Article

• The EMBO Journal (1999) 18, 1506 - 1515
• doi:10.1093/emboj/18.6.1506

Sac1p plays a crucial role in microsomal ATP transport, which is distinct from its function in Golgi phospholipid metabolism

Kai-Ute Kochendörfer¹, Angela R. Then¹, Brian G. Kearns², Vytas A. Bankaitis² and Peter Mayinger¹

1. Zentrum für Molekulare Biologie, Universität Heidelberg, Im Neuenheimer Feld 282, D-69120 Heidelberg, Germany
2. Department of Cell Biology, University of Alabama at Birmingham, Birmingham, AL 35294-0005, USA

Correspondence to:

Peter Mayinger, E-mail: mayingerp@sun0.urz.uni-heidelberg.de

Received 26 October 1998; Accepted 19 January 1999; Revised 19 January 1999

• Keywords:

  • inositol,
  • microsomal ATP transport,
  • phospholipid metabolism,
  • protein trafficking,
  • Sac1p

Introduction

ATP transport into the endoplasmic reticulum (ER) is a prerequisite for multiple reactions inside the ER lumen. Many steps in ER protein processing are catalyzed by luminal ATP-dependent factors. In yeast, the post-translational transfer of secretory protein precursors can only occur if a sufficient level of ATP is present inside the ER lumen (Mayinger and Meyer, 1993; Mayinger et al., 1995). The ATP dependence of protein translocation can be attributed mainly to the function of Kar2p, the yeast homolog of the mammalian BiP protein (Normington et al., 1989; Rose et al., 1989). This chaperone has two ATP-dependent functions in the post-translational mode of translocation. First, it mediates the transfer of bound precursors into the translocation pore via interaction with Sec63p (Lyman and Schekman, 1997). Subsequently, the transport of precursors through the translocation pore is catalyzed by Kar2p and ATP in a mechanism in which Kar2p may function as 'molecular ratchet' or as a 'molecular motor' (Schatz and Dobberstein, 1996). The hsp70 homologs Ssi1p and Cer1p also appear to play roles in post-translational translocation in yeast (Baxter et al., 1996; Hamilton and Flynn, 1996). In additon, Kar2p and other ATP-dependent chaperones are crucial for the proper folding of translocated secretory proteins inside the ER lumen and ensure that only correctly folded proteins leave the ER and proceed along the secretory pathway toward their final destination within the cell (Gething and Sambrook, 1992; Simons et al., 1995). Finally, the discovery of ATP-dependent factors such as Lhs1p as important components of the heat shock response in yeast stipulates the presence of sufficient levels of ATP in the ER lumen for thermotolerance (Saris et al., 1997).

Identification and characterization of components that are required for efficient uptake of cytosolic ATP are therefore crucial to understand how ATP-dependent processes in the ER lumen are linked to the dynamic function of the ER. Although, a number of mammalian microsomal ATP transport systems have been discovered and analyzed, the molecular identity of these transporters or of components linked to ATP transport remained elusive until now (Clairmont et al., 1992; Guillen and Hirschberg, 1995; Abeijon et al., 1997). Recently, ATP transport activity from yeast microsomes was characterized in vitro in reconstituted proteoliposomes and it was shown that ATP uptake into yeast ER is mediated by an ATP/ADP antiport system (Mayinger et al., 1995). Furthermore, the partial purification of transport activity led to the identification of the ER and Golgi membrane protein Sac1p as a crucial component in ATP transport. Microsomal membranes isolated from strains harboring a deletion in the SAC1 gene (sac1) were found to be deficient in ATP transport activity. Only low levels of ATP transport could be recovered when detergent extracts obtained from the microsomal membranes of a sac1 strain were reconstituted in the in vitro assay, while overexpression of Sac1p caused an increased capacity for microsomal ATP transport (Mayinger et al., 1995). These findings would have been consistent with Sac1p representing the authentic ATP transporter.

The identification of Sac1p as a component in microsomal ATP transport was, however, surprising for two reasons. First, the SAC1 gene already had been implicated in two other cellular processes by genetic analysis. Loss of the SAC1 gene leads to suppression of certain act1 mutant alleles (Novick et al., 1989) and bypasses the requirement for the phosphatidylinositol transfer protein Sec14p for cell viability (Cleves et al., 1989). sac1 mutants also display alterations in cell wall structure and lose structural integrity at alkaline pH (Novick et al., 1989; Boyum and Guidotti, 1997). Secondly, the primary sequence of Sac1p shows no homology to the structure of known solute carrier proteins. Instead, a major portion of Sac1p is homologous to the N-terminal domain of the mammalian presynaptic phosphatidylinositol 5'-phosphatase synaptojanin (Majerus, 1996; McPherson et al., 1996). Recently, three yeast phosphatidylinositol 5'-phosphatases were identified which also contain an N-terminal Sac1p domain (Srinivasan et al., 1997; Stolz et al., 1998a, b), but neither the exact biological function of these proteins nor the role of the N-terminal Sac1p homology domain is known. In further studies, it could be demonstrated that sac1 cells and cells expressing certain mutant alleles of SAC1, such as sac1-22, display specific changes in phospholipid metabolism which are instrumental for 'bypass Sec14p' (Kearns et al., 1997; M.P.Rivas, B.G. Kearns, S.Guo, Z.Xie, M.C.Sekar, K.Hosaka, S.Kagiwada, J.D.York and V.A.Bankaitis, submitted).

To address the question of how Sac1p participates in microsomal ATP transport, we examined the ability of pure recombinant Sac1p to catalyze ATP uptake into proteoliposomes. We find that Sac1p itself does not function as an ATP transporter, whereas loss of the SAC1 gene results in ATP transport defects and ER-specific defects in protein translocation and protein folding. We conclude that Sac1p, rather than being the ATP transporter itself, is a regulatory or accessory factor required for efficient ATP transport.

Expression of GST–Sac1p in sac1 cells

To purify Sac1p to homogeneity, we inserted the SAC1 gene into a yeast episomal (2) vector next to the 5' end of glutathione S-transferase (GST), placing the expression of this chimeric fusion protein under control of the inducible GAL1/10 promoter (Mitchell et al., 1993). Between the GST-coding region and the SAC1 gene a linker was present containing five glycine residues and a thrombin cleavage site to allow the recovery of recombinant Sac1p without the GST tag (Figure 1A). For all subsequent experiments, we used a sac1 background with the reg1-501 mutation, which eliminates glucose repression of GAL expression (Hovland et al., 1989). GST–Sac1p expression could, therefore, be stimulated directly by addition of 2% galactose to the growth medium. Examination of cell extracts by SDS–PAGE and Western analysis revealed the appearance of an 89 kDa band after galactose induction which was recognized by a monoclonal antibody that is specific for the Sac1p protein (Figure 1B). This molecular weight is consistent with that expected for the GST–Sac1p fusion protein. For comparison, a 67 kDa protein band is present when the SAC1 gene is constitutively overexpressed in sac1 cells using an episomal 2 vector with the endogenous SAC1 promoter (Whitters et al., 1993) (Figure 1B). The overexpressed GST–Sac1p in sac1 cells is 40 times more abundant compared with the levels of Sac1p in wild-type cells. Cell fractionation and analysis of the distribution of GST–Sac1p within the different fractions revealed that essentially all the GST–Sac1p is membrane bound and co-localizes with the ER marker Kar2p (data not shown).

Figure 1.

Expression of a GST–Sac1p fusion protein. (A) Structure of the GST–SAC1 gene fusion. The GST–SAC1 expression vector was created by cloning an XbaI fragment encompassing the entire SAC1 coding region into the XbaI site of pEGKT (Mitchell et al., 1993). In the resulting vector pPM102, the SAC1 gene is positioned next to a linker encoding a thrombin cleavage site directly adjacent to the structural gene for GST under GAL1/10 control (Mitchell et al., 1993). (B) Expression of GST–Sac1p in sac1 cells. Expression of the GST–Sac1p fusion protein was induced by supplementing the growth medium with 2% galactose for 7 h. Subsequently, the cells were lysed and aliquots of the extracts were analyzed by SDS–PAGE and Western blotting. As a control for the expression, extracts from cells bearing a 2 plasmid with a genomic fragment containing the entire SAC1 gene including its endogenous promoter were also analyzed.

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Expression of GST–Sac1p complements the reduced ATP transport in sac1 cells

Disruption of the SAC1 gene causes several growth phenotypes. sac1 cells are cold-sensitive, inositol auxotrophs, and the doubling time for growth in rich media is approximately twice as long as wild-type doubling times (Cleves et al., 1989; Whitters et al., 1993). To determine whether the GST–Sac1p fusion protein is functional, we tested sac1 cells containing the GST–SAC1 plasmid for galactose-dependent complementation of these phenotypes. As shown in Figure 2A, the galactose-induced expression of GST–Sac1p confers viability to sac1 cells grown at 15°C or cultivated on inositol-free medium. In addition, the doubling time for growth of these cells on galactose-containing media is indistinguishable from that of wild-type cells (not shown). We conclude from this complementation analysis that the expressed recombinant GST–Sac1p can functionally substitute for the wild-type protein. Since it was most important to our study that the GST–Sac1p can completely fulfill the function that wild-type Sac1p plays in ATP uptake, we tested microsomal ATP transport directly. Microsomal membranes from galactose-induced sac1 cells containing the GST–SAC1 plasmid were prepared and tested for their ability to transport ATP. It was shown previously that microsomes from sac1 cells have greatly reduced ATP transport activity (Mayinger et al., 1995). In contrast, the GST–Sac1p-containing microsomes exhibit a four times higher capacity for ATP uptake than wild-type microsomes (Figure 2B). For comparison, the level of GST–Sac1p in these microsomes is 40 times higher than the amount of Sac1p in wild-type microsomes (data not shown), which indicates that either some of the expressed GST–Sac1p is inactive or that the stimulating effect of GST–Sac1p in microsomal ATP transport is saturable.

Figure 2.

Expression of a GST–Sac1p fusion protein in sac1 cells complements the specific phenotypes associated with a loss of SAC1. (A) sac1 cells containing the GST–SAC1 fusion on a 2 plasmid were plated on selective plates containing 2% galactose, and the complementation of the cold-sensitivity and inositol auxotrophy phenotypes was analyzed. (B) Microsomes from wild-type cells and from sac1 cells expressing GST–Sac1p were prepared according to Rothblatt and Meyer (1986) and assayed for specific uptake of ATP. ATP transport was measured as described in Materials and methods. Data are from three independent measurements.

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Purification of recombinant Sac1p by affinity chromatography

Having ascertained that GST–Sac1p functions in microsomal ATP uptake in vitro, we next asked whether purified recombinant Sac1p would be sufficient for catalyzing ATP transport in reconstituted proteoliposomes. For the purification of Sac1p, we used microsomes prepared from sac1 cells expressing the GST–Sac1p fusion protein. All fractions obtained during the purification procedure were analyzed by SDS–PAGE, which is depicted in Figure 3. After solubilization with Triton X-100 and 250 mM NaCl (Figure 3, lane 2), microsomal extracts were incubated with glutathione–agarose beads. The supernatant after quick centrifugation was significantly depleted of the GST–Sac1p fusion protein (Figure 3, lane 3). This adsorption step was followed by three washes (Figure 3, lanes 4 and 5). Then, the glutathione–agarose beads containing the bound GST–Sac1p were divided into two aliquots and either treated with thrombin, to cleave off the Sac1p portion of the fusion protein (Figure 3, lane 6, thrombin-SN, and lane 7, thrombin-beads), or the bound GST–Sac1p was eluted by boiling the beads with electrophoresis sample buffer (Figure 3, lane 8, SDS). These data show that GST–Sac1p binds specifically to glutathione–agarose beads. Thrombin cleavage of the linker between the GST protein and Sac1p results in essentially pure recombinant Sac1p that differs from wild-type Sac1p only by the presence of seven additional amino acids on its N-terminus. Using these conditions for purification, we did not observe other proteins in these fractions that could be potential interacting components of Sac1p. In rare cases, in which additional protein species were found in the thrombin-cleaved fraction, the contaminating components could be identified as breakdown products of Sac1p which were recognized by Sac1p-specific antibodies (not shown).

Figure 3.

Purification of recombinant Sac1p by affinity adsorption on glutathione–agarose. Microsomes containing high levels of GST–Sac1p were solubilized in 3% Triton X-100, 250 mM NaCl, 10 mM HEPES pH 7.4. The extract was centrifuged to obtain a pellet fraction (lane 1, PE) and a supernatant fraction containing the solubilized GST–Sac1p (lane 2, SN). The supernatant was diluted 1:1 with water and then mixed with glutathione–agarose beads. After 30 min incubation, the beads were washed three times with 1% Triton X-100, 125 mM NaCl, 10 mM HEPES pH 7.4 (lanes 4 and 5; only the first and the last wash fractions are shown in the figure). The beads containing the bound GST–Sac1p were resuspended in 0.5% Triton X-100, 150 mM NaCl, 2.5 mM CaCl₂, 50 mM Tris–HCl, pH 8.0, 0.1% -mercaptoethanol, treated with thrombin and centrifuged to obtain a supernatant fraction containing the recombinant Sac1p (lane 6, thrombin-SN) and a pellet fraction containing the GST portion of the fusion protein which remained tightly bound to the glutathione–agarose beads (lane 7, thrombin-PE). Alternatively, an aliquot of the beads containing the bound GST–Sac1p was solubilized directly in electrophoresis buffer (lane 8, SDS).

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Reconstitution and functional characterization of purified recombinant Sac1p

To analyze the function of pure recombinant Sac1p in microsomal ATP transport, we reconstituted the purified protein into proteoliposomes and examined ATP transport activities. Two questions were addressed in these experiments. First, whether the purified recombinant Sac1p is still active. Secondly, whether the purified protein itself functions as an ATP transporter. To answer the first question, sac1 microsomes were solubilized with Triton X-100 and reconstituted with or without addition of a fraction containing pure recombinant Sac1p. In both cases, the reconstitution was performed in the presence of 10 mM ATP to obtain proteoliposomes loaded with exchangable nucleotide substrate. In previous experiments, it was shown that ATP transport into reconstituted proteoliposomes follows a nucleotide antiport mechanism and can, therefore, only be measured when ADP or ATP is present inside the liposomes (Mayinger et al., 1995). If extracts from sac1 microsomes were reconstituted, only a very low level of ATP uptake could be observed (Figure 4), which is consistent with our previous finding (Mayinger et al., 1995). However, when purified recombinant Sac1p was reconstituted together with extracts of sac1 microsomes, ATP transport activity was almost fully restored, indicating that the recombinant Sac1p is functional and sufficient to repair the ATP transport defect associated with sac1 microsomes. Next, we wanted to determine whether the purified recombinant Sac1p itself can function as a specific ATP antiporter in the reconstituted system. Accordingly, we prepared proteoliposomes containing only the purified protein. Analysis by SDS–PAGE and Western blotting confirmed that the purified recombinant Sac1p was incorporated efficiently into the phospholipid vesicles (not shown). Significantly, these proteoliposomes were completely inactive in ATP transport (Figure 4). These results provide strong evidence that Sac1p is necessary for effective ATP uptake into reconstituted proteoliposomes, yet cannot catalyze ATP transport alone.

Figure 4.

Reconstitution of the recombinant Sac1p. Proteoliposomes were prepared as described in Materials and methods, using detergent extracts from wild-type microsomes or from sac1 microsomal membranes, with or without addition of 5 g of purified recombinant Sac1p. Alternatively, proteoliposomes were prepared using only a fraction containing 5 g of the purified recombinant Sac1p. The reconstitution mixtures were supplemented with 10 mM ATP to create liposomes loaded with exchangable nucleotide substrate. ATP uptake measurements were conducted as described in Materials and methods. Data are from three independent measurements.

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The microsomal ATP transport defect in sac1 causes a folding defect in the ER

The collective biochemical data suggest that Sac1p is not an ATP transporter but rather an essential co-factor or regulator of microsomal ATP transport. To test whether such a function is relevant for live yeast cells, we examined how the ATP transport defect in sac1 mutants manifests itself in vivo. The deficiency in microsomal ATP uptake that is seen in sac1 microsomes should lower the ATP/ADP ratio inside the ER lumen, which in turn would affect ATP-dependent steps in this compartment. We tested this by a pulse–chase analysis of ER-dependent protein trafficking. As previously observed, a complete loss of the SAC1 gene causes a substantial defect in procarboxypeptidase Y (proCPY) transport from the ER to the Golgi complex (Mayinger et al., 1995) (Figure 5A). Since the function of the ER chaperone Kar2p is ATP-dependent, an in vivo ATP transport defect should result in increased levels of unfolded secretory proteins in the ER lumen, thus affecting the net rate of transport out of the ER. To detect such a deficiency in protein folding in sac1 cells, we examined the association of Kar2p with proCPY in the ER in vivo by metabolic labeling and immunoprecipitation. As shown in Figure 5A, almost no Kar2p can be co-immunoprecipitated with CPY-specific antibodies during the transit of proCPY through the ER. In sac1 cells, a substantially higher proportion of Kar2p was associated with proCPY which correlates with the accumulation of proCPY in the ER. This finding indeed points towards a malfunction of Kar2p in sac1 cells in vivo. These data are also in line with the previous observation that in wild-type cells only a very small portion of Kar2p can be co-immunoprecipitated with CPY-specific antibodies, while particular kar2 mutants with a defect in the ATPase domain show significantly higher amounts of Kar2p associated with proCPY (Te Heesen and Aebi, 1994; Simons et al., 1995).

Figure 5.

The microsomal ATP transport defect in sac1 cells leads to a malfunction of Kar2p. (A) Cells from wild-type and sac1 cells were grown to mid-log phase. The cells were pulse-labeled with [³⁵S]methionine and [³⁵S]cysteine for 5 min at 25°C. The reactions were chased with non-labeled amino acids for the indicated times. The cells were lysed in the presence of NaN₃ and apyrase to prevent dissociation of polypeptide-bound Kar2p. The extracts were immunoprecipitated with anti-CPY polyclonal antibodies. The precipitated material was analyzed by SDS–PAGE and fluorography or by probing with anti-Kar2p antibodies to determine the portion of Kar2p associated with proCPY in the ER (p1, ER form of CPY; p2, Golgi form of CPY; m, mature CPY). (B) Cells from wild-type and sac1 cells were grown to mid-log phase. Three equal aliquots of each strain were incubated further for 2.5 h either at 25°C, at 25°C after addition of 10 mM DTT, or at 15°C. The cells were lysed and the extracts were analyzed by SDS–PAGE and Western blotting using anti-Kar2p polyclonal antibodies. The band below the Kar2p band, which appears in the DDT-treated samples, is a Kar2p breakdown product. (C) Standard genetic techniques were used to create double mutants with appropriate genotypes and containing pAT2 (SAC1 URA3) or pAT2 plus pPM103 (sac1-22 LYS2). The strains were streaked out on plates with or without FOA and incubated at 25°C for 5 days.

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The accumulation of unfolded proteins in the ER leads to activation of a signaling pathway that activates the transcription of resident ER proteins. This up-regulation of ER-resident chaperones is known as the unfolded protein response (UPR) (Cox and Walter, 1996; Shamu, 1998). Therefore, we compared the protein levels of Kar2p in wild-type and sac1 cells. As a control, the cells were incubated with the reducing agent dithiothreitol (DTT) which is known to induce the UPR (Figure 5B). Since sac1 cells cannot grow at 15°C, we also tested the influence of temperature on Kar2p levels in these cells (Figure 5B). Consistent with published results (Cox et al., 1993; Kohno et al., 1993), DTT induces a dramatic increase in the levels of Kar2p in wild-type cells, while a shift to 15°C leads to only slightly elevated amounts of Kar2p. In contrast, sac1 cells show a high steady-state level of Kar2p even without DTT (Figure 5B). Furthermore, addition of DTT does not lead to a further increase in Kar2p levels in these cells. These results indicate that the UPR in sac1 cells is constitutively switched on and that the level of Kar2p cannot be stimulated further by the addition of reducing agents. This finding confirms that unfolded proteins accumulate in the ER lumen of sac1 cells, which may be a consequence of reduced supply of ATP to this compartment.

We further reasoned that the intimate link between ATP transport and Kar2p function in the ER lumen would also become apparent by specific genetic interactions between sac1 and kar2 mutant alleles. We performed this analysis using a sac1 allele, the sac1-22 allele, which has no deficiency in microsomal ATP transport (Kearns et al., 1997), the kar2-159^ts allele, which is defective in the release of bound polypeptides, and the kar2-1^ts allele, which has a defect in polypeptide binding (Te Heesen and Aebi, 1994). To generate the appropriate genetic combinations, we transformed both kar2 mutant strains with pAT2, a plasmid bearing a wild-type SAC1 gene and the URA3 auxotrophic marker, or with both pAT2 and pPM103, a plasmid containing the sac1-22 allele and the LYS2 marker. Then, we disrupted the genomic copy of SAC1 in these strains and plated the cells on media with or without addition of 5'-fluoroorotic acid (FOA). Orotidine-5'-phosphate decarboxylase, the product of the URA3 gene, catalyzes the last step in uracil biosynthesis and is essential for converting FOA to 5'-fluorouracil, which is toxic to yeast cells (Boeke et al., 1987). Therefore, only cells which are viable without the pAT2 (URA3 SAC1) plasmid can grow on FOA-containing plates, while negative genetic interactions between the sac1 and kar2 alleles become visible by reduced growth on such media. As shown in Figure 5C, the kar2-159^ts sac1 as well as the kar2-1^ts sac1 double mutant strains are not viable on FOA plates at 25°C and, therefore, show a strong synthetically lethal phenotype. Importantly, the sac1-22 allele on a plasmid is able to rescue the synthetic phenotype. The significance of this finding is discussed below.

sac1-22 strains show wild-type microsomal ATP transport and CPY trafficking

Some aspects of the pleitropic phenotype displayed by sac1 cells are based on an altered inositol phospholipid metabolism, which may complicate interpretation of our results. This aberrant inositol phospholipid metabolism in sac1 cells obviates the essential Sec14p requirement for Golgi secretory function (Cleves et al., 1989; Kearns et al., 1997; M.P.Rivas, B.G.Kearns, S.Guo, Z.Xie, M.C.Sekar, K.Hosaka, S.Kagiwada, J.D.York and V.A.Bankaitis, submitted). Consequently, one could argue that the defects in ER to Golgi trafficking, apparent in sac1 cells are an indirect consequence of alterations in inositol phospholipids, which may cause a general disturbance of intracellular membrane function. It was therefore crucial to dissect genetically and biochemically the effects that are based on an altered inositol phospholipid metabolism from the consequences of impaired ATP transport in the ER.

Recently, we identified a particular mutant allele, sac1-22, which displays the same aberrant Golgi inositol phospholipid metabolism as sac1 strains, but fully complements the in vitro ATP transport defect that is associated with a complete loss of the SAC1 gene function (Kearns et al., 1997) (Figure 6A). We therefore analyzed ER to Golgi transport of proCPY in sac1-22 cells. As depicted in Figure 6B, ER to Golgi trafficking in sac1-22 cells is indistinguishable from that in wild-type cells, which is consistent with the wild-type levels of ATP transport that can be observed in sac1-22 microsomes (Figure 6A). This finding provides strong additional evidence that the ER-specific defects in protein trafficking in sac1 cells are a direct consequence of impaired microsomal ATP transport which causes the malfunction of luminal ATP-dependent factors and is not related to changes in Golgi inositol phospholipid metabolism. This argument is strengthened by the observation that the sac1-22 allele is also capable of complementing the synthetically lethal phenotype of various kar2 sac1 double mutants (Figure 5C).

Figure 6.

The sac1-22 mutant has normal microsomal ATP transport and CPY trafficking. (A) Microsomes from sac1-22 and wild-type cells were prepared according to Rothblatt and Meyer (1986) and assayed for microsomal ATP transport as described in Materials and methods. Data are from three independent measurements. (B) Cells from wild-type and sac1-22 strains were grown to mid-log phase. The cells were pulse-labeled with [³⁵S]methionine and [³⁵S]cysteine for 5 min at 25°C. The reactions were chased with non-labeled amino acids for the indicated times. The cells were lysed and the extracts were immunoprecipitated with anti-CPY polyclonal antibodies. The precipitated material was analyzed by SDS–PAGE and fluorography (p1, ER form of CPY; p2, Golgi form of CPY; m, mature CPY).

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sac1 shows specific synthetic interactions with genes coding for ATP-requiring translocon components

Since kar2 alleles and sac1 are synthetically lethal, we wanted to investigate the genetic link between ATP transport and ATP-dependent reactions in the ER in more detail. We speculated that the ATP transport deficiency in sac1 strains would also produce a synthetic phenotype when combined with mutant alleles of genes required for ATP-dependent steps in protein translocation. To test for such genetic interactions, we used plasmid pAT2 (SAC1 URA3) and plasmid pAT3 (sac1-22 TRP1). Appropriate mutant strains containing the sec61-2^ts, sec62-1^ts and sec63-1^ts alleles were crossed with ATY300, a sac1::HIS3 strain. The diploids were transformed with the plasmid pAT2 (SAC1 URA3) alone or with both plasmid pAT2 and plasmid pAT3 (sac1-22 TRP1). Tetrads from the respective transformed diploids were dissected and replica plated on rich medium and on FOA plates to select against pAT2 (SAC1 URA3). In cases where a significant number of spores did not grow on FOA plates, the segregation of the lethal phenotype of at least 18 tetrads was analyzed to confirm that these particular double mutants were not viable (not shown). Synthetic lethality at all temperatures was observed when the sec61-2^ts or the sec63-1^ts allele was combined with sac1, whereas a sec62-1^ts sac1 strain was viable at 25 or 30°C and displayed only the ts and cs phenotype of the respective sec62 and sac1 alleles (Figure 7). The synthetic lethal double mutants containing pAT2 (SAC1 URA3) plus pAT3 (sac1-22 TRP1) were also analyzed on FOA plates. Importantly, all synthetic lethal combinations with sac1 were able to grow on FOA when the sac1-22 allele was present on a plasmid (Figure 7). These results show that the synthetic interactions of sac1 with sec61-2^ts and sec63-1^ts are based on the diminished ATP transport that is associated with sac1 and are not connected with the inositol phospholipid phenotype which is present in both sac1 and sac1-22 alleles.

Figure 7.

sac1 alleles display synthetic lethality with mutations in genes coding for ATP-dependent factors in protein translocation. Standard genetic techniques were used to create strains with the appropriate genotypes containing either pAT2 (SAC1 URA3) or both pAT2 and pAT3 (sac1-22 TRP1), as indicated in the figure. The strains were streaked out on plates with or without FOA and incubated at 25°C for 5 days.

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The results presented confirm the notion that Sac1p is intimately associated with microsomal ATP transport in yeast. To analyze this aspect in detail, we used a GST-tagged version of Sac1p. The recombinant GST–Sac1p complemented all defects that are present in a sac1 strain. However, the biochemical data obtained with pure recombinant Sac1p do not support a hypothesis in which Sac1p represents the ATP transporter itself. The purified recombinant Sac1p was unable to confer ATP transport activity to reconstituted proteoliposomes but could complement the impaired ATP transport in reconstituted detergent extracts from sac1 microsomes when present in the reconstitution mixture. Therefore, Sac1p is likely to be an essential co-factor for efficient ATP uptake into yeast microsomes and does not represent the authentic transporter. How does Sac1p interact with the ATP transport machinery? One possibility is that Sac1p regulates ATP transport via downstream factors or by generating second messengers that bind to the ATP transporter. Phosphoinositides would be obvious candidates for such signals, though our results indicate that ATP transport is not affected by the accumulation of inositol phospholipids in membranes of sac1 mutants. Alternatively, Sac1p may interact directly with components of the ATP transport system in the ER membrane. Our previous finding that Sac1p is a prominent constituent of a purified fraction enriched in ATP transport activity (Mayinger et al., 1995) is in favor of such a possibility. Additionally, it was shown before that Sac1p localizes both to Golgi and to ER membranes (Whitters et al., 1993), where it potentially could interact with the ATP antiport system.

There is substantial evidence that Sac1p plays an important role in regulating phospholipid metabolism. Loss of the SAC1 gene causes inositol auxotrophy and leads to alterations in inositol phospholipid homeostasis (Whitters et al., 1993; Kearns et al., 1997; M.P.Rivas, B.G.Kearns, S.Guo, Z.Xie, M.C.Sekar, K.Hosaka, S.Kagiwada, J.D.York and V.A.Bankaitis, submitted). Aberrant phospholipid metabolism has been proposed to bypass either directly or indirectly the essential cellular requirement for SEC14, which encodes the yeast phosphatidylinositol transfer protein (Bankaitis et al., 1990; Cleves et al., 1991a; Kearns et al., 1997; M.P.Rivas, B.G.Kearns, S.Guo, Z.Xie, M.C.Sekar, K.Hosaka, S.Kagiwada, J.D.York and V.A.Bankaitis, submitted). It was therefore an a priori possibility that the ATP transport defect that is associated with loss of the SAC1 gene is an indirect consequence of this effect. Strong evidence for a direct role of Sac1p in ATP transport in vivo comes from the analysis of the sac1-22 missense allele. sac1-22 strains exhibit the same dramatic changes in Golgi inositol phospholipid metabolism that are seen in a sac1 strain, but have wild-type levels of microsomal ATP transporter activity. The use of the sac1-22 allele enabled us to link microsomal ATP transport directly to protein translocation and folding in the ER in vivo. sac1 displays synthetic lethality with sec61^ts, sec63^ts and kar2^ts alleles, while sac1-22 on a low copy plasmid rescued these synthetic lethal phenotypes. Importantly, a sec62 sac1 combination is viable. This finding is consistent with a mechanism of protein translocation in which Sec61p, Sec63p and Kar2p participate in a functional complex that is strictly dependent on luminal ATP when engaged in the translocation of precursor proteins across the ER membrane (Panzner et al., 1995; Lyman and Schekman, 1997). In contrast, Sec62p functions in a step that does not require luminal ATP. Recent data indicate that Sec62p, together with Sec71p and Sec72p, represents the binding site for precursors of secretory proteins on the ER membrane (Sanders et al., 1992; Lyman and Schekman, 1997). This binding of secretory precursors to the ER membrane was shown previously to be independent of ATP (Sanz and Meyer, 1989).

The requirement for a critical level of luminal ATP for protein translocation and protein folding and the importance of Sac1p in maintaining such a concentration of luminal ATP is substantiated further by our in vivo analysis of CPY trafficking in sac1 cells. It was demonstrated previously that Kar2p is an essential factor directly linked to translocation and folding of CPY in the ER lumen (Te Heesen and Aebi, 1994; Simons et al., 1995). Kar2p is a member of the hsp chaperone family and binds transiently to unfolded portions of translocated proteins in an adenine nucleotide-dependent cycle. In this mechanism, ATP binding to Kar2p is needed for the release of bound substrate proteins. Accordingly, one would predict that a drop in the ADP/ATP ratio in the ER lumen should lead to an increased life time of Kar2p-containing complexes and also to a decreased level of free Kar2p in the ER lumen. Our analysis of protein trafficking in sac1 cells shows that a significant portion of proCPY remains associated with Kar2p in the ER lumen, thus slowing the exit of proCPY out of the ER. In addition, the unfolded protein response pathway (Cox and Walter, 1996; Shamu, 1998) is constitutively activated in sac1 strains. These data provide strong additional evidence that the luminal ATP/ADP ratio in sac1 cells is reduced to such a level that the function of ATP-dependent luminal chaperones becomes severely compromised.

Though it is not possible at this point to assemble all these aspects into a unifying model, the collective biochemical and genetic data presented here underscore the importance of the SAC1 gene in microsomal ATP transport and prove that this function is distinct from the role of Sac1p in regulating Golgi phospholipid metabolism. The question of how Sac1p may function in both ATP transport and Golgi phospholipid metabolism assumes particular importance, since these results show a possible connection between metabolic energy supply in the ER and Golgi function. Since Sac1p is a constituent of both ER and Golgi membranes, this protein may be involved in compartment-specific signaling events potentially via inositol phospholipids. Generation of transport vesicles is a highly regulated process that also depends on specific changes in the membrane distribution of specific types of phospholipids (Cleves et al., 1991b; Alb et al., 1996; Ktistakis et al., 1996; Chen et al., 1997). In addition, there is increasing evidence that phosphoinositides such as PtdIns(4)P, PtdIns(3)P and PtdIns(4,5)P₂ have important roles in regulating specific intracellular membrane trafficking events (De Camilli et al., 1996). The function of Sac1p in modulating inositol phospholipid metabolism is counter to the role of Sec14p in vesicle budding, since Sac1p defects rescue Golgi secretory function and cell viability in Sec14p-deficient yeast. (Cleves et al., 1989; Whitters et al., 1993; Kearns et al., 1997; M.P.Rivas, B.G.Kearns, S.Guo, Z.Xie, M.C.Sekar, K.Hosaka, S.Kagiwada, J.D.York and V.A.Bankaitis, submitted).

Changes in flux through the secretory pathway also alter the turnover of secretory proteins in the ER and Golgi. It is known that overexpression of secretory proteins in yeast causes increased synthesis of ER-resident proteins and phospholipids, which are both regulated by UPR (Cox et al., 1997; Menzel et al., 1997). Therefore, the level of ATP needed in the ER lumen has to be adapted to the respective size of the ER, which can vary substantially (Koning et al., 1996; Cox et al., 1997; Menzel et al., 1997) depending on the different levels of translocating secretory proteins and folding substrates passing through the ER lumen. Sac1p may therefore be a factor involved in coordinating ER and Golgi secretory competence in response to the supply of ATP to the ER. Finally, the homology of the N-terminal portion of Sac1p to the non-catalytic domain of a number of mammalian and yeast inositol-5'-phosphatases (McPherson et al., 1996; Kearns et al., 1997; Stolz et al., 1998a, b) provides additional evidence that Sac1p is involved in the regulation of intracellular membrane traffic by inositol phospholipids. The identification of upstream and downstream elements that interact with Sac1p and the elucidation of the role of Sac1p in inositol phospholipid signaling will be required to define more precisely the exact function that Sac1p plays in these processes.

Plasmids, strains, media and reagents

The plasmid pPM102 was created by ligating a 2.4 kb XbaI fragment bearing SAC1 (Cleves et al., 1989) into pEGKT (URA3, 2) (Mitchell et al., 1993), next to the structural gene for GST, which is under control of the GAL1/10 promoter. The plasmid pPM103 is pRS317 (LYS2 CEN) (Sikorski and Hieter, 1989), containing the sac1-22 allele. The plasmid pAT2 is YCplac33 (URA3 CEN) (Gietz and Sugino, 1988), containing the SAC1 gene. pAT3 is YCplac22 (TRP1 CEN) (Gietz and Sugino, 1988), containing the sac1-22 allele.

The strain ATY300 (MATaura3-52 his3-200 ade2-101 lys2-801 met- can^Rreg1-501 LEU2::GAL1-lacZ sac1::HIS3) was generated by deleting the SAC1 gene in wild-type strain YM1862 (MATaura3-52 his3-200 ade2-101, lys2-801 met-can^Rreg1-501 LEU2::GAL1-lacZ) (Hovland et al., 1989). For construction of double mutants with the sac1 allele, ATY300 was crossed with a sec61-2 strain (Y0098; gift from S.Jentsch, Heidelberg, Germany), sec62-1 strain (RSY529; gift from R.Schekman, Berkeley, CA) and sec63-1 strain (J51-5c; gift from D.Meyer, Los Angeles, CA). The diploids were transformed with pAT2 or with both pAT2 and pAT3. Double mutant derivatives with relevant genotypes were obtained by tetrad analysis. Synthetic interactions were analyzed by plating on 5'-FOA plates. sac1 kar2 double mutants were constructed by transforming kar2-159 and kar2-1 strains (YG0041, YG0044; gift from M.Aebi, Zürich, Switzerland) with pAT2 or with both pAT2 and pPM103. Subsequently, the SAC1 gene in these strains was disrupted to generate sac1 kar2-159 and sac1 kar2-1 strains, each containing a wild-type copy of SAC1 or an additional sac1-22 copy on respective plasmids to allow plasmid shuffling.

Yeast complete media (YPD), Hartwell's complete media (HC) and yeast minimal media (SD) were prepared as described (Adams et al., 1997). Yeast minimal media that lack inositol were prepared according to Klig et al. (1985). Complementation studies, meiotic segregation analyses, plasmid shuffling and gene disruptions were performed according to standard genetic techniques (Adams et al., 1997). GST–agarose, protein A–Sepharose, thrombin, Sephadex G-150 and phenylmethylsulfonyl fluoride (PMSF) were purchased from Sigma. SM2 Bio Beads were obtained from Bio-Rad. Zymolyase was obtained from ICN. Nucleotides were obtained from Boehringer Mannheim. Radiochemicals were purchased from Amersham. Polyclonal antiserum against Kar2p was a gift from G.Brodsky (University of Pennsylvania, Pittsburg, PA).

Purification of recombinant Sac1p

The strain ATY300 was transformed with pPM102 to obtain ATY300–3. Cells were grown to mid-logarithimic phase in SD medium containing 2% glucose. Expression of GST–Sac1p was induced directly by addition of 2% galactose to the medium since the strain ATY300-3 contained the reg1-501 mutation which abolishes glucose repression of GAL1/10 (Hovland et al., 1989). The cultures were incubated for an additional 6–8 h and cells were harvested by centrifugation. Isolation of microsomal membranes was performed according to Rothblatt and Meyer (1986). The GST–Sac1p fusion protein was solubilized by incubating the membranes (1 mg/ml) for 15 min on ice in 250 l of solubilization buffer containing 3% Triton X-100, 250 mM NaCl, 20 mM HEPES pH 7.4, 1 mM PMSF and 10 g/ml CLAP. After clearing the detergent extract by centrifugation at 20 000 g for 10 min, the supernatant was diluted 1:1 with water, mixed with 200 l of 50% glutathione–agarose suspended in 125 mM NaCl, 10 mM HEPES, pH 7.4 and incubated with gentle agitation for 30 min at 4°C. The beads were washed three times with 1 ml of 1% Triton X-100, 125 mM NaCl, 10 mM HEPES, pH 7.4. and then incubated in 0.5% Triton X-100, 150 mM NaCl, 2.5 mM CaCl₂, 50 mM Tris–HCl pH 8.0, 0.1% -mercaptoethanol and 0.5 U of thrombin for 20 min at room temperature. Thrombin digestion was stopped by addition of 1 mM PMSF and the beads were removed by centrifugation. The supernatant, containing the pure recombinant Sac1p, was concentrated by centrifugal filter units (Ultrafree MC, Millipore).

Preparation of reconstituted proteoliposomes

Reconstitution of detergent extracts of microsomal membranes was carried out essentially as described in Mayinger et al. (1995). Phosphatidylcholine/cholesterol (9:1) liposomes were prepared by sonication of a phosphatidyl/cholesterol suspension (100 mg/ml total lipid, in 100 mM MOPS, pH 7.4) in an ice bath. For reconstitution of the recombinant Sac1p, 10 g/ml of purified protein was supplemented with a total amount of 48 mg/ml Triton X-100. To this solution, 24 mg/ml of the lipid mixture was added in small portions. After the solution became translucent, internal substrate nucleotides were added to a final concentration of 10 mM and the mix was incubated for 30 min on ice. Proteoliposomes were generated by removal of detergent accomplished by repeated passage over Bio Beads SM2 columns (1 g of SM2 beads/24 mg of Triton X-100). The proteoliposomes were separated from non-reconstituted material by gel filtration on a Sephadex G-150 column.

ATP transport assay

ATP transport into yeast microsomes was determined as described previously (Mayinger and Meyer, 1993). ATP uptake into reconstituted proteoliposomes was determined similarly as in Mayinger et al. (1995). A 300 l aliquot of liposome suspension containing 10 mM ATP as counter substrate was mixed rapidly with 50 M [¹⁴C]ATP at 25°C. The exchange was stopped at different time points by rapid passage of 50 l aliquots over 150 mg of wet Dowex 8 (chloride form, 100–200 mesh) in small glass columns. Free nucleotides are bound rapidly by the strong anion exchange resin, while liposomes are not retained. The columns were washed once with 100 l of 100 mM MOPS, pH 7.4. Radioactivity in the combined eluate and wash was quantified by scintillation counting. Non-specific association of ATP with proteoliposomes was determined by conducting the transport assay in the presence of 0.5% Triton X-100 and by measuring ATP uptake at 0°C. All transport assays were performed in the presence of 100 M carboxyatractyloside, a specific inhibitor of the mitochondrial ADP/ATP carrier, to rule out any influence by mitochondrial contaminants.

Metabolic labeling

Yeast strains were grown at 30°C in minimal medium (SD) to mid-log phase. Cells at 1.25 OD₆₀₀ were harvested, washed with SD and resuspended in 1.3 ml of SD. The cells were incubated at 25°C for 15 min and then labeled with 0.5 mCi of Trans ³⁵S-label for 5 min. Subsequently 1.3 ml of 2 chase solution was added (final concentration: 0.03% methionine, 0.03% cysteine and 0.03 M ammonium sulfate). Aliquots were removed at different time intervals and quenched on ice with 10 mM NaN₃. The samples were pelleted and washed in phosphate-buffered saline. Glass bead lysis and denaturing immunoprecipitations were conducted as described earlier (Mayinger et al., 1995). For native immunoprecipitations, cell pellets were resuspended in 250 l of 2% CHAPS, 200 mM NaCl, 50 mM HEPES pH 7.4 and lysed with 200 g of glass beads in the presence of 1 mM NaN₃ and apyrase (1 U) to prevent dissociation of polypeptide-bound Kar2p. The extracts were immunoprecipitated with anti-CPY polyclonal antibodies. The precipitated material was analyzed by SDS–PAGE and fluorography or by probing with anti-Kar2p antibodies to determine the portion of Kar2p associated with proCPY in the ER.

We thank Jeffrey L.Brodsky for Kar2p antibodies; David I.Meyer, Randy Schekman, Stefan Jentsch, Markus Aebi, Jörg Höhfeld and Robert J.Deschenes for yeast strains and plasmids; and Teresa Nicolson, Gerlinde Konrad and Bernhard Dobberstein for critically reading the manuscript. We also acknowledge the technical assistance of Corinna Weidt and Barbara Schell. B.G.K. and V.A.B. were supported by National Institutes of Health (NIH) grant GM44530. The research was supported by a grant from the Deutsche Forschungsgemeinschaft to P.M. (Ma 1363/3-1).

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