthase). At 60 min, both ATPases were found predominantly in fractions 9-13, although a smaller peak of Pma1{Delta}901 was still associated with ER-containing fractions. Only at the 120-min time point did the peak of HA-tagged Pma1{Delta}901 co-locate with that of wild-type ATPase (Fig. 7). In all of the gradients, vanadate-sensitive ATPase activity was detected only in fractions 10-15 (not shown), confirming that these more dense fractions contained the active, plasma membrane-bound enzyme preformed prior to induction of tagged ATPase. Any HA-tagged ATPase synthesized during the 15-min induction period would be expected to make a negligible contribution to total measured ATPase activity.

Pma1{Delta}901 Is Degraded in the Vacuole—Although Pma1{Delta}901 displayed the ability to reach the plasma membrane in the present study (Fig. 3), it was much less stable than the wild-type control (Fig. 2). Thus, there was good reason to think that the two ATPases might ultimately be shipped at different rates to the endosomal/vacuolar system for degradation.

To test this idea, wild-type Pma1p, Pma1{Delta}901, and several other truncated ATPases were introduced via centromeric plasmids into strain W303-1B and its derivative W303-1B pep4{Delta}, which lacks vacuolar aspartyl protease activity. Protein profiles for transiently expressed wild-type, Pma1{Delta}686, and Pma1{Delta}901 ATPases are illustrated in Fig. 8A. Pma1{Delta}686, a mutant arrested in the ER, was not protected by the pep4{Delta} mutation. On the other hand, Pma1{Delta}901 ATPase was protected completely for at least 5 h after induction, implicating the vacuole as the primary site of degradation for this ATPase. At the 3-h time point in the pep4{Delta} background, immunofluorescence analysis showed both wild-type and Pma1{Delta}901 H+-ATPases at the plasma membrane (Fig. 8B). Undigested Pma1{Delta}901 could also be seen in intracellular aggregates localized within vacuolar membranes, which were visualized with antibody against the 60-kDa subunit of yeast vacuolar H+-ATPase.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, a series of truncations have been used to examine the role of the C-terminal membrane domain in the stability of yeast Pma1 H+-ATPase and in the delivery of the ATPase to the plasma membrane. Unlike previous work on P-type ATPases, where a typical approach has been to express fusion constructs either in vitro (37) or in heterologous cell backgrounds (38, 39), the transient expression system used in this study has enabled us to track the behavior of the Pma1 H+-ATPase as a non-fusion protein in its native yeast background, thereby avoiding any complications that might have been caused by adding an unrelated protein sequence or by relying upon the membrane-trafficking machinery of an unrelated cell type.


Figure 7
View larger version (18K):
[in this window]
[in a new window]
  		FIGURE 7.
Abnormal trafficking of Pma1{Delta}901 ATPase. BMY401 and BMY4017 expressing HA-tagged WT and Pma1{Delta}901 ATPases, respectively, were harvested at 15, 60, or 120 min post-induction, and the cell lysates were separated on linear (20-60%) sucrose gradients (see "Experimental Procedures"). A sample from each gradient fraction was immunoblotted to detect HA-tagged proteins. Anti-HA signal is shown as follows: wild type (open squares), Pma1{Delta}901 (solid circles). The inset shows chemiluminescent image of immunoblot to detect the ER membrane marker protein Dpm1p. c, control sample of total cell lysate.

 
Contribution of the C-terminal Membrane Domain to Pma1 H+-ATPase Stability—The results of the present study show that stepwise truncations from the C terminus toward the middle of the ATPase lead to significant defects in stability and membrane trafficking. The most drastic change was seen in the shortest product, Pma1{Delta}452, which contained only half of the cytoplasmically exposed N (nucleotide-binding) domain. This polypeptide became arrested in perinuclear ER-derived structures and was shipped to the proteasome for degradation (t1/2 = 70 min). Adding back the rest of the N domain caused the next mutant in the series, Pma1{Delta}686, to be sequestered in a different set of ER-derived structures, from which it underwent slower proteasomal degradation (t1/2 = 116 min). Presumably, the cytoplasmic domain of Pma1{Delta}686 folded more tightly than that of Pma1{Delta}452, masking degradation signals that may be exposed in the shorter construct and slowing delivery to the proteasomal machinery. This idea is supported by the results of an earlier study in which the large cytoplasmic domain of yeast Pma1 H+-ATPase, when expressed by itself as a soluble protein in E. coli, folded well enough to bind TNP-ATP with high affinity (40); similar findings have been reported for the large cytoplasmic domain of mammalian sarcoplasmic reticulum Ca2+-ATPase (41, 42) and Na+,K+-ATPase (43). The slower degradation of Pma1{Delta}686 compared with Pma1{Delta}452 is also consistent with recent findings by Taxis et al. (44) on a series of modular protein substrates. In these constructs, the presence of a well folded cytoplasmic domain increased the time required for clearance of a membrane-anchored protein, due to the need for additional chaperones and accessory proteins to unfold the substrate prior to proteasomal degradation.


Figure 8
View larger version (21K):
[in this window]
[in a new window]
  		FIGURE 8.
Pma1{Delta}901 ATPase is degraded in the vacuole. A, samples were taken from cultures grown and induced as described in the legend to Fig. 2. HA-tagged wild-type (WT) and mutant ATPases were expressed transiently in either the W303-1B parental background (open symbols) or strain W303-1B pep4{Delta} lacking aspartyl protease (proteinase A) activity (solid symbols). Total protein extracts were made at the time points indicated, separated by SDS-PAGE, immunoblotted using polyclonal anti-HA antibodies, and detected by 125I-protein A and autoradiography. Each point represents the average of at least three determinations. Error bars have been omitted for clarity, but S.E. values were typically less than 20%. Top panel, wild type; middle panel, Pma1{Delta}686; bottom panel, Pma1{Delta}901. B, from time course experiments in A, cell volumes equivalent to an A600 nm of ~5.0 units were processed for double label immunofluorescence as described under "Experimental Procedures." In this experiment, monoclonal anti-HA antibody and an FITC-conjugated secondary antibody were used to detect HA-tagged plasma membrane H+-ATPase (left-hand panels), whereas a rabbit antibody against the 60-kDa vacuolar H+-ATPase subunit and a Texas Red-conjugated secondary antibody decorated vacuolar membranes (middle panels). Merged images are shown in the right-hand panels. Images show wild-type (top), Pma1{Delta}686 (middle), and Pma1{Delta}901 (bottom) ATPases at 180 min after induction of expression in strain W303-1B pep4{Delta}. Bar,5 µm.

 
Further improvement in ATPase stability occurred when transmembrane helices M5-M10 were added back in constructs Pma1{Delta}718 through Pma1{Delta}881. Despite the fact that all of these mutant polypeptides were arrested in punctate ER-derived structures and ultimately degraded by the proteasome, they displayed steadily increasing half-lives from 152 to 288 min. Thus, it appears that individual helices from M5 through M10 contribute in a cumulative fashion to the stability of the Pma1 polypeptide. By contrast, a previous study by Béguin et al. (45) pointed to a more complex relationship between C-terminal helices and protein stability in the case of Na+,K+-ATPase. When truncated constructs of the Xenopus laevis {alpha}1 subunit were expressed in a Xenopus oocyte system and assayed by immunoprecipitation, a polypeptide containing only M1-M4 and ~85% of the large cytoplasmic domain was stable, but the addition of M5 and M6 caused it to disappear completely during a 48-h chase, with no sign of residual proteolytic fragments. The authors traced this effect to a pair of Pro residues in the short extracellular loop between M5 and M6 (Pro799-Leu-Pro), which appeared to act as a degradation signal under the conditions of their experiments. (It may be significant that there are no prolines in the M5-M6 loop of Pma1 H+-ATPase.) Further along in the same Na+,K+-ATPase {alpha}1 sequence, Béguin et al. (45) found evidence of a stabilizing effect when constructs containing M7 and M8 were co-expressed with the beta1 subunit, reflecting the association of beta1 with the extracytoplasmic M7-M8 loop. Worth noting in this respect is the fact that yeast Pma1p has no beta subunit and appears to undergo membrane insertion and trafficking without the need for a "helper" protein of this kind. Indeed, yeast Pma1p may be a better model for the vast majority of P-type ATPases that lack auxiliary subunits as well as for other single subunit membrane proteins. Overall, our data showing a gradual increase in stability from Pma1{Delta}718 to Pma1{Delta}881 lend support to the idea advanced by Vashist and Ng (46) that the number of transmembrane segments in an integral membrane protein may influence the time required to extract misfolded forms of the protein from the lipid bilayer prior to proteasomal degradation.

A Plasma Membrane Targeting Sequence in the C-terminal Tail of Pma1 H+-ATPase—Of the truncation mutants analyzed in the present study, only Pma1{Delta}901 successfully exited the ER and reached the cell surface, where it split ATP at nearly one-half of the wild-type rate in GM cells and was able to support growth. This result effectively defines a 20-amino acid sequence between Ala881 and Glu901, stretching from the end of M10 to the midpoint of the C-terminal tail, which can target the ATPase to the plasma membrane. It should be noted, however, that the truncated ATPase was not completely normal, since it displayed aberrant kinetics, heightened sensitivity to trypsinolysis in vitro, and rapid delivery in vivo to the vacuole for degradation. Thus, even the last 18 residues of the C-terminal tail play a clear role in protein folding and function.

Among other P2-type ATPases, most membrane-targeting studies have concentrated on the closely related Na+,K+- and H+,K+-ATPases. Here the picture is complicated, both by the presence of a beta subunit and by the use of polarized cells to identify signals for basolateral versus apical targeting. Na+,K+-ATPase is typically located in the basolateral membrane of epithelial cells, whereas H+,K+-ATPase is mobilized to the apical surface of gastric parietal cells, where it pumps acid into the stomach lumen. To map the determinants of sorting, Dunbar et al. (47) constructed a set of chimeras between Na+,K+- and H+,K+-ATPase {alpha} subunits and tracked their behavior upon transfection into LLC-PK1 renal epithelial cells. The results were unexpectedly complex; substituting M4 of the H+,K+-ATPase {alpha} subunit for the equivalent Na+,K+-ATPase sequence was sufficient to redirect the chimera to the apical surface, but the same effect could be produced by incorporating the sequences flanking both ends of M4. Moreover, an earlier study (48) showed that the beta subunit of H+,K+-ATPase was delivered upon transfection to the basolateral surface of Madin-Darby canine kidney (MDCK) cells but to the apical surface of LLC-PK1 cells, indicating that identical targeting sequences can be "read" differently in two different cell types. By contrast with the intricacies of the results for Na+,K+- and H+,K+-ATPases, Pma1 H+-ATPase offers a relatively straightforward way to identify targeting sequences in a P2-type ATPase, uncomplicated by the presence of a beta subunit or the need to distinguish between two alternative membrane destinations.

Looking beyond the P2-type ATPase family, one finds many examples of membrane proteins in which the C-terminal region dictates delivery to the plasma membrane. For example, C-terminal tyrosine-containing motifs (e.g. NPXY) and dileucine motifs have long been known to direct proteins to the basolateral surface of polarized cells (reviewed in Ref. 49), and a C-terminal dileucine has recently been shown to play this role for Menkes protein, a copper-transporting P1-type ATPase, during expression in MDCK cells (50). Less is known about motifs that underlie the apical targeting of proteins, such as the cystic fibrosis transmembrane conductance regulator, sodium/phosphate cotransporter (NaPi II), and GABA transporter (GAT-3). Once again, however, there is strong evidence for localization of the signal to the C terminus (51-55), and recent work has narrowed the responsible region to 39 residues in the case of rhodopsin (56) and 30 residues in the case of megalin (57) during expression in MDCK cells. Every indication is that these targeting events are achieved by interaction of the responsible sequence with PDZ domain-containing proteins (reviewed in Ref. 58).

Interestingly, the 20-amino acid sequence between Ala881 and Glu901 of the yeast Pma1 H+-ATPase contains none of the C-terminal motifs previously associated with membrane protein targeting. Instead, it seems plausible that the highly charged nature of this sequence, which includes six positive and three negative residues, promotes interactions with other proteins that direct transport to the plasma membrane. This will be a fertile area for future study, with the long range aim of using the well recognized experimental advantages of yeast to map the molecular mechanism of targeting in detail.


   FOOTNOTES
 
* This work was supported by National Institutes of Health Research Grant GM15761. 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

1 To whom correspondence should be addressed: Dept. of Genetics, Yale University School of Medicine, 333 Cedar St., New Haven, CT 06510. Tel.: 203-737-1770; Fax: 203-737-1771; E-mail: carolyn.slayman{at}yale.edu.

2 The abbreviations used are: HA, hemagglutinin; ER, endoplasmic reticulum; TNP-ATP, 2',3'-O-(2,4,6-trinitrophenyl)-adenosine-5'-triphosphate; FITC, fluorescein isothiocyanate; GM, glucose-metabolizing; CS, carbon-starved. Back


   ACKNOWLEDGMENTS
 
We thank Thierry Ferreira for his work with the conditional expression system and other members of the Slayman laboratory for helpful discussions and support. We thank Daniele Urban-Grimal for providing yeast strains WG4a, WG4a-11/22a, W303-1B, and W303-1B pep4{Delta} and Mark Rose for providing antibody to Kar2p.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Morsomme, P., Slayman, C. W., and Goffeau, A. (2000) Biochim. Biophys. Acta 1469, 133-157[Medline] [Order article via Infotrieve]
  2. Serrano, R., Kielland-Brandt, M. C., and Fink, G. R. (1986) Nature 319, 689-693[CrossRef][Medline] [Order article via Infotrieve]
  3. Lutsenko, S., and Kaplan, J. H. (1995) Biochemistry 34, 15607-15613[CrossRef][Medline] [Order article via Infotrieve]
  4. Moller, J. V., Juul, B., and le Maire, M. (1996) Biochim. Biophys. Acta 1286, 1-51[Medline] [Order article via Infotrieve]
  5. Axelsen, K. B., and Palmgren, M. G. (1998) J. Mol. Evol. 46, 84-101[CrossRef][Medline] [Order article via Infotrieve]
  6. DeWitt, N. D., dos Santos, C. F., Allen, K. E., and Slayman, C. W. (1998) J. Biol. Chem. 273, 21744-21751[Abstract/Free Full Text]
  7. Wang, Q., and Chang, A. (1999) EMBO J. 18, 5972-5982[CrossRef][Medline] [Order article via Infotrieve]
  8. Petrov, V. V., Padmanabha, K. P., Nakamoto, R. K., Allen, K. E., and Slayman, C. W. (2000) J. Biol. Chem. 275, 15709-15716[Abstract/Free Full Text]
  9. Chang, A., and Fink, G. R. (1995) J. Cell Biol. 128, 39-49[Abstract/Free Full Text]
  10. Gong, X., and Chang, A. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 9104-9109[Abstract/Free Full Text]
  11. Ferreira, T., Mason, A. B., Pypaert, M., Allen, K. E., and Slayman, C. W. (2002) J. Biol. Chem. 277, 21027-21040[Abstract/Free Full Text]
  12. Serrano, R. (1988) Biochim. Biophys. Acta 947, 1-28[Medline] [Order article via Infotrieve]
  13. Richter-Ruoff, B., Wolf, D. H., and Hochstrasser, M. (1994) FEBS Lett. 354, 50-52[CrossRef][Medline] [Order article via Infotrieve]
  14. Thomas, B. J., and Rothstein, R. (1989) Cell 56, 610-630
  15. Nakamoto, R. K., Rao, R., and Slayman, C. W. (1991) J. Biol. Chem. 266, 7940-7949[Abstract/Free Full Text]
  16. Sherman, F., Hicks, J. B., and Fink, G. R. (1986) Methods in Yeast Genetics: A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
  17. Ito, H., Fukuda, Y., Murata, K., and Kimura, A. (1983) J. Bacteriol. 153, 163-168[Abstract/Free Full Text]
  18. Wang, G., Tamas, M., Hall, M. J., Pascal-Ahuir, A., and Perlin, D. S. (1996) J. Biol. Chem. 271, 25438-25445[Abstract/Free Full Text]
  19. Volland, C., Garnier, C., and Haguenauer-Tsapis, R. (1992) J. Biol. Chem. 267, 23767-23771[Abstract/Free Full Text]
  20. Laemmli, U. K. (1970) Nature 227, 680-685[CrossRef][Medline] [Order article via Infotrieve]
  21. Miranda, M., Allen, K. E., Pardo, J. P., and Slayman, C. W. (2001) J. Biol. Chem. 276, 22485-22490[Abstract/Free Full Text]
  22. Monk, B. C., Kurtz, M. B., Marrinan, J. A., and Perlin, D. S. (1991) J. Bacteriol. 173, 6826-6836[Abstract/Free Full Text]
  23. Lowry, O. H., Roseborough, N. J., Farr, A. L., and Randall, R. J. (1951) J. Biol. Chem. 193, 265-275[Free Full Text]
  24. Ambesi, A., Allen, K. E., and Slayman, C. W. (1997) Anal. Biochem. 251, 127-129[CrossRef][Medline] [Order article via Infotrieve]
  25. Kuhlbrandt, W., Zeelen, J., and Dietrich, J. (2002) Science 297, 1692-1696[Abstract/Free Full Text]
  26. Mason, A. B., Kardos, T. B., and Monk, B. C. (1998) Biochim. Biophys. Acta 1372, 261-271[Medline] [Order article via Infotrieve]
  27. Benito, B., Moreno, E., and Lagunas, R. (1991) Biochim. Biophys. Acta 1063, 265-268[Medline] [Order article via Infotrieve]
  28. Harris, S. L., Na, S., Zhu, X., Seto-Young, D., Perlin, D. S., Teem, J. H., and Haber, J. E. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 10531-10535[Abstract/Free Full Text]
  29. Egner, R., Rosenthal, F. E., Kralli, A., Sanglard, D., and Kuchler, K. (1998) Mol. Biol. Cell 9, 523-543[Abstract/Free Full Text]
  30. Loayza, D., Tam, A., Schmidt, W. K., and Michaelis, S. (1998) Mol. Biol. Cell 9, 2767-2784[Abstract/Free Full Text]
  31. Rose, M. D., Misra, L. M., and Vogel, J. P. (1989) Cell 57, 1211-1221[CrossRef][Medline] [Order article via Infotrieve]
  32. Heinemeyer, W., Gruhler, A., Mohrle, V., Mahe, Y., and Wolf, D. H. (1993) J. Biol. Chem. 268, 5115-5120[Abstract/Free Full Text]
  33. Shearer, A. G., and Hampton, R. Y. (2004) J. Biol. Chem. 279, 188-196[Abstract/Free Full Text]
  34. Schlesser, A., Ulaszewski, S., Ghislain, M., and Goffeau, A. (1988) J. Biol. Chem. 263, 19480-19487[Abstract/Free Full Text]
  35. Portillo, F. (2000) Biochim. Biophys. Acta 1469, 31-42[Medline] [Order article via Infotrieve]
  36. Ambesi, A., Pan, R. L., and Slayman, C. W. (1996) J. Biol. Chem. 271, 22999-23005[Abstract/Free Full Text]
  37. Lin, J., and Addison, R. (1995) J. Biol. Chem. 270, 6935-6941[Abstract/Free Full Text]
  38. Beggah, A. T., Beguin, P., Bamberg, K., Sachs, G., and Geering, K. (1999) J. Biol. Chem. 273, 24921-24931
  39. Jahn, T. P., Schulz, A., Taipalensuu, J., and Palmgren, M. G. (2002) J. Biol. Chem. 277, 6353-6358[Abstract/Free Full Text]
  40. Capieaux, E., Rapin, C., Thines, D., Dupont, Y., and Goffeau, A. (1993) J. Biol. Chem. 268, 21895-21900[Abstract/Free Full Text]
  41. Moutin, M.-J., Cuillel, M., Rapin, C., Miras, R., Anger, M., Lompré, A.-M., and Dupont, Y. (1994) J. Biol. Chem. 269, 11147-11154[Abstract/Free Full Text]
  42. Moutin, M.-J., Rapin, C., Miras, R., Vinçon, M., and Dupont, Y. (1998) Eur. J. Biochem. 251, 682-690[Medline] [Order article via Infotrieve]
  43. Gatto, C., Wang, A. X., and Kaplan, J. H. (1998) J. Biol. Chem. 273, 10578-10585[Abstract/Free Full Text]
  44. Taxis, C., Hitt, R., Park, S.-H., Deak, P. M., Kostova, Z., and Wolf, D. (2003) J. Biol. Chem. 278, 35903-35913[Abstract/Free Full Text]
  45. Béguin, P., Hasler, U., Staub, O., and Geering, K. (2000) Mol. Biol. Cell 11, 1657-1672[Abstract/Free Full Text]
  46. Vashist, S., and Ng, D. (2004) J. Cell Biol. 165, 41-52[Abstract/Free Full Text]
  47. Dunbar, L. A., Aronson, P. L., and Caplan, M. J. (2000) J. Cell Biol. 148, 769-778[Abstract/Free Full Text]
  48. Roush, D. L., Gottardi, C. J., Hussein, Y. N., Roth, M. G., and Caplan, M. J. (1998) J. Biol. Chem. 273, 26862-26869[Abstract/Free Full Text]
  49. Muth, T. R., and Caplan, M. J. (2003) Annu. Rev. Cell Dev. Biol. 19, 333-366[CrossRef][Medline] [Order article via Infotrieve]
  50. Greenough, M., Pase, L., Voskoboinik, I., Petris, M. J., O'Brien, A. W., and Camakaris, J. (2004) Am. J. Physiol. 287, C1463-C1471
  51. Moyer, B. D., Denton, J., Karlson, K. H., Reynolds, D., Wang, S., Mickle, J. E., Milewski, M., Cutting, G. R., Guggino, W. B., Li, M., and Stanton, B. A. (1999) J. Clin. Invest. 104, 1353-1361[Medline] [Order article via Infotrieve]
  52. Moyer, B. D., Duhaime, M., Shaw, C., Denton, J., Reynolds, D., Karlson, K. H., Pfeiffer, J., Wang, S., Mickle, J. E., Milewski, M., Cutting, G. R., Guggino, W. B., Li, M., and Stanton, B. A. (2000) J. Biol. Chem. 275, 27069-27074[Abstract/Free Full Text]
  53. Milewski, M. I., Mickle, J. E., Forrest, J. K., Stafford, D. M., Moyer, B. D., Cheng, J., Guggino, W. B., Stanton, B. A., and Cutting, G. R. (2001) J. Cell Sci. 114, 719-726[Abstract]
  54. Gisler, S. M., Stagljar, I., Traebert, M., Bacic, D., Biber, J., and Murer, H. (2001) J. Biol. Chem. 276, 9206-9213[Abstract/Free Full Text]
  55. Muth, T. R., Ahn, J., and Caplan, M. J. (1998) J. Biol. Chem. 273, 25616-25627[Abstract/Free Full Text]
  56. Chuang, J.-Z., and Sung, C.-H. (1999) J. Cell Biol. 142, 1245-1256
  57. Takeda, T., Yamazaki, H., and Farquhar, M. G. (2003) Am. J. Physiol. 284, C1105-C1113
  58. Caplan, M. J. (2003) Am. J. Physiol. 284, C1101-C1104

Add to CiteULike CiteULike   Add to Complore Complore