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. 2003 Feb;2(1):143–149. doi: 10.1128/EC.2.1.143-149.2003

Glucose Regulation of Saccharomyces cerevisiae Cell Cycle Genes

Laura L Newcomb 1, Jasper A Diderich 1,†, Matthew G Slattery 2, Warren Heideman 1,2,*
PMCID: PMC141163  PMID: 12582131

Abstract

Nutrient-limited Saccharomyces cerevisiae cells rapidly resume proliferative growth when transferred into glucose medium. This is preceded by a rapid increase in CLN3, BCK2, and CDC28 mRNAs encoding cell cycle regulatory proteins that promote progress through Start. We have tested the ability of mutations in known glucose signaling pathways to block glucose induction of CLN3, BCK2, and CDC28. We find that loss of the Snf3 and Rgt2 glucose sensors does not block glucose induction, nor does deletion of HXK2, encoding the hexokinase isoenzyme involved in glucose repression signaling. Rapamycin blockade of the Tor nutrient sensing pathway does not block the glucose response. Addition of 2-deoxy glucose to the medium will not substitute for glucose. These results indicate that glucose metabolism generates the signal required for induction of CLN3, BCK2, and CDC28. In support of this conclusion, we find that addition of iodoacetate, an inhibitor of the glyceraldehyde-3-phosphate dehydrogenase step in yeast glycolysis, strongly downregulates the levels CLN3, BCK2, and CDC28 mRNAs. Furthermore, mutations in PFK1 and PFK2, which encode phosphofructokinase isoforms, inhibit glucose induction of CLN3, BCK2, and CDC28. These results indicate a link between the rate of glycolysis and the expression of genes that are critical for passage through G1.

Yeast in common laboratory cultures undergo repeated growth cycles in which cells are inoculated into growth medium, begin to proliferate, deplete the medium, and cease or dramatically slow proliferation. This cycle has been repeated for countless generations in the laboratory and in domestic culture. In many media, a critical element is glucose. In rich medium containing glucose, proliferation is very rapid during log-phase fermentative growth, but it slows dramatically when glucose is exhausted. Cells then enter a plateau phase in which G1 is prolonged and proliferation ceases, followed by a slow post-log phase of oxidative growth (48, 51, 52). Rapid log-phase proliferation can be restored by the addition of glucose to a post-log culture.

Glucose is known to affect a variety of processes in Saccharomyces cerevisiae. These include glucose repression of genes used in growth on alternative carbon sources and the induction of genes needed for glucose transport and protein synthesis (6, 17, 26, 49). Indeed, glucose has a profound effect on the transcription of yeast genes. In pioneering work with cDNA microarrays, DeRisi et al. demonstrated that ca. 2,000 genes are regulated by the change from fermentative growth on glucose to oxidative growth at the diauxic shift (9). Many of the expected changes in gene expression were observed in these experiments: genes encoding glycolytic enzymes were downregulated as glucose was exhausted, whereas the expression of genes involved in oxidative metabolism increased. Although the regulation of some of these genes by glucose has been well studied, the mechanism by which glucose regulates many genes remains unknown.

Although the precise mechanisms by which yeast cells sense the presence of glucose are still unclear, several signaling pathways for glucose sensing have been identified in yeast. The RGT2 and SNF3 genes encode proteins that resemble hexose transporters in structure. These proteins also have long cytoplasmic tails that are required for signal generation. Rgt2 and Snf3 do not transport glucose but instead initiate signals in response to glucose that in turn activate a pathway that allows the Rgt1 transcription factor to upregulate glucose transporter expression (34, 35). It is presumed that binding of glucose to these cell surface proteins produce signals that allow the cell to synthesize the transport proteins needed for glucose uptake.

An additional pathway directing transcriptional changes in response to glucose involves stimulation of adenylyl cyclase and an increase in intracellular cyclic AMP (cAMP). The Ras-cAMP pathway includes the GTP-binding Ras proteins, encoded by RAS1 and RAS2, the guanine nucleotide exchange factor Cdc25, and the Ras GTPase-activating proteins, Ira1 and Ira2, and adenylyl cyclase, encoded by CYR1. This pathway plays a key role in activating adenylyl cyclase, since both Ras and Cdc25 proteins are essential for basal adenylyl cyclase activity and cell viability (24). It has been argued that the Ras pathway is not directly involved in glucose signaling to adenylyl cyclase (7, 46). Instead, a G-protein-coupled receptor is proposed to initiate the glucose signal that increases cAMP (29). GPR1, encoding the receptor, was cloned as a prey in two-hybrid studies by using Gpa2 as bait (29, 56, 57). Strains expressing constitutively active Gpa1 bypass the requirement for Gpr1 in cAMP stimulation (41). The G protein, Gpa2, was cloned based on homology with mammalian heterotrimeric Gα proteins and is necessary for the glucose-specific increase in cAMP (7).

Ultimately, glucose activation of adenylyl cyclase leads to activation of the cAMP-dependent protein kinase A (PKA) encoded by the genes TPK1, TPK2, and TPK3. The activity of Rap1, a transcriptional activator of genes encoding ribosomal proteins and proteins required for glycolysis, increases upon activation of PKA by cAMP (28). In addition, PKA targets other transcription factors such as Msn2 and Msn4, to downregulate expression of stress response element (STRE)-controlled genes in the absence of glucose (43). Upon addition of glucose the increase in PKA activity inactivates these transcription factors, leading to a decrease in expression of STRE-controlled genes (18).

Once inside the cell, glucose activates another pathway involved in repression of genes not needed during growth on glucose. In this pathway, the product of the HXK2 gene, encoding a hexokinase involved in the initial stage of glucose metabolism, produces a signal that regulates Snf1, a member of the AMP-activated protein kinase family. Snf1 in turn regulates the Mig1 repressor protein (25, 27). In this pathway, the presence of glucose is thought to be sensed by hexokinase II (23, 30).

Nutrient signals are also carried by the Tor phosphatidylinositol 3-kinases encoded by TOR1 and TOR2. This pathway is the target for the drug rapamycin and is involved in signaling glucose and nitrogen limitation. The TOR pathway regulates a diverse set of processes, including ribosome biogenesis, transport of nitrogen sources, and nitrogen-regulated gene expression (45).

In order for nutrient signals to regulate cellular proliferation, these signals must be in some way connected to the cell cycle machinery. Nutrient availability affects the passage from G1 into S phase: G1 becomes prolonged at the diauxic shift, and cells cease progress through G1 altogether as nutrients are depleted. Movement from G1 into S phase at Start is mediated by a peak in expression of two G1 cyclins encoded by CLN1 and CLN2 (20, 39, 54). The timing of this event is in turn regulated by an additional G1 cyclin CLN3 (12, 44, 47), and an unrelated protein encoded by BCK2 (10, 15, 32). The effects of Cln3 are dependent on the cyclin-dependent protein kinase Cdc28, whereas the effects of Bck2 are largely Cdc28 independent (53). Changes in the expression level of CLN3 and BCK2 affect the length of G1: overexpression of either CLN3 or BCK2 shortens G1, the deletion of either lengthens G1, and the deletion of both genes leaves the cell barely able to proceed through G1 at all.

The connections between nutrient-sensing pathways such as those described above and cell cycle regulatory components have not been completely defined. Connections that have been established include effects of the Tor and cAMP pathways on the translation of the Cln3 G1 cyclin (1, 21, 37). Grr1, a ubiquitin-protein ligase plays a role in both glucose signaling and G1 cyclin stability (2). In addition nitrogen starvation affects CLN3 transcription (19, 36) and Cln3 translation (16). Glucose also increases the transcription of CLN3 in a process that requires a set of repeated elements upstream of the CLN3 open reading frame that are binding sites for the Azf1 protein (33). Little is known about nutrient regulation of BCK2 and CDC28; however, we have shown that these transcripts are upregulated when glucose is added to post-log-phase cells (55). In this report we characterize the glucose induction of CLN3, BCK2, and CDC28 mRNA. Induction of these genes requires the transport and metabolism of glucose but does not require the cAMP-, Tor-, RGT2/SNF3-, or HXK2-mediated nutrient signaling pathways. Our results suggest that a pathway that monitors glucose metabolism regulates transcription of cell cycle regulatory genes.

MATERIALS AND METHODS

Yeast strains and growth conditions.

The media used were YEP (1% yeast extract, 2% peptone) and S (0.67% yeast nitrogen base) with the indicated carbon source at 2% (42). The term “post-log” refers to cultures grown for 2 to 3 days in yeast extract-peptone-dextrose (YEPD) to an optical density at 660 nm (OD660) of ca. 5 to 7. Rapamycin treatment was as described in (8). Budding indexes are the average of triplicate determinations of 100 cells each. Strains are listed in Table 1.

TABLE 1.

S. cerevisiae strains

Strain Genotype Source or reference
DS10 MATahis3-11,15 leu2-3,112 lys1 lys2 ura3-52 trp1Δ 4
DM15 cln3Δ in DS10 21
TL1 bck2Δ::KanMX in DS10 This study
BY4741 MATahis3Δ1. leu2Δ0 met15Δ0 ura3Δ0 Research Genetics
MC996A MATahis 3-11,15 leu 2-3,112 ura3-52 38
KY73 MATahis 3-11,15 leu 2-3,112 ura3-52 hxt1Δ::HIS3::hxt4Δ hxt5Δ::LEU2 hxt2Δ::HIS3 hxt3Δ::LEU2::hxt6Δ hxt7Δ::HIS3 gal2Δ 31
DTY123 MATahis 4-539 ura 3-52 lys2-801 suc2 D. Thiele
DTY124 MATahis 4-539 ura 3-52 lys2-801 suc2 hxk2Δ::URA3 D. Thiele
FM392 MATα his3Δ1 leu2Δ0 ura3Δ0 lys2Δ0 M. Johnston
YM6370 MATα his3Δ1 leu2Δ0 ura3Δ0 lys2Δ0 snf3Δ::hisG-URA3-hisG rgt2Δ::KanMX M. Johnston
ENYWA MATα ura3-52 leu2-3,112 trp1-289 his3-delta1 MAL2-8c MAL3 SUC3 3
EBY.81 MATα ura3-52 leu2-3,112 trp1-289 his3-delta1 MAL2-8c MAL3 SUC3 pfk1Δ::URA3 3
EBY.82 MATα ura3-52 leu2-3,112 trp1-289 his3-delta1 MAL2-8c MAL3 SUC3 pfk2Δ::URA3 3
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RNA preparation and Northern blotting.

Total yeast RNA was isolated as described previously (13) from cells growing in the indicated medium at an OD660 of ∼1. The RNA samples were separated by formaldehyde gel electrophoresis and transferred to a Gene Screen Plus membrane (New England Nuclear). To ensure uniform loading and transfer of RNA, ethidium bromide was added to the samples, and the ethidium-stained rRNA was visualized on the blots under UV illumination. Blots were also probed with a radiolabeled 0.6-kb SacI fragment from U2 to confirm uniform loading. Northern blots were probed with a 3-kb HinDIII fragment of BCK2, a 1-kb BamHI/HinDIII fragment of CDC28, or a 1.8-kb BamHI fragment from CLN3. All probes were radiolabeled to a specific activity of 109 cpm/μg by using [α-32P]dCTP (3,000 μCi/nmol) and random priming. Blots were analyzed by using a Molecular Dynamics Phosphorimager SI and Molecular Dynamics ImageQuant V1.2 software.

RESULTS

Glucose induction of cell cycle mRNAs.

When post-log-phase cells are reinoculated into fresh glucose medium, the mRNAs encoding CLN3, BCK2, and CDC28 are induced (Fig. 1A). After the initial induction, we often observe unexplained periodic fluctuations in the levels of these messages: the mRNA is first induced, levels then transiently decrease before increasing again. This is especially true of CLN3 mRNA but was also observed in several experiments with CDC28 and BCK2 mRNA.

FIG. 1.

FIG. 1.

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Glucose induction of CLN3, BCK2, and CDC28 mRNAs and return to proliferative growth. (A) Wild-type (DS10) cells were grown overnight to post-log phase in YEPD medium and then transferred to fresh YEPD at an OD660 of 0.8. Samples were collected at the indicated time points for RNA preparation and Northern blotting. U2 RNA was used as a loading and transfer control. (B) Cells were treated as described above, and bud counts on triplicate samples were performed at the indicated times after transfer. At least 300 cells were counted per time point for each strain.

The induction of these messages in response to fresh glucose medium precedes a return to proliferative growth. We have previously shown that deletion of CLN3 delays the initial bud emergence as cells resume growth (55). To determine whether BCK2 plays a similar role, we tested the effect of BCK2 deletion on the time course of bud emergence after the addition of fresh glucose medium (Fig. 1B). As previously observed, deletion of CLN3 produced a delay in bud emergence in response to fresh glucose medium of about an hour. In contrast, BCK2 deletion had no effect on the time course of this process. We conclude that, although CLN3 plays a role in the rapid return to mitotic growth in response to fresh medium, BCK2 does not play a role that cannot be completely served by the presence of a normal copy of CLN3. Deletion of CDC28 was not tested in this experiment because CDC28 is an essential gene.

Glucose signaling.

We found that induction of CLN3, BCK2, and CDC28 required the transport of glucose across the cell membrane and was not observed in a strain that lacks functional hexose transporters (Fig. 2). In this experiment, a strain in which the HXT1-7 and GAL2 genes encoding hexose transport proteins have been deleted (31), was grown in YEP-2% glycerol-ethanol and transferred to fresh glucose medium. In wild-type cells, glucose produced a substantial increase in the CLN3, BCK2, and CDC28 mRNAs. This was markedly decreased in the mutant strain. These results indicate that glucose transport into the cell is required for the observed message induction. In contrast to wild-type cells, glucose addition did not produce an increase in the number of budded cells in the mutant strain (not shown).

FIG. 2.

FIG. 2.

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Glucose transport is required for induction of cell cycle messages. Wild-type (MC996A) and isogenic hxt1-7Δ gal2Δ (KY73) cells were grown in YEP-2% glycerol-ethanol and transferred to fresh YEPD at an OD660 of 1.0. Samples were collected at the indicated time points for RNA preparation and Northern blotting.

RGT2 and SNF3 encode cell surface glucose sensors (34, 35). To determine whether Rgt2 or Snf3 carry glucose signals that induce CLN3, BCK2, and CDC28, we compared the response to glucose between a wild-type and an rgt2Δ snf3Δ strain. The levels of cell cycle mRNAs in the mutant strain were identical to those observed in the wild-type control strain, indicating that the glucose signals generated by these proteins are not necessary for the response (Fig. 3).

FIG. 3.

FIG. 3.

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The Snf3 and Rgt2 glucose sensors are not required for glucose induction. Wild-type (WT; FM392) and snf3Δ rgt2Δ (YM6370) cells were grown in YEP-2% ethanol to post-log phase. Cells were transferred to fresh YEPD media at an OD660 of 0.8. Samples were collected at the indicated time points for RNA preparation and Northern blotting.

Inhibition of the Tor pathway with rapamycin did not affect the induction of CLN3, BCK2, and CDC28 (Fig. 4), in agreement with previous results showing that the Tor pathway is involved in regulating Cln3 translation but not mRNA levels (1, 5).

FIG. 4.

FIG. 4.

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Rapamycin blockade of the Tor pathway does not prevent induction of cell cycle genes by glucose. Wild-type (DS10) cells were grown overnight in YEPD to post-log phase and rapamycin (200 ng/ml, final concentration) or vehicle (ethanol) alone was added. The samples were incubated for 20 min and then transferred to fresh YEPD at an OD660 of 0.8, maintaining the previous rapamycin concentration. Samples were collected at the indicated time after transfer to fresh medium for RNA preparation and Northern blotting.

Carbon source metabolism and gene induction.

Responses to glucose are also mediated by the hexokinase encoded by HXK2. We tested an hxk2Δ strain to determine whether the loss of Hxk2 function prevents the induction of the CLN3, BCK2, and CDC28 messages by glucose medium (Fig. 5A). We found that the loss of HXK2 decreases the transcriptional response to a degree but does not block the response.

FIG. 5.

FIG. 5.

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Hxk2 signaling and glucose induction of CLN3, BCK2, or CDC28. (A) Cells from a wild-type (DTY123) and an isogenic hxk2Δ strain (DTY124) were grown overnight in YEPD to post-log phase. Cells were transferred to fresh YEPD medium at an OD660 of 0.8. Samples were collected at the indicated time after transfer to fresh medium for RNA preparation and Northern blotting. (B) Wild-type cells were grown in YEPD until they had exhausted glucose from the medium (36 h after inoculation, at an OD660 of 6), and then either d-glucose or 2-deoxy glucose was added to produce a final concentration of 2%. Control cells received an equivalent volume of water. Cells were incubated for the indicated time and then were collected for RNA blotting as described in Materials and Methods.

One possible explanation for this is that Hxk2 activity is necessary for some portion of glucose induction of CLN3, BCK2, and CDC28. To examine this further, we tested the ability of 2-deoxy glucose, an analog of glucose that can be transported into the cell and phosphorylated in the initial step of glycolysis but cannot be further metabolized. Addition of 2-deoxy glucose failed to induce CLN3, BCK2, and CDC28 (Fig. 5B). Whereas 2-deoxy glucose is a substrate for Hxk2 and allows the generation of Hxk2-mediated glucose signals (14), it fails to support glycolysis. It is known that loss of HXK2 produces cells with impaired fermentation rates (11). Since 2-deoxy glucose does not support glycolysis, and HXK2 deletion reduces glycolysis, a possible model is that the transcription of CLN3, BCK2, and CDC28 are linked to the rate of glycolysis. To test this, we used iodoacetate, an inhibitor of the glyceraldehyde-3-phosphate dehydrogenase step in glycolysis (50). Addition of iodoacetate reversed the glucose induction of CLN3, BCK2, and CDC28 (Fig. 6). These results suggest that glucose produces its effects by stimulating glycolysis.

FIG. 6.

FIG. 6.

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Inhibition of glycolysis by iodoacetate downregulates CLN3, BCK2, or CDC28 mRNAs. Wild-type (DS10) cells were grown overnight in YEPD to post-log phase. A sample was collected, and the remaining cells were inoculated in fresh YEPD at 0.8 OD660 with iodoacetate at the indicated concentration (mM) and incubated for 1 h at 30°. Samples were then collected for RNA preparation and Northern blotting.

Because iodoacetate has effects on multiple enzyme systems, the effects of iodoacetate on cell cycle transcript levels might not be specific to inhibition of glycolysis. To produce a more specific effect, we tested the effect of deletions in PFK1 and PFK2, encoding isoforms of phosphofructokinase, catalyzing an early step in glycolysis. Deletion of either PFK1 or PFK2 markedly decreased induction of CLN3, BCK2, and CDC28 by glucose (Fig. 7). These results indicate that it is the metabolism of glucose that generates the signal that activates these genes.

FIG. 7.

FIG. 7.

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Deletion of PFK1 or PFK2 blocks glucose induction of cell cycle genes. Wild-type (ENYWA-1A), pfk1Δ (EBY.81), and pfk2Δ (EBY.82) were grown overnight in YEPD. The cells were inoculated in fresh YEPD at an OD660 of 0.8, and samples were collected at the indicated time for RNA preparation and Northern blotting.

We have previously demonstrated that the levels of Cln3 protein and Cln3/Cdc28 kinase activity are higher in cells growing in fermentable carbon sources than in cells grown in nonfermentable carbon sources (21, 36). However, CLN3 expression appeared to be affected by the lag phase between inoculation of cells into the nonfermentable medium and the beginning of active growth. To examine this in more detail, we measured the levels of CLN3, BCK2, and CDC28 mRNAs in wild-type cells during growth after inoculation into fresh medium containing either glucose or ethanol as the carbon source (Fig. 8). As expected, glucose produced higher levels of all three messages than ethanol medium. However, for both carbon sources, the mRNA levels changed over time. This was especially apparent for CLN3 mRNA, which peaked shortly before the period of rapid growth.

FIG. 8.

FIG. 8.

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Growth in different carbon sources. Wild-type (BY4741) cells were grown overnight to post-log phase in YEPD. Cells were transferred to fresh YEPD or YEP-2% ethanol media at an OD660 of 0.2. Samples were taken for culture density measurement (A) and RNA preparation (B) at the times indicated.

DISCUSSION

S. cerevisiae strains have descended through innumerable cycles of nutrient excess and restriction. This demands that yeast cells regulate the cell cycle so that proliferation is rapid in the presence of rich nutrients and yet ceases during periods of nutrient limitation. Nutrients such as glucose must therefore generate signals that are in some way connected to the gene products that regulate cell cycle progression. We find that CLN3, BCK2, and CDC28 mRNAs are all increased when post-log-phase cells are transferred to fresh glucose medium. Our results indicate that some process in glycolysis regulates genes that regulate cell cycle progression.

Coregulation of CLN3, CDC28, and BCK2.

Although glucose upregulates CLN3, BCK2, and CDC28, we noticed differences in the expression patterns of the three genes. Glucose addition consistently had the greatest effects on CLN3 mRNA, whereas the effects on BCK2 were generally the least pronounced of the three messages examined. In addition, although CLN3 and BCK2 message levels increased as growth on ethanol got under way, CDC28 message levels did not. These differences indicate that, although all three messages appear to be affected by a process in glycolysis, they may not be regulated by the same mechanism.

Glucose signals.

Several distinct pathways are involved in normal yeast responses to glucose. Some of these probably involve the interaction of glucose with cell surface receptors, whereas others require the import of glucose into the cell. These include the Gpr1/Gpa2/cAMP pathway, the Tor pathway, the Snf3 and Rgt2 homologs of hexose transporters, and the Hxk2 hexokinase. None of these signal transduction pathways appear to play an important role in glucose induction of CLN3, BCK2, or CDC28 mRNAs, since the message levels are unaffected by the loss of any of these pathways. This was shown by our experiments with HXK2, SNF3, and RGT3 deletion mutants, and rapamycin blockade of the Tor pathway, along with previously published work involving the Tor pathway (22), and the cAMP pathway (36, 40).

On the other hand, glucose import and metabolism are clearly required for the induction of CLN3, BCK2, and CDC28 mRNAs. The process is blocked in a strain lacking glucose transport genes, and decreased in an hxk2Δ strain in which fermentation is slowed (11). Furthermore, 2-deoxy glucose, which cannot enter glycolysis, does not substitute for glucose. CLN3 is induced by a variety of fermentable carbon sources, suggesting that fermentation produces the signal that regulates CLN3 transcription (36). Addition of iodoacetate, an inhibitor of glyceraldehyde-3-phosphate dehydrogenase produced an almost complete reversal of the glucose induction of the genes examined. Finally, deletion of the genes encoding phosphofructokinase blocked glucose induction of CLN3, BCK2, and CDC28.

Together, our results indicate that these transcripts are linked to carbon source metabolism. It is intriguing that nonfermentable carbon sources such as ethanol can stimulate an increase in CLN3 mRNA. Although this effect is considerably smaller than the effect of glucose, it may indicate that the signal is generated at a point in common between fermentable and nonfermentable carbon source metabolism.

Much of the study of glucose effects on S. cerevisiae has focused on the repression of genes involved in alternate carbon source metabolism and the induction of genes encoding glucose transport and ribosomal proteins. The landmark microarray experiments by DeRisi et al. examining changes in transcript abundance as yeast cells exhaust the glucose in the medium at the diauxic shift demonstrated the profound effects that glucose has on gene expression in yeast (9). It is not yet clear whether the known glucose-sensing pathways can account for this large-scale regulation. Our results point to an uncharacterized pathway by which glycolysis generates a signal that regulates the mRNA levels for a set of cell cycle genes. Whether this process is restricted to the control of the cell cycle or is also involved in other glucose responses remains to be determined.

Acknowledgments

We thank Mark Johnston and Dennis Thiele for providing yeast strains. We acknowledge the valuable assistance of Chateen Carbonara, Stacey Guillen, Tracy Laabs, and Aaron Miller.

This work was supported by grant number 9982537 from the National Science Foundation.

REFERENCES

  • 1.Barbet, N. C., U. Schneider, S. B. Helliwell, I. Stansfield, M. F. Tuite, and M. N. Hall. 1996. TOR controls translation initiation and early G1 progression in yeast. Mol. Biol. Cell 7:25-42. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Barral, Y., S. Jentsch, and C. Mann. 1995. G1 cyclin turnover and nutrient uptake are controlled by a common pathway in yeast. Genes Dev. 9:399-409. [DOI] [PubMed] [Google Scholar]
  • 3.Boles, E., J. Heinisch, and F. K. Zimmermann. 1993. Different signals control the activation of glycolysis in the yeast Saccharomyces cerevisiae. Yeast 9:761-770. [DOI] [PubMed] [Google Scholar]
  • 4.Boorstein, W. R., and E. A. Craig. 1990. Regulation of a yeast HSP70 gene by a cAMP responsive transcriptional control element. EMBO J. 9:2543-2553. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Cardenas, M. E., N. S. Cutler, M. C. Lorenz, C. J. Di Como, and J. Heitman. 1999. The TOR signaling cascade regulates gene expression in response to nutrients. Genes Dev. 13:3271-3279. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Carlson, M. 1999. Glucose repression in yeast. Curr. Opin. Microbiol. 2:202-207. [DOI] [PubMed] [Google Scholar]
  • 7.Colombo, S., P. Ma, L. Cauwenberg, J. Winderickx, M. Crauwels, A. Teunissen, D. Nauwelaers, J. H. de Winde, M. F. Gorwa, D. Colavizza, and J. M. Thevelein. 1998. Involvement of distinct G-proteins, Gpa2 and Ras, in glucose- and intracellular acidification-induced cAMP signalling in the yeast Saccharomyces cerevisiae. EMBO J. 17:3326-3341. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Crespo, J. L., T. Powers, B. Fowler, and M. N. Hall. 2002. The TOR-controlled transcription activators GLN3, RTG1, and RTG3 are regulated in response to intracellular levels of glutamine. Proc. Natl. Acad. Sci. USA 99:6784-6789. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.DeRisi, J., I. Iyer, and P. Brown. 1997. Exploring the metabolic and genetic control of gene expression on a genomic scale. Science 278:680-686. [DOI] [PubMed] [Google Scholar]
  • 10.Di Como, C. J., H. Chang, and K. T. Arndt. 1995. Activation of CLN1 and CLN2 G1 cyclin expression by BCK2. Mol. Cell. Biol. 15:1835-1846. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Diderich, J. A., L. M. Raamsdonk, A. L. Kruckeberg, J. A. Berden, and K. Van Dam. 2001. Physiological properties of Saccharomyces cerevisiae from which hexokinase II has been deleted. Appl. Environ. Microbiol. 67:1587-1593. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 12.Dirick, L., T. Bohm, and K. Nasmyth. 1995. Roles and regulation of Cln-Cdc28 kinases at the start of the cell cycle of Saccharomyces cerevisiae. EMBO J. 14:4803-4813. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Ellwood, M. S., and E. A. Craig. 1984. Differential regulation of the 70K heat shock gene and related genes in Saccharomyces cerevisiae. Mol. Cell. Biol. 4:1454-1459. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Entian, K. D., and K. U. Frohlich. 1984. Saccharomyces cerevisiae mutants provide evidence of hexokinase PII as a bifunctional enzyme with catalytic and regulatory domains for triggering carbon catabolite repression. J. Bacteriol. 158:29-35. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Epstein, C. B., and F. R. Cross. 1994. Genes that can bypass the CLN requirement for Saccharomyces cerevisiae cell cycle START. Mol. Cell. Biol. 14:2041-2047. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Gallego, C., E. Gari, N. Colomina, E. Herrero, and M. Aldea. 1997. The Cln3 cyclin is downregulated by translational repression and degradation during the G1 arrest caused by nitrogen deprivation in budding yeast. EMBO J. 16:7196-7206. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Gancedo, J. M. 1998. Yeast carbon catabolite repression. Microbiol. Mol. Biol. Rev. 62:334-361. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Garreau, H., R. N. Hasan, G. Renault, F. Estruch, E. Boy-Marcotte, and M. Jacquet. 2000. Hyperphosphorylation of Msn2p and Msn4p in response to heat shock and the diauxic shift is inhibited by cAMP in Saccharomyces cerevisiae. Microbiology 146:2113-2120. [DOI] [PubMed] [Google Scholar]
  • 19.Gasch, A. P., P. T. Spellman, C. M. Kao, O. Carmel-Harel, M. B. Eisen, G. Storz, D. Botstein, and P. O. Brown. 2000. Genomic expression programs in the response of yeast cells to environmental changes. Mol. Biol. Cell 11:4241-4257. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Hadwiger, J. A., C. Wittenberg, H. E. Richardson, M. de Barros Lopes, and S. I. Reed. 1989. A family of cyclin homologs that control the G1 phase in yeast. Proc. Natl. Acad. Sci. USA 86:6255-6259. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Hall, D. D., D. D. Markwardt, F. Parviz, and W. Heideman. 1998. Regulation of the Cln3/Cdc28 kinase by cAMP in Saccharomyces cerevisiae. EMBO J. 17:4370-4378. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Hardwick, J. S., F. G. Kuruvilla, J. K. Tong, A. F. Shamji, and S. L. Schreiber. 1999. Rapamycin-modulated transcription defines the subset of nutrient-sensitive signaling pathways directly controlled by the Tor proteins. Proc. Natl. Acad. Sci. USA 96:14866-14870. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23.Hohmann, S., J. Winderickx, J. H. de Winde, D. Valckx, P. Cobbaert, K. Luyten, C. de Meirsman, J. Ramos, and J. M. Thevelein. 1999. Novel alleles of yeast hexokinase PII with distinct effects on catalytic activity and catabolite repression of SUC2. Microbiology 145:703-714. [DOI] [PubMed] [Google Scholar]
  • 24.Ishikawa, T., I. Uno, and K. Matsumoto. 1986. The role of cAMP in controlling yeast cell division. Bioessays 4:52-56. [DOI] [PubMed] [Google Scholar]
  • 25.Jiang, R., and M. Carlson. 1996. Glucose regulates protein interactions within the yeast SNF1 protein kinase complex. Genes Dev. 10:3105-3115. [DOI] [PubMed] [Google Scholar]
  • 26.Johnston, M. 1999. Feasting, fasting and fermenting. Glucose sensing in yeast and other cells. Trends Genet. 15:29-33. [DOI] [PubMed] [Google Scholar]
  • 27.Johnston, M., and M. Carlson. 1992. Regulation of carbon and phosphate utilization: the molecular and cellular biology of the yeast Saccharomyces. Gene Expr. 2:193-281.1450661 [Google Scholar]
  • 28.Klein, C., and K. Struhl. 1994. Protein kinase A mediates growth-regulated expression of yeast ribosomal protein genes by modulating RAP1. Mol. Cell. Biol. 14:1920-1928. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Kraakman, L., K. Lemaire, P. Ma, A. W. Teunissen, M. C. Donaton, P. Van Dijck, J. Winderickx, J. H. de Winde, and J. M. Thevelein. 1999. A Saccharomyces cerevisiae G-protein coupled receptor, Gpr1, is specifically required for glucose activation of the cAMP pathway during the transition to growth on glucose. Mol. Microbiol. 32:1002-1012. [DOI] [PubMed] [Google Scholar]
  • 30.Kraakman, L. S., J. Winderickx, J. M. Thevelein, and J. H. De Winde. 1999. Structure-function analysis of yeast hexokinase: structural requirements for triggering cAMP signalling and catabolite repression. Biochem. J. 343(Pt. 1):159-168. [PMC free article] [PubMed] [Google Scholar]
  • 31.Kruckeberg, A. L., L. Ye, J. A. Berden, and K. van Dam. 1999. Functional expression, quantification and cellular localization of the Hxt2 hexose transporter of Saccharomyces cerevisiae tagged with the green fluorescent protein. Biochem. J. 339:299-307. [PMC free article] [PubMed] [Google Scholar]
  • 32.Levine, K., K. Huang, and F. R. Cross. 1996. Saccharomyces cerevisiae G1 cyclins differ in their intrinsic functional specificities. Mol. Cell. Biol. 16:6794-6803. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Newcomb, L. L., D. D. Hall, and W. Heideman. 2002. AZF1 is a glucose-dependent positive regulator of CLN3 transcription in Saccharomyces cerevisiae. Mol. Cell. Biol. 22:1607-1614. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Ozcan, S., J. Dover, and M. Johnston. 1998. Glucose sensing and signaling by two glucose receptors in the yeast Saccharomyces cerevisiae. EMBO J. 17:2566-2573. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Ozcan, S., J. Dover, A. G. Rosenwald, S. Wölfl, and M. Johnston. 1996. Two glucose transporters in Saccharomyces cerevisiae are glucose sensors that generate a signal for induction of gene expression. Proc. Natl. Acad. Sci. USA 93:12428-12432. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Parviz, F., and W. Heideman. 1998. Growth-independent regulation of CLN3 mRNA levels by nutrients in Saccharomyces cerevisiae. J. Bacteriol. 180:225-230. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Polymenis, M., and E. V. Schmidt. 1997. Coupling cell division to cell growth by translational control of the G1 cyclin CLN3 in yeast. Genes Dev. 11:2522-2531. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Reifenberger, E., E. Boles, and M. Ciriacy. 1997. Kinetic characterization of individual hexose transporters of Saccharomyces cerevisiae and their relation to the triggering mechanisms of glucose repression. Eur. J. Biochem. 245:324-333. [DOI] [PubMed] [Google Scholar]
  • 39.Richardson, H. E., C. Wittenberg, F. Cross, and S. I. Reed. 1989. An essential G1 function for cyclin-like proteins in yeast. Cell 59:1127-1133. [DOI] [PubMed] [Google Scholar]
  • 40.Robertson, L. S., H. C. Causton, R. A. Young, and G. R. Fink. 2000. The yeast A kinases differentially regulate iron uptake and respiratory function. Proc. Natl. Acad. Sci. USA 97:5984-5988. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Rolland, F., J. H. De Winde, K. Lemaire, E. Boles, J. M. Thevelein, and J. Winderickx. 2000. Glucose-induced cAMP signalling in yeast requires both a G-protein coupled receptor system for extracellular glucose detection and a separable hexose kinase-dependent sensing process. Mol. Microbiol. 38:348-358. [DOI] [PubMed] [Google Scholar]
  • 42.Sherman, F. 1991. Getting started with yeast. Methods Enzymol. 194:3-21. [DOI] [PubMed] [Google Scholar]
  • 43.Smith, A., M. P. Ward, and S. Garrett. 1998. Yeast PKA represses Msn2p/Msn4p-dependent gene expression to regulate growth, stress response and glycogen accumulation. EMBO J. 17:3556-3564. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Stuart, D., and C. Wittenberg. 1995. CLN3, not positive feedback, determines the timing of CLN2 transcription in cycling cells. Genes Dev. 9:2780-2794. [DOI] [PubMed] [Google Scholar]
  • 45.Thomas, G., and M. N. Hall. 1997. TOR signalling and control of cell growth. Curr. Opin. Cell Biol. 9:782-787. [DOI] [PubMed] [Google Scholar]
  • 46.Trevillyan, J. M., and M. L. Pall. 1979. Control of cyclic adenosine 3′,5′-monophosphate levels by depolarizing agents in fungi. J. Bacteriol. 138:397-403. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Tyers, M., G. Tokiwa, and B. Futcher. 1993. Comparison of the Saccharomyces cerevisiae G1 cyclins: Cln3 may be an upstream activator of Cln1, Cln2, and other cyclins. EMBO J. 12:1955-1968. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48.Vivier, M. A., M. G. Lambrechts, and I. S. Pretorius. 1997. Coregulation of starch degradation and dimorphism in the yeast Saccharomyces cerevisiae. Crit. Rev. Biochem. Mol. Biol. 32:405-435. [DOI] [PubMed] [Google Scholar]
  • 49.Warner, J. R. 1999. The economics of ribosome biosynthesis in yeast. Trends Biochem. Sci. 24:437-440. [DOI] [PubMed] [Google Scholar]
  • 50.Webb, J. 1966. Enzyme and metabolic inhibitors, vol. III. Academic Press, Inc., New York, N.Y.
  • 51.Werner-Washburne, M., E. Braun, G. C. Johnston, and R. A. Singer. 1993. Stationary phase in the yeast Saccharomyces cerevisiae. Microbiol. Rev. 57:383-401. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.Werner-Washburne, M., E. L. Braun, M. E. Crawford, and V. M. Peck. 1996. Stationary phase in Saccharomyces cerevisiae. Mol. Microbiol. 19:1159-1166. [DOI] [PubMed] [Google Scholar]
  • 53.Wijnen, H., and B. Futcher. 1999. Genetic analysis of the shared role of CLN3 and BCK2 at the G1-S transition in Saccharomyces cerevisiae. Genetics 153:1131-1143. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54.Wittenberg, C., K. Sugimoto, and S. I. Reed. 1990. G1-specific cyclins of S. cerevisiae: cell cycle periodicity, regulation by mating pheromone, and association with the p34CDC28 protein kinase. Cell 62:225-237. [DOI] [PubMed] [Google Scholar]
  • 55.Wu, M., L. Newcomb, and W. Heideman. 1999. Regulation of gene expression by glucose in Saccharomyces cerevisiae: a role for ADA2 and ADA3/NGG1. J. Bacteriol. 181:4755-4760. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 56.Xue, Y., M. Batlle, and J. P. Hirsch. 1998. GPR1 encodes a putative G protein-coupled receptor that associates with the Gpa2p Gα subunit and functions in a Ras-independent pathway. EMBO J. 17:1996-2007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 57.Yun, C. W., H. Tamaki, R. Nakayama, K. Yamamoto, and H. Kumagai. 1997. G-protein coupled receptor from yeast Saccharomyces cerevisiae. Biochem. Biophys. Res. Commun. 240:287-292. [DOI] [PubMed] [Google Scholar]

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