Báo cáo khoa học: Transcriptional responses to glucose at different glycolytic rates in Saccharomyces cerevisiae ppt

Transcriptional responses to glucose at different glycolytic rates in Saccharomyces cerevisiae Karin Elbing 1 , Anders Sta ˚ hlberg 1 , Stefan Hohmann 2 and Lena Gustafsson 1 1 Department of Chemistry and Bioscience-Molecular Biotechnology, Chalmers University of Technology, Go ¨ teborg, Sweden; 2 Department of Cell and Molecular Biology-Microbiology, Go ¨ teborg University, Go ¨ teborg, Sweden The addition of glucose to Saccharomyces cerev isiae cells causes reprogramming of gene expression. Glucose is sensed by membrane receptors as well as (so far elusive) intracellular sensing mechanisms. T he availability of four yeast s trains that display d ifferent hexose uptake capacities allowed us to study glucose-induced effects at different glycolytic rates. Rapid glucose responses were observed i n all strains able to take up glucose, consistent with intracellular sensing. The degree of long-term responses, however, clearly correlated with the glycolytic rate: glucose-stimulated expression of genes e ncoding enzymes o f the lower part of glycolysis showed an almost linear correlation with the glycolytic rate, while expression levels of genes encoding gluconeogenic enzymes and invertase (SUC2) showed an inverse correla- tion. Glucose control of SUC2 expression is mediated by the Snf1-Mig1 pathway. Mig1 dephosphorylation upon glucose addition is known to lead to repression of target genes. Mig1 was initially dephosphorylated upon glucose addition in all strains able t o take up glucose, but remained dephospho- rylated only at high glycolytic rates. Remarkably, transient Mig1-dephosphorylation was accompanied by t he repres- sion of SUC2 expression at high glycolytic rates, but sti- mulated SUC2 expression at low glycolytic rates. This suggests that Mig1-mediated repression can be overruled by factors mediating induction via a low glucose signal. A t low and moderate glycolytic rates, Mig1 was partly dephos- phorylated both in the presence of phosphorylated, active Snf1, and unphosphorylated, inactive Snf1, indicating that Mig1 was actively phosphorylated and dephosphorylated simultaneously, suggesting independent control of both processes. Taken together, it appears that glucose addition affects t he expression of SUC2 as well as Mig1 activity by both Snf1-dependent and -independent mechanisms that can now be dissected and r esolved as e arly and l ate/sustained responses. Keywords: Saccharomyces cerevisiae; Mig1; Snf1;glucose repression; glucose signal. Addition of glucose to Saccharomyces cerev isiae cells growing in the absence of glucose causes an extensive reprogramming of gene expression and metabolism. These changes affect c hromatin s tructure, t ranscription, mRNA stability, translation and post-translational modifications [1–4]. A range of d ifferent signalling p athways, including, among others, the Snf1–Mig1 pathway, the Snf3–Rgt2 pathway and the Ras-cAMP pathway [5], are r esponsible for these effects. Glucose sensing appears to occur at different levels. While membrane-localized receptors (Gpr1, Snf3, Rgt2) h ave been reported, other pathways appear to be controlled by so far elusive intracellular signals and sensors. In this work we focus on such effects previously reported to probably be the result of intracellular sensing/ signalling. We have addressed the question of how signalling and its output are affecte d by different glycolytic r ates at identical extracellular conditions. Our data show that even seemingly simple responses can b e dissected into different components with potentially different underlying mecha- nisms. This study focused on the effects on mRNA levels of different sets of genes. One such set are genes encoding enzymes of glycolysis. While expression of genes encoding enzymes operating in both glycolysis and gluconeo genesis usually remain constitutive [6,7], expression of genes for enzymes specific to the lower part of glycolysis is stimulated upon glucose addition [8]. The underlying signalling pathway is not understood. However, it has been reported that stimulated expression requires glucose metabolism through the upper part of glycolysis [9]. On the other hand, expression of genes encoding enzymes specific for gluconeogenesis, respiration, or the uptake and utilization of alternative carbon sources, is efficiently repressed b y glucose [4]. Glucose repression is a c omplex p rocess involving differ- ent regulators affecting different subsets of genes. Best studied is the Snf1–Mig1 pathway, which is involved in the (de)repression of genes encoding enzymes needed for the utilization of alternative carbon sources as well as for gluconeogenesis and respiration. The p rotein kinase Snf1 is activated by phosphorylation a t low/no glucose [10]. Recently, three protein kinases – Elm1, Tos3 and Pak1 [11–13] – were identified that seem to mediate Snf1 activation. It is unclear how these kinases are c ontrolled, Correspondence to K. Elbing, Department of Chemistry and Bioscience-Molecular Biotechnology, Chalmers University of Technology, PO Box 462, 405 30 Go ¨ teborg, Sweden. Fax: +46 31 773 25 99, Tel.: +46 31 773 25 81, E-mail: Karin.Elbing@molbiotech.chalmers.se Abbreviations:DAPI,4¢,6-diamidino-2-phenylindole dihydro- chloride; HA, haemagglutinin; QPCR, quantitative PCR. (Received 6 August 2004, revised 21 October 2004, accepted 22 October 2004) Eur. J. Biochem. 271, 4855–4864 (2004) Ó FEBS 2004 doi:10.1111/j.1432-1033.2004.04451.x but it appears that the hexokinases, Hxk1 and Hxk2, may play some role in this process [14–17]. In addition, a decreased Glc7 phosphatase activity may also contribute to Snf1 activation, as has been s hown by deletion s tudies of REG1 by Treitel et al. and McCartney et al. [ 10,18]. Also, protein interactions, as well as carbon source-dependent phosphorylation of Reg1, may effect Reg1/Glc7 activity [19,20]. An active Snf1 phosphorylates at least four sites in the transcriptional rep ressor Mig1. M ig1 phosphorylation causes the majority of t he protein to exit the nucleus [21]. Recent data, however, suggests t hat phosphorylation-medi- ated altered interaction with the two co-repressors Cyc8 (Ssn6) and Tup1 on target promoters i s the primary cause for the switch between repression and derepression [22]. Time-course analyses suggested that the process of glucose repression consists of a short- and a long-term response (minutes and hours, respectively) [23,24]. Those could be d istinguished on the basis of their different requirements f or sugar kinases, suggesting different signal- ling pathways. While long-term glucose repression required Hxk2, for short-term repression any of the three sugar kinases, Hxk1, Hxk 2 or Glk1, was sufficient [23,24]. It should be noted that Hxk2 does not have a unique role in glucose repression, as often c laimed in the literature, but that Hxk1 also contributes to glucose and, in particular, to fructose repression [24]. Earlier studies showed a correlation between glucose consumption rate and glucose repression [25–27]. Our previously reported series o f strains, in which sugar uptake is mediated by the individual expression of different native and chimeric hexose transporters [28,29], display a wide spectrum of glucose uptake rates. These strains a re therefore useful for investigating the effects of different glycolytic rates on glucose-induced signalling pathways. For this study we have chosen four strains, which represents t he full range o f glycolytic r ates: a wild- type, with a high glycolytic rate; a HXT-null strain, which does not take up glucose owing to the d eletion of all known hexose transporter (HXT) genes; a strain expres- sing Hxt7 as the sole s ugar transporter, which displays relatively high sugar uptake rates; and a strain that expresses Hxt-Tm6*, a chimera of Hxt1 and Hxt7. Hxt- Tm6* mediates low uptake rates and, for that reason, the strain does not produce ethanol also in the presence of high external sugar levels [28,29]. Materials and methods Strains The strains u sed are liste d in Table 1 and all derive from CEN.PK2-1C MATa leu2-3 122 ura3-52 trp1-289 his3-D MAL2-8 c SUC2 hxt12D [30]. KOY.PK2-1C83 (wild-type) is the prototrophic version of the CEN.PK2-1C s train [28]. In KO Y.VW100P (HXT-null), all known hexose transport- ers have been deleted and an expression cassette h as been introduced in the HXT3-6-7 locus [28]. KOY.HXT7P (HXT7)andKOY.TM6*P(HXT-TM6*) have HXT7 and the chimera HXT-TM6*, respectively, cloned into this expression cassette [28,29]. Plasmid pRS316 carrying either SNF1 [10] or MIG1 [31] tagged with the haemagglutinin (HA) epitope at the C-termini w as transformed into the KOY.PK2-1C82, KOY.HXT7, KOY.TM6* and KOY.VW100 strains, which are isogenic to the strains listed above e xcept that they contain the ura3-52 marker. The resulting transform- ants are h ence prototrophic. For Mig1-GFP localization, plasmid BM3315 [21] was transformed into the s ame strains. Cultures Cells were precultured at 30 °Cfor48hin50mLof complete minimal medium [32], supplemented with 1% (v/v) ethanol. Fermentors containing 1.5 L of minimal medium (5· concentrated) were inoculated to an attenuance (D), at 610 nm, of 0.05. Conditions were maintained constant at 30 °C, 1500 r.p.m. and pH 5.0. Off gas was maintained at 0.75 LÆmin )1 by using a mass flow regulator. Gas was passed through a condenser to avoid evaporation. Carbon dioxide production and oxygen consumption were measured on-line (type 1308; Bruel and Kjaeer, Naerum, Denmark). At a D 610 of 1 to 1.5, glucose was added to a final c oncentration of 5% and samples were taken at 1, 5, 10, 15, 20, 30 and 60 min as well as at residual glucose concentrations of 1.5–2.5%. For the HXT-null st rain, samples were taken in the ethanol consumption phase following glucose addition. Biochemical determinations and consumption rates Glucose and ethanol were measured in the s upernatant (1 min at 16 060 g) using enzymatic combination kits (Roche). Several samples were taken during logarithmic growth on glucose, and t he specific glucose consumption rate was determined at a specific time-point. Quantitative PCR (QPCR) Samples for RNA extraction were taken into ice-cold water. RNA was extracted, treated with DNase, and c hecked for purity by agarose-gel electrophoresis. Samples were pre- pared [28] and normalized against the quotient between the levels of the ACT1 and IPP1 mRNAs. The lowest value for each gene was set to 1. The standard deviation of t he QPCR is < 0.25 cycles and at least two independent fermentations were performed. Duplicate samples from each fermentation w ere analysed. Protein extracts and Western blot analysis Cells were harvested and proteins extracted as described in McCartney et al. [10]. For the detection of Mig1-HA, samples were separated by PAGE on 7.5% (w/v) SDS gels and blotted onto nitrocellulose membranes. Membranes were blocked at room temperature for 1 h in TTBS [TBS containing 0.1% (v/v) Tween-20] containing 3% (w/v) BSA, washed three t imes (5 min each wash) in TTBS, incubated at 4 °C f or 3 h with HA mAb (1 : 1000) (Amersham) i n TTBS containing 3% (w/v) BSA, washed three times (5 min each wash) in TTBS, and incubated for 1 h at room temperature with secondary anti-mouse immunoglobulin (1 : 5000 dilution) in TTBS containing 3% (w/v) BSA. The membrane was washed three times ( 5 min each wash) in 4856 K. Elbing et al. (Eur. J. Biochem. 271) Ó FEBS 2004 TTBS prior to detection by chemiluminescence using ELC plus (Amersham). Snf1-HA samples were dialysed against buffer overnight [150 m M NaCl, 1% (v/v) Triton X-100, 0.5% (w/v) deoxycholate, 50 m M Tris/HCl, pH 8.00, supplemented with 50 m M sodium fluoride and 5 m M sodium pyrophosphate], and 400 mg of total protein was used for immunoprecipitation of Snf1-HA [10]. The preci- pitate was dissolved in SDS sample buffer, separated by PAGE on a 7.5% (w/v) SDS gel, blotted onto nitrocellulose membrane and phospho-Snf1 was detected by using the a-PT210 antibody, as described by McCartney et al.[10]. As a c ontrol for equal loading, membranes were stripped and the HA epitope on Snf1 was detected by a monoclonal anti-HA immunoglobulin, as described above. Phosphatase treatment For phosphatase treatment of Mig1, 50 lg of total protein extract was precipitated with 10% (w/v) trichloroacetic acid and sedimented for 30 min at 4 °C. The sediments were washed twice w ith ice-cold 100% acetone for 15 min and centrifuged for 15 min between each wash, air-dried, resuspended in 82 lLofH 2 O containing 10 lLof10· phosphatase buffer and 8 U calf intestine alkaline phospha- tase (Roche), and incubated a t 37 °C for 1 h. Samples were again precipitated with trichloroacetic acid, resuspended i n SDS sample buffer, boiled for 5 min and electrophoresed. Gels were blotted and proteins detected, as described above (in Western blot analysis), for M ig1-HA detection. Determination of invertase activity Cells were grown in Erlenmeyer flasks containing 2· minimal medium [32] supplemented w ith 5% ( w/v) glucose to a D 610 of 1, then harvested by centrifugation. P rotein extracts and m easurements of invertase activity were performed as described previously [33]. Microscopy Localization of Mig1-GFP was visu alized by using a GFP filter on a Leica DMRXA microscope. DNA was stained by 4 0 ,6-diamidino-2-phenylindole dihydrochloride (DAPI) (1 lgÆmL )1 )for10minat30°Cafterwhichthe cells were quickly washed three times in growth media. Results Four strains displaying different glycolytic rates The wild-type, HXT7 and HXT-TM6* strains display high (15.8 mmol g )1 Æh )1 ), intermediate (10.7 mmol g )1 Æh )1 )andlow(3.5mmolg )1 Æh )1 ) g lucose consumption rates, respectively [28,29]. The HXT-null strain neither takes up glucose nor grows with glucose as the sole carbon source [34] (Fig. 1). In order to follow glucose- induced responses, the yeast strains were grown in the presence of 1% (v/v) ethanol to a D 610 of 1, pulsed w ith glucose to a final concentration of 5 %, and sampled over a period of 1 h as well as in the subsequent glucose consumption phase (Fig. 1). After the glucose pulse, the wild-type and HXT7 strains displayed a clear biphasic growth with an initial respiro-fermentative phase where ethanol was produced (Fig. 1) a nd a subsequent respirat- ory phase where this ethanol was then consumed (data not shown). In the HXT-TM6* strain, glucose is only respired, as described previously [28,29]. Following glucose addition the HXT-TM6* strain initially consumed glucose Table 1. Saccharomyces cerevisiae strains. Strain Genotype Source or reference KOY.PK2-1C83 (wild-type) MATa MAL2-8 c SUC2 Prototrophic [28] KOY.PK2-1C82 MATa MAL2-8 c SUC2 ura 3-52 Auxotrophic: this study KOY.VW100P (HXT-null) MATa MAL2-8 c SUC2 hxt17D ura3-52 gal2 D ::loxP stl1 D::loxP agt1 D::loxP ydl247w D::loxP yjr160c D::loxP hxt13 D::loxP hxt15 D::loxP hxt16 D::loxP hxt14 D::loxP hxt12 D::loxP hxt9 D ::loxP hxt11 D::loxP hxt10 D::loxP hxt8 D::loxP hxt514 D::loxP hxt2 D::loxP hxt367 D::loxP Prototrophic [28] Integration cassette at former HXT367 site containing the truncated, constitutive promoter of HXT7 [46], the KlURA3 open reading frame for counter selection, and the HXT7 terminator KOY.VW100 As KOY.VW100P but the KlURA3 in the integration cassette has been replaced with the KanMX Auxotrophic: this study KOY.HXT7P (HXT7) KOY.VW100P Integration into the cassette: HXT7prom-HXT7-HXT7term, ura3-52::URA3 Prototrophic [29] KOY.TM6*P (HXT-TM6*) KOY.VW100P Integration into the cassette: HXT7prom-TM6*-HXT7term, ura3-52::URA3 Prototrophic [28] KOY.HXT7 KOY.VW100P Integration into the cassette: HXT7prom-HXT7-HXT7term, ura3-52 Auxotrophic: this study KOY.TM6* KOY.VW100P Integration into the cassette: HXT7prom-TM6*-HXT7term, ura3-52 Auxotrophic: this study Ó FEBS 2004 Glucose response in S. cerevisiae (Eur. J. Biochem. 271) 4857 and e thanol simultaneously, a nd once ethanol was deple- ted it continued to catabolize g lucose (Fig. 1). The HXT- null strain continued consuming ethanol, leaving glucose unconsumed. Short-term response to glucose addition Using QPCR we monitored the response to glucose of four glucose-induced genes encoding enzymes of the lower part of glycolysis (TPI1, PGK1, PDC1 and ADH1), of three glucose-repressed genes encoding enzymes in gluconeogen- esis and the glyoxylate cycle (FBP1, MDH2, ADH2), as well as of the glucose-repressed SUC2 (invertase) gene. In wild- type cells, expression of all four glycolytic genes was strongly stimulated by glucose, reaching a plateau after about 30 min ( Fig. 2). Expression of t hese genes was not stimulated at all in the HXT-null strain, o r rather diminished inthecaseofPGK1 and TPI1. The strains expressing Hxt7 and Hxt-TM6* as sole hexose transporter showed intermediate levels of stimulation, which differed in a gene-specific manner (Fig. 2). Generally, it appeared that the degree of induction correlated approximately with the glycolytic rate (measured as the glucose consumption rate) of the strains. The mRNA level of the gluconeogenic and glyoxylate cycle genes, FBP1, ADH2 and MDH2, was rapidly diminished following glucose addition in all strains able to take up glucose. In the HXT-null strain, the mRNA of all these g enes transiently increased and the n e ither plateaued or decreased. The expression level of SUC2 diminished in the wild-type yeast and in the strain e xpressing HXT7, w hile it did not respond to glucose addition in the HXT-null strain. In the HXT-TM6* strain, expression of SUC2 was transiently stimulated. Long-term glucose response In order to study the long-term glucose response, samples from cells growing exponentially with glucose w ere t aken when 1.5–2.5% of glucos e was still pres ent in the culture medium (indicated in Fig. 1). For th e HXT-null strain, samples were taken 5–8 h after g lucose addition when the strain was still consuming ethanol. For the glucose-induced glycolytic genes TPI1, PGK1, PDC1 and ADH1, the long-term expression level showed an approximately linear correlation with the glycolytic rate, especially for PGK1 and PDC1 genes (Fig. 3). In the HXT- null strain, expression levels of TPI1, PDC1 an d ADH1 did not differ from those of cells growing in the presence of ethanol only, while the mRNA level of PGK1 was threefold lower. Expression of the gluconeogenic genes FBP1 and MDH2 was strongly repressed by 5% glucose in the wild- type an d HXT7 strains and repressed to a lower extent i n the HXT-TM6* strain (Table 2). Expression of FBP1 and MDH2 was unaffected by glucose in the HXT-null strain. Expression of ADH2 was strongly repressed i mmediately after glucose addition and remained repressed in the wild- type and HXT7 strains. In the HXT-TM6* strain, however, ADH2 remained repressed during the phase of glucose/ ethanol co-consumption (data not shown), but when ethanol was depleted and the strain only consumed glucose, ADH2 became fully derepressed (Table 2). The reason for this behaviour is unclear. Expression of SUC2 was repressed twofold in the wild-type yeast, slightly increased in the HXT7 strainandstimulatedfourfoldintheHXT-TM6* strain during growth on g lucose. In the HXT-null strain, expression of SUC2 did not seem to respond to glucose (Table 2). I n agreement with mRNA levels, invertase activity measurements with glucose-grown cells showed increased activity in the HXT7 and HXT-TM6*strains, while activity remained at a low level in the wild-typ e yeast. The HXT-null strain, which was grown on ethanol supple- mented with 5% glucose, displayed an intermediate level of activity (Fig. 4). Fig. 1. Cu ltu re profiles. Measurements of glucose (gÆL )1 )(r), ethanol (gÆL )1 )(h) and attenuance (D 610 )(m) for the wild-type, HXT7, HXT- TM6*andHXT-null strains following glucose a ddition at 0 h to cells grown on ethanol. The bracket indicates samples taken during the first 60 min after glucose ad dition, and the arro ws specify t he time-points for sampling during growth on glucose. 4858 K. Elbing et al. (Eur. J. Biochem. 271) Ó FEBS 2004 Snf1 and Mig1 phosphorylation in the wild-type yeast, and in HXT7 , HXT-TM6* and HXT-Null strains Because of the interesting expression pattern of SUC2,we investigated the state of the glucose repression signalling pathway by monitoring the phosphorylation patterns of Mig1 and Snf1. Snf1 is activated by phosphorylation at low/ no glucose [10,35], and phosphorylation on the critical T210 residue can be monitored by using a specific antibody [10]. Active Snf1 phosphorylates the repressor Mig1 on multiple sites, lead ing t o d erepression of target genes, such as SUC2 [18,36,37]. Mig1 phosphorylation can be visualized as a mobility shift by using HA-tagged Mig1 and immunoblot- ting. The short-term response was studied by monitoring the electrophoretic migration of Mig1 following the addition of glucose t o e thanol-grown cells (the same conditions as in Figs 1 and 2). In ethanol-grown cells, M ig1 appeared as a ladder of bands (Fig. 5A), indicating that the protein was phosphorylated to a different extent and was partially inactive as a repressor. Interestingly, Mig1 from cells growing in the presence of 0.05% glucose migrated as a single slow band, indicating that under these conditions Mig1 is fully phosphorylated and inactive. This fits with the observation that SUC2 expression is much higher in cells growing in the presence of low glucose levels than in ethanol medium ([38], o wn unpublished data). Mig1 from cells growing with 5% g lucose, on t he other hand, migrated as a single fast band of fully dephosphorylated and hence actively repressing Mig1 (Fig. 5 A, see also Fig. 6 ). Interestingly, in all glucose-utilizing strains, the addition of glucose to ethanol-grown cells caused a c ollapse of the Mig1 ladder to the unph osphorylated (actively repressing) form. Only in the HXT-null strain was the band pattern largely unaltered. W hile Mig1 remained unphosphorylated in the wild-type yeast throughout the time course of the experiment, it appeared to be rephosphorylated in the HXT7 and HXT-TM6* strain towards the end of the time course. As it appeared that the level of Mig1 increased during the time course of the experiment, we performed QPCR analysis of MIG1 gene expression (Fig. 5B). Indeed, Fig. 2. Gene expression analysis and quantitative PCR (QPCR) analysis. Diagram of central metabolism to indicate the position of t he relevant enzymes in metabolism. mRNA levels were determined for TPI1, PGK1, PDC1, ADH1, FBP1, MDH2, ADH2 and SUC2 for the wild-type (j), HXT7 (h), HXT-TM6*(m)andHXT-null (s) strains. Cells were grown in 1% ethanol, glucose was added at 0 h to a final concentration of 5% and samples were t aken during t he first hour after glucose addition. O ne representative result is shown. Ó FEBS 2004 Glucose response in S. cerevisiae (Eur. J. Biochem. 271) 4859 expression of MIG1 was stimulated upon glucose addition, in accordance with recently published data [39]. Stimulation of expression inversely correlated with the glycolytic rate and, interestingly, was apparent even in the HXT-null strain. To monitor the long-term glucose response, the four strains were grown in the presence of a high (5%) concentration of g lucose to a D 610 of 1.0. A sample was shifted to a low (0.05%) concentration of glucose as a control, and the phosphorylation state of Snf1 and the mobility pattern of Mig1 were analysed (Fig. 6). Mig1 from wild-type cells migrated as the a pparently fully phosphor- ylated form on the low concentration of glucose and as the dephosphorylated form on the high concentration of glucose (Fig. 6A). Migration of this latter band did not change upon treatment with alkaline phosphatase, confirm- ing that it represents the fully dephosphorylated form (Fig. 6C). Snf1 was largely unphosphorylated in wild-type cells growing in a high concentration of glucose, while the level of phosphorylated Snf1 was increased in cells shifted to a low concentration of glucose. In the HXT-null strain, Mig1 migrated at an intermediate rate (high glucose) or as a diffuse ladder ( low glucose), and Snf1 was phosphorylated under both conditions. In the HXT7-expressing strain, Snf1 was (as in the wild-type) unphosphorylated when grown on a high concentration of glucose, whereas Mig1 was partially phosphorylated (Fig. 6A,B), as also illustrated by the fact that the Mig1-band migrated more quicly after phosphatase treatment (Fig. 6C). In the HXT-TM6* strain, Snf1 was strongly phosphorylated in cells growing in conditions of both high and low glucose, consistent with a fully glucose- derepressed s tate o f the cell. Interestingly, it appeared that Mig1 assumed an intermediate level of phosphorylation in the HXT-TM6* strain on high glucose (Fig. 6A,B). When comparing the three strains able to take up glucose, it appeared that the phosphorylation of Mig1 correlated well with the glycolytic rate, wh ereas Snf1 pho sphorylation did not (Fig. 6A,6B). A good correlation was also seen of the glycolytic rate, apparent phosphorylation state of Mig1, and its subcellular localization. Dephosphorylated Mig1, for example i n glu- cose-grown wild-type cells, has been reported to concentrate in the nucleus, and this was also observed i n the present study (Fig. 7). Mig1 from HXT7-expressing cells showed increased nuclear localization, although not as strongly as in the wild-type. In HXT-TM6*, as well as in HXT-null cells, Mig1 was localized diffusely throughout the c ell after the glucose pulse. I n the latter two strains, DAPI staining did not clearly reveal the nucleus owing to a high abundance o f mitochondria, which is consistent with the respiratory metabolism of these strains. Discussion In this study we have used yeast strains with a very broad range of glycolytic rates to study glucose-induced responses while maintaining identical growth conditions as well as high external glucose concentrations. The results confirm previous reports in that the signalling pathways studied here are triggered inside the c ell rather Fig. 3. Correlation of expression levels and glucose consumption rates. Plot of the relative fold change of the TPI1, PGK1, PDC1 and ADH1 genes in the wild-type , HXT7, HXT-TM6*andHXT-null strains during glucose growth as compared to ethanol growth vs. the glucose consumption rate. The names of th e strains are indicated above t he graph to show which strain displayed which glucose consumption rate. Error bars sho w standard deviation of the relative fold change from four independent measurements. Table 2. Fold changes during 5% glucose growth as compared to ethanol growth for glucose-repressed genes. Significantly repressed genes are indicated in bold italic; significantly induced genes are shown in bold. Data for the wild-type (WT) an d TM6* strains were previously published in Otterstedt et a l.[28]. Gene WT HXT7 HXT-TM6* HXT-null FBP1 )63 )71 )13 )1.5 MDH2 )120 )77 )7.9 )1.3 ADH2 )105 )120 )1.0 )2.0 SUC2 )2.2 1.5 3.5 1.3 Fig. 4. Relationship between invertase activity and glucose consumption rate. Plo t of specific invertase activity of the wild-type, HXT7, HXT- TM6*andtheHXT-null strai ns during growth on 5% glucose vs. glucose consumption rate. Error bars show t he standard deviation of invertase activity from at least three independent measurements. 4860 K. Elbing et al. (Eur. J. Biochem. 271) Ó FEBS 2004 than by plasma membrane-localized receptors. This was first illustrate d by the fact that the HXT-null strain, which does not take up glucose, also does not respond to glucose addition. We only observed t wo potentially relevant devi- ations: expression levels of gluconeogenic genes transiently increased upon glucose addition to the HXT-null strain, and the expression l evel of MIG1 was moderately stimulated. These effects could be caused either by minute amounts of glucose diffusing into cells of the HXT-null strain or to signalling pathways sensing external glucose, such as the Gpr1-PKA pathway. That signalling is t riggered inside the cells is further indicated by the fact that different glucose consumption, and h ence glycolytic rates, caused a different signalling output. The actual signal(s) and sensing mecha- nisms still remain to be identified, but strains like those used here will certainly be useful in such studies. We observed an a lmost perfect correlation between the apparent glycolytic rate and t he degree of induction of glycolytic gene expression. This is consistent with previous chemostat studies of the CEN.PK strain cultured at different glycolytic rates within the respiro-fermentative phase, i.e. high dilution rates [40]. Interestingly, a ll glu cose- consuming strains responded equally quickly to glucose addition and the difference was manifested as different amplitudes of expression. This suggests that the – so far elusive – sensing mechanism somehow monitors quantita- tive differences of the glycolytic rate. Similarly, expression of gluconeogenic genes was repressed in all three glucose-consuming strains equally quickly. Hence, consistent with previous studies, repression of these genes is very sensitive to glucose [41]. However, gluconeogenic genes were repressed to a much lesser extent in HXT-TM6* cells growing in the presence of high glucose levels, suggesting t he inte resting s cenario that HXT-TM6* cells co-express glycolytic and gluconeogenic enzymes. Potential futile cycling is n ot likely as a higher biomass is obtained in the HXT-TM6* strain as compared to wild-type yeast [28]. Moreover, the alcohol dehydrogenases seem to be regulated in an interesting way in this strain. Expression of ADH2, which encodes the glucose-repressed alcohol dehy- drogenase responsible for ethanol consumption, was strongly repressed in HXT-TM6* cells during glucose/ ethanol co-consumption. It is possible, that the enzyme encoded by the glycolytic ADH1, whose expre ssion w as stimulated fourfold under these conditions (data not shown) takes over the role of Adh2. Once ethanol was depleted and the strain grew solely on glucose, ADH2 expression was again derepressed to the same level as before glucose addition. The glycolytic rate was identical during glucose consumption and glucose/ethanol co-consumption in the HXT-TM6* cells (data not shown). The expression of SUC2, a classical model for a glucose- regulated gene , appeared particularly interesting, as it showed very different responses in the f our strains. Fig. 5. Mig1 gel mobility pattern in response t o glucose addition. Glu- cose was add ed at 0 h to a final concentration of 5%. (A) The phos- phorylation level of Mig1 was estimated as a band-shift. Samples (30 lg) from wild-type cells grown at high (5%) or low (0.05%) concetrations of glucose were loaded as a comparison. Slow migration indicates fully phosphorylated and fast migration fully dephosph or- ylated Mig1 (see Fig. 6B for phosphatase-treated controls). A total of 60 lg o f extract was l oaded for wild-type, HXT7, HX T-TM6*and HXT-null strains. (B) mRNA expression of Mig1 during the first hour after glucose addition, as determined by quantitative P CR (QPCR). Wild-type (j), HXT7 (h), HXT-TM6*(m), HXT-null strains (s). Fig. 6. Mig1 and Snf1 phosphorylation in glucose-growing cells. Strains were grown in 5% glucose (H) and shifted to 0.05% glucose (L) for 2h.TheHXT-null strain was grown in 1% ethanol supplemented with 5% glu cose ( H) and shifted to 0.05% glucose (L) for 2 h. (A) The migration pattern of Mig1. A t otal of 60 lgofextractwasloadedin each lane. (B) Detection of phosphorylated Snf1 by using an antibody specific for Snf1 phosphorylated at T210. The haemagglutinin (HA) signalwasusedasaloadingcontrol.(C)Treatmentofextractswith alkaline phosphatase as a control for the Mig1 phosphorylation state. A total of 50 lg of total protein from the wild-type, HXT7 and HXT- TM6* strains were incubated with and without calf intestine alkaline phosphatase (AP). Un treated wild-type samples were loaded as migration comparisons. Ó FEBS 2004 Glucose response in S. cerevisiae (Eur. J. Biochem. 271) 4861 Employing strains expressing different hexose transporters or a given transporter a t different levels, it has previously been observed that there is a goo d correlation between the apparent glycolytic rate and the degree of long-term glucose repression [42–44]. This is confirmed here, although the picture is complicated by the fact that expression of SUC2 is stimulated by low glucose levels ([38], own data). Stimulated SUC2 expression upon glucose ad dition in the HXT-TM6* strain illustrates that the glucose repression signalling system perceives a Ôlow glucoseÕ signal, despite the fact that the external gluco se level is high. The derepressed state of this strain is confirmed by a high level of phosphorylation o f Snf1. In order to achieve complete glucose repression, the wild-type glycolytic rate seems to be required because even the HXT7 strain, which displayed  two-thirds of the wild- type rate, did not fully repress SUC2 expression. Expression of SUC2 and gluconeogenic genes is con- trolled by the Snf1 kinase and th e Mig1 repressor. Gluconeogenic genes are also controlled by the Sn f1- dependent Cat8 and Sip4 a ctivators. Monitoring Snf1 and Mig1 phosphorylation revealed some unexpected observa- tions that will require further investigation. Perhaps most perplexing is the observation that Mig1 becomes rapidly dephosphorylated upon glucose addition in the HXT-TM6* strain while, at the same time, the expression level of SUC2 strongly increases. This is in clear contradiction to the current view that dephosphorylated, nuclear Mig1 represses SUC2 expression. This observation suggests that the system which mediates induction of SUC2 at a low glycolytic rate is able to overcome Mig1-mediated r epression. Another surprising observation concerns the only partial phosphory- lation of Mig1 in the HXT-TM6* strain growing at high glucose levels, despite the fact that Snf1 is strongly phosphorylated. Partial phosphorylation of Mig1 is also seen in the HXT7 strain at high glucose, even though Snf1 is unphosphorylated. T his i s n ot caused by the strain being unable to dephosphorylate Mig1, as this species is observed transiently upon glucose addition. This observation sug- gests that the phosphorylation state of Mig1 is not only controlled by Snf1-dependent phosphorylation but, obviously, also by dephosphorylation, which is mediated by the Glc7-Reg1 system [18]. If indeed the observed Mig1 phosphorylation pattern is caused by simultaneous phos- phorylation/dephosphorylation, these two processes might be controlled by different signallin g mechanisms. The fact that one distinct Mig1 band is observed under these conditions further suggests that certain phosphorylation sites are used preferentially, which will be tested in the future. The interplay between th e t wo processes apparently allows fine-tuning of the M ig1 phosphorylation level. An almost linear correlation between Mig1 activity and sites phosphorylated by Snf1 has been observed [21]. Future work, for which the strains u sed here w ill be instrumental, will address t he precise m echanisms controlling Mig1 activity and their interplay with the factor(s) m ediating induction by low glucose. It has previously been proposed that the establishment of glucose repression can be dissected into a short-term and a long-term response. That proposal was based on d ifferent roles of the sugar kinases: the hxk2D mutant displayed short-term glucose r epression but was unable t o maintain repression [24]. In a similar way, the HXT7 and HXT-TM6* strains displayed short-term Mig1 dephosphorylation (sup- posedly activating the repressor, although stimulated SUC2 expression was observed, see a bove) but subsequently Mig1 became rephosphorylated. Although unlikely, we cannot exclude that in our experiment this biphasic behaviour is caused by properties of the single hexose transporters Fig. 7. Mig1-GFP localization in the wild-type, HXT7 and HXT-TM6* strains growing on 5% glucose and in the HXT-null strain grown on 1.5% ethanol supplemented with 5% glucose. BF, bright field DAPI: staining with DAPI to determine the location of the nucleus. In HXT-TM6*andHXT-null cells the position of the nucleus is difficult to determine owing to the abundance of mitochondria. 4862 K. Elbing et al. (Eur. J. Biochem. 271) Ó FEBS 2004 expressed in these cells. Both Hxt7 and Hxt-TM6* are high- affinity glucose transporters, which in wild-type cells are active at low/no glucose and inactivated in medium containing a h igh concentration of glucose [45]. H ence, it may be that during adaptation to glucose, the levels of active transporters diminish, although quantification of the transporter mRNA of the chimeras shows identical expres- sion during growth on ethanol and glucose (data not shown). Ano ther interpretation for the biph asic behaviour is, like in the hxk2D mutant, the initial, acute response and the late s ustained response are governed by different regulatory systems. In that scenario, the initial response seems to be more sensitive to glucose, while the sustained response would require higher glucose levels. Acknowledgements We ackn owledge Martin S chmidt and Arle Kruckeberg f or critical reading of the manuscript. We thank Martin Schmidt for the Mig1-HA and Snf1-HA plasmids and the Snf1 a-PT210 antibody. We also thank Mark Johnston for the Mig1-GFP plasmid. This work was supported by the European Commission ( contract BIO4-CT98-0562) a s well as grants from the Swedish N ational Energy Administration (P1009-5), the Swedish Council for Forestry and Agricultural Research (52.0609/ 97) and Swedish Rese arch Council (621-2001-1988) to Lena Gustafs- son. Stefan Hohmann holds a research position from t he Swed ish Research Council. References 1. Winderickx, J.G. & Taylor. P.M. (2004) Nutrient Induced Responses in Eukaryotic Cells, 1st edn. Springer-Verlag, Heidelberg. 2. Scheffler, I.E., de la Cruz, B.J. & Prieto, S.(1998)Control of mRNA turnover as a m echanism of gluc ose repression in Saccharomyces cerevisiae. Int. J. Biochem. Cell. Biol. 30, 1175–1193. 3. Sudarsanam,P.,Cao,Y.,Wu,L.,Laurent,B.C.&Winston,F. (1999) The nucleosome remodeling complex, Snf/Swi, is r equired for the maintenance of transcription in vivo and is partially redundant with the histone acetyltransferase, Gcn5. EMBO J. 18, 3101–3106. 4. Gancedo, J.M. (1998) Yeast carbon catabolite r epressio n. Microbiol. Mol. Biol. Rev. 62, 3 34–361. 5. Rolland, F., Winderickx, J. & Thevelein, J.M. (2002) Glucose- sensing and -signalling mechanisms in yeast. FEM Yeast Res. 2, 183–201. 6. Chambers, A., Packham, E.A. & Graham, I.R. (1995) Control of glycolytic gen e e xpressio n in the budding yeast (Saccharomyces cerevisiae). Curr. Genet. 29, 1–9. 7. Nilsson, A., Norbeck, J., Oelz, R., Blomberg, A. & Gustafsson, L. (2001) Fermentative capacity after cold storage of baker’s yeast is dependent on the initial physiological state but not correlated to the levels o f g lycolytic enzymes. Int. J. Fo od M ic robiol. 71,111– 124. 8. Moore,P.A.,Sagliocco,F.A.,Wood,R.M.&Brown,A.J.(1991) Yeast glycolytic mRNAs are differentially regulated. Mol. Cell. Biol. 11, 5330–5337. 9. Boles, E., Heinisch, J. & Zimmermann, F.K. (1993) Diffe rent signals control the activation of glycolysis in the yeast Saccharo- myces cerevisiae. Yeast 9, 761–770. 10. McCartney, R.R. & Schmidt, M.C. (2001) Regulation of S nf1 kinase. Activation requires phosphorylation of threonine 210 by an upstream kinase as well as a distinct step mediated by the Snf4 subunit. J. Biol. Chem. 276, 36460–36466. 11. Hong, S.P., Leiper, F.C., Woods,A.,Carling,D.&Carlson,M. (2003) Activation of yeast Snf1 and mammalian AMP-activated protein kinase by upstream kinases. Proc. Natl Acad. Sci. USA 100, 8839–8843. 12. Nath, N., McCartney, R.R. & Schmidt, M.C. (2003) Yeast Pak1 kinase associates with and activates Snf1. Mol. Cell. Biol. 23, 3909–3917. 13. Sutherland, C .M., Hawley, S.A., Mc Cartney, R.R., L eech, A., Stark,M.J.,Schmidt,M.C.&Hardie,D.G.(2003)Elm1pisoneof three upstream kinases for the Saccharomyces cerevisiae SNF1 complex . Curr. Biol. 13, 1299–1305. 14. Hohmann, S., Winderickx, J., de Win de, J.H., Valckx, D ., Cob- baert, P., Luyten, K., de Meirsman,C.,Ramos,J.&Thevelein, J.M. (1999) Nove l alleles of yeast h exokinase PII with distinct effects on catalytic activity and catabolite repression of SUC2. Microbiology 145, 703–714. 15. Kraakman, L.S., Winderickx, J., Thevelein, J.M. & De Winde, J.H. (1999) Structure-function analysis of yeast hexokinase: structural requirements for triggering cAMP signalling and cata- bolite repression. Biochem. J. 343, 159–168. 16. Mayordomo, I. & Sanz, P. (2001) Hexokinase PII: structural analysis and glucose signalling in the yeast Saccharomyces cerevisiae. Yeast 18, 923–930. 17. Ahuatzi, D., Herrero, P., De La Cera, T. & Moreno, F. (2004) The glucose regulated nuclear localization of hexokinase 2 in Saccharomyce s c erevisia e is Mig1 dependent. J. Biol. Chem. 279, 14440–14446. 18. Treitel, M.A., Kuchin, S. & C arlson, M. (1998) Snf1 p rotein kinase regulates phosphorylation of the Mig1 repressor in Saccharomyces cerevisiae. Mol. Cell. Biol. 18, 6273–6280. 19. Dombek, K.M., Kacherovsky, N. & Young, E.T. (2004) The Reg1-interacting proteins, Bmh1, Bm h2, Ssb1, and S sb2, have roles in m aintaining glucose repression in Saccharomyces cerevisiae. J. Biol. Chem. 279, 39165–39174. 20. Sanz, P., Alms, G.R., Haystead, T.A. & Carls on, M. (2000) Regulatory interactions between the Reg1-Glc7 protein phos- phatase and the Snf1 protein kinase. Mol. Cell. Biol. 20, 1321– 1328. 21. De Vit, M.J., W addle, J.A. & Johnston, M. (1997) Regulated nuclear translocation of the Mig1 glucose repressor. Mol. Biol. Cell 8, 1603–1618. 22. Papamichos-Chronakis, M., G ligoris, T. & Tzamarias, D. (2004) The Snf1 kinase controls glucose repression in yeast by modulat- ing interactions betwee n the Mig1 repressor and the C yc8-Tup1 co-repressor. EMBO Report 5, 368–372. 23. Sanz, P., Nieto, A. & Prieto, J.A. (1996) Glucose repression m ay involve processes with different s ugar kinase requirements. J. Bacteriol. 178, 4721–4723. 24. De Winde, J .H., Crauwels, M., Hohmann, S., Thevelein, J.M. & Winderickx, J. (1996) Differential requirem ent of the yeast su gar kinases for sugar sensing in estab lishing the catabolite-repressed state. Eur. J. Bioc hem. 241, 633–643. 25. Herwig, C., Chetreanu, F., Niederberger, P., Marson, I. & van Stockar, U. (2002) Quantitative analysis of t he impact of HXK2 and REG1 deletion in Saccharomyces cerevisiae on invertase expression and respiration. Enzyme Microb. Te chnol. 31, 505– 515. 26. Ye L., Kruckeberg, A.L., Berden, J.A. & van Dam, K. (1999) Growth and glucose repression are controlled by glu cose transport in Saccharomyces cerevisiae cells containing only one glucose transporter. J. Bacteriol. 18 1, 4673–4685. 27. Blank, L.M. & Sauer, O . (2004) TCA cycle activity in Saccharo- myces cerevisiae is a function of environmentally determined specific growth and glucose uptake r ates. Microbiology 150, 1085– 1093. Ó FEBS 2004 Glucose response in S. cerevisiae (Eur. J. Biochem. 271) 4863 28. Otterstedt,K.,Larsson,C.,Bill,R.M.,Sta ˚ hlberg, A., Boles, E., Hohmann, S. & Gustafsson, L. (2004) Switching the mode of metabolism in the yeast Saccharomyces cerevisiae. EMBO Report 5, 532–537. 29. Elbing, K., Larsson, C., Bill, R.M.,Albers,E.,Snoep,J.L.,Boles, E., Hohmann, S. & Gustafsson, L. ( 2004) The role of hexose transport in control of glycolytic flux in Saccharomyces cerevisiae. Appl. Environ. Microbiol. 70, 5323–5330. 30. van Dijken, J.P., Bauer, J., Brambilla, L., Duboc, P., Francois, J.M., Gancedo, C., Giuseppin, M.L., Heijnen, J.J., Hoare, M., Lange, H.C., Madden, E.A., Niederberger, P., Nielsen, J., Parrou, J.L.,Petit,T.,Porro,D.,Reuss,M.,vanRiel,N.,Rizzi,M., Steensma, H.Y., Verrips, C.T ., Vindelov, J. & Pronk, J.T. (2000) An interlaboratory c omparison o f physiological and genetic properties of four Saccharomyces cerevisiae strai ns. Enzyme Microb. Technol. 26, 706–714. 31. Schmidt, M.C. & McCartney, R .R. (2000) Beta-subunits of Snf1 kinase are required for kinase funct ion and substrate definition. EMBO J. 19, 4936–4943. 32. Verduyn, C., Postma, E., Scheffers, W.A. & Van Di jken, J.P. (1992) Effect of benzoic acid on metabolic fluxes in ye asts: a continuous-culture study on the regulation of respiration and alcoholic fermentation. Yeast 8, 501–517. 33. Goldstein, A. & Lampen, J.O. (1975) Beta- D -fructofuranoside fructohydrolase from yeast. Methods Enzymol. 42, 504–511. 34. Wieczork e, R ., Krampe, S., Weierstall, T., Freidel, K., Hollen- berg, C.P. & Boles, E. (1999) Concurrent knock-out of at least 20 transporter genes is required to block uptake of hexoses in Sac- charomyces cerevisiae. FEBS Lett. 464, 123–128. 35. Wilson, W.A., Hawley, S.A. & Hardie, D.G. (1996) Glucose repression/derepression in budding yeast: SNF1 protein kinase is activated by phosphorylation under derepressing conditions, a nd this correlates with a high AMP : ATP ratio. Curr. Biol. 6, 1426– 1434. 36. Ostling, J. & Ronne, H. (1998) Negative control of the Mig1p repressor by Snf1p-dependent phosphorylation in the absenc e o f glucose. Eur. J. Biochem. 252, 162–168. 37. Smith, F.C., Davies, S.P., Wilson, W.A., Carling, D. & Hardie, D.G. (1999) The SNF1 kinase complex from Saccharomyces cerevisiae phosphorylates the transcriptional repressor protein Mig1p in vitro at four sites within or near regulatory domain 1. FEBS Lett. 453, 219–223. 38. Ozcan, S., Vallier, L.G., Flick, J.S., Carlson, M. & Johnston, M . (1997) Expression of the SUC2 gene of Saccharomyces cerevisiae is induced by low levels of glucose. Yeast 13, 127–137. 39. Kaniak, A ., Xue, Z., Macool, D., Kim, J .H. & Johnston, M. (2004) Regulatory network connecting two glucose signal trans- duction pathways in Saccharomyces cerevisiae. Eukaryot. Cell 3, 221–231. 40. van Hoek, P., van Dijken, J.P. & Pronk, J.T. (2000) Regulation of fermentative capacity an d levels of glycolytic enzym es in chemo- stat cult ures o f Saccharomyces cerevisia e. Enzyme Microb. Tech- nol. 26, 7 24–736. 41. Yin, Z., Hatto n, L. & Brown, A .J. ( 2000) D ifferential p ost-tran- scriptional regulation of yeast mRNAs in respo nse to high and low glucose concentrations. Mol. Microbiol. 35, 553–565. 42. Sierkstra, L.N., Sillje, H.H., Verbakel, J.M. & Verrips, C.T. (1993) The glucose-6-phosphate-isome rase reaction is essential for n or- mal glucose repression in Saccharomyces cerevisiae. Eur. J. Bio- chem. 214, 121–127. 43. Ronne, H. (1995) Glucose repression in fungi. Trends Genet. 11, 12–17. 44. Reifenberger, E., Boles, E. & Ciriacy, M . (1997) Kinetic char- acterization of individual hexose transporters of Saccharomyces cerevisiae and t heir rela tion to the triggering mechanisms of g lu- cose repression. Eur. J. Biochem. 245, 324–333. 45. Diderich, J.A., S chepper, M., van Hoe k, P., Luttik, M.A., van Dijken, J.P., Pronk, J.T., Klaassen, P., Boelens, H.F., de Mattos, M.J., van Dam, K. & Kruckeberg, A.L. (1999) Glu- cose uptake kinetics and transcript ion o f HXT genes in c hemostat cultures of Saccharomyces cerevisiae. J. Biol. Chem. 274, 15350– 15359. 46. Hauf, J ., Zimmermann, F.K. & Muller, S. (2000) Simultaneous genomic overexpression of seven glycolytic enzymes in the yeast Saccharomyces cerevisiae. Enzyme Microb. Technol. 26, 688– 698. 4864 K. Elbing et al. (Eur. J. Biochem. 271) Ó FEBS 2004 . Transcriptional responses to glucose at different glycolytic rates in Saccharomyces cerevisiae Karin Elbing 1 , Anders Sta ˚ hlberg 1 ,. previous chemostat studies of the CEN.PK strain cultured at different glycolytic rates within the respiro-fermentative phase, i.e. high dilution rates [40]. Interestingly,