Eukaryotic cell proliferation is controlled by growth factors and essential nutrients. In their absence, cells may enter into a quiescent state (G0). In Saccharomyces cerevisiae, the conserved protein kinase A (PKA) and rapamycin‐sensitive TOR (TORC1) pathways antagonize G0 entry in response to carbon and/or nitrogen availability primarily by inhibiting the PAS kinase Rim15 function. Here, we show that the phosphate‐sensing Pho80–Pho85 cyclin–cyclin‐dependent kinase (CDK) complex also participates in Rim15 inhibition through direct phosphorylation, thereby effectively sequestering Rim15 in the cytoplasm via its association with 14‐3‐3 proteins. Inactivation of either Pho80–Pho85 or TORC1 causes dephosphorylation of the 14‐3‐3‐binding site in Rim15, thus enabling nuclear import of Rim15 and induction of the Rim15‐controlled G0 program. Importantly, we also show that Pho80–Pho85 and TORC1 converge on a single amino acid in Rim15. Thus, Rim15 plays a key role in G0 entry through its ability to integrate signaling from the PKA, TORC1, and Pho80–Pho85 pathways.
Eukaryotic signal transduction pathways convert extracellular cues such as growth factors, hormones, and/or nutrients to intracellular signals which, following proper integration, determine the decision between cell proliferation and entry into a nondividing resting state, termed quiescence (G0). In the yeast Saccharomyces cerevisiae, the PKA and TORC1 signaling pathways, which positively regulate cell proliferation in response to nutrient availability, are key determinants of G0 entry (Thevelein and de Winde, 1999; Gray et al, 2004; Martin and Hall, 2005). As a consequence, inactivation of either of these major pathways results in a G0‐like growth arrest phenotype, even in the presence of abundant nutrients. Recent evidence suggests that the signal flow through both pathways is integrated at various levels. For instance, TORC1 depletion was proposed to favor the formation of an inactive nuclear PKA/Bcy1 holoenzyme (Schmelzle et al, 2004). In addition, both PKA and TORC1 converge on common target proteins, such as the stress response transcription factors Msn2/4, as well as Rim15 (Schneper et al, 2004). Rim15 is a distinct member of the PAS kinase family, which functions as a key controller of many aspects of the G0 program (including G1 cell cycle arrest, glycogen and trehalose synthesis, and activation of Gis1‐ and/or Msn2/4‐dependent transcription (e.g. of HSP26 and GRE1; Reinders et al, 1998; Pedruzzi et al, 2000, 2003; Cameroni et al, 2004).
Entry into a G0‐like state can also be triggered by phosphate starvation (Lillie and Pringle, 1980), albeit the corresponding regulatory mechanisms are largely unknown (Gray et al, 2004). The primary nutrient‐signaling kinase that orchestrates the phosphate starvation response is Pho85, a CDK that associates with a family of 10 cyclins, each of which can potentially direct Pho85 to different target substrates (Carroll and O'Shea, 2002). The best‐studied partner of Pho85 is the cyclin Pho80. It has previously been shown that the Pho80–Pho85 cyclin–CDK complex regulates the phosphate starvation response by controlling both the localization and activity of the transcription factor Pho4. Pho4, in turn, activates transcription of phosphate acquisition genes (Carroll and O'Shea, 2002). Interestingly, several studies indicate that Pho85 negatively controls the expression of an additional set of genes (including glycogen and trehalose synthesis, oxidoreductive stress, and protein folding genes) that are typically induced under glucose‐limiting conditions prior to entry into G0 (DeRisi et al, 1997; Timblin and Bergman, 1997; Ogawa et al, 2000; Carroll et al, 2001; Nishizawa et al, 2004). These findings suggest that proper execution of the G0 program also includes integration of the Pho85‐mediated phosphate signal.
The work presented here pinpoints Rim15 as a key target of the Pho85 signaling pathway. Results from physiologic, genetic and biochemical studies indicate that Rim15 is a Pho80–Pho85 kinase substrate and that the resulting phosphorylation of Rim15 favors its cytoplasmic retention via association with the 14‐3‐3 protein Bmh2. Our data further demonstrate that this regulation is important for proper control of G0 entry in response to phosphate availability. Interestingly, inactivation of either Pho80–Pho85 or TORC1 leads to dephosphorylation of the 14‐3‐3 binding site in Rim15, suggesting that both signaling pathways converge on a single amino acid in Rim15 to regulate its nucleocytoplasmic distribution.
Results and discussion
Protein kinase activity of Rim15 is critical for its nuclear export
As part of our continued efforts to define in detail the molecular mechanism of Rim15 function, we asked whether the Rim15 kinase activity itself plays a role in its subcellular localization. To address this question, we analyzed the localization of kinase‐inactive GFP‐Rim15K823Y and GFP‐Rim15C1176Y fusion proteins (see Materials and methods for further details) in rim15Δ cells. We found that GFP‐Rim15K823Y and GFP‐Rim15C1176Y, but not GFP‐Rim15, accumulated in the nuclei of rapamycin‐treated or glucose‐limited rim15Δ cells (Figure 1A). Furthermore, when such rapamycin‐treated or glucose‐limited rim15Δ mutant cells were respectively re‐supplied with nutrient‐rich medium or released from the rapamycin block (both in the presence of the translation inhibitor cycloheximide), GFP‐Rim15K823Y and GFP‐Rim15C1176Y, but not GFP‐Rim15, remained predominantly in the nucleus (data not shown). The simplest interpretation of these results is that the kinase activity of Rim15 is required for its efficient export from the nucleus. Moreover, from a collection of importin β‐related, nuclear‐transport‐receptor mutants tested, only the msn5Δ mutant showed both GFP‐Rim15C1176Y and GFP‐Rim15K823Y constitutively localized in the nucleus (Figure 1B). Thus, our data indicate that the kinase activity of Rim15 is needed for its efficient, Msn5‐driven nuclear export and show that GFP‐tagged kinase‐inactive Rim15 variants are useful tools in the study of nutrient‐regulated cytoplasmic retention and/or nuclear import of Rim15.
Cytoplasmic retention of GFP‐Rim15 depends on its association with 14‐3‐3 proteins
Rim15 is a distant member of the NDR family of AGC kinases, which share the unique feature of harboring an insert of at least 30 amino acids between the protein kinase subdomains VII and VIII (Tamaskovic et al, 2003; Cameroni et al, 2004). Although the function of this insert segment is not well understood, at least in the case of NDR1, it is known to contain a nonconsensus nuclear localization signal (Tamaskovic et al, 2003). Interestingly, the corresponding kinase insert in Rim15 contains neither a consensus nor a nonconsensus NDR1 nuclear localization signal, but instead harbors (flanking amino‐acid residue Thr1075 [T1075]) the single high‐stringency 14‐3‐3‐binding motif in Rim15 (RSXpS/TXP; Figure 2A; Yaffe et al, 2001).
Several observations point to a role for the yeast 14‐3‐3 proteins Bmh1/2 in the nutrient‐controlled nuclear exclusion of Rim15, possibly by a similar mechanism as previously described for the transcription factors Msn2/4 and the DYRK‐family kinase Yak1 (Beck and Hall, 1999; Moriya et al, 2001). First, when fused to GST, the kinase insert of Rim15 (Rim15KI) bound Bmh2‐HA3 in co‐immunoprecipitation (co‐IP) experiments (Figure 2B). Importantly, this interaction was largely disrupted by introducing a T1075A mutation in Rim15KI, which abolishes the presumed phosphothreonine (pT) residue critical for regulation of 14‐3‐3‐protein–protein interactions (Yaffe et al, 2001). Second, while GFP‐Rim15C1176Y was rarely found in the nuclei of exponentially growing cells (i.e. in <1% of the cells), introduction of the T1075A mutation or deletion of the entire kinase insert in GFP‐Rim15C1176Y caused the corresponding fusion proteins to constitutively localize in the nuclei of 40 or 68% of the cells, respectively (Figure 2C). The slightly more robust constitutive nuclear localization of GFP‐Rim15C1176Y/ΔKI suggests that additional domains in the kinase insert of Rim15 may participate in the cytoplasmic retention of Rim15 (see below). Third, the switch from galactose to glucose‐rich medium of a strain in which the only source of Bmh protein was provided by BMH2 under the control of the GAL1 promoter resulted in the expected depletion of Bmh2 (Figure 2D), and was paralleled by the nuclear accumulation of GFP‐Rim15C1176Y (Figure 2E). In a strain carrying a genomic wild‐type copy of RIM15, this nuclear localization resulted in the Rim15‐dependent transcriptional activation of GRE1 and HSP26 (Figure 2F; notably, Bmh protein depletion caused a weak, transient activation of HSP26 even in the absence of Rim15, indicating that Bmh proteins regulate HSP26 transcription in part also via a Rim15‐independent process). Interestingly, in this context, we found that 27 of the 76 genes that were reported to be induced following Bmh1/2 depletion (Ichimura et al, 2004), were also included in the set of genes that required Rim15 for induction at the diauxic shift (Cameroni et al, 2004). Taken together, our findings indicate that the association between Bmh1/2 proteins and the pT1075 14‐3‐3‐binding site in Rim15 can account for the cytoplasmic retention of GFP‐Rim15 in exponentially growing cells.
The Pho80–Pho85 cyclin–CDK complex phosphorylates T1075 in Rim15
The T1075 amino acid of Rim15 is not only a critical residue of the 14‐3‐3 binding site, but is also part of a proposed Pho85 consensus phophorylation sequence S/TPXI/L (Figure 2A; O'Neill et al, 1996). This suggested to us that Pho85 may phosphorylate T1075 in Rim15. In support of this assumption, we found that Pho85 physically interacted with Rim15 in co‐IP experiments (Figure 3A), and that Pho85, but not kinase‐inactive Pho85E53A (Nishizawa et al, 1999), phosphorylated Rim15KI in the presence of the cyclin Pho80 (see Materials and methods for identification of Pho80; Figure 3B). In addition, Pho80–Pho85‐mediated phosphorylation was significantly reduced (to 46.0%±6.6) by the introduction of the T1075A mutation in Rim15KI (Figure 3B), indicating that T1075 is a Pho80–Pho85 target in vitro (notably, the residual phosphorylation level in Rim15KI‐T1075A suggests the presence of an additional site[s] in Rim15KI that may be targeted in vitro by Pho80–Pho85). To assess whether Pho80–Pho85 indeed phosphorylates T1075 in Rim15 within cells, we raised antibodies that were highly specific for a pT1075‐containing peptide, as indicated by their ability to recognize GST‐Rim15, but not GST‐Rim15T1075A, or phosphatase‐treated GST‐Rim15 (Figure 3C). Using these specific anti‐pT1075 antibodies, we found that phosphorylation of T1075 in Rim15‐myc13 depends largely on the presence of Pho85 (Figure 3D). Thus, the T1075 residue in Rim15 is also a likely Pho80–Pho85 target in vivo. Taken together with our finding that loss of Pho85 or Pho80 (but not of Pcl1/Pcl2) resulted in the constitutive nuclear localization of GFP‐Rim15C1176Y in 71% and 58% of the cells, respectively (Figure 3E), our data suggest a model in which the Pho80–Pho85 kinase promotes the cytoplasmic retention of Rim15 in exponentially growing cells principally through phosphorylation of the T1075 residue of the 14‐3‐3 site in Rim15.
The Pho80–Pho85 cyclin–CDK complex antagonizes the Rim15‐dependent G0 program
To address the physiological importance of our findings, we next examined the role of Pho85, if any, in the Rim15‐dependent G0 phenotypes (Reinders et al, 1998; Pedruzzi et al, 2003). We observed that loss of Pho80 or Pho85 significantly enhanced GRE1‐lacZ induction (Figure 4A) and trehalose synthesis (Figure 4B) particularly following glucose limitation. Notably, the corresponding higher induction levels, which in the case of pho85Δ mutant cells also correlated with a slightly enhanced G0 survival (Figure 4C), depended to a large extent on the presence of Rim15 (Figure 4A–C). Similarly, glycogen hyperaccumulation in pho85Δ mutants (Timblin et al, 1996) was largely dependent on the presence of Rim15 (Figure 4D). Together with our data presented above, these genetic results suggest a model in which the Pho80–Pho85 cyclin–CDK complex antagonizes the Rim15‐dependent G0 program by regulating the nuclear exclusion of Rim15.
We also found that the levels of both Pho80 and Pho85 drop significantly in the late post‐diauxic shift phase (data not shown), suggesting that the Pho80–Pho85 kinase modulates the Rim15‐dependent G0 program in response to phosphate levels as long as residual carbon sources are available. This conclusion is confirmed by the findings that phosphate starvation resulted in the dephosphorylation of Rim15‐pT1075 (Figure 4E), nuclear accumulation of GFP‐Rim15C1176Y (Figure 4F), and, as recently published, Rim15 target gene expression (Swinnen et al, 2005). Thus, our physiologic, biochemical and genetic data are consistent with a model in which Rim15 represents a physiologically important Pho80–Pho85 cyclin–CDK target.
TORC1 also impinges on pT1075 in Rim15
We have previously shown that the nuclear accumulation of Rim15 is antagonized by the TORC1 pathway, and likely involves Rim15 hyper‐phosphorylation (Pedruzzi et al, 2003). During our analyses of the various kinase‐inactive Rim15 proteins, we observed that their electrophoretic mobility depended on the kinase activity of Rim15 itself and, therefore, likely reflects the autophosphorylation state of Rim15 (data not shown). A simple model could be that rapamycin‐induced hyper‐phosphorylation of Rim15 is a consequence, rather than the cause, of Rim15 nuclear import and may be due to the activation of Rim15 in the presumably low PKA nuclear environment (Griffioen and Thevelein, 2002; Pedruzzi et al, 2003).
How then does TORC1 regulate the nucleocytoplasmic distribution of Rim15? In this respect, we observed that rapamycin treatment caused rapid dephosphorylation of pT1075 in Rim15 (Figure 4E), indicating that TORC1, like Pho80–Pho85, regulates the cytoplasmic retention of Rim15 primarily by controlling the phosphorylation state of its 14‐3‐3‐binding site. The fact that rapamycin treatment does not significantly change the expression pattern of genes controlled by the Pho80–Pho85‐regulated Pho4 transcription factor (e.g. PHO5, PHO11, and PHO12; Hardwick et al, 1999) argues against TORC1 acting upstream of Pho80–Pho85. Thus, TORC1, rather than impinging on Pho80–Pho85, may instead inhibit the activity of a pT1075‐targeting phosphatase(s) and/or activate an alternative T1075‐targeting kinase (Figure 5). In this context, our previous findings indicated that the known TORC1 effectors, including the type 2A‐related protein phosphatase Sit4 and possibly also the type 2A protein phosphatases Pph21 and Pph22 (Di Como and Arndt, 1996; Jiang and Broach, 1999), may not be required for rapamycin‐induced nuclear accumulation of Rim15 (Pedruzzi et al, 2003). Thus, unless these protein phosphatases redundantly control Rim15 function, our data suggest the existence of an additional, yet unidentified TORC1 effector(s) that regulates the phosphorylation state of T1075 in Rim15. Lastly, we would like to point out that TORC1 and Pho80–Pho85 localize to the cytoplasm and nucleus, respectively (Kaffman et al, 1998; Harris and Lawrence, 2003), and these kinases may therefore act on different pools of Rim15 (Figure 5). Our observation that rapamycin treatment and phosphate starvation act synergistically to cause a more complete depletion of GFP‐Rim15C1176Y from the cytoplasm supports this model (Figures 1A and 4F).
A final point remaining to be addressed is our previous finding that nuclear exclusion of Rim15 also requires the yeast PKB/Akt homolog Sch9. Analysis of the kinase insert sequence of Rim15 reveals three sites (S1004, S1044, and T1096), each of which is very similar to the consensus site phosphorylated by PKB/Akt (RXRXXS/T; Alessi et al, 1996) and conforms to a low‐stringency 14‐3‐3‐binding site (Yaffe et al, 2001). Given the dimeric nature of 14‐3‐3 proteins, it is possible that the kinase insert of Rim15 may engage in binding both monomeric subunits within a single 14‐3‐3 protein dimer, as previously shown to be the case for other proteins (Yaffe, 2002). Since our preliminary data indicate that Sch9 can phosphorylate Rim15KI in vitro, it will be of interest to determine whether Sch9 phosphorylates an additional 14‐3‐3 site in Rim15KI, which may act in concert with the 14‐3‐3‐binding site flanking amino‐acid residue T1075 to mediate tandem 14‐3‐3 binding.
In conclusion, we have identified a new regulatory mechanism by which the availability of phosphate modulates the G0 program in yeast. We propose that the PAS kinase Rim15, in addition to integrating carbon and/or nitrogen source signals via the PKA, TORC1, and Sch9 nutrient‐sensory kinases (Pedruzzi et al, 2003), also integrates information on the availability of phosphate via the Pho80‐Pho85 cyclin–CDK complex to properly orchestrate the G0 program, a key developmental process in eukaryotic cells.
Materials and methods
Yeast strains, media and genetic techniques
The rim15Δ strains IP31 and CDV115 and their corresponding wild‐type strains KT1960/1 and W303‐1A, respectively, have been described earlier (Thomas and Rothstein, 1989; Pedruzzi et al, 2003). The msn5Δ strain YBL029, which is isogenic to PAY20 and W303‐1A (Blondel et al, 1999), was a kind gift of Dr M Peter. Polymerase chain reaction (PCR)‐based gene deletions (pcl1Δ∷kanMX2, pcl2Δ∷kanMX2, pho80Δ∷kanMX2, pho85Δ∷kanMX2, and rim15Δ∷kanMX2 transformed into KT1960 and/or KT1961) were carried out as described (Longtine et al, 1998). The corresponding single mutants (all in same isogenic background) were used to construct IP48‐3C (pho85Δ), CDV201‐3B (pho85Δ rim15Δ), CDV237‐10C (pho80Δ), CDV237‐7A (pho80Δ rim15Δ), and CDV237‐9D (pcl1Δ pcl2Δ rim15Δ) by a series of repeated combinatorial mating and sporulation of the resulting diploids. Gene deletions were confirmed by PCR with gene‐specific primers. PCR‐based deletion of RIM15 (using a rim15Δ∷kanMX2 cassette) in strain SL1470 (bmh1Δ bmh2Δ [pGAL1‐BMH2]; described in Gelperin et al, 1995) yielded strain CDV235. The isogenic strains BY236 (wild type), BY714 (pcl1Δ pcl2Δ pcl5Δ pcl9Δ clg1Δ), BY708 (pcl1Δ pcl2Δ pcl5Δ pcl9Δ), BY637 (pcl1Δ pcl2Δ pcl5Δ clg1Δ), BY634 (pcl1Δ plc2Δ), and BY490 (pho80Δ) have been previously described (Measday et al, 1997). Strains were grown at 30°C in standard rich medium with 2% glucose (YPD) or synthetic medium with 2% glucose (SD), 4% galactose (SGal), or 2% raffinose (SRaf) as carbon source (Burke et al, 2000). High‐ and no‐phosphate media were described earlier (Kaffman et al, 1998). Rapamycin was added to the media at a final concentration of 2 μg ml−1. Standard yeast genetic manipulations were used (Burke et al, 2000). DNA was stained with 4,6‐diamidino‐2‐phenylindole (DAPI), which was added to the cultures (4 h prior to fluorescence microscopy) at a concentration of 1 μg ml−1.
Full‐length Rim15, Rim15K823Y, Rim15C1176Y, Rim15C1176Y/T1075A, and Rim15C1176Y/ΔKI (in which the kinase insert domain encompassing amino acids 952–1138 is deleted; ΔKI) were expressed as GFP‐tagged versions under the control of the constitutive ADH1 promoter from low copy number plasmids pFD846, pFD1008, pFD633, pVW1017, and pVW1068, respectively. In addition, full‐length Rim15 and Rim15T1075A were expressed as GST‐tagged versions under the control of the GAL1 promoter from the high copy number plasmids pNB566 and pLC824, respectively. Finally, full‐length Rim15 was also expressed as myc13‐tagged version under the control of the TDH3 promoter from high copy number plasmids pVW904. Plasmids expressing epitope‐tagged variants of full‐length Rim15 were constructed with the QuickChange Site‐Directed Mutagenesis Kit (Stratagene) using appropriate primers that introduced the K823Y‐, C1176Y‐, and/or T1075A‐encoding mutations in the corresponding parent plasmids (pFD846 and pNB566). To express a GST‐tagged version of the kinase insert of Rim15 (Rim15KI) under the control of the tetO7 promoter, an NotI–SalI fragment, containing the 672 nucleotides downstream of and including the GST start codon (Reinders et al, 1998), and a PCR‐generated SalI–PstI fragment, encoding amino‐acid residues 944 to 1149 of Rim15, were coligated with NotI–PstI‐cut pCM184 (Garí et al, 1997), thus creating pVW900 (tetO7‐GST‐RIM15KI). Plasmid pVW902 (tetO7‐GST‐RIM15KI‐T1075A) was obtained by site‐directed mutagenesis (see above) of RIM15KI in pVW900 using appropriate primers that introduced the T1075A‐encoding mutation. For expression of GST‐Rim15KI in Escherichia coli (BL21; Stratagene), a PCR‐generated BamHI–EcoRI fragment, encoding amino‐acid residues 944 to 1149 of Rim15, was ligated into BamHI–EcoRI‐digested pGEX3X (Amersham) to yield pVW995 (Ptac‐GST‐RIM15KI). Plasmid pVW997 (Ptac‐GST‐RIM15KI‐T1075A) was obtained by site‐directed mutagenesis (see above) of RIM15KI in pVW995 using appropriate primers that introduced the T1075A‐encoding mutation. Bmh2‐HA3 was expressed under the control of its own promoter from either a high‐copy number plasmid (pTB419), which was kindly provided by Dr M Hall (Beck and Hall, 1999), or a low copy number plasmid (pCDV994) that was created by cloning the BamHI–HindIII‐digested, BMH2‐HA3 carrying fragment of pTB419 at the BamHI–HindIII sites of pLC921, a YCplac33 (Gietz and Sugino, 1988) version that carries the natMX4 cassette‐containing EcoRI–HindIII fragment of pAG25 (Goldstein and McCusker, 1999) in its polylinker region. Full‐length Tpk1, Tps1 and Pho85 were expressed as HA2‐tagged versions under the control of the GAL1 promoter from plasmids pCDV503 (GAL1‐HA2‐TPK1), pAR502 (GAL1‐HA2‐TPS1) and pIP774 (GAL1‐HA2‐PHO85), respectively. Full‐length Bud14 was expressed as HA3‐tagged version under its own promoter from plasmid pFD662 (BUD14‐HA3). In addition, full‐length Pho85 was also expressed as a HA2‐tagged version under the control of the tetO7 promoter from plasmid pVW883 (tetO7‐HA2‐PHO85). Plasmid pVW884 (tetO7‐HA2‐PHO85E53A) was obtained by site‐directed mutagenesis (see above) of PHO85 in pVW883 using appropriate primers to introduce the kinase‐inactivating E53A‐encoding mutation. The integrative pLS9‐GRE1‐lacZ plasmid (pIP490) was constructed by cloning a PCR‐generated EcoRI fragment, containing the 778 nucleotides upstream of the GRE1 start codon, into the EcoRI site of pLS9 (Sarokin and Carlson, 1986). Plasmids YCpIF2‐GST and p2466, which allow expression of GST‐Pho80 under the control of the CUP1 promoter, have been described earlier (Reinders et al, 1998; Tan et al, 2003).
Kinase‐inactivating K823Y and C1176Y mutations in Rim15
The K823Y substitution replaces the invariant lysine in kinase subdomain II of Rim15 with tyrosine and results in an ATP‐binding deficient version of Rim15 (Vidan and Mitchell, 1997; Reinders et al, 1998). During detailed analyses of our original GFP‐Rim15 clone (pFD633; Pedruzzi et al, 2003), we discovered that it acquired a fortuitous point mutation (G3527A) that changes the cysteine residue at position 1176 to a tyrosine (C1176Y), located immediately downstream of the almost invariant glycine of the kinase catalytic sub‐domain IX (Hanks et al, 1988). Subsequent experiments indeed revealed that the C1176Y mutation results in inactivation of the Rim15 protein kinase activity (data not shown). Consistent with our previous results, GFP‐Rim15C1176Y accumulates in the nuclei of glucose‐limited and rapamycin‐treated rim15Δ cells (Figure 1A; Pedruzzi et al, 2003). However, the presence of a functional (genomic or plasmid‐expressed) Rim15 kinase efficiently prevents the observed nuclear accumulation of Rim15 (data not shown). While these new findings do not affect our previous conclusions (Pedruzzi et al, 2003), they provide a basis for the present study of cytoplasmic retention and/or nuclear import of GFP‐Rim15.
Identification of the Pho80 cyclin
We expected that loss of the Pho85 partner cyclins, either alone or in combination, should phenocopy the loss of Pho85 with respect to hyper‐induction of Rim15‐dependent genes. Indeed, we found that loss of the cyclin Pho80 (in strain BY490), but none of various combinations of other cyclin mutations (including pcl1Δ pcl2Δ pcl5Δ pcl9Δ clg1Δ [BY714], pcl1Δ pcl2Δ pcl5Δ pcl9Δ [BY708], pcl1Δ pcl2Δ pcl5Δ clg1Δ [BY637], and pcl1Δ plc2Δ [BY634]), resulted in hyperactivation of HSP26, SSA3, and GRE1 transcription when assayed during the early diauxic shift phase (data not shown).
GST pull‐down, immunoprecipitation, immunoblot analyses, and phospho‐specific antibodies
To perform coprecipitation experiments between Rim15 and Pho85, strain KT1960 was cotransformed with pNB566 or YCpIF2‐GST (expressing GST‐Rim15 or GST under the GAL1 promoter, respectively) and either pIP774, pCDV503, pAR502, or pFD662, which express HA2‐Pho85, HA2‐Tpk1, HA2‐Tps1, or Bud14‐HA3, respectively. Induction of GAL1‐driven expression and cell lysis were essentially performed as described (Lenssen et al, 2005). GST and GST‐tagged Rim15 were purified from clarified extracts using glutathione sepharose 4B beads (Amersham Biosciences). Bound proteins were eluted with sample buffer (5 min, 95°C) and subjected to standard immunoblot analysis for detection of coprecipitated HA2‐Pho85, HA2‐Tpk1 and HA2‐Tps1. For co‐IP experiments between Bmh2 and Rim15KI or Rim15KI‐T1075, strain W303‐1A was cotransformed with pTB419 expressing Bmh2‐HA3 under the control of its own promoter and pVW900 or pVW902, which express GST‐Rim15KI or GST‐Rim15KI‐T1075A, respectively, under the control of the tetO7 promoter. To allow expression of the tetO7‐controlled genes, cells were grown for at least six generations in exponential growth phase (OD600<1.0) in the absence of doxycycline. Subsequently, cells were lysed (see above) and HA‐tagged Bmh2 proteins were purified from clarified extracts with the protein G‐agarose IP kit (Roche Diagnostics GmbH) following the manufacturer's instructions using monoclonal mouse anti‐HA antibodies (HA.11; Covance). Bound proteins were eluted (as above) and subjected to standard immunoblot analysis for detection of co‐precipitated GST‐Rim15KI and GST‐Rim15KI‐T1075A. Dephosphorylation of GST‐Rim15 (purified from whole‐cell extracts; see above) was carried out by a 30‐min incubation at 30°C with 1 U of λ‐phosphatase (Biolabs, NewEngland). In control reactions, phosphatase inhibitors (10 mM NaF, 10 mM Na‐orthovanadate, 10 mM p‐NO2‐phenylphosphate, 10 mM β‐glycerophosphate, and 10 mM Na‐pyrophosphate) were added. Antibodies against Rim15 phosphorylated on T1075 were raised against a phosphorylated synthetic peptide (S‐R‐S‐S‐pT‐P‐P‐L‐A‐N‐P‐T; where pT represents phosphothreonine 1075 of Rim15), adsorbed with the unphosphorylated form of the peptide, and affinity‐purified with the phosphorylated peptide by Eurogentec.
Protein kinase assays and quantification of substrate phosphorylation
To assay in vitro phosphorylation of Rim15 by Pho85, HA2‐Pho85 (pVW883) and HA2‐Pho85E53A (pVW884), fusion proteins were expressed in (following growth for at least six generations in exponential growth phase in the absence of doxycycline) and purified from rim15Δ pho85Δ (CDV201‐3B) cells. Following this, cells were disrupted by vortexing in lysis buffer (50 mM Tris–HCl, pH 7.5, 0.15 M NaCl, 0.5 mM EDTA, 0.1% NP‐40, 10% glycerol, 1 mM PMSF, 1 mM DTT, one tablet of Complete Protease Inhibitor Cocktail [CPIC; Roche Diagnostics GmbH] per 50 ml, and phosphatase inhibitors [see above] in the presence of acid‐washed glass beads. HA‐tagged Pho85 and Pho85E53A proteins were purified as outlined above for the HA‐tagged Bmh2 protein. Kinase assays were performed with HA2‐Pho85‐ and HA2‐Pho85E53A‐bound beads at 30°C for 30 min in kinase buffer (50 mM Tris–HCl, pH 7.5, 20 mM MgCl2, 1 mM DTT, 1 mM ATP, one tablet of CPIC per 50 ml, and 10 μCi γ‐ATP) containing 50 ng of GST‐Pho80 (purified from rim15Δ pho85Δ (CDV201‐3B) cells; see also Tan et al, 2003) and the indicated Rim15‐derived substrates (purified from E. coli). Reactions were stopped by adding SDS‐gel loading buffer and boiling for 5 min and then subjected to SDS–PAGE. Gels were dried and exposed to X‐ray film. Substrate phosphorylation levels were quantified using a PhosphorImager (Cyclone Phosphor System; PerkinElmer) and analysed with OptiQuant Image Analysis software (Packard). Digital images of immunoblots were acquired with a CanoScan LiDE scanner (Canon) and Photoshop 7.0 (Adobe) and densitometric analysis of protein bands was done with OptiQuant Image Analysis software.
We thank Drs B Andrews, J Cannon, M Hall, and S Lemmon for providing yeast strains, plasmids, and/or anti‐Bmh2 antibodies, R Bisig for technical assistance, and Drs C Georgopoulos and R Loewith for critical reading of the manuscript. CDV is supported by the Swiss National Science Foundation grant 631‐062731.00 and the Canton of Geneva.
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