Pmr1, a Golgi Ca2+/Mn2+-ATPase, is a regulator of the target of rapamycin (TOR) signaling pathway in yeast

  1. Gina Devasahayam*,
  2. Danilo Ritz†,
  3. Stephen B. Helliwell†,‡,
  4. Daniel J. Burke§, and
  5. Thomas W. Sturgill*,
  1. Departments of *Pharmacology and
  2. §Biochemistry and Molecular Genetics, University of Virginia Health Sciences Center, 1300 Jefferson Park Avenue, Charlottesville, VA 22908; and
  3. †Division of Biochemistry, Biozentrum, University of Basel, CH-4056 Basel, Switzerland

  1. Edited by Craig B. Thompson, University of Pennsylvania, Philadelphia, PA, and approved September 25, 2006 (received for review May 30, 2006)

 
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Abstract

The rapamycin·FKBP12 complex inhibits target of rapamycin (TOR) kinase in TORC1. We screened the yeast nonessential gene deletion collection to identify mutants that conferred rapamycin resistance, and we identified PMR1, encoding the Golgi Ca2+/Mn2+-ATPase. Deleting PMR1 in two genetic backgrounds confers rapamycin resistance. Epistasis analyses show that Pmr1 functions upstream from Npr1 and Gln-3 in opposition to Lst8, a regulator of TOR. Npr1 kinase is largely cytoplasmic, and a portion localizes to the Golgi where amino acid permeases are modified and sorted. Nuclear translocation of Gln-3 and Gln-3 reporter activity in pmr1 cells are impaired, but expression of functional Gap1 in the plasma membrane of a pmr1 strain in response to nitrogen limitation is enhanced. These two phenotypes suggest up-regulation of Npr1 function in the absence of Pmr1. Together, our results establish that Pmr1-dependent Ca2+ and/or Mn2+ ion homeostasis is necessary for TOR signaling.

The TOR (target of rapamycin) protein kinase in Saccharomyces cerevisiae exists in two complexes called TORC1 and TORC2, and rapamycin inhibits only TORC1 (1). Rapamycin-treated cells behave as if they are starved for nitrogen (for reviews, see refs. 2 and 3), implicating TORC1 in growth regulation in response to nutrients. Both incubation of cells in poor nitrogen sources or addition of rapamycin to cells in rich medium induce rapid and overlapping changes in gene expression and activity of nutrient transporters. Both stimuli cause derepression of genes in pathways for ammonia and scavenging nitrogen compounds. Derepression is caused by activation of GATA transcription factors Gln-3 and/or Gat1 (2, 3). Highly specific amino acid transporters are replaced with general amino acid transporter (Gap1). Ammonium transporters (e.g., MEP2; ref. 4) as well as transporters that scavenge nitrogen (e.g., DAL5) are also derepressed. Inhibiting TORC1 derepresses the transcription factor Gln-3 and induces its translocation to the nucleus (5). In the nucleus, Gln-3 induces a subset of nitrogen-repressed genes, including GAP1 and MEP2 (2).

The Ser/Thr-protein kinase encoded by NPR1 (6) positively regulates expression of Gap1 and Mep2 posttranscriptionally; reciprocally, Npr1 negatively regulates specific amino acid transporters (e.g., Tat2) (7, 8). Dephosphorylation of Npr1 in response to rapamycin or nitrogen starvation requires phosphatase Sit4 (9), as does the nuclear translocation of Gln-3 (5). Npr1 negatively regulates Gln-3 in rich medium, but this process is Sit4-independent (10). GAP1 is minimally expressed under nutrient-rich conditions, and any synthesized Gap1 is sorted from the Golgi to the vacuole for destruction in a ubiquitin-dependent pathway (8, 11). In npr1Δ cells, plasma membrane sorting of Gap1 no longer occurs, and the permease is sorted constitutively from the Golgi to the vacuole (8). Npr1 is also required to stabilize Gap1 at the plasma membrane in nitrogen-poor media (8). Npr1 is thought to regulate the sorting and expression of Mep2 similarly to Gap1 (12, 13).

Here, we identify pmr1 in a screen of the genome-deletion strain set in S. cerevisiae for rapamycin resistance. Pmr1 is a Golgi-localized ATPase that transports Ca2+ and Mn2+ ions from the cytoplasm into the Golgi, and thus it functions in the secretory pathway also used by permeases (14). We present data suggesting that PMR1 is a negative regulator of TORC1 activity. We show that Pmr1 modulates the localization of the Gln-3 transcription factor even in the absence of Npr1, suggesting that TOR may regulate Gln-3 localization in an Npr1-independent fashion. We show that Gap1 is localized to the plasma membrane and that it is active in cells lacking Pmr1, suggesting that Golgi-associated Npr1 activity is elevated in a pmr1 mutant. Finally, we show that Npr1 is cytoplasmic and that a portion localizes to the Golgi.

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Results

Deleting PMR1 Confers Rapamycin Resistance.

We identified a pmr1 mutant in a genome-wide screen for rapamycin resistance by using the haploid-deletion collection (≈4,500 nonessential deletion mutants). We scored for rapamycin resistance after growing all deletion mutants individually on YPD medium containing 20 ng/ml and 100 ng/ml rapamycin. We identified fpr1 (FKBP12) in the screen, and it confers rapamycin resistance as expected (15) (Fig. 1 A). In total, 18 mutants were found to be resistant on medium containing up to 100 ng/ml rapamycin, including pmr1Δ (Table 2, which is published as supporting information on the PNAS web site). PMR1 (YGL167C) overlaps HUR1 (YGL168W), and pmr1Δ eliminates the function of both genes (Fig. 1 A). Rapamycin resistance was recessive in a heterozygous diploid (not shown), and it was complemented in a haploid by a YCp PMR1 plasmid (lacking the HUR1 promoter) but not by a YCp HUR1 plasmid (Fig. 1 B). Therefore, rapamycin resistance in the pmr1Δ strain results from a loss of Pmr1. PMR1 deleted in two genetic backgrounds, BY4741 and TB50a, conferred rapamycin resistance (Fig. 1 A). A pmr1Δ ura3Δ strain (YSH3) isogenic with BY4741 was also rapamycin-resistant (not shown).

Fig. 1.
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Fig. 1.

Deletion of pmr1Δ confers rapamycin resistance. (A) The pmr1Δ mutant is rapamycin-resistant in yeast strains BY4741 (YGD3) and TB50a (LJ25-1A). Wild-type and pmr1Δ cells were grown to mid-logarithmic phase, serially diluted, and spotted onto YPD or YPD plus 100 ng/ml rapamycin. The HUR1 gene overlaps PMR1 at the 3′ end. (B) The rapamycin resistance of pmr1Δ (YGD3) is complemented by pPMR1 (pKC21) but not by pHUR1 (HUR1-pR316). Cells were streaked onto SC-ura and SC-ura containing 20 ng/ml rapamycin. (C) The pmr1Δ yeast strains YR122 and YR1234 [derivatives of W303 (YR98) and AA255 (YR401)] are rapamycin-hypersensitive. Cells were grown to mid-logarithmic phase, serially diluted, and spotted on YPD and YPD containing 20 ng/ml rapamycin. (D) Rapamycin-hypersensitive strains of pmr1Δ (YR122 and YR1234) (14, 16) become resistant when transformed with SSD1-V allele in pRS316 (19). Cells were grown to mid-logarithmic phase, serially diluted, and spotted on YPD and YPD containing 20 ng/ml rapamycin.


Interestingly, two pmr1 strains, YR122 and YR1234 (14, 16), in different genetic backgrounds (W303 and an S288-C derived background AA255) were rapamycin-hypersensitive compared with wild-type (Fig. 1 C). W303 contains the loss-of-function ssd1-d mutation (17). SSD1 is named for suppressor of SIT4 deletion (18), and Sit4 functions in TORC1 signaling (5, 9). SSD1-V modulates phenotypes associated with deletion of TCO89, which encodes a component of TORC1 (19). To determine whether the rapamycin hypersensitivity was the result of SSD1, both W303- and AA255-derived strains were transformed with a YCp SSD1-V plasmid (19). SSD1-V conferred rapamycin resistance similar to strain LJ25–1A (pmr1Δ in TB50a) (Fig. 1 D). We conclude that variation in rapamycin sensitivity between these strains is due, at least in part, to alleles of SSD1.

PMR1 Is a Negative Regulator of TORC1.

Deletion of NPR1 or GLN3 increases resistance to rapamycin (5, 9, 20), and this increase was confirmed by isolation of npr1Δ as well as gln3Δ in our screen (Table 2). Rapamycin resistance of the pmr1Δ npr1Δ double mutant was the same as the resistance of either single mutant (Fig. 2 C), suggesting that Pmr1 functions in the same pathway as Npr1 and Gln-3 with respect to rapamycin resistance. Excess expression of either Npr1 or Gln-3, by using YEp plasmids, partially suppressed the rapamycin resistance of pmr1Δ (Fig. 2 A and B). A YEp plasmid encoding the kinase-dead allele of Npr1 (Npr1-kd) failed to suppress the rapamycin resistance of pmr1Δ, showing that Npr1 kinase activity is necessary for suppression. These data suggest that PMR1 functions within the TOR pathway, upstream from both NPR1 and GLN3. We suspect that there is partial suppression of rapamycin resistance because both Npr1 and Gln-3 are subject to inhibitory posttranslational modifications, even when the proteins are overexpressed.

Fig. 2.
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Fig. 2.

PMR1 regulates TORC1 signaling. (A) Overexpression of Npr1 kinase (pAS103) (9), but not kinase-dead mutant (D579E, D565G) (35) suppresses rapamycin resistance of pmr1Δ (YGD3). Cells were grown to mid-logarithmic phase, serially diluted, and spotted on SC-ura and SC-ura containing 100 ng/ml rapamycin. (B) Overexpression of GLN3 (pHAC181-GLN3) suppresses rapamycin resistance of pmr1Δ (YGD3). PMR1 is in a YCP plasmid, pHAC111. Cells were spotted on SC-leu and SC-leu containing 100 ng/ml rapamycin. (C) Double mutant npr1Δ pmr1Δ (YGD12) rapamycin resistance is the same as pmr1Δ (YGD3). Cells were grown to mid-logarithmic phase, serially diluted, and spotted on YPD and YPD containing 20 ng/ml rapamycin. (D) Double mutant pmr1Δ lst8-1 (YGD5) has wild-type rapamycin resistance. Cells were grown to mid-logarithmic phase, serially diluted, and spotted on YPD and YPD containing 20 ng/ml rapamycin. Plates were incubated at 30°C for 2 days and 23°C for 5 days. (E) LST8 (pRS313-LST8) complements pmr1Δ lst8-1 and lst8-1 (YGD5, YGD4) mutant phenotype. Cells were spotted on SC-his medium (without and with 100 ng/ml rapamycin) and incubated for 7 days at 23°C.


We tested a pmr1Δ lst8-1 double mutant to determine how PMR1 functioned relative to TOR complexes. Lst8 is a Golgi-localized, essential WD40 repeat containing protein that functions as a positive regulator in both TORC1 and TORC2 (21). lst8-1 is a hypomorphic allele (partially functional) that is rapamycin-hypersensitive (Fig. 2 D) (22). The pmr1Δ lst8-1 double mutant reciprocally suppressed the rapamycin phenotypes of the lst8-1 and the pmr1Δ single mutants. The double mutant had wild-type rapamycin sensitivity (Fig. 2 D). The rapamycin hypersensitivity of lst8-1 is the result of Pmr1 because removing Pmr1 restores wild-type sensitivity (Fig. 2 D). The reciprocal suppression was caused by LST8 because the pmr1Δ lst8-1 strain transformed with a YCP LST8 plasmid was rapamycin-resistant, like a pmr1Δ mutant (Fig. 2 E). The reciprocal suppression suggests that Lst8 and Pmr1 function in opposition with respect to the rapamycin response. We infer that Pmr1 opposes Lst8 in TORC1 signaling and that it is a negative regulator of the TOR signaling pathway.

We used two measures of the Gln-3 transcription factor to show that TORC1 activity was increased in vivo in pmr1Δ cells. In wild-type cells, during rapamycin treatment, Gln-3 is translocated into the nucleus (7). Therefore Gln-3 localization is a measure of TORC1 activity. When TORC1 activity is high, Gln-3 is cytoplasmic; and when TORC1 activity is low, Gln-3 is nuclear. We analyzed the cellular localization of GFP-tagged Gln-3 in wild-type and pmr1Δ cells. In wild-type cells, Gln-3-GFP is cytoplasmic, and it is translocated into the nucleus during rapamycin treatment as expected (Fig. 3 A Top). Gln-3-GFP was largely cytoplasmic in the pmr1Δ cells when grown in the absence of rapamycin (Fig. 3 A second from Top, large arrow). During rapamycin treatment, Gln-3-GFP was incompletely translocated into the nucleus in pmr1Δ cells (Fig. 3 A second from Top, small arrow). Gln-3-GFP localization was quantified (n = 50–75) and categorized as diffuse (cytoplasmic), partially nuclear, and completely nuclear (Fig. 3 B). Gln-3 was predominantly in the nucleus in 60% of wild-type cells and in 20% of pmr1Δ cells after rapamycin treatment. These data suggest that there is increased TORC1 activity in a pmr1Δ mutant. To confirm that there was less active Gln-3-dependent transcription in pmr1Δ cells, we used Mep2 as a reporter for Gln-3 activity. Mep2 is a member of a family of ammonium transporters, and it is a high-affinity NH4 + transporter, and MEP2 expression is directly regulated by Gln-3 (4). We used a MEP2-LacZ fusion as a reporter of Mep2 expression, and we found that it was induced in the presence of rapamycin in wild-type cells, when Gln-3 was nuclear. However, it was not induced by rapamycin in pmr1Δ cells, suggesting less Gln-3 activity in the nucleus (Fig. 3 C). The Gln-3 localization and Mep2 expression data suggest that there is partial TORC1 activity in pmr1Δ cells treated with rapamycin, consistent with the genetic data that Pmr1 is an inhibitor of TORC1.

Fig. 3.
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Fig. 3.

Loss of Pmr1 suppresses Gln-3 activation. (A) Gln-3 is partially translocated into the nucleus in pmr1Δ (YGD3) and in pmr1Δ npr1Δ (YGD12) in rapamycin (30 min). Wild-type (BY4741), pmr1Δ (YGD3), npr1Δ (YGD11), and pmr1Δ npr1Δ (YGD12) yeast cells containing pGFP-Gln-3 were visualized by differential interference contrast (DIC) microscopy, DAPI, and GFP. Gln-3 is nuclear in BY4741 (wild-type; WT) in rapamycin. The large arrow in pmr1Δ (−rapamycin) points to a cell with cytoplasmic localization, and the smaller arrow (+rapamycin) points to a cell with partial nuclear localization. (B) Quantification of Gln-3-GFP localization (nuclear, partially nuclear, or cytoplasmic) in wild-type (BY4741), pmr1Δ (YGD3), npr1Δ (YGD11), and pmr1Δ npr1Δ (YGD12) cells with and without 100 ng/ml rapamycin (30 min). Fifty to seventy-five cells were counted, and the graph is the percent of cells showing different localizations. (C) Gln-3-dependent MEP2 expression is reduced in rapamycin in pmr1Δ (YGD3) compared with wild-type and in pmr1Δ npr1Δ (YGD12) cells compared with npr1Δ (YGD11). Each value is an average of duplicates.


Nuclear localization of Gln-3 is negatively regulated by Npr1 (10). Gln-3-GFP was constitutively localized to the nucleus in npr1Δ cells, confirming that Npr1 is required to keep Gln-3 in the cytoplasm. Gln-3 was predominantly nuclear in npr1Δ cells when grown in rich medium in the absence of rapamycin (Fig. 3 A third from Top). We analyzed the localization of Gln-3-GFP in cells lacking Npr1 and Pmr1. Interestingly, Gln-3 was mostly cytoplasmic in pmr1Δ npr1Δ cells when grown in rich medium in the absence of rapamycin, suggesting that nuclear localization is not completely dependent on Npr1. This result agrees with recent publications showing that the effect of Npr1 on nuclear localization of Gln-3 can be modulated depending on the nitrogen source (13, 23). The amount of nuclear Gln-3 increased in response to rapamycin in pmr1Δ npr1Δ cells (Fig. 3 A Bottom). We confirmed the response by using the MEP2-LacZ fusion reporter. MEP2 transcription was high in npr1Δ cells correlating with high levels of Gln-3 in the nucleus (Fig. 3 C). The amount of transcription was greatly reduced in pmr1Δ npr1Δ cells (Fig. 3 C). The phenotype of the double mutant as assayed by Gln-3-GFP localization and MEP2-LacZ suggests that pmr1Δ is epistatic to npr1Δ (Fig. 3 and Table 3, which is published as supporting information on the PNAS web site). We interpret this result to mean that increased TORC1 activity in the absence of Pmr1 can sequester Gln-3 to the cytoplasm in an npr1Δ mutant, and this sequestration can be partially overcome by rapamycin.

PMR1 Regulates Gap1 Sorting.

Gap1 protein sorting in the trans-Golgi is regulated in response to the nitrogen source. In poor-nitrogen conditions such as urea, Gap1 is sorted to the plasma membrane; and in rich-nitrogen conditions, such as glutamate or ammonia, Gap1 is routed to the vacuole, where it undergoes ubiquitylation and is degraded (24). Mutations in ubiquitylation genes (bul1, bul2, rsp5 or doa4) cause Gap1 to be sorted to the plasma membrane, suggesting polyubiquitylation as a sorting signal for Gap1 (11, 24). In npr1Δ cells, plasma membrane sorting of Gap1 no longer occurs, and the permease is sorted from the Golgi to the vacuole (8).

Npr1 kinase regulates amino acid permease sorting and expression (8, 9). When TORC1 is active, Npr1 activity is low. We expected that Npr1 activity would be low in pmr1Δ because Pmr1 negatively regulates TORC1. Therefore, we expected that there would be little Gap1 at the plasma membrane. We localized Gap1-GFP and found that there was increased sorting and stabilization of Gap1 at the plasma membrane and a marked reduction of the intracellular pool of Gap1-GFP in pmr1Δ cells in urea-containing medium (>90%, n = 100) (Fig. 4 A). We measured Gap1 activity by citrulline uptake, and we confirmed that Gap1 activity was significantly higher in pmr1Δ cells compared with wild-type cells (Fig. 4 B). These data are consistent with hyperactive Npr1, causing altered sorting and stabilization of the permease and suggesting that Pmr1 may negatively regulate Npr1 activity in the Golgi.

Fig. 4.
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Fig. 4.

Pmr1 regulates Gap1 sorting. (A) Gap1-GFP is present almost exclusively at the plasma membrane in pmr1Δ (YSH3) cells. (B) 14C-citrulline uptake rate is higher in pmr1Δ (YSH3) cells compared with wild-type (YSH1). (C) Npr1-GFP partially colocalizes in Golgi. Yeast cells (TB50a) containing YCp Npr1-GFP and YIp Sec7p-dsRed (YGD18) were visualized by using differential interference contrast (DIC) microscopy, green (GFP), blue (DAPI), and red (dsRed) filters. A magnified image of one of the cells is in the lower part of the panel, and the arrows identify Npr1-GFP (green) that colocalizes with Sec7-dsRed (red). The merge of the magnified image is on the right. (D) Model of Pmr1 function in the TORC1 cell growth regulatory pathway.


Npr1-GFP Colocalizes with Golgi.

The intracellular localization of Npr1 is unknown. De Craene et al. (ref. 8 and unpublished observations) suggested that Npr1 was cytoplasmic with punctate intracellular structures. We localized Npr1 in wild-type cells by coexpressing Npr1-GFP and Sec7-dsRed (25). NPR1-GFP complemented rapamycin resistance of an npr1Δ mutant (not shown), indicating that it was a functional fusion protein. Npr1-GFP was diffusely cytoplasmic, with concentration in an intracellular compartment that appeared punctate. Sec7 is a peripheral protein of the late Golgi, and it cycles between a vesicle coat complex and soluble cytoplasmic form (26). Npr1-GFP stained diffusely in the cytoplasm, but the portion that was punctate was colocalized with Sec7-dsRED in 80% of the cells, n = 100 (Fig. 4 C Bottom, arrows). Images of cells containing Sec7-dsRED and photographed through 470- to 525-nm filters (GFP) showed no detectable signal, confirming that we were detecting Npr1-GFP localized in the Golgi (Fig. 5, which is published as supporting information on the PNAS web site). The localization of Npr1-GFP was unaffected by rapamycin treatment (data not shown). Npr1-GFP also localized in the Golgi in pmr1Δ cells, and localization was not altered by rapamycin treatment (not shown). Thus, our data establish a pool of Npr1 in close association with Golgi positioned to phosphorylate and/or interact with Gap1 in transit or to be regulated by Pmr1.

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Discussion

We identify PMR1 as a gene whose loss of function causes resistance to rapamycin. Pmr1 is required for proper function of the secretory pathway, N- and O-linked glycosylation of proteins, protein sorting, and endoplasmic reticulum-associated protein degradation (16). PMR1 genetically interacts with secretory pathway Ras-like small GTPases such as YPT1 (14) and the Rab7 homolog YPT7 (27). PMR1 deletion mutants secrete heterologous proteins at levels 5- to 50-fold higher than wild-type cells (14).

Our studies place Pmr1 in TORC1 signaling, shown schematically in a model (Fig. 4 D). The rapamycin resistance of pmr1Δ can be explained because TORC1 is negatively regulated by Pmr1. We found that Gap1 was stabilized and Gln-3 inhibited in pmr1 strains, findings that suggest up-regulation of Npr1 activity. We had expected that Npr1 activity would be low in pmr1 because Pmr1 negatively regulates TORC1, and TORC1 inhibits Sit4-dependent activation of Npr1 (9). Loss of Pmr1 may increase Npr1 activity in the Golgi by a mechanism independent of Sit4; alternatively, changes in the concentration of ions in the secretory compartments may bypass the requirement for Npr1 for Gap1 stabilization. The epistasis analyses between NPR1 and PMR1 suggest that the nitrogen starvation signals required for derepression of Gln-3 are transduced by Pmr1 through TORC1 and Npr1.

We identify a pool of Npr1 that colocalizes with Golgi. The localization of Tor1 has been studied in yeast by biochemical fractionation (28) and by immunogold electron microscopy (29). Tor1 is peripherally associated with membranes with a portion in membranes having density that is reported to be similar to plasma membrane and a portion in a distinct, unidentified pool (28). The localization of mTOR has been controversial, but mTOR may be associated with the endoplasmic reticulum and Golgi (30). It is thought that Gap1 is phosphorylated en route to the plasma membrane by Npr1, which protects Gap1 from degradation (31). We propose that Pmr1 has a dual role in negatively regulating TORC1 activity and negatively regulating Npr1 in the Golgi.

Npr1 is a key regulator in yeast, but it has no identified homolog by similarity searches in mammals. Localizing Npr1 to the Golgi is an important step toward identifying a functional homolog of Npr1 in mammalian cells. Protein kinase D can associate with Golgi or the plasma membrane, and it may be a candidate for a functional homolog of Npr1 in man (32). Furthermore, proteomic approaches are identifying a limited number of protein kinases associated with the Golgi (33).

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Materials and Methods

Strains.

For a complete list and genotypes of yeast strains used in this work, see Table 1 (most strains are congenic with BY4741; some strains are congenic with TB50a, including LJ25-1A, YGD18, and YGD21). The EUROSCARF systematic deletion collection for nonessential genes is in BY4741. To delete PMR1 in TB50a, pmr1Δ::kanMX4 fragment was amplified from genomic DNA of BY4741 pmr1Δ [primers PMR1-KOF (5′-GACTTCGCCTCGTTTTGG-3′) and PMR1-G418-KO (5′-GCACGAACAATGTTTAACTTATGCTCAGGCGTAGCACGAGCAAAAATATTATCGATGAATTCGAGCTCG-3′)] and transformed into TB50, and tetrads were dissected. pmr1Δ npr1Δ was made by crossing LJ21-6D with YGD10, and tetrads were dissected. YGD10 (npr1Δ::natMX4) was made by transforming YGD11 from the deletion set with BglII–SacI-digested pAG25 (34). pmr1Δ npr1Δ is G418- and NAT-resistant. CKY526 (22) (gift from Chris Kaiser, Massachusetts Institute of Technology, Cambridge, MA) has the lst8-1 mutation (L300S) in S288C. pmr1Δ lst8-1 double mutants were made by crossing LJ21-6D with CKY526; tetrads were dissected, and the rapamycin phenotype was complemented with pRS313-LST8. The double mutants contain the lst8-1 mutation at the LST8 locus, verified by DNA sequencing of PCR products.

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Table 1.

Yeast strains used in this study


Plasmids.

pHUR1 is the HUR1 gene (≈1.1 kb) cloned into the PstI–SalI sites of pRS316. HUR1 is PCR-amplified from yeast genomic DNA with 5′ end PstI primer (HUR1-F) 5′-AAAACTGCAGTCATTGCGAAGGCGATG-3′ and 3′ end SalI primer (HUR1-R2) 5′-ACGCGTCGACGGTGTGGAACCTGTTGATCATG-3′. The PCR product was digested with PstI–SalI. HA-tagged NPR1 and NPR1-KD were in pRS426 (SacI–KpnI sites) and were subcloned from pHA-NPR1 and p416, respectively (35) (gift from Yu Jiang, University of Pittsburgh, Pittsburgh, PA). The KD mutant is mutated at catalytic site residues D565G, D579E. GLN3 was subcloned into pHAC181 (BamHI–PstI) from pTB400 (gift from Michael Hall, Biozentrum, Basel, Switzerland). Npr1 and Gln-3 fusion proteins are expressed from pGFP33 (gift from Michael Hall), and they were cloned from wild-type BY4741 yeast genomic DNA by PCR (NPR1, 2.9-kb XbaI–SalI fragment; GLN3, 2.9-kb PstI–SphI fragment). The primers were: NPR1-GFP forward, 5′-GCTCTAGATGCACGAAAAGCTGTACGAG-3′; NPR1-GFP reverse, 5′-ACGCGTCGACTTGATTATTTTGCTTTTTCT-3′; GLN3-GFP forward, 5′-AACTGCAGCCAAAGAAGAGGACCTCG-3′; GLN3-GFP reverse, 5′-CGGCATGCTATACCAAATTTTAACCAATC-3′. YIplac204-T/C-Sec7-DsRed.T4 (25) (gift from Ben Glick, University of Chicago, Chicago, IL) was linearized with Bsu36I for integration into the trp1 locus of TB50a. pKC21 is PMR1 in pRS316 (gift from Kyle Cunningham, The Johns Hopkins University, Baltimore, MD). PMR1 was amplified from S. cerevisiae genomic DNA with KpnI–SphI primers (≈3.8 kb) and cloned into pHAC111 (gift from Michael Hall). The primers were: PMR1 forward, 5′-GGGGTACCATCAGTGTGAGTACACTGC-3′; and PMR1 reverse, 5′-ACATGCATGCAACATTTGAGAA- ATACGTTGAGTC-3′). LST8 was cloned from yeast genomic DNA with BamHI primers into pRS313 (36). The primers were: GD110 (LST8 forward, 5′-CACCAGGATCCAAAGCTTGGACCACC-3′; GD111 (LST8 reverse), 5′-TGCAAGGATCCGGTTCAATAGTAGATATTAT-3′. SSD1-V (JK9–3da allele) is in pRS316 (CEN, URA3) (pPL092; gift from Ted Powers, University of California, Davis, CA) (19).

Media.

Rapamycin (Sigma, St. Louis, MO) was in 90% ethanol/10% Tween 20. Yeast medium was prepared according to standard methods (37). BY4741 diploids were sporulated in 1% potassium acetate and 0.005% zinc acetate liquid medium for 7–10 days. Yeast transformations and tetrad dissections were done as described in refs. 37 and 38. pmr1Δ lst8-1 double mutants were grown on rapamycin-containing plates for 5 days at 23°C, and they were serially diluted 5-fold, from a starting A 600 of 1.0. Yeast cultures were serially diluted 10-fold from a starting A 600 of 1.0 or 0.5, and plates were incubated at 30°C for 3–4 days. To measure MEP2 expression, WT and pmr1Δ cells containing YCpMEP2-LacZ (4) were grown in SC-ura overnight and then in YPD to A 600 0.1–0.4; they were then divided, and one half was treated with rapamycin (100 ng/ml) for 30 min. β-Galactosidase assays with o-nitrophenyl-β-d-galactopyranoside as substrate were performed as described (Yeast Protocol Handbook, Clontech, Mountain View, CA) and β-galactosidase units were calculated with time in hours.

Microscopy.

For Npr1 and Gln-3 localization, yeast cells were visualized by using an Axiovert 200 inverted fluorescence photomicroscope (Zeiss, Gottingen, Germany) equipped with a Hamamatsu digital CCD camera with DICIII, Endow GFP (exciter, 470 ± 20 nm; emitter, 525 ± 25 nm), DAPI (exciter, 360 ± 20 nm; emitter, 460 ± 25 nm) or dsRed (exciter, 546 ± 6 nm; emitter, 605 ± 37.5 nm) filters (Chroma Technology Corp., Brattleboro, VT), with a 100× oil immersion plan-apochromat objective (Zeiss). Images were taken with Openlab 3.1.4 software (Improvision, Lexington, MA). Yeast cells were fixed in 4% paraformaldehyde/3.4% sucrose; they were then washed and resuspended in potassium phosphate/sorbitol buffer. Cells were vortexed for 2 min at high speed before being placed on acid-washed, polylysine-coated glass slides. Vectashield mounting medium containing DAPI (Vector Laboratories, Burlingame, CA) was used with 22 × 22-1 cover glass (finest; Fisher, Pittsburgh, PA). For Golgi labeling, Sec7-dsRed fusion was integrated in yeast at trp1 locus. Cells were grown in SC-ura medium.

For Gap1p localization, yeast strains (YSH1, YSH3) expressing Gap1-GFP (pCK230; gift from Chris Kaiser) were grown overnight in minimal medium containing urea as a nitrogen source. One milliliter of the culture was harvested at an A 600 between 0.1 and 0.4 and resuspended in ≈20 μl of the same medium containing 10 mM sodium azide. The resuspension (2.4 μl) was dropped on a glass slide, and 2.4 μl of 1.6% low-melt agarose was added and mixed before the coverslip was placed. All images were taken on an Axiophoto fluorescence microscope (objective lens 100×; NA 1,1) (Zeiss) with identical exposure times.

Amino Acid Uptake.

For Gap1p activity, yeast strains were cultured in SD medium containing 0.2% urea as the sole nitrogen source. Cultures were grown to an A 600 of 0.1- 0.4 and collected by filtration. Amino acid uptake assays were performed as described in ref. 24.

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Acknowledgments

We thank Michael Hall along with Robbie Loewith and Ben Glick for advice, reagents, and comments on the manuscript and Linghuo Jiang and Shuang Niu for the screen identifying pmr1. We are indebted to many (see Materials and Methods) for kindly providing reagents. We are grateful to Hans Rudolph for sharing unpublished data on rapamycin hypersensitivity of YR122 and YR1234. We thank Chris Yellman, Thurl Harris, and the Burke and Sturgill laboratory members for useful suggestions and reagents and Katrina Clines for technical assistance. This work was supported by National Institutes of Health Grants GM62890 (to T.W.S.) and GM40334 (to D.J.B.).

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Footnotes

  • To whom correspondence should be addressed. E-mail: tws7w{at}virginia.edu

  • Author contributions: G.D., S.B.H., D.J.B., and T.W.S. designed research; G.D., D.R., and S.B.H. performed research; G.D., S.B.H., D.J.B., and T.W.S. analyzed data; and G.D., S.B.H., D.J.B., and T.W.S. wrote the paper.

  • ↵ ‡Present address: Developmental and Molecular Pathways, Novartis Institutes for Biomedical Research, Novartis AG, CH 4002 Basel, Switzerland.

  • The authors declare no conflict of interest.

  • This article is a PNAS direct submission.

  • Abbreviation:
    TOR,
    target of rapamycin.
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