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Journal of Molecular Biology, Vol. 302, No. 1, Sep 2000, pp. 103-120

The Saccharomyces cerevisiae homologue  YPA1  of the mammalian  phosphotyrosyl phosphatase activator  of protein phosphatase  2A  controls progression through the  G1  phase of the yeast cell cycle

Christine Van Hoof¹, Veerle Janssens¹, Ivo De Baere¹, Johannes H. de Winde², Joris Winderickx², Françoise Dumortier², Johan M. Thevelein², Wilfried Merlevede¹ and Jozef Goris<sup>1

1</sup> Afdeling Biochemie, Faculteit Geneeskunde, Katholieke Universiteit Leuven, Herestraat 49, B-3000, Leuven, Belgium
² Laboratorium voor Moleculaire Celbiologie, Katholieke Universiteit Leuven, Kardinaal Mercierlaan 92, B-3001, Leuven-Heverlee, Belgium

Received 18 April 2000;  revised 21 July 2000;  accepted 25 July 2000. ; Available online 25 March 2002.

ABSTRACT

The Saccharomyces cerevisiae gene YPA1 encodes a protein homologous to the phosphotyrosyl phosphatase activator, PTPA, of the mammalian protein phosphatase type 2A (PP2A). In order to examine the biological role of PTPA, we disrupted YPA1 and characterised the phenotype of the ypa1Δ mutant. Comparison of the growth rate of the wild-type strain and the ypa1Δ mutant on glucose-rich medium after nutrient depletion showed that the ypa1Δ mutant traversed the lag period more rapidly. This accelerated progression through "Start" was also observed after release from -factor-induced G1 arrest as evidenced by a higher number of budding cells, a faster increase in CLN2 mRNA expression and a more rapid reactivation of Cdc28 kinase activity. This phenotype was specific for deletion of YPA1 since it was not observed when YPA2, the second PTPA gene in budding yeast was deleted. Reintroduction of YPA1 or the human PTPA cDNA in the ypa1Δ mutant suppressed this phenotype as opposed to overexpression of YPA2. Disruption of both YPA genes is lethal, since sporulation of heterozygous diploids resulted in at most three viable spores, none of them with a ypa1Δ ypa2Δ genotype. This observation indicates that YPA1 and YPA2 share some essential functions. We compared the ypa1Δ mutant phenotype with a PP2A double deletion mutant and a PP2A temperature-sensitive mutant. The PP2A-deficient yeast strain also showed accelerated progression through the G1 phase. In addition, both PP2A and ypa1Δ mutants show similar aberrant bud morphology. This would support the notion that YPA1 may act as a positive regulator of PP2A in vivo.

Author Keywords: phosphotyrosyl phosphatase activator (PTPA); G1/S transition; -factor; rapamycin; cell growth

Abbreviations: PTPA, phosphotyrosyl phosphatase activator; PP2A, protein phosphatase 2A; 5-FOA, 5-fluoro-orotic acid; MAP kinase, mitogen-activated protein kinase; TOR, target of rapamycin; PK-A, protein kinase A; YPD, yeast extract, peptone and glucose medium; YPGal, yeast extract and peptone medium supplemented with galactose; SD medium, synthetic medium supplemented with glucose; EB, extraction buffer; DTT, dithiothreitol; PMSF, phenylmethane-sulphonylfluoride

INTRODUCTION

The phosphotyrosyl phosphatase activator, PTPA, has been identified as a 37 kDa cellular protein that stimulates the phosphotyrosyl phosphatase (PTPase) activity of the dimeric holoenzyme of protein phosphatase type 2A (PP2A_D) in vitro, suggesting that PP2A can act as a non-classical dual specificity phosphatase [Cayla et al 1990]. PTPA-activated PP2A has a distinct in vitro substrate specificity compared to "classical" protein phosphotyrosyl phosphatases [Agostinis et al 1996], suggesting that PTPA-activated PP2A might dephosphorylate a selected group of tyrosyl phosphorylated substrates in vivo. PTPA is a ubiquitous protein, identified in many species using biochemical [Cayla et al 1990 and Van Hoof et al 1994] as well as molecular biological procedures [Cayla et al 1994 and Van Hoof et al 1998]. It is present in highly differentiated mammalian tissues as well as in unicellular eukaryotic organisms like budding yeast. However, the biological function of the PTPA-activated PTPase activity of PP2A remains elusive. The Ser/Thr phosphatase activity of PP2A is known to be involved in many cellular functions such as cell growth, cell differentiation, cell division and cell transformation (for reviews see [Mayer-Jaekel and Hemmings 1994, Mumby and Walter 1993, Van Hoof et al 1996 and Wera and Hemmings 1995]). The biological role of budding yeast PP2A has been elucidated by genetic analysis of mutants deficient for PP2A activity. Also in yeast, PP2A seems to have multiple functions in essential cellular events such as bud emergence, cell cycle regulation and cell wall integrity (for a review see [Stark 1996]). Two functionally redundant genes, PPH21 and PPH22, encode the catalytic subunit of PP2A. Strains with a single deletion of one of the two genes display a wild-type phenotype, whereas double disruption of the two PP2A genes causes a severe growth defect [Ronne et al 1991 and Sneddon et al 1990].

The Saccharomyces cerevisiae PTPA gene, YPA1, is located on chromosome IX (ORF YIL 153w). It encodes a protein, 38 % identical to human PTPA and it has a C-terminal amino acid extension relative to vertebrate PTPA. In vitro, the purified bacterially expressed [Van Hoof et al 1998] as well as the native [Van Hoof et al 1994] yeast protein display PTPA activity towards mammalian PP2A, indicating that budding yeast contains a functional homologue of mammalian PTPA. Alignment of the amino acid sequences of PTPA from yeast to human revealed several regions that are highly conserved. Screening of the protein sequence database with these conserved regions revealed the presence of a homologue of Ypa1 in yeast, encoded by YPA2 (ORF YPL 152w), located at chromosome XVI [Van Hoof et al 1998]. Ypa2 is 27 and 36 % identical to Drosophila and vertebrate PTPA, respectively, but only 25 % with Ypa1. Ypa1 and Ypa2 are the most divergent proteins among all PTPA proteins identified so far. Ypa2 also contains a C-terminal extension relative to vertebrate PTPA but it shows no homology to the extension of Ypa1. The role of these extensions for PTPA’s function is not clear. However, they seem to be unrelated to PTPA activity at least in vitro [Van Hoof et al 1998]. Database screening using the conserved regions as template revealed also a PTPA homologue in Schizosaccharomyces pombe (accession number: Z98980). Therefore, the highly conserved boxes found by alignment of PTPA from different species represent new motifs for an essential cellular function. Moreover, deletion of these conserved regions in the bacterially expressed rabbit protein abolishes PTPA activity towards PP2A and one of these deletions results in a dominant negative PTPA protein [Van Hoof et al 1998].

The mechanisms controlling the budding yeast mitotic cell cycle have been studied in great detail (for a review see [Lew et al 1997]). The G1 phase commitment to cell division, "Start", is separated in two phases: Start A being the nutrient and growth checkpoint, where the availability of nutrients, a critical rate of protein synthesis and growth of the cells up to a critical cell size is required to proceed cell division. In budding yeast the Ras/cAMP pathway [Tokiwa et al 1994] and the TOR pathway [Barbet et al 1996] regulate this control step. Start B is the replication and proliferation checkpoint, where the availability of the G1 cyclins regulated by transcriptional activation mechanisms is the critical determinant to execute Start [Stuart and Wittenberg 1995]. At this cell cycle checkpoint the decision of conjugation by activation of the mating pheromone pathway prevents the haploid cells from entering a new mitotic cycle by a G1 arrest, followed by the induction of genes required for morphological changes (shmoo formation) during mating [McKinney et al 1993].

Here we report the phenotypic analysis of the ypa1Δ mutant in order to understand the role of PTPA (as potential regulator of PP2A) in yeast. We demonstrate that PTPA apparently functions in the G1 phase of the yeast cell cycle and report genetic evidence for interaction with PP2A.

RESULTS

Viability, morphology and sporulation behaviour of the ypaΔ strains

Neither deletion of YPA1 nor of YPA2 in the haploid W303-1A strain affected viability at 30 °C on YPD plates. However, in liquid YPD medium the ypa1Δ mutant, but not the ypa2Δ mutant, showed moderate temperature sensitivity with a slower growth rate at 37 °C (Figure 1). About 25 % of the budding cells in the ypa1Δ strain had an aberrant bud morphology (elongated or pear-shaped bud), whereas single deletion of YPA2 had no effect on cell and bud morphology (data not shown).

 

Figure 1. Temperature sensitivity of ypa deletion mutants. Wild-type cells (♦) and ypa1Δ () and ypa2Δ () deletion mutants were inoculated in YPD medium. The growth rate of the different strains at 37 °C was monitored at A = 600 nm in a Bioscreen cell density meter. Because the increase in absorbance is not linear with the increase in cell density for values above 1, cell density is expressed in arbitrary units.

 

Double disruption of both YPA genes appeared to be lethal, since no transformants could be obtained by transformation of the ypa1Δ strain with the ypa2::TRP1 linearized fragment or by transformation of the ypa2Δ strain with the ypa1::URA3 linearized fragment. To investigate further the potential lethality of the double ypa1Δ ypa2Δ mutant, we generated heterozygous diploids, carrying one deleted ypa1::HIS5 and one disrupted ypa2::TRP1 allele by transforming a diploid W303-1A strain with these disrupted genes as described in Materials and Methods. After sporulation and tetrad analysis of 44 tetrads (see Table 1) none of the spores were able to grow on synthetic medium in which His and Trp were omitted. This result indicates that ypa1 ypa2 double deletion is indeed lethal. Microscopic examination of the ypa1Δ ypa2Δ spores showed that they form microcolonies of about 100 cells. Apparently, the ypa1Δ ypa2Δ spores can germinate, but the cells fail to divide further after a few cell cycles. To confirm the lethality of the ypa1Δ ypa2Δ double mutant, we performed a plasmid-loss experiment with the double deletion mutant containing the YPA1 gene on a plasmid. For this purpose, the YPA1 gene was reintroduced in the heterozygous ypa1Δ/YPA1 ypa2Δ/YPA2 diploid strain under control of the GAL1 promotor in the pYES2 plasmid, which contains the URA3 marker (Invitrogen). Dissection of 36 tetrads yielded eight asci with four viable spores, four asci containing spores with the genotype of a wild-type strain, a single ypa1Δ and ypa2Δ mutant and a double ypa1Δ ypa2Δ mutant, all containing YPA1 on the pYES2 plasmid. After selection for loss of GAL1-YPA1 on the URA3 pYES2 plasmid by growth on SGal Ura⁺ medium containing 1 mg/ml 5-fluoro-orotic acid (5-FOA) [Boeke et al 1987], almost all of the spores with a wild-type, single ypa1Δ or ypa2Δ genotype lost the YPA1 gene on the URA3 containing plasmid and were able to grow on SGal Ura⁺ supplemented with 5-FOA. In contrast, none of the double ypa1Δ ypa2Δ spores, were able to produce colonies on SGal Ura⁺ + 5-FOA medium, suggesting that loss of the functional YPA1 gene on the URA3 plasmid is lethal for the double deletion strain.

 

 

Table 1. Meiotic segregation of ypa1::HIS5 and ypa2::TRP1 after sporulation of the diploid YPA1/ypa1Δ YPA2/ypa2Δ strain
 

 

The ypa1Δ mutant has altered growth kinetics

When the growth profile of the ypa1Δ mutant was compared to that of a wild-type W303-1A strain with the URA3 marker gene, the ypa1Δ mutant exhibited a shorter lag-phase (Figure 2(a) and entered the resting state more rapidly (Figure 2(b) when stationary phase cells were inoculated in fresh YPD and incubated for 24 hours at 30 °C. After 11 hours of growth, cells of the ypa1Δ mutant were already accumulating with a 1N DNA content as measured by FACS analysis, indicating that these cells were arresting cell proliferation. At that time, the cells of the wild-type strain still displayed a 1N and 2N DNA distribution, indicating that they were still actively proliferating (Figure 3). Three possible explanations for the growth defect of the ypa1Δ mutant after the rapid fermentative growth phase were tested. First, the glucose might be more rapidly metabolised, leading to a more rapid exhaustion of this nutrient and growth arrest. This does not seem to be the case: both the wild-type and the ypa1Δ strain metabolise glucose equally well (Figure 2(c). Second, the ypa1Δ mutant may be unable to grow under gluconeogenic/respiratory conditions with ethanol as carbon source. The ypa1Δ mutant was able to grow on ethanol as non-fermentable carbon source although with a moderate decrease in growth rate compared to the wild-type strain (data not shown). Third, the respiratory pathway might be hampered. To test this, both strains were grown in YPD in the presence of antimycin, an inhibitor of the respiratory pathway. Following this treatment, the gluconeogenic/respiratory growth phase on ethanol was clearly abolished in the wild-type strain. For the ypa1Δ mutant, the antimycin effect was much less pronounced, and the moment of growth arrest and entrance into stationary phase cells coincided with that of the wild-type strain in the presence of antimycin (Figure 2(d). These results indicate that the altered growth profile of the ypa1Δ mutant is not due to a defect in glucose fermentation, but rather to a failure in proper gluconeogenic/respiratory growth on ethanol.

 

Figure 2. Growth curves, glucose consumption and antimycin response of the different strains in YPD medium at 30 °C. (a) and (b) Growth rate of ypa1Δ (), ypa2Δ (*), and ypa1Δ mutants with YPA1 (), YPA2 () or human PTPA (×) reintroduced on a high-copy yeast expression vector (pXL2) compared to the growth profile of the wild-type strain (♦). (c) Glucose consumption of the WT (♦) and ypa1Δ () strains during exponential growth. (d) Growth profile of the wild-type (, ♦) and ypa1Δ mutant (•, ) in the presence (/•) and absence (♦/) of antimycin (2 g/ml), respectively.

 

 

Figure 3. FACS analysis of the DNA content of the wild-type strain and the ypa1Δ mutant grown for 11 hours in YPD medium at 30 °C.

 

The phenotype of a shorter lag phase for the ypa1Δ mutant was suppressed by introducing overexpressing plasmids carrying the YPA1 gene or the human PTPA cDNA (Figure 2(a), but not by overexpressing YPA2. On the other hand, the faster entrance into G0 of the ypa1Δ strain is suppressed by reintroduction of YPA1, but not by reintroduction of HPTPA or YPA2 (Figure 2(b). Also YPA1ΔC, lacking the sequence encoding the C-terminal extension of Ypa1 relative to vertebrate PTPA, did not complement for the faster entrance into G0, whereas it did suppress the phenotype of a shorter lag phase (data not shown). Therefore, YPA1 seems to be specifically involved in two distinct events during vegetative growth and the C-terminal tail of Ypa1 is essential for its regulatory role in the gluconeogenic/respiratory growth pathway.

The ypa1Δ mutant is less responsive to -factor-induced G1 arrest

Since the ypa1Δ mutant has a shorter lag-period, entering the proliferating phase more rapidly, we reasoned that Ypa1 might function in progression through Start in G1. To examine this hypothesis in more detail, we arrested exponentially growing wild-type and ypa1Δ cells in G1 with -factor. Addition of -factor to MATa cells results in three major responses [Herskowitz 1995]. (1) Activation of a MAP kinase cascade, leading to transcriptional activation of genes required for mating such as FUS1 [McCaffrey et al 1987] and FAR1 [Chang and Herskowitz 1990]. (2) Cell cycle arrest in late G1, preventing cells from progressing into S phase by inactivation of the Cln/Cdc28 kinase activity. This inhibition is accomplished by repression of CLN1 and CLN2 transcription and by inhibition of the upstream Cln3/Cdc28 kinase activity [Jeoung et al 1998] through direct binding of Far1 [Chang and Herskowitz 1990]. (3) Alteration of cellular morphology, resulting in an elongated, pear-shaped cell, also called "shmoo" by engagement of genes such as FUS1.

We have determined the sensitivity to -factor and the release from G1 arrest in the ypa1Δ mutant by following these responses. Shmoo formation (morphological response to -factor) and budding index (proliferating activity) were determined and FUS1 and CLN2 mRNA expression were quantified. Apparently, deletion of YPA1 makes cells more resistant to -factor, since at all concentrations of -factor used, more budding and less shmoo cells were observed (Figure 4(a). Northern blot analysis showed that these observations coincide with lower FUS1 mRNA expression and higher expression of CLN2 mRNA in ypa1Δ cells, which is most apparent upon addition of 0.5 M -factor (Figure 5), whereas YPA1 mRNA expression was not altered upon -factor addition (data not shown). These results indicate that the ypa1Δ mutant is less sensitive to -factor-induced G1-arrest and that the difference with the wild-type strain is more pronounced at lower concentrations of -factor. Therefore, 1 M -factor was used in all further experiments. Moreover, transformation of the ypa1Δ strain with a vector carrying YPA1 or HPTPA cDNA, largely overcame the G1 arrest defect of the ypa1Δ mutant in response to -factor as determined by microscopic analysis (Figure 4(c) and quantification of the CLN2 mRNA expression (data not shown). These results indicate that the lower response to -factor-induced G1-arrest is specific for YPA1.

 

Figure 4. -Factor sensitivity of wild-type and ypaΔ strains. Budding index (a) and shmoo formation (b) two hours after addition of different concentrations of -factor to wild-type () and ypa1Δ strains (&z.sqne;). Budding index (c) and shmoo formation (d) after addition of 1 M -factor to the wild-type (), ypa1Δ (&z.sqne;) and ypa2Δ strains () and after reintroduction of YPA1 (&z.sqsw;) or human PTPA () in the ypa1Δ strain.

 

 

Figure 5. Northern blot analysis of -factor-regulated transcription of FUS1 and CLN2. Wild-type (WT) and ypa1Δ mutant (ypa1Δ) strains were treated with 0.5 or 5 M -factor for three hours. 10 g RNA was loaded in each lane and after transfer, the blot was hybridized with FUS1, CLN2 and actin probes. The constitutively expressed actin gene was used as control for RNA loading. FUS1 and CLN2 transcription are quantified as the % of maximal mRNA expression relative to the actin mRNA level of the same sample and the numbers are indicated underneath each lane.

 

The lower response to -factor of the ypa1Δ mutant is due to an accelerated progression through the G1/S transition point after -factor-induced G1 arrest

We have investigated the response to -factor-induced G1-arrest of the ypa1Δ mutant in more detail. FACS analysis clearly shows that at the time of maximal morphological response to -factor (two hours after addition of 1 M -factor), most wild-type cells had a 1N DNA content, indicative of a G1 arrest, whereas the ypa1Δ mutant contained a large pool of replicating cells with a 2N DNA content (Figure 6). Nevertheless, Cdc28 kinase activity was as rapidly inactivated in the ypa1Δ mutant as in the wild-type strain upon -factor addition. However, this transient inactivation was faster relieved in the ypa1Δ mutant (after two hours when -factor is not yet removed) compared to the wild-type strain (Figure 7), suggesting that activation of the mating pathway as such is not affected by YPA1 deletion, but that the -factor-induced G1 arrest is more rapidly released, resulting in faster progression into S phase. This faster recovery from -factor-induced G1 arrest is reflected in the rapid increase in CLN2 mRNA expression during -factor treatment of ypa1Δ cells (Figure 5). Taken together, we can conclude that deletion of YPA1 overcomes G1 arrest induced by -factor more rapidly, indicating a role for YPA1 as negative regulator of the cell cycle checkpoint Start.

 

Figure 6. FACS analysis of the DNA content of wild-type strain (WT) and ypa1Δ mutant (ypa1Δ). -Factor (1 M) was added to exponentially growing cells and the DNA content of propidium iodide-stained cells was determined at the indicated times.

 

 

Figure 7. Cdc28 kinase activity of -factor-treated wild-type (♦) and ypa1Δ mutant () strains. -Factor (1 M) was added to exponentially growing cultures and the cells were further incubated at 30 °C for three hours. Subsequently, -factor was removed by washing and resuspension of the cells in fresh YPD medium. The time of -factor removal is indicated by an arrow. Samples were taken every hour in the presence of -factor and every 30 minutes after release of -factor-induced G1 arrest.

 

A PP2A-depleted yeast strain resembles the ypa1Δ mutant in its -factor-induced G1-arrest defect

Since PTPA is known to be a regulator of mammalian PP2A, we examined the response to -factor of the PP2A deletion mutant, H336, which carries a deletion of the PPH21 gene and has the PPH22 gene expressed from the GAL1 promotor. Shift of this mutant from galactose to glucose containing medium causes repression of PPH22, hence leading to the complete absence of PP2A activity. This PP2A deletion mutant also showed a higher budding index and less shmoo formation in response to -factor (Figure 8 and Figure 9). Also maximal induction of FUS1 was lower compared to the wild-type strain (Figure 9(c), while the CLN2 mRNA level increased to the same extent as in the ypa1Δ mutant, two hours after addition of -factor. At that time the CLN2 mRNA level of wild-type cells was still much lower (Figure 9(d). Cdc28 kinase activity of the three strains was completely inactivated in response to -factor (Figure 9(e). It was reactivated 30 minutes faster in the ypa1Δ mutant compared to the wild-type strain, simultaneously with the faster increase in CLN2 mRNA expression. However, Cdc28 kinase activity remained very low in the PP2A-depleted strain, although CLN2 mRNA expression increased as fast as in the PTPA deletion mutant (Figure 9(d). This very low Cdc28 kinase activity in the PP2A deletion mutant might be related to its severe growth defect.

 

Figure 8. Microscopic visualisation of cells of the wild-type strain (WT), ypa1Δ mutant and PP2A deficient strain (H336) after incubation with 1 M -factor for two hours. Bar represents 15 m.

 

 

Figure 9. Morphological changes, changes in the FUS1 and CLN2 mRNA expression and changes in Cdc28 kinase activity in response to -factor of the wild-type strain (), the ypa1Δ mutant (&z.sqne;) and the PP2A deficient strain, H336 (). (a) Budding index and (b) shmoo formation as well as (c) FUS1 and (d) CLN2 mRNA expression was followed during -factor-induced G1 arrest. Budding and shmoo forming cells were also counted during recovery from -factor-induced G1 arrest by refreshing the medium. (e) Cdc28 kinase activity of -factor-treated cells of the wild-type strain (♦), ypa1Δ mutant () and PP2A deletion mutant (). -Factor (1 M) was added to exponentially growing cultures and incubation was continued at 30 °C for two hours. -Factor was removed (indicated by an arrow) by washing the cells and resuspending them in fresh YPD medium. Samples were taken every 30 minutes in the presence of -factor and after release of -factor-induced G1 arrest.

 

We also examined this -factor-induced G1 arrest in a temperature-sensitive mutant of PP2A, DEY142-1C [Evans and Stark 1997]. Cells were pre-grown at the permissive temperature of 24 °C and two hours before addition of -factor shifted to the restrictive temperature of 37 °C. The morphological changes caused by addition of -factor (unbudded cells and shmoo formation) were similar to that of the ypa1Δ mutant and the PP2A deletion mutant (data not shown). In this experiment also FAR1 mRNA expression was followed after addition of -factor. The level of FAR1 transcription in the PP2A- and YPA1-deficient strains was not significantly different from that of the wild-type strain (Figure 10(a). However, a faster reactivation of CLN2 mRNA expression and Cdc28 kinase activity was observed in the Ypa1 and PP2A-depleted strains after an initial inactivation in response to -factor (Figure 10(b).

 

Figure 10. Comparison of biochemical changes in response to -factor of the wild-type strain (), ypa1Δ mutant (&z.sqne;) and PP2A temperature-sensitive mutant, DEY142-1C (). (a) FAR1 and (b) CLN2 mRNA expression were followed during -factor-induced G1 arrest. (c) Cdc28 kinase activity of -factor-treated cells of the wild-type strain (♦), ypa1Δ mutant () and PP2A temperature-sensitive mutant, DEY142-1C (•). Exponentially growing cultures were incubated at 30 °C (WT, ypa1Δ) or 37 °C (DEY 142) for three hours in the presence of 1 M -factor. Samples were taken every 30 minutes in the presence of -factor and after release of -factor-induced G1 arrest. A control experiment revealed similar budding and shmoo indexes of the wild-type strain at 30 °C and 37 °C.

 

In summary, the phenotypes of the ypa1Δ mutant and the PP2A loss of function mutant after -factor-induced G1 arrest are very similar in several aspects. This tends to indicate that PTPA acts as a physiological regulator of PP2A in vivo.

The ypa1Δ mutant is released more rapidly from G1 arrest induced by nutrient depletion

So far, the results indicate a role for YPA1 in the G1/S transition after -factor-induced G1 arrest, and not in the -factor-induced activation of the mating pathway. To substantiate these data we examined the behaviour of a ypa1Δ mutant after release from G1 arrest caused by nutrient depletion instead of -factor addition. As shown in Figure 11(a), ypa1Δ cells resumed budding or proliferation after 90 minutes whereas the wild-type strain resumed budding only after 180 minutes. The faster growth rate of the ypa1Δ mutant as evidenced by the faster increase in optical density of a stationary phase culture inoculated in fresh YPD medium (Figure 2(a) is apparently due to accelerated initiation of proliferation. This faster progression through Start is reflected in a more rapid increase in CLN2 transcription (Figure 11(b) and a faster reactivation of Cdc28 kinase activity. The wild-type strain reached the same Cdc28 kinase activity only 120 minutes later (Figure 11(c). These observations confirm the accelerated progression through Start of the ypa1Δ mutant compared to the wild-type strain and show that this acceleration is independent of the treatment used to induce the G1 arrest.

 

Figure 11. Release from G1 arrest induced by nutrient depletion. (a) Budding index, (b) CLN2 mRNA expression and (c) Cdc28 kinase activity were monitored after dilution of the wild-type (/♦) and ypa1Δ (&z.sqne;/) strains to A₆₀₀ = 0.2 in fresh YPD medium. Samples were taken during 4.5 hours every 90 minutes to determine budding index and every 30 or 60 minutes to quantify CLN2 mRNA expression and assay Cdc28 kinase activity. (d) Rapamycin sensitivity of the wild-type and ypa1Δ strain. Cells were diluted to an A₆₀₀ = 0.1 and spotted on YPD plates containing 0.1 g/ml or 0.5 g/ml rapamycin. The rapamycin-YPD plates were incubated at 30 °C for three days before being photographed.

 

Further support for Ypa1 affecting nutrient control of G1 progression is obtained by the difference in rapamycin sensitivity between ypa1Δ and wild-type cells. Rapamycin arrests cells in G1 at the same cell cycle checkpoint (Start A) as nutrient depletion by interference with the TOR pathway, a signalling pathway coupling initiation of translation to G1 progression in response to nutrient availability [Barbet et al 1996]. Figure 11(d) shows that the ypa1Δ mutant is much more resistant to rapamycin compared to the wild-type strain, suggesting that YPA1 might control G1 progression by interference with the TOR pathway.

The negative regulation of Start is a function specific for Ypa1 and cannot be accomplished by Ypa2

Analysis of the ypa2Δ mutant for the same phenotypic traits as investigated for the ypa1Δ mutant, revealed a similar growth profile as the wild-type strain (Figure 2(a). Overexpression of YPA2 in the ypa1Δ mutant did not suppress the growth related ypa1Δ phenotypes (Figure 2(a), and, unlike the ypa1Δ mutant, the ypa2Δ mutant showed no temperature sensitivity at 37 °C (Figure 1). Furthermore, the ypa2Δ mutant behaved in the same way as the wild-type strain upon -factor-induced G1 arrest index (Figure 4(c). These results indicate that YPA1 and YPA2 encode non-redundant proteins and that only YPA1 is involved in yeast cell proliferation and gluconeogenic/respiratory growth. However, double deletion of both YPA genes resulted in lethality. The double deletion strain was unable to grow at 30 °C or 37 °C. Therefore, the contribution of Ypa2 to the total Ypa activity could be too small to exhibit a detectable phenotype when only YPA2 is deleted, but results in a complete loss of Ypa activity when also YPA1 is deleted, leading to a lethal phenotype. This hypothesis is corroborated by the ratio found for both transcripts. At the transcriptional level, the YPA1/YPA2 mRNA ratio is 4:1 in exponentially growing wild-type cells (data not shown). However, quantification by specific detection of both YPA gene products at the protein level will be required to determine the precise Ypa1/Ypa2 ratio.

DISCUSSION

General properties of the ypa1Δ mutant

In order to elucidate the biological role of the budding yeast PTPA homologue, YPA1, we examined a haploid ypa1Δ mutant for any phenotype(s) indicative of specific functions of Ypa in the yeast cell. The ypa1Δ mutant was viable and showed some temperature sensitivity when grown in YPD at 37 °C. Furthermore, the ypa1Δ strain displayed aberrant bud morphology, indicating a role for YPA1 in bud emergence. ypa1Δ strains of opposite mating types were able to mate and the resulting ypa1Δ diploid strain produced viable spores displaying normal germination.

Deletion of YPA1 causes accelerated progression through Start and faster release from -factor-induced G1 arrest

The faster progression of the ypa1Δ mutant through the lag-phase was a first indication that deletion of the YPA1 gene causes accelerated progression through Start. We used -factor to induce G1 arrest in the wild-type and ypa1Δ strain and examined different cell cycle-related parameters known to be affected by addition of -factor. After the initial decline in proliferating activity in response to -factor, measured as a decrease in the budding index, CLN2 mRNA expression and complete inactivation of the Cdc28 kinase, the ypa1Δ mutant resumed cell proliferation and passed through Start more rapidly than the wild-type strain. These results indicate that YPA1 plays a negative role in passage through the G1/S transition.

YPA1 seems not to be involved in activation of the mating pathway

The ypa1Δ mutant showed significantly less shmoo forming cells in response to -factor, a reduced extent of FUS1 induction and a subsequent more rapid decrease in FUS1 mRNA. On the other hand FAR1 mRNA expression was very similar in the ypa1Δ mutant and the wild-type strain. From these observations, the ypa1Δ mutant seems less responsive to -factor. However, activation of the pheromone pathway itself in response to -factor does not seem to be affected. This is supported by the following results: (1) CLN2 mRNA expression dropped as fast and to the same extent as in wild-type cells, (2) complete inactivation of the Cdc28 kinase activity was observed and occurred with the same kinetics in both strains, (3) FAR1 as well as FUS1 transcription were both induced in response to -factor and in the case of FAR1 no difference in the rate or level of transcription was observed in comparison with the wild-type strain. These responses occurred 30 to 60 minutes after addition of -factor (Figure 6 and Figure 10) and preceded the morphological responses such as arrest of bud emergence and formation of shmoo cells, which occurred only 90–120 minutes after -factor-induced G1 arrest (Figure 4). Thus, the lower drop in budding index as well as the reduced number of shmoo forming cells can be explained by a faster recovery from -factor-induced G1 arrest, as evidenced by the fact that CLN2 transcription and Cdc28 kinase activity of the ypa1Δ mutant already increased again to the critical level required for resumption of cell division at the moment that wild-type cells still exhibited the maximal morphological responses to -factor. We suggest that cells of the ypa1Δ mutant recover faster from -factor-induced G1 arrest and initiate progression through Start already before they start to show the morphological response to mating factor.

The faster recovery from the -factor-induced G1 arrest might be due to a faster densensitization of the mating pathway, for instance by an altered regulation of the endogenous BAR1 gene product, an extracellular protease responsible for the degradation of -factor [Sprague and Herskowitz 1981]. However, BAR1 mRNA expression in response to -factor was even higher in wild-type as in ypa1Δ cells (unpublished observations). Since PTPA is known to function as an activator of a phosphatase, deletion of YPA1 might result in a permanent hyperphosphorylation of components of the pheromone pathway. For example, hyperphosphorylation of the C-terminal domain of the -factor receptor results in a faster desensitisation of the receptor [Reneke et al 1988] and phosphorylation of Far1 results in its degradation and passage through Start [McKinney et al 1993]. Furthermore, PP2A is known to be an in vivo inactivator of MAP kinase cascades [Alessi et al 1995].

Alternatively, the faster progression through G1 after -factor-induced G1 arrest might be independent of signalling in the mating pathway and would rather be caused by an effect on the regulation of the G1/S transition in general. This possibility was suggested by the accelerated progression of the ypa1Δ mutant through the lag-phase upon refeeding of nutrient-depleted cells and by monitoring distinct cell cycle parameters upon release from G1 arrest caused by nutrient depletion (Figure 11). Also in this case the ypa1Δ mutant showed a faster increase in proliferating activity accompanied by a more rapid induction of CLN2 transcription and accelerated reactivation of Cdc28 kinase activity. Therefore, YPA1 does not seem to be involved specifically in the mating pheromone pathway but rather in a more general system regulating the G1/S transition.

Potential targets of YPA1 in its function as negative regulator of Start

CLN3 is known to be a dose-dependent regulator of Start. Unlike Cln1 and Cln2, Cln3 is not regulated at the transcriptional level, but by proteolysis and phosphorylation [Tyers et al 1992]. Therefore, Ypa1 as a regulator of PP2A might be involved in the regulation of Cln3 dephosphorylation. Dephosphorylation of Cln3 is known to eliminate most of the kinase activity associated with the Cln3-Cdc28 kinase complex [Tyers et al 1992]. This dephosphorylation event is not responsible for dissociation of Cln3 from Cdc28 nor for its degradation, but might be responsible for inactivation of the Cln3-Cdc28 kinase complex after Start, resulting in the subsequent decrease in CLN2 transcription [Stuart and Wittenberg 1995]. In this way deletion of YPA1 might abolish this dephosphorylation event, causing faster progression through Start.

Moreover, PTPA as regulator of PP2A might affect the TOR pathway. A component of this pathway, Tap42 has been demonstrated to be associated with the catalytic subunits of PP2A [Di Como and Arndt 1996]. A temperature-sensitive mutation in the TAP42 gene causes G1 arrest and confers rapamycin resistance. Rapamycin causes the dissociation of Tap42 from Sit4 and Pph21/22 by inhibition of the TOR kinase which phosphorylates Tap42 in vivo, thereby promoting the association of Tap42 with Pph21/22 [Jiang and Broach 1999]. The Tap42-Pph21/22 complex might have altered activity towards substrates involved in promoting protein synthesis [Nananoshi et al 1998 and Chung et al 1999]. Tpd3 and Cdc55 inhibit the association of Tap42 with Pph21/22 by direct competition and dephosphorylation of Tap42 [Jiang and Broach 1999]. Tap42 is not associated with Sit4 and Pph21 when cells are arrested in G0 by nutrient starvation, while Tap42/Sit4 complex formation is greatly enhanced upon nutrient refeeding of stationary phase cultures, suggesting that binding of Tap42 to Sit4 and Pph21 is required for cells to enter the mitotic cell cycle. Interestingly, deletion of YPA1, and to a very weak extent also deletion of YPA2, caused higher resistance to rapamycin (Figure 11(d) and [Rempola et al 2000]), which was even more pronounced when the YPA1 gene was deleted in a PP2A single deletion mutant (unpublished observations). Therefore, the YPA1 gene might be involved in the rapamycin-sensitive TOR signalling pathway to control the Start cell cycle checkpoint. Since the rapamycin-resistant phenotype of the ypa1Δ mutant is more severe in a PP2A single disruption genetic background, Ypa1 might function in this cell cycle control pathway through its action on PP2A. However, unlike TAP42, YPA1 seems to act as a negative regulator of cell cycle progression. Therefore, Ypa1 might inhibit Tap42/Pph21 complex formation through its interaction with PP2A. This would result in a change in PP2A activity towards specific substrates such as Tap42 itself and prevent budding yeast from resuming cell division.

YPA1 is implicated in the control of the diauxic shift or gluconeogenic/respiratory growth pathway

Comparison of the growth profile of the ypa1Δ mutant with that of the wild-type strain demonstrated that the ypa1Δ strain enters more rapidly into stationary phase. Deletion of YPA1 does not change the rate of exponential growth on glucose during the fermentative growth phase. However, although the ypa1Δ mutant was able to grow on non-fermentable carbon sources, its respiratory growth was clearly slower than that of the wild-type strain. This result suggests a participation of YPA1 in some mitochondrial function in budding yeast. Alternatively, YPA1 might be involved in a mechanism controlling the diauxic shift, where budding yeast reprograms its metabolic and biosynthetic machinery and switches from fermentation to respiration [De Winde et al 1997]. In this context it is important to note that the ypa1Δ mutant fails to derepress stress response genes such as CTT1 (unpublished observations) when glucose becomes limited in the growth medium. The potential function of YPA1 in the control of the diauxic shift is an important issue for further investigation since the mechanism triggering this event in budding yeast remains largely unknown.

The C-terminal tail of Ypa1 seems to be important for its role in controlling diauxic shift or respiratory growth, since introduction of the human PTPA cDNA lacking the sequence encoding this C-terminal extension or introduction of YPA1ΔC or YPA2, were unable to overcome this ypa1Δ phenotype. The C-terminal extension of Ypa2 is the most divergent region between the two Ypa proteins [Van Hoof et al 1998].

The two YPA genes appear to have different functions in budding yeast

Deletion of YPA2, did not result in any phenotypic trait conferred by YPA1 deletion and the growth behaviour of the ypa2Δ mutant is very similar to that of the wild-type strain. These results indicate that the second PTPA homologue in yeast is neither involved in regulation of cell cycle progression nor in controlling diauxic shift or respiratory growth. In addition, [Ramotar et al 1998] recently reported that YPA1, but not YPA2 is involved in the repair of DNA damage caused by oxidative stress. Also [Rempola et al 2000] found by functional analysis of the two yeast PTPA homologues that deletion of YPA1 caused pleiotropic phenotypes, while the phenotypes for ypa2Δ were less severe or non-existent. Moreover, overexpression of YPA2 is unable to suppress the ypa1Δ phenotypes. Thus, unlike PP2A, which is encoded by two functionally overlapping genes, the two YPA genes are at least for some functions non-redundant.

On the other hand, double disruption of both genes is lethal, indicating that there is some functional redundancy between the YPA genes. Ypa2 might be a structural homologue of Ypa1 with different functions which can still confer some Ypa1-complementing activity in Ypa1-deficient cells. Alternatively, Ypa1 and Ypa2 are functionally overlapping proteins and because Ypa2 is much less abundant, it is only responsible for a small fraction of total Ypa activity. This explains the absence of phenotypic defects in the ypa2Δ mutant.

The lethality of the ypa1Δ ypa2Δ double mutant is confirmed by the results described by [Rempola et al 2000], while [Ramotar et al 1998] reported the existence of a viable haploid ypa1Δ ypa2Δ double mutant. This discrepancy might be explained by the use of different genetic backgrounds (W303-1A versus DBY747) since [Rempola et al 2000] reported that the presence of the SSD1-v allele (SSD1-d2 in W303-1A) renders the double mutant viable. Otherwise, suppressor mutations might have occurred in the haploid ypa1Δ mutant transformed with ypa2Δ as reported earlier when deletion of two genes is lethal [Ronne et al 1991].

Indication for a physiological interaction between PTPA and PP2A

There is some evidence that Ypa1 can function autonomously and independently of PP2A in budding yeast. First, the C-terminal extension of Ypa1 is essential for its role in the gluconeogenetic/respiratory pathway and human PTPA cannot suppress this ypa1Δ mutant phenotype, suggesting that this function is unrelated to the intrinsic PTPA activity of Ypa1 and therefore also independent of PP2A. In addition, Ypa1 but not PP2A is involved in the repair mechanism of oxidative DNA damage [Ramotar et al 1998]. However, we have also obtained some evidence for a Ypa-PP2A interaction in yeast. Comparison of the behaviour of the PP2A loss-of-function mutant and the ypa1Δ mutant after -factor-induced G1 arrest, showed that both strains released faster from the G1 arrest resulting in an accelerated progression through the G1 phase. However, no reactivation of Cdc28 kinase was observed in the PP2A double deletion mutant. A possible explanation for this difference might be that PP2A acts as a positive regulator of G2 cyclin/Cdc28 kinase activity, probably through its dephosphorylating activity [Lin and Arndt 1995]. Deletion of PP2A causes a G2 delay, creating a pool of cells, which are arrested as large budding cells, displaying no basal Cdc28 kinase activity. Only the cells that progress through this G2 block, are sensitive to -factor-induced G1 arrest. To overcome these limitations, we have used the temperature-sensitive PP2A mutant, DEY142-1C, to monitor Cdc28 kinase activity. This mutant displays a less severe phenotype than the constitutively depleted PP2A double deletion mutant [Evans and Stark 1997]. Although this strain showed the same accelerated reactivation of Cdc28 kinase activity after its inactivation by the -factor-induced G1 arrest compared to wild-type cells, we failed to measure any Cdc28 kinase activity in a temperature-sensitive PP2A mutant (DEY172-2B) restrictive at a lower temperature [Evans and Stark 1997], or in a PP2A mutant that is also depleted for the PP2A-like PPH3 gene (DEY214) (data not shown). From these results, we can conclude that the phenotypes of the ypa1Δ mutant and the PP2A loss-of-function mutant after -factor-induced G1 arrest are very similar, suggesting a related function for both genes in negative regulation of progression through Start. This supports the notion that yeast PTPA (YPA) may act as a positive regulator for some function of PP2A in vivo.

MATERIALS AND METHODS

Strains, media and growth conditions

The Saccharomyces cerevisiae strains used in this study are listed in Table 2. All strains are isogenic with the wild-type strain W303-1A. The URA3 marker gene was inserted in this wild-type strain to avoid the effects of the marker during the phenotypic analysis of the ypa1Δ mutant. Construction of the ypa1Δ mutant and ypa2Δ mutant as well as the strains overexpressing YPA1, YPA2 or human PTPA (HPTPA) cDNA are described further. Strain H336 is a conditional PP2A deletion mutant, containing a deletion of the PPH21 gene and the PPH22 gene under the control of the galactose-inducible and glucose-repressible GAL1 promotor [Ronne et al 1991]. The temperature-sensitive PP2A mutants, DEY142-1C and DEY214, contain a temperature-sensitive mutation in PPH22 and a deletion of PPH21. The DEY214 temperature-sensitive mutant has an additional deletion of the PP2A-like PPH3 gene [Evans and Stark 1997].

 

 

Table 2. Yeast strains and plasmids used in this study
 

 

Cells were grown in YPD (1 % (w/v) yeast extract, 2 % (w/v) bacterial peptone and 2 % (w/v) glucose) except for the PP2A-deficient strain H336, which was precultured on YP medium supplemented with 2 % (w/v) galactose (YPGal). The synthetic minimal medium (SD) used for selection of the appropriate auxotrophic mutation and for preculture of the strains used in complementation experiments with the overexpression strains contained 0.67 % (w/v) yeast nitrogen base, 2 % glucose and 0.002 % of each auxotrophic amino acid required for growth. Growth was monitored by measuring the absorbance at 600 nm of cultures grown in complete YPD medium for 24 hours. Stationary phase cultures were diluted to A₆₀₀ = 0.2 for determination of the growth curves.

Construction of the haploid strains with disruption of the YPA1 or YPA2 gene

The YPA1 clone containing the complete open reading frame [Van Hoof et al 1998] was digested with SalI to interrupt the open reading frame at position 415. The URA3 marker gene was recovered from the YpDU plasmid [Berben et al 1991] by PCR with oligonucleotides containing the SalI restriction site at their 5′ end. The URA3 gene was then inserted in the digested and interrupted YPA1 construct. Digestion of the resulting plasmid with XhoI and HindIII generated a linearized fragment containing the disrupted YPA1 gene. This linear fragment was used to transform the wild-type W303-1A strain using the one-step gene disruption procedure [Rothstein 1983]. Disruption of the YPA1 gene was confirmed by Southern blot and PCR analysis.

The complete open reading frame encoded by the YPA2 gene [Van Hoof et al 1998] was digested with BglII to interrupt the open reading frame at position 294. The TRP1 marker gene was recovered from the YpDW plasmid [Berben et al 1991] by digestion with BamHI and inserted in the digested and interrupted YPA2 construct. A linearized fragment containing the disrupted YPA2 gene was generated by digesting the resulting plasmid with NcoI and BamHI. This linear fragment was used to transform the wild-type W303-1A strain to obtain a single ypa2Δ mutant. Southern blot and PCR analysis was used to confirm proper disruption of YPA2. All attempts made to obtain the double ypa1Δ ypa2Δ mutant, by introducing a ypa2Δ in the ypa1Δ mutant or vice versa were unsuccessful, apparently because the haploid double deletion mutant is lethal (see Results).

Construction of plasmids overexpressing YPA1, YPA1ΔC, YPA2 and the human PTPA cDNA

Plasmids containing the complete coding sequence of the yeast and human PTPAs were digested with EcoRI and BamHI. YPA1ΔC (YPA1 with deletion of the C-terminal extension) was obtained by PCR with a sense oligonucleotide, containing the YPA1 start codon and a EcoRI restriction site, and an antisense oligonucleotide, containing a stop codon at position 961 of YPA1 followed by a BamHI restriction site. This resulted in a truncated YPA1 with the same length as HPTPA. In order to obtain efficient overexpression of these DNA fragments, they were cloned into a multi-copy vector containing the phosphoglycerate kinase (PGK1) yeast promotor. This vector, pXKL2 (kindly provided by K. Luyten, Leuven, Belgium), is derived from the YEplac181 multi-copy vector [Gietz and Sugino 1988].

Construction of the heterozygous diploid YPA1/ypa1Δ YPA2/ypa2Δ mutant

A second YPA1 disrupted gene was created by insertion of the Schizosaccharomyces pombe his5⁺ marker gene into the YPA1 gene. Two oligonucleotides each containing 60 bp corresponding to the start and the end of the YPA1 gene and both prolonged with the 5′ sense and 3′ antisense his5⁺ tags, respectively were synthesized and used as primers in a PCR reaction with the pFA6a-HIS5MX6 plasmid [Lontine et al 1998] as template. The PCR product obtained was used to transform diploid W303-1A MATa/MAT cells to a HIS⁺ strain, since the S. pombe his5⁺ gene complements the S. cerevisiae HIS3 gene, but avoids false positives generated by gene conversion. Simultaneously, the diploid W303-1A strain was cotransformed with the ypa2::TRP1 construct described above, in order to obtain a diploid heterozygous YPA1/ypa1Δ YPA2/ypa2Δ mutant. Heterozygous deletion mutants were selected on SD His⁻/Trp⁻ plates and the presence of both wild-type and disrupted alleles for both YPA genes was confirmed by PCR with a set of primers for both genes. After sporulation of the diploid transformants on agar plates (1.5 %, w/v) containing 1 % (w/v) potassium-acetate (pH=6) at 24 °C for several days, tetrad analysis was performed using a micromanipulator (Singer) and lyticase (Aldrich) to digest the asci. After germination of the separated ascospores on fresh YPD plates or in the case of introduction of GAL1-YPA1 on the pYES2 plasmid on YPGal plates, the genotype of the segregants was examined by replicating the colonies on specific media. The presence of the deletions was inferred from the prototrophies determined by the genetic marker genes and confirmed by PCR. Negative selection against GAL1-YPA1 on the pYES2 plasmid with a URA3 marker was performed by growing the segregants in liquid SGal Ura⁺ medium (synthetic minimal medium supplemented with 2 % galactose and 0.1 % uracil), and spreading the cell suspension in a 1/5000 dilution on SGal Ura⁺ agar plates. Colonies were replicated to SGal Ura⁺ containing 1 mg/ml 5-fluoro-orotic acid (5-FOA) and evaluated for loss of the GAL1-YPA1 gene by their ability to grow on this selection medium.

Cell cycle inhibition at Start

Overnight cultures were diluted in YPD to an A₆₀₀ of 0.15 and grown to early exponential phase. At that time, cultures were diluted to an A₆₀₀ of 0.25 and -factor was added at a concentration of 1 M to YPD acidified with HCl to pH 4 [Elion et al 1990]. -Factor was removed by resuspending centrifuged cultures in fresh YPD medium. Nutrient depletion was performed by growing the strains on YPD medium for 48 hours until the cultures contained only small unbudded cells as determined by microscopic analysis, indicating that the strains were arrested in G1. These stationary phase cultures were diluted to an A₆₀₀ = 0.5 in fresh YPD medium. Samples were taken every 30 minutes for determination of the optical density of the cultures at 600 nm, microscopic analysis of the budding index, Northern blot analysis of CLN2 mRNA expression and biochemical analysis of Cdc28 kinase activity.

The PP2A double deletion mutant, H336, was initially grown to exponential phase in YPGal and subsequently incubated overnight in YPD in order to ascertain that all PP2A had disappeared. The temperature-sensitive PP2A mutant, DEY142-1C, was grown at 24 °C until exponential phase and switched to 37 °C in order to inactivate PP2A two hours prior to the addition of cell cycle inhibitors.

RNA extraction and Northern blot analysis

Five ml of culture was taken at the indicated times and RNA was prepared and loaded on a 1 % (w/v) agarose gel, transferred to a Hybond-N membrane and hybridized as described by [Crauwels et al 1996]. Actin, CLN2, FAR1 and FUS1 DNA probes to hybridize the Northern blots were all prepared by PCR with oligonucleotides synthesised according to the appropriate sequences found in the yeast genome database as primers and yeast genomic DNA as template. The Northern blot was quantitatively analysed by using a phospho-imager (FUJIX BAS1000) and the BASMac program (Macintosh). The expression level of the mRNA of interest was quantified relative to the intensity of the hybridization signal of the constitutively expressed actin mRNA of the same sample.

Cdc28 kinase activity assay

Cultures of 10 ml were harvested at the indicated times, centrifuged, washed and resuspended in 50 l cold extraction buffer (EB: 80 mM -glycerophosphate (pH 7.3), 20 mM EGTA, 15 mM MgCl_2, 1 mM DTT) containing protease inhibitors (2 g/ml leupeptin, 2 g/ml pepstatin and 1 mM PMSF) and 1 mM vanadate. Cells were broken by vigorously vortexing with glass beads and the lysate was cleared by centrifugation for ten minutes at 4 °C in a microcentrifuge at 13,000 g. 10 l of p13-Sepharose beads were added to 40 l cell-free extract in order to affinity-purify the Cdc28 kinase and incubated overnight at 4 °C on a rotating wheel. Next day, the affinity complex was washed three times with EB buffer prior to assay of the Cdc28 kinase activity with Histone H1 as substrate as described [Derua et al 1997].

Microscopy and flow cytometry

Culture samples of 1 ml were taken at the indicated times and resuspended in 100 l fixation buffer (0.125 M KH₂PO₄ (pH 6.5) and 3.7 % formaldehyde) to fix the cells prior to microscopy. To determine the budding index and shmoo formation in response to -factor, 200 cells were counted at each time point. For flow cytometry 1 ml culture samples were treated as described [Butler et al 1991] using a Becton Dickinson FACScan and data were analysed using the Lysis II software.

ACKNOWLEDGEMENTS

We thank Maria Veeckmans for expert technical assistance, Valère Feytons for synthesising the oligonucleotides and Jan Morren for his help with the layout of the Figures. H. Ronne and M. Stark kindly provided the PP2A-deficient strains.

This work was supported by the Fonds voor Wetenschappelijk Onderzoek (F.W.O.), the Geconcerteerde Onderzoeks Acties (G.O.A.), the Human Frontier Science Program and the European Commission (E.C.) Human Capital and Mobility Program and Biomed2. C.V.H. is a postdoctoral fellow of the F.W.O.

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