Cell cycle-dependent transcription in yeast: promoters ... - Nature

Oncogene (2005) 24, 2746–2755

& 2005 Nature Publishing Group All rights reserved 0950-9232/05 $30.00 www.nature.com/onc

Cell cycle-dependent transcription in yeast: promoters, transcription factors, and transcriptomes Curt Wittenberg1 and Steven I Reed*,2 1

Department of Molecular Biology, MB-3, The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, CA 92037, USA; 2Department of Molecular Biology, MB-7, The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, CA 92037, USA

In the budding yeast, Saccharomyces cerevisiae, a significant fraction of genes (>10%) are transcribed with cell cycle periodicity. These genes encode critical cell cycle regulators as well as proteins with no direct connection to cell cycle functions. Cell cycle-regulated genes can be organized into ‘clusters’ exhibiting similar patterns of regulation. In most cases periodic transcription is achieved via both repressive and activating mechanisms. Finetuning appears to have evolved by the juxtaposition of regulatory motifs characteristic of more than one cluster within the same promoter. Recent reports have provided significant new insight into the role of the cyclin-dependent kinase Cdk1 (Cdc28) in coordination of transcription with cell cycle events. In early G1, the transcription factor complex known as SBF is maintained in a repressed state by association of the Whi5 protein. Phosphorylation of Whi5 by Cdk1 in late G1 leads to dissociation from SBF and transcriptional derepression. G2/M-specific transcription is achieved by converting the repressor Fkh2 into an activator. Fkh2 serves as a repressor during most of the cell cycle. However, phosphorylation of a cofactor, Ndd1, by Cdk1 late in the cell cycle promotes binding to Fkh2 and conversion into a transcriptional activator. Such insights derived from analysis of specific genes when combined with genome-wide analysis provide a more detailed and integrated view of cell cycle-dependent transcription. Oncogene (2005) 24, 2746–2755. doi:10.1038/sj.onc.1208606

1983) allowed for the testing of a large number of genes for cell cycle regulation of transcription. Not surprisingly, many genes involved in specific cell cycle processes were found to be expressed maximally during specific cell cycle intervals where synthesis of encoded products could be rationalized in terms of function (Wittenberg and Reed, 1991). Based on single-gene analyses, it could be predicted that a significant but limited set of genes encoding proteins directly involved in periodic cell cycle functions would be subject to cell cycle regulation. However, more recently, large-scale array analysis of the yeast transcriptome has revealed that a much larger set of genes (minimally 800 or >10% of protein-encoding genes) exhibit cell cycle regulation and that many of the encoded proteins are not ostensibly linked to cell cycle functions (Cho et al., 1998; Spellman et al., 1998). This presumably reflects a more intricate and subtle relationship between cellular physiology and cell cycle progression than has been appreciated. Furthermore, highresolution scrutiny has revealed that even within socalled clusters of coordinately-regulated genes, there are significant differences observable between individual members, suggesting individualized optimization of transcriptional programming according to function (Cho et al., 1998; Spellman et al., 1998). In this article, we will review advances based on detailed analysis of transcriptional control of individual genes, but also discuss the implications of findings based on genomewide analysis.

Keywords: transcription; cell cycle; cyclin-dependent protein kinase The G1 gene cluster

Introduction With the advent of modern molecular biology, some 30 years ago, it became possible to efficiently analyse transcription patterns of individual genes. In yeast, the convergence of relatively straightforward cell cycle synchronization protocols (Amon, 2002) with rapid and efficient mutational cloning techniques (Struhl, *Correspondence: SI Reed; E-mail: [email protected]

Cell cycle initiation during G1 phase is the consequence of transcriptional activation of greater than 200 genes, many of which participate in events specifically associated with cell cycle progression (Cho et al., 1998; Spellman et al., 1998). The primary cis-acting regulators of the G1-specific gene cluster are targeted by two alternative heterodimeric transcription factors comprised of unique DNA-binding components, Swi4 (SBF) or Mbp1 (MBF), and a common component, Swi6, that participates in transactivation (Breeden, 1996). The optimal binding site for MBF is called the Mlu Cell cycle Box (MCB), due to the presence of an

Cell cycle-dependent transcription in yeast C Wittenberg and SI Reed

2747

MluI restriction site in the consensus sequence, and that for SBF is the Swi4 Cell cycle Box (SCB) (Breeden, 1996; Lee et al., 2002; Kato et al., 2004). The target genes for SBF and MBF have been traditionally defined by the presence of redundant SCB and MCB sequences in their promoters. Functionally, SBF and MBF targets, respectively, fall roughly into two classes. MBF targets include those involved in the control or execution of DNA replication and repair (POL2, CDC2, RNR1, CLB5/6), whereas SBF targets include those involved in cell morphogenesis, spindle pole body duplication and other growth-related functions (CLN1/2, PCL1/2, GIN4, FKS1/2). This apparent differentiation into distinct functional classes may have adaptive significance due to regulatory differences between the two transcription factors. However, each group also includes many members that do not fall neatly into these categories. Consequently, it is possible that this separation is merely a reflection of the evolutionary history of these transcriptional regulatory modules (see below). Whereas the optimal binding sites for SBF and MBF are distinct, there is considerable overlap in their specificity both in terms of the DNA sequence requirements for binding and the occurrence of such sites within promoters. Consequently, there is considerable ambiguity concerning the role of each factor in regulating specific genes (Partridge et al., 1997). This is apparent from the global analysis of the binding of the specific transcription factors to promoters (Iyer et al., 2001; Simon et al., 2001; Lee et al., 2002) and may explain the modest effect of inactivation of SBF or MBF on the expression of G1-specific genes (Koch et al., 1993; de Bruin and Wittenberg, unpublished). Regulation of G1-specific transcription G1-specific transcription is promoted, in large part, by G1 cyclin-associated CDK activity. In fact, deregulation of that process was the basis for the discovery of the G1 cyclin Cln3 (Cross, 1988; Nash et al., 1988). Mutations leading to stabilization of Cln3 result in premature expression of G1-specific genes and, thereby, small cell size and resistance to G1-phase arrest by mating pheromone. Any of the three G1 cyclins (Cln1, Cln2, or Cln3) in conjunction with Cdk1 (Cdc28) were later shown to be sufficient to mobilize Cdk1-dependent activation of G1-specific transcription (Cross and Tinkelenberg, 1991; Dirick and Nasmyth, 1991; Marini and Reed, 1992). However, whereas Cln1- and Cln2containing CDK complexes are sufficient for transcriptional activation, Cln3/Cdk1 is the primary activator under physiological conditions (Tyers et al., 1993; Dirick et al., 1995; Stuart and Wittenberg, 1995). Consistent with that role, Cln3 is largely nuclear whereas the other G1 cyclins are distributed to the nucleus and cytoplasm (Miller and Cross, 2000; Edgington and Futcher, 2001). Although Cln3 is not essential for viability, it becomes essential in cells lacking Bck2, a protein of unknown molecular function but that is required for basal transcriptional activation

of at least a portion of SBF and MBF target genes, including CLN1 and CLN2 (Epstein and Cross, 1994; Di Como et al., 1995; Wijnen and Futcher, 1999). The identification of a central role for Cln3/Cdk1 in transcriptional activation (Cross and Tinklenberg, 1991; Dirick and Nasmyth, 1991), and the finding that Cdk1 is likely to act directly rather than via its effect on the accumulation of other proteins (Marini and Reed, 1992), has led to a protracted search for CDK targets involved in activation. Swi6 and Swi4 are established targets in vitro and Swi6 is known to be phosphorylated in a Cdk1-dependent manner in vivo (Sidorova et al., 1995; Ubersax et al., 2003; Geymonat et al., 2004). Elimination of Cdk sites by mutation leads to a defect in nuclear import of Swi6 (Sidorova et al., 1995; Geymonat et al., 2004). However, those mutations appear not to have an effect on the timing, extent of transcriptional activation or Cdk dependence of transcriptional activation, suggesting that phosphorylation of Swi6 is not rate limiting for access to the nucleus. The lack of a functional link between phosphorylation of the transcription factors and transcriptional activation has led to the suggestion that Cdk1 acts indirectly via phosphorylation of another factor (Wijnen et al., 2002), perhaps a previously unrecognized transcriptional activator or repressor. Although a number of proteins have been shown to interact with Swi6 (e.g., Hrr25 (Ho et al., 1997), Skn7 (Morgan et al., 1995), and Stb1 (Costanzo et al., 2003)), none appears to play a significant role in the Cln/Cdk-dependent activation of transcription. Recently, the search for additional factors regulating G1 transcription led to the discovery of Whi5 as an SBF-associated transcriptional repressor (Costanzo et al., 2004; de Bruin et al., 2004). Inactivation of Whi5, previously shown to cause cell cycle initiation at a small cell size (Jorgensen et al., 2002; Zhang et al., 2002), suppresses the requirement for Cln3 in the absence of Bck2 (Costanzo et al., 2004; de Bruin et al., 2004). Consistent with that phenotype, inactivation of WHI5 leads to premature activation of G1-specific transcription during the G1 phase. The Whi5 protein associates with G1-specific promoters in an SBF-dependent manner and is released from DNA coincident with transcriptional activation. It is phosphorylated at Cdk consensus sites in vivo and in vitro and can be phosphorylated in vitro by both G1- and B-type cyclin-associated Cdk1. Consistent with a role for Cdk1 in activation of SBF-dependent transcription, phosphorylation of SBF/Whi5 complexes in vitro promotes dissociation of the complex. The regulation of SBF by Whi5 bears striking resemblance to the regulation of the metazoan G1/S transcription factor, E2F, by the retinoblastoma tumor suppressor protein, Rb (Dyson, 1998; Nevins, 2001) (Figure 1). This is despite the absence of detectable primary structure homology between the protein components of the two transcriptional regulatory systems, suggesting that they evolved independently. Although inactivation of Whi5 leads to premature expression of G1-specific genes and, thereby, premature cell cycle initiation, the strongly periodic expression of Oncogene


2748 Repressed complex Y Whi5 Swi6 E A Swi4 S T

Activated complex

Cln3/Cdk1 Swi6 Swi4 P

Cln1 & 2, Clb5 & 6 and S phase genes

P

Whi5

G1 M E T A Z O A N

Rb E2F

S CycD/Cdk4/6 E2F

Cyclin E & A and S phase genes

P P

Rb

Figure 1 Regulation of G1-specific transcription by cyclindependent protein kinase in yeast and metazoans. In yeast, Cln3associated Cdk1 phosphorylates Whi5, promoting its release from SBF and leading to derepression of SBF-dependent transcription. Activation of SBF-dependent genes leads to accumulation of G1and S-phase cyclins which promote S-phase entry. Accumulation of Cln1- and Cln2-associated Cdk1 may provide additional regulation of Whi5 and SBF. In metazoans cyclin D/Cdk4/6 phosphorylates Rb, relieving its inhibition of E2F/DP1 and leading to derepression of G1/S-phase gene expression. Activation of E2F-responsive genes leads to the accumulation of cyclins E and A, which promote Sphase entry. Accumulation of cyclin E/Cdk2 further phosphorylates Rb, contributing to the activation of E2F-responsive genes

both SBF- and MBF-dependent genes in whi5 mutants belies the existence of other mechanisms regulating this gene family (de Bruin et al., 2004). This is, in part, a consequence of the periodic accumulation of Clb2 (see below), which antagonizes the binding of Swi4 to SBFdependent promoters (Amon et al., 1993; Siegmund and Nasmyth, 1996) leading to transcriptional repression. Although it is not known precisely how localization of Swi6 is regulated, it is likely that both SBF and MBF levels are influenced by the lack of availability of nuclear Swi6 outside the G1 phase (Sidorova et al., 1995; Queralt and Igual, 2003). However, it is unclear whether limitation of nuclear Swi6 is a factor in the periodic expression of target genes in whi5 mutants. It has been suggested that nucleocytoplasmic cycling of Swi6 is a requirement for transcriptional activation (Queralt and Igual, 2003). Finally, Cln3/Cdk can promote transcriptional activation even in the absence of Whi5 (de Bruin and Wittenberg, unpublished). It is possible that this activation involves the direct phosphorylation of Swi6 by Cln/Cdk. Consistent with that notion, the loss of Cdk target sites on Whi5 and Swi6 appears to be lethal (Costanzo et al., 2004). Despite the capacity of Whi5 to bind to MBF under some conditions, activation of MBF-dependent genes appears largely independent of Whi5. Strikingly, MBFdependent genes remain dependent upon Cln/Cdk for activation when Whi5 is inactive. Thus, whereas SBFdependent genes are strongly derepressed in cells lacking both G1 cyclin and Whi5 function (de Bruin et al., 2004), MBF-dependent genes fail to be expressed (de Bruin and Wittenberg, unpublished). In contrast, like SBF-dependent genes, MBF-dependent genes are fully activated when Cln1 and Cln2 functions are retained. Oncogene

This suggests that Cln/Cdk1 activates MBF via a Whi5independent mechanism. MBF activation may involve direct phosphorylation of Swi6 or phosphorylation of another MBF-associated protein. Although Stb1 appears to activate MBF and to be a target of Cln1/2/ Cdk1, phosphorylation does not appear to be required for its activity (Costanzo et al., 2003). The primary determinant of the timing of G1-specific transcription is the activity of Cln3/Cdk1 toward the inactive SBF and MBF transcription factors (Tyers et al., 1993; Dirick et al., 1995; Stuart and Wittenberg, 1995). As such, it is also the primary determinant of cell size. Several regulatory mechanisms appear to play a role in determining Cln3/Cdk1 activity. First, the CLN3 gene is subject to cell cycle-regulated expression as a member of the M/G1 phase gene cluster (McInerny et al., 1997; MacKay et al., 2001) (see below). In addition, CLN3 gene expression and protein abundance are induced by glucose (Hall et al., 1998; Parviz et al., 1998). Next, CLN3 mRNA must be efficiently translated. That process appears to be subject to both transcript-specific and general translational controls (Gallego et al., 1997; Polymenis and Schmidt, 1997). Recent evidence suggests a rather surprising regulatory mechanism whereby CLN3 mRNA and Cdk1 are tethered together in the cytoplasm by the Whi3 protein, perhaps as a mechanism governing complex assembly and transport (Gari et al., 2001; Wang et al., 2004). Furthermore, Cln3 is known to be a highly unstable protein targeted for degradation by an SCF ubiquitin ligase (Tyers et al., 1992). However, the details of that regulation are unclear. Finally, CLN3 appears to be the target of a still mysterious free-running oscillator that underlies the timing of cell cycle events (Haase and Reed, 1999).

The S-phase gene clusters Two clusters of genes characterized by the histone genes and the MET genes, respectively, are expressed immediately following the wave of the G1-specific transcription as cells progress into S phase (Cho et al., 1998; Spellman et al., 1998). The nine histone genes comprise the entire histone cluster. Their expression is tightly coordinated and slightly earlier than the MET gene cluster (Cho et al., 1998; Spellman et al., 1998). Based on genome-wide ChIP analysis, the histone gene promoters are bound by SBF and possibly MBF (Iyer et al., 2001; Simon et al., 2001; Ng et al., 2002), consistent with the presence of putative Swi4-binding motifs in their promoters (Osley et al., 1986; Lee et al., 2002; Kato et al., 2004). In addition, each of the histone gene promoters binds to one or both of the transcriptional repressors, Hir1 and Hir2 (Sherwood et al., 1993), which may act by recruitment of the RSC chromatin remodeling complex (Ng et al., 2002). Binding of the forkhead transcription factors Fkh1/2 (see below) has also been documented. Although this constellation of regulators suggests that histone promoters are G1-specific genes


2749

that are constrained in their expression by the action of transcriptional repressors, inactivation of Hir1 and Hir2 renders the genes constitutive rather than simply shifting the timing of their expression (Sherwood et al., 1993). Thus, it remains unclear whether expression outside the G1/S is mediated by SBF (positive control) or whether Hir proteins are responsible for the difference in the timing of transcription relative to other members of the G1-specific gene cluster (negative control). The mechanism by which Hir proteins are linked to the cell cycle is also unclear. One reasonable suggestion is that the Hir proteins are direct CDK targets, like their mammalian homolog HirA (Hall et al., 2001), and that phosphorylation regulates their cell cycle-dependent function. However, neither yeast repressor contains a full consensus Cdk phosphorylation site nor are they known to be direct targets for Cdk1. The functional significance of S-phase-specific expression of genes of the MET cluster remains enigmatic (but see below). The members of this family differ substantially both in degree of cell cycle-dependent fluctuation and their precise timing. Many members of the family appear to be targeted by the Met4, Met 28, Met32 and Cbf1 transcription factors. All of these factors are clearly associated with the transcriptional induction of MET genes in response to low adenosyl methionine (Thomas and Surdin-Kerjan, 1997). However, whether those factors, or some other regulator, mediate cell cycle-dependent expression is unclear.

Regulation of the Clb2 cluster Transcriptional array analysis has defined a group of 35 yeast genes that are transcribed roughly from the end of S phase until nuclear division (Cho et al., 1998; Spellman et al., 1998). This set of genes has been termed the ‘Clb2 cluster’ based on the CLB2 mitotic cyclin gene, which had previously been shown to exhibit these mRNA kinetics (Ghiara et al., 1991; Surana et al., 1991). Other members of this family include CLB1 (another mitotic cyclin gene), CDC5 (the yeast polo-like kinase homolog), CDC20 (a mitotic specificity factor for the APC protein–ubiquitin ligase), and SWI5 and ACE2 (transcription factors required for late M-early G1specific gene expression). Interestingly, it was through analysis of the SWI5 promoter that insight into regulation of this gene cluster was first obtained. A protein complex known as Swi5 factor (SFF) was shown to be capable of binding to specific elements in the SWI5 promoter (Lydall et al., 1991). More recent work on SWI5 and other cluster genes, notably CLB2, has revealed that SFF sites are binding sites for members of the forkhead family of transcription factors (Pic et al., 2000; Kumar et al., 2000). Yeast contains several members of the ubiquitous forkhead family of transcription factors, classified based on their forkhead (winged-helix) DNA-binding domains. Two of these, Fkh1 and Fkh2, have been shown to be capable of binding to SFF sites in vitro and in vivo

(Kumar et al., 2000; Pic et al., 2000; Hollenhorst et al., 2001). Deletion of either FKH1 or FKH2 does not severely disrupt periodic expression of Clb2 cluster genes. However, simultaneous disruption of both genes leads to constitutive basal transcription that is uncoupled from the cell cycle, suggesting that these transcription factors are functionally redundant (Zhu et al., 2000; Hollenhorst et al., 2001). Yet, in vivo analysis of the CLB2 and other Clb2 cluster promoters suggests that SFF sites are for the most part occupied by Fkh2 and not Fkh1 (Koranda et al., 2000; Hollenhorst et al., 2001). This is likely explained by the presence of binding sites for the MADS-box transcription factor Mcm1 in Clb2 cluster promoters and the cooperativity between Fkh2 and Mcm1 binding, a property not shared with Fkh1 (Hollenhorst et al., 2001). The binding of Fkh2 and Mcm1 to Clb2 cluster promoters does not explain the periodic expression of these genes since occupancy by these factors is not cell cycle regulated. The key to cell cycle regulation is Ndd1, a protein that serves as a co-activator with Fkh2 (Loy et al., 1999; Koranda et al., 2000) (Figure 2). Ndd1 is expressed periodically during S phase and is turned over during mitosis (Loy et al., 1999). Without an apparent DNA-binding domain, Ndd1 transactivation activity depends on binding to the forkheadassociated (FHA) domain of Fkh2 (Koranda et al., 2000). The timing of this binding, which correlates with transcription of Clb2 cluster genes, is mediated by Cdk1-dependent phosphorylation of Ndd1 (Reynolds et al., 2003). The primary cyclin responsible is Clb2 itself (Reynolds et al., 2003), leading to an apparent paradox: if activation of Clb2 transcription by Ndd1 depends on Clb2, how is the process initiated? Two simple nonexclusive models provide a potential explanation. First, the basal level of Clb2 expression might be sufficient to prime the process, which would then rapidly accelerate due to a positive feedback loop. Second, the process could be primed by another B-type cyclin, not under Ndd1 control, for example, Clb3 or Clb4. Termination of Clb2 cluster transcription presumably occurs when Ndd1 is degraded during mitosis (Loy et al., 1999). Interestingly, whereas FKH1 and FKH2 are not essential (the double deletion is viable), Ndd1 is essential (Loy et al., 1999). However, deletion of FKH2 renders Ndd1 nonessential (Koranda et al., 2000; Reynolds et al., 2003). This strongly suggests that, in mediating the periodic expression of Clb2 cluster genes, Fkh2 possesses both repressive and transactivating activities, with Ndd1 binding serving as a functional switch (Figure 2). Deletion analysis of Fkh2 has suggested a mechanism for Ndd1-mediated relief from repression by Fkh2. Compared to Fkh1, Fkh2 contains a carboxy-terminal extension of approximately 280 amino acids. Deletion of this extension, similarly to deletion of the entire FKH2 gene, renders Ndd1 nonessential (Reynolds et al., 2003). Therefore, this Cterminal domain of Fkh2 presumably critical to repression can be neutralized by conformational changes associated with Ndd1 binding. How the C-terminus of Fkh2 establishes repression of Clb2 cluster genes and Oncogene


2750 SBF/MBF

?

Ndd1

Clb2/Cdk1

APC proteasome Ndd1

FHA

Mcm1

Fkh2

Repressor complex

G1/S

Fkh2

Repressor complex

S/G2

P FHA

FHA

Mcm1

Ndd1

Mcm1

Fkh2

Activator complex

G2/M

FHA

Mcm1

Fkh2

Repressor complex

M/G1

Figure 2 Regulation of Clb2 cluster genes. Clb2 cluster genes are characterized by binding sites for Mcm1 and Fkh2 that are occupied continuously through the cell cycle. Genetic evidence suggests that this minimal complex is repressive. Activation depends on the coactivator Ndd1, which is synthesized during during late G1 and/or S phase, possibly through the action of SBF or MBF. However, binding of Ndd1 to Fkh2 depends on phosphorylation by the Clb2/Cdk1 kinase complex during G2. Genetic data suggest that binding of phosphorylated Ndd1 to the FHA domain of Fkh2 leads to a conformational change that converts Fkh2 from a repressor to a transcriptional activator. APC-mediated ubiquitylation and degradation of Ndd1 during mitosis restores the complex to its repressive state, terminating transcription of Clb2 cluster genes

how Ndd1 antogonizes this remains to be determined. In addition, the mechanism whereby, in the absence of Fkh2 and Ndd1, Fkh1 mediates cell cycle-regulated periodic expression of Clb2 cluster genes is not understood (Reynolds et al., 2003). Presumably, a cell cycleregulated Fkh1 co-activator other than Ndd1 must be involved, but this protein has not yet been identified. The respective functions of Fkh1 and Fkh2 in regulation of Clb2 cluster transcription are further complicated by the finding that these proteins have general nonredundant roles in transcriptional elongation of a broad spectrum of yeast genes (Morillon et al., 2003). Although mutation of FKH1 and/or FKH2 leads to defects in phosphorylation of the large subunit of RNA polymerase II, in polymerase movement from the 50 –30 ends of ORFs, and in transcriptional termination, the molecular basis for this is not known. Nevertheless, post-initiation transcriptional functions of Fkh1 and Fkh2 might contribute to Clb2 cluster periodic expression. Interestingly, an Fkh1/Fkh2 homolog in mammalian cells, FoxM1, has been shown to have a role in transcription of the cyclin B1 gene (the Clb2 homolog) as well as genes encoding other mitotic regulators (Leung et al., 2001; Wang et al., 2002). Therefore, the role of transcription factors containing forkhead DNAbinding domains and their cognate DNA-binding sites may constitute an ancient and highly conserved regulatory motif for late cell cycle transcription.

Regulation of M-G1 genes A large number of yeast genes are expressed from M phase to early G1 (Cho et al., 1998; Spellman et al., 1998). Many of these are required for G1 functions (e.g., MCM proteins involved in prereplication complex assembly, transcription factors required for G1 gene expression and proteins involved in the yeast mating response, which occurs during G1 (Cho et al., 1998; Oncogene

Spellman et al., 1998)). However, some proteins with no ostensible G1-specific function (e.g., genes of the PHO regulon) also exhibit this expression pattern (Cho et al., 1998; Spellman et al., 1998). The largest group of genes exhibiting M-G1 periodicity are those driven by the MADS-box transcription factor, Mcm1 (Nurrish and Treisman, 1995; Shore and Sharrocks, 1995). Unlike the Clb2 cluster of genes that also require Mcm1 binding, the so-called MCM cluster genes do not contain sites for Fkh1/2 binding. As expected, therefore, deletion of FKH1 and FKH2 has no effect on these genes (Zhu et al., 2000). Instead, most of these genes contain a site for binding the homeodomain repressors Yox1 and Yhp1 (Pramila et al., 2002) in close proximity to a palindromic site capable of binding an Mcm1 dimer, known as an ECB for ‘early cell cycle box’ (McInerny et al., 1997) (although there is some disagreement on this issue; see Horak and Snyder, 2002). In addition, Yox1 and Yhp1 can bind directly to Mcm1 (Pramila et al., 2002), presumably promoting cooperativity. It is the occupancy of these repressive sites that blocks Mcm1-mediated transcription through most of the cell cycle, and it is the periodic transcription of the genes encoding these presumably unstable repressors that determines their functional interval. Yox1 is expressed in mid-G1 through early S phase (Spellman et al., 1998; Horak and Snyder, 2002; Pramila et al., 2002), whereas Yhp1 appears to be expressed later in the cell cycle (Cho et al., 1998; Spellman et al., 1998), leaving an interval from M phase to early G1 where these repressors are absent. During this window, Mcm1 can apparently drive transcription of these genes without DNA binding of additional activating proteins. The second class of M-G1 expressed genes is referred to as the Sic1 cluster. Interestingly, periodic expression of these genes is affected by deletion of FKH1 and FKH2 (Zhu et al., 2000). Yet, Fkh1 and Fkh2 do not bind the promoters of these genes. The basis for the indirect requirement for FKH genes is the assignment of transcription factors Swi5 and Ace2 to the Clb2 cluster


2751

(Nasmyth et al., 1987; Spellman et al., 1998). Genes of the Sic1 cluster contain related consensus sites that bind Ace2 and Swi5 (Zhu et al., 2000). The relationship between Ace2 and Swi5, however, is complex. Although both transcription factors recognize the same set of sites on target genes, their effect on such genes can vary from activation to repression (Dohrmann et al., 1996; McBride et al., 1999). The explanation for this differential behavior appears to reside in the ability of these factors to bind co-activators and co-repressors that are also targeted to genes of the Sic1 cluster. For example, the gene encoding the endonuclease HO involved in mating type switching appears to be specific for Swi5, presumably because it binds cooperatively with the homeodomain protein Pho2, which also binds to the HO promoter (McBride et al., 1999). Ace2 cannot interact with Pho2. Conversely, the CTS1 gene, which is specifically activated by Ace2, contains a putative repressor-binding site that prevents activation by Swi5 but not Ace2 (McBride et al., 1999; Doolin et al., 2001). A third group of genes that are activated during the M-G1 window are those that are normally induced by mating pheromone (the MAT cluster in Spellman et al., 1998). It has been shown that the periodic expression of these genes depends on a DNA sequence known as the pheromone response element (PRE) and a PRE-binding transcription factor, Ste12 (Oehlen et al., 1996). Under conditions of receptor-mediated activation of the pheromone response pathway, the MAP kinase homolog Fus3 promotes activation of Ste12 by phosphorylating and inactivating two co-repressors of Ste12 known as Dig1 and Dig2 (Bardwell et al., 1998; van Drogen et al., 2001; Breitkreutz et al., 2003; Kusari et al., 2004). However, whether Dig1/Dig2 targeting by Fus3 is involved in M-G1 activation of Ste12 is not known. One plausible idea is that genes encoding elements ratelimiting in the signal transduction pathway, many of which have PRE elements themselves, also contain other cell cycle-regulated elements, which would prime the system and set up a positive feedback loop. In effect, an increase in basal signaling would then increase the transcription of rate-limiting signaling genes, further amplifying the basal signal. Intriguingly, STE2, the alpha-factor receptor gene at the apex of the pheromone response pathway, contains a hybrid promoter (see below), with PRE elements (driven by Ste12) and ECBs (driven by Mcm1) (Hwang-Shum et al., 1991). It is therefore possible that transcription of STE2 driven by Mcm1 initiates an increase in basal signaling down the pheromone response pathway leading to increased Ste12-mediated STE2 transcription. In effect, Mcm1 priming of STE2 transcription would set up a Ste12driven positive feedback loop affecting all PRE-containing promoters. A fourth group of M-G1-activated genes are surprisingly those of the PHO regulon (Cho et al., 1998; Spellman et al., 1998). These genes encoding proteins involved in scavenging and transporting phosphate are coordinately induced by phosphate starvation (Lenburg and O’Shea, 1996). The phosphate-sensing pathway ultimately impinges on a cyclin-dependent kinase Pho85

by activating the Cdk inhibitor Pho81 under lowphosphate conditions (Lenburg and O’Shea, 1996). Inactivation of Pho85 allows nuclear retention of transcription factor Pho4, which, alone or in concert with the homeodomain protein Pho2, drives transcription of more than 20 PHO regulon genes (Komeili and O’Shea, 1999). M/G1 activation of PHO regulon genes requires the Pho85 inhibitor Pho81 and transcription factors Pho2 and Pho4, indicating that the phosphate sensing pathway is involved (Neef and Kladde, 2003). Further analysis has suggested that the timing is a result of periodic storage and depletion of polyphosphate pools, ultimately leading to low intracellular phosphate levels at the end of the cell cycle that trigger the phosphate starvation response (Neef and Kladde, 2003).

Hybrid promoters A significant number of cell cycle-regulated genes do not precisely fit the criteria used to define any of the major clusters. In most cases, this is because the promoters of these genes contain binding sites that are characteristic of more than one cluster (Kato et al., 2004). Although the majority of promoters activated specifically during G1 are bound by SBF, MBF, or both, it is clear that many of those genes are also bound by other transcription factors including the fork-head transcription factors Fkh1 and Fkh2, and Stb1 (Lee et al., 2002). As the data supporting these interactions is derived largely from genome-wide analysis of asynchronous populations, their impact on the pattern of expression of those genes remains unclear. Establishing whether the binding of these factors has regulatory significance or whether they act simultaneously or sequentially awaits analysis in synchronized populations of wild-type and mutant cells. Hybrid promoters have also been observed in the Clb2 cluster. For example, CDC20 is grouped in the Clb2 cluster based upon its pattern of transcription in wild-type cells and its well-documented Fkh2-binding sites (Spellman et al., 1998; Hollenhorst et al., 2001). However, CDC20 differs from most other Clb2 cluster genes in several important respects. First, the interval of mRNA accumulation is more restricted than most Clb2 cluster genes, with peak expression coming relatively late (Zhu et al., 2000). Second, CDC20 maintains periodic transcription even in an fkh1 fkh2 null background, although the kinetics of expression are shifted to an M/G1 pattern characteristic of the MCM cluster (Zhu et al., 2000). Analysis of the sequences surrounding the CDC20 Mcm1-binding sites (ECBs) reveals at least one consensus and possibly several for binding of the Yox1 repressor (Pramila et al., 2002). Thus, transcription of CDC20 is most likely positively regulated at the G2–M boundary by Fkh2 in conjunction with Ndd1 and Mcm1, but negatively regulated by Yox1 and Yhp1 until the M phase. Another example of a hybrid promoter is that of the FAR1 gene. Far1, a protein required for both cell cycle Oncogene


2752

arrest and polarized growth in response to mating pheromone, accumulates from mitosis through G1 in non-mating cells (Oehlen et al., 1996). Examination of the FAR1 promoter reveals both an ECB, presumably regulated by Mcm1, Yox1, and Yhp1, and PREs regulated by Ste12 (Oehlen et al., 1996). Elimination of Mcm1 results in loss of mitotic expression of FAR1, whereas elimination of Ste12 results in loss of G1 expression (Oehlen et al., 1996). Thus, utilizing two transactivating systems allows FAR1 expression to be extended over a broader cell cycle interval than would be permitted by either system alone. The gene encoding the alpha-factor pheromone receptor STE2 appears to be regulated in a similar fashion (Hwang-Shum et al., 1991; Spellman et al., 1998). Cell cycle regulation of genes not ostensibly involved in cell cycle functions Arguably, virtually all metabolic and structural functions in microbial organisms either directly or indirectly impinge on proliferation. Since the success of most microorganisms is measured in terms of utilizing available resources to outcompete other organisms, the link between nutrient acquisition, nutrient assimilation, energy metabolism, and cell cycle control should not be surprising. In some cases, the periodic nature of gene expression can be rationalized in this context. For example, genes of the PHO regulon are induced at the M–G1 boundary (Cho et al., 1998; Spellman et al., 1998). Most of the proteins encoded by PHO regulon genes are directly involved in scavenging phosphate from the environment and internalizing it. Since the availability of phosphate is one of the determinants that yeast cells use to make a decision on initiating a new cell cycle during G1, perhaps expression of PHO regulon genes at the M/G1 boundary, thereby mobilizing all available environmental phosphate, constitutes a means of optimizing the reliability of this decision-making process. The MET regulon genes serve as another example. These genes, encoding enzymes of the methionine biosynthetic pathway, are normally induced in response to methionine depletion in the environment (Thomas and Surdin-Kerjan, 1997). However, they are also induced periodically during the S phase (see above) (Cho et al., 1998; Spellman et al., 1998). The most likely rationale for this pattern is that methionine is the precursor for S-adenosyl methionine, the one-carbon donor essential for the biosynthesis of dTTP required for genome replication during the S phase. Other periodically expressed genes are more difficult to rationalize. The gene encoding the plasma membrane ATPase PMA1 is expressed periodically during G2, similarly to Clb2 cluster genes (Cho et al., 1998; Spellman et al., 1998). Pma1 functions as an ATPdriven proton efflux pump, presumably regulating intracellular pH, a parameter not obviously linked to cell cycle processes. However, Pma1 has also been shown to be required for maintaining intracellular Ca2 þ levels (Withee et al., 1998). Therefore, an increase Oncogene

in PMA1 expression during G2 is most likely associated with potentiation of a transient increase in intracellular Ca2 þ required for mitosis in many organisms (Whitaker, 1997). In conclusion, it appears that cell cycle regulation of transcription of genes not directly linked to cell cycle functions can usually be rationalized in the context of supporting or potentiating cell cycle functions indirectly. Maintaining cell cycle organization: interactions between transcriptional programs Sequential waves of transcription are critical for maintaining the organization of cell cycle events. The genome-wide analysis of transcription factor binding, gene expression arrays, and bioinformatics analysis of promoter motifs along with gene-specific studies of transcriptional regulation have led to the visualization of the network of regulatory interactions that promote and maintain cell cycle-regulated transcription (Futcher, 2002; Lee et al., 2002; Kato et al., 2004). Our current understanding of those interactions is depicted in Figure 3. Whereas many of these regulatory interactions are established by experimental observation, some are simply inferred from genome-wide location analysis and require experimental confirmation. Integration of the cell cycle-dependent transcriptional regulatory network appears to be imposed via three general mechanisms. First, the gene encoding a

Figure 3 Integration of transcriptional regulatory networks during the cell cycle. This diagram depicts the regulatory interactions occurring between cell cycle-specific gene clusters as discussed in the text. Transcription factors are denoted by ovals, cell cycle regulatory proteins are denoted by circles, and transcriptional repressors are denoted by squares. Dashed lines represent gene expression and solid lines represent direct regulatory interactions. Gray areas indicate intervals of transcription factor activity


2753

transcription factor (activator or repressor) can be directly regulated in the context of the cell cycledependent transcription program. For example, NDD1, encoding an essential component of the active form of the Fkh2 transcription complex, is a member of the G1-specific gene cluster (Spellman et al., 1998; Loy et al., 1999) and is required for activation of the Clb2 gene cluster (Loy et al., 1999; Koranda et al., 2000). Similarly, SWI4, which encodes the DNA-binding component of the G1-specific transcriptional activator SBF, is expressed as a member of the M/G1 gene cluster (McInerny et al., 1997; Spellman et al., 1998). The second potential mode of coupling is via cell cycle-dependent expression of an enzymatic activity required for regulation of transcription of another gene cluster. Regulated expression of cyclin genes provides the best examples of this mechanism. Expression of the CLB2 gene, the canonical member of the G2 phase gene family, leads to activation of Clb2/Cdk1 which binds to Swi4 and, thereby, inactivates SBF (Amon et al., 1993; Siegmund and Nasmyth, 1996). The same Cdk complex phosphorylates Ndd1 to activate expression of the Clb2 cluster (Reynolds et al., 2003). Mcm1-dependent expression of CLN3 during M/G1 provides another example of this form of regulation (McInerny et al., 1997; Pramila et al., 2002). The activation of Cln3/Cdk1 promotes phosphorylation and inactivation of Whi5, thereby activating G1-specific transcription (Costanzo et al., 2004; de Bruin et al., 2004). Finally, a third observed mechanism involves the regulation of one cluster of genes by a transcriptional repressor expressed as a member of another gene cluster. Perhaps the best example of this form of regulation is the restriction of Mcm1-mediated gene expression to M/G1 by the transcriptional repressors Yox1 and Yhp1, which are expressed during G1/S and G2/M phases, respectively. Yox1 is most likely a target of SBF (Spellman et al., 1998; Horak and Snyder, 2002; Pramila et al., 2002).

Conclusions Yeast genomic analysis has provided a wealth of data, including the observation that a large number of genes are transcribed according to a cell cycle program (Cho et al., 1998; Spellman et al., 1998). While the fact that expression of a significant number of genes is regulated in this fashion is not surprising, perhaps the magnitude of this cohort (>10%) is. Whether this is a phenomenon related specifically to budding yeast and its lifestyle remains to be determined. Indeed recent genome-wide analysis of cell cycle transcription in fission yeast (S. pombe) indicates that a much lower percentage of genes are cell cycle regulated in that organism (136 compared to 800) (Rustici et al., 2004). Furthermore, cell cycle regulation of transcription appears less complex and relegated to a limited interval. Whereas in S. cerevisiae, waves of transcription around the cell cycle are linked, forming a closed circuit (see above), in S. pombe, linkage

appears to be minimal and most of the cell cycle is devoid of cell cycle-regulated transcription (Rustici et al., 2004). Why two yeasts inhabiting similar environmental niches should exhibit such disparate transcriptional programs is difficult to rationalize, but suggests the advantages gained by extensive cell cycle programming of transcription can be achieved by other regulatory strategies. Interestingly, it was found that a core set of approximately 40 genes exhibited cell cycle regulation of transcription in both organisms, suggesting that this constitutes a set of genes for which cell cycle regulation is critical rather than simply advantageous (Rustici et al., 2004). On the other hand, genome-wide analysis of transcription through the cell cycle in human cells indicates that minimally 800 genes are cell cycle-regulated, similarly to S. cerevisiae (Cho et al., 2001; Whitfield et al., 2002). Although, as expected, most of the 40 or so core genes that exhibit cell cycle regulation in both S. cerevisiae and S. pombe are also cell cycle regulated in proliferating human cells, the human data suggest that, as in S. cerevisiae, many functions only peripherally related to the cell cycle at best are part of the cell cycle transcriptional program, indeed constituting the majority of cell cycleregulated genes (Cho et al., 2001; Whitfield et al., 2002). These processes include signal transduction, mRNA metabolism, protein translation, energy metabolism, ionic homeostasis and many other categories. Therefore, it would appear that transcriptionally toggling diverse cellular processes and functions to the cell cycle program is the rule rather than the exception. In addition to the pervasiveness of cell cycle-regulated transcription, there are interesting mechanistic parallels that imply that conservative pressures are at work. The recent discovery of Whi5 as a G1-specific transcriptional repressor belies a mechanistic conservation of transcriptional regulatory mechanisms. Although Whi5 shares no recognizable sequence homology to any of the, socalled, ‘pocket proteins’ (pRb, p107 and p130), as a repressor of G1-specific transcription that is antagonized by G1 cyclin/Cdk, it can be considered a functional analog arising through convergent evolution (Figure 1). Similarly, the utilization of forkhead domain transcription factors for G2-specific transcription in both yeasts and metazoans underscores the conservative tendancy of mechanisms governing cell cycle-regulated transcription. As yeast lacks some of the extensive redundancy that plagues the study of transcriptional processes in animal cells, it will likely continue to prove a useful model for the dissection of mechanisms governing cell cycle-mediated transcriptional control. Acknowledgements We thank all of the investigators whose research has contributed to our understanding of cell cycle-regulated transcription in yeast, and apologize to those who have been overlooked in the interest of brevity. We thank Rob de Bruin for comments on the manuscript and members of the TSRI Cell Cycle Group for their interest and support. Research concerning cell cycle-regulated transcription in our own laboratories is supported by funding from the National Institutes of Health. Oncogene


2754 References Amon A. (2002). Methods Enzymol., 351, 457–467. Amon A, Tyers M, Futcher B and Nasmyth K. (1993). Cell, 74, 993–1007. Bardwell L, Cook JG, Zhu-Shimoni JX, Voora D and Thorner J. (1998). Proc. Natl. Acad. Sci. USA, 95, 15400–15405. Breeden L. (1996). Curr. Top. Microbiol. Immunol., 208, 95– 127. Breitkreutz A, Boucher L, Breitkreutz BJ, Sultan M, Jurisica I and Tyers M. (2003). Genetics, 165, 997–1015. Cho RJ, Campbell MJ, Winzeler EA, Steinmetz L, Conway A, Wodicka L, Wolfsberg TG, Gabrielian AE, Landsman D, Lockhart DJ and Davis RW. (1998). Mol. Cell, 2, 65–73. Cho RJ, Huang M, Campbell MJ, Dong H, Steinmetz L, Sapinoso L, Hampton G, Elledge SJ, Davis RW and Lockhart DJ. (2001). Nat. Genet., 27, 48–54. Costanzo M, Nishikawa JL, Tang X, Millman JS, Schub O, Breitkreuz K, Dewar D, Rupes I, Andrews B and Tyers M. (2004). Cell, 117, 899–913. Costanzo M, Schub O and Andrews B. (2003). Mol. Cell. Biol., 23, 5064–5077. Cross FR. (1988). Mol. Cell. Biol., 8, 4675–4684. Cross FR and Tinklenberg AH. (1991). Cell, 65, 875–883. de Bruin RA, McDonald WH, Kalashnikova TI, Yates III J and Wittenberg C. (2004). Cell, 117, 887–898. Di Como CJ, Chang H and Arndt KT. (1995). Mol. Cell. Biol., 15, 1835–1846. Dirick L, Bohm T and Nasmyth K. (1995). EMBO J., 14, 4803–4813. Dirick L and Nasmyth K. (1991). Nature, 351, 754–757. Dohrmann PR, Voth WP and Stillman DJ. (1996). Mol. Cell. Biol., 16, 1746–1758. Doolin MT, Johnson AL, Johnston LH and Butler G. (2001). Mol. Microbiol., 40, 422–432. Dyson N. (1998). Genes Dev., 12, 2245–2262. Edgington NP and Futcher B. (2001). J. Cell Sci., 114, 4599– 4611. Epstein CB and Cross FR. (1994). Mol. Cell. Biol., 14, 2041– 2047. Futcher B. (2002). Curr. Opin. Cell. Biol., 14, 676–683. Gallego C, Gari E, Colomina N, Herrero E and Aldea M. (1997). EMBO J., 16, 7196–7206. Gari E, Volpe T, Wang H, Gallego C, Futcher B and Aldea M. (2001). Genes Dev., 15, 2803–2808. Geymonat M, Spanos A, Wells GP, Smerdon SJ and Sedgwick SG. (2004). Mol. Cell. Biol., 24, 2277–2285. Ghiara JB, Richardson HE, Sugimoto K, Henze M, Lew DJ, Wittenberg C and Reed SI. (1991). Cell, 65, 163–174. Haase SB and Reed SI. (1999). Nature, 401, 394–397. Hall C, Nelson DM, Ye X, Baker K, DeCaprio JA, Seeholzer S, Lipinski M and Adams PD. (2001). Mol. Cell. Biol., 21, 1854–1865. Hall DD, Markwardt DD, Parviz F and Heideman W. (1998). EMBO J., 17, 4370–4378. Ho Y, Mason S, Kobayashi R, Hoekstra M and Andrews B. (1997). Proc. Natl. Acad. Sci. USA, 94, 581–586. Hollenhorst PC, Pietz G and Fox CA. (2001). Genes Dev., 15, 2445–2456. Horak CE and Snyder M. (2002). Funct. Integr. Genomics, 2, 171–180. Hwang-Shum JJ, Hagen DC, Jarvis EE, Westby CA and Sprague Jr GF. (1991). Mol. Gen. Genet., 227, 197–204. Iyer VR, Horak CE, Scafe CS, Botstein D, Snyder M and Brown PO. (2001). Nature, 409, 533–538. Jorgensen P, Nishikawa JL, Breitkreutz BJ and Tyers M. (2002). Science, 297, 395–400. Oncogene

Kato M, Hata N, Banerjee N, Futcher B and Zhang MQ. (2004). Genome Biol., 5, R56. Koch C, Moll T, Neuberg M, Ahorn H and Nasmyth K. (1993). Science, 261, 1551–1557. Komeili A and O’Shea EK. (1999). Science, 284, 977–980. Koranda M, Schleiffer A, Endler L and Ammerer G. (2000). Nature, 406, 94–98. Kumar R, Reynolds DM, Shevchenko A, Goldstone SD and Dalton S. (2000). Curr. Biol., 10, 896–906. Kusari AB, Molina DM, Sabbagh Jr W, Lau CS and Bardwell L. (2004). J. Cell Biol., 164, 267–277. Lee TI, Rinaldi NJ, Robert F, Odom DT, Bar-Joseph Z, Gerber GK, Hannett NM, Harbison CT, Thompson CM, Simon I, Zeitlinger J, Jennings EG, Murray HL, Gordon DB, Ren B, Wyrick JJ, Tagne JB, Volkert TL, Fraenkel E, Gifford DK and Young RA. (2002). Science, 298, 799–804. Lenburg ME and O’Shea EK. (1996). Trends Biochem. Sci., 21, 383–387. Leung TW, Lin SS, Tsang AC, Tong CS, Ching JC, Leung WY, Gimlich R, Wong GG and Yao KM. (2001). FEBS Lett., 507, 59–66. Loy CJ, Lydall D and Surana U. (1999). Mol. Cell. Biol., 19, 3312–3327. Lydall D, Ammerer G and Nasmyth K. (1991). Genes Dev., 5, 2405–2419. MacKay VL, Mai B, Waters L and Breeden LL. (2001). Mol. Cell. Biol., 21, 4140–4148. Marini NJ and Reed SI. (1992). Genes Dev., 6, 557–567. McBride HJ, Yu Y and Stillman DJ. (1999). J. Biol. Chem., 274, 21029–21036. McInerny CJ, Partridge JF, Mikesell GE, Creemer DP and Breeden LL. (1997). Genes Dev., 11, 1277–1288. Miller ME and Cross FR. (2000). Mol. Cell. Biol., 20, 542–555. Morgan BA, Bouquin N, Merrill GF and Johnston LH. (1995). EMBO J., 14, 5679–5689. Morillon A, O’Sullivan J, Azad A, Proudfoot N and Mellor J. (2003). Science, 300, 492–495. Nash R, Tokiwa G, Anand S, Erickson K and Futcher AB. (1988). EMBO J., 7, 4335–4346. Nasmyth K, Seddon A and Ammerer G. (1987). Cell, 49, 549– 558. Neef DW and Kladde MP. (2003). Mol. Cell. Biol., 23, 3788– 3797. Nevins JR. (2001). Hum. Mol. Genet., 10, 699–703. Ng HH, Robert F, Young RA and Struhl K. (2002). Genes Dev., 16, 806–819. Nurrish SJ and Treisman R. (1995). Mol. Cell. Biol., 15, 4076– 4085. Oehlen LJ, McKinney JD and Cross FR. (1996). Mol. Cell. Biol., 16, 2830–2837. Osley MA, Gould J, Kim S, Kane MY and Hereford L. (1986). Cell, 45, 537–544. Partridge JF, Mikesell GE and Breeden LL. (1997). J. Biol. Chem., 272, 9071–9077. Parviz F, Hall DD, Markwardt DD and Heideman W. (1998). J. Bacteriol., 180, 4508–4515. Pic A, Lim FL, Ross SJ, Veal EA, Johnson AL, Sultan MR, West AG, Johnston LH, Sharrocks AD and Morgan BA. (2000). EMBO J., 19, 3750–3761. Polymenis M and Schmidt EV. (1997). Genes Dev., 11, 2522– 2531. Pramila T, Miles S, GuhaThakurta D, Jemiolo D and Breeden LL. (2002). Genes Dev., 16, 3034–3045. Queralt E and Igual JC. (2003). Mol. Cell. Biol., 23, 3126– 3140.


2755 Reynolds D, Shi BJ, McLean C, Katsis F, Kemp B and Dalton S. (2003). Genes Dev., 17, 1789–1802. Rustici G, Mata J, Kivinen K, Lio P, Penkett CJ, Burns G, Hayles J, Brazma A, Nurse P and Bahler J. (2004). Nat. Genet., 36, 809–817. Sherwood PW, Tsang SV and Osley MA. (1993). Mol. Cell. Biol., 13, 28–38. Shore P and Sharrocks AD. (1995). Eur. J. Biochem., 229, 1– 13. Sidorova JM, Mikesell GE and Breeden LL. (1995). Mol. Biol. Cell, 6, 1641–1658. Siegmund RF and Nasmyth KA. (1996). Mol. Cell. Biol., 16, 2647–2655. Simon I, Barnett J, Hannett N, Harbison CT, Rinaldi NJ, Volkert TL, Wyrick JJ, Zeitlinger J, Gifford DK, Jaakkola TS and Young RA. (2001). Cell, 106, 697–708. Spellman PT, Sherlock G, Zhang MQ, Iyer VR, Anders K, Eisen MB, Brown PO, Botstein D and Futcher B. (1998). Mol. Biol. Cell, 9, 3273–3297. Struhl K. (1983). Nature, 305, 391–397. Stuart D and Wittenberg C. (1995). Genes Dev., 9, 2780–2794. Surana U, Robitsch H, Price C, Schuster T, Fitch I, Futcher AB and Nasmyth K. (1991). Cell, 65, 145–161. Thomas D and Surdin-Kerjan Y. (1997). Microbiol. Mol. Biol. Rev., 61, 503–532. Tyers M, Tokiwa G and Futcher B. (1993). EMBO J., 12, 1955–1968.

Tyers M, Tokiwa G, Nash R and Futcher B. (1992). EMBO J., 11, 1773–1784. Ubersax JA, Woodbury EL, Quang PN, Paraz M, Blethrow JD, Shah K, Shokat KM and Morgan DO. (2003). Nature, 425, 859864. van Drogen F, Stucke VM, Jorritsma G and Peter M. (2001). Nat. Cell Biol., 3, 1051–1059. Wang H, Gari E, Verges E, Gallego C and Aldea M. (2004). EMBO J., 23, 180–190. Wang X, Kiyokawa H, Dennewitz MB and Costa RH. (2002). Proc. Natl. Acad. Sci. USA, 99, 16881–16886. Whitaker M. (1997). Prog. Cell Cycle Res., 3, 261–269. Whitfield ML, Sherlock G, Saldanha AJ, Murray JI, Ball CA, Alexander KE, Matese JC, Perou CM, Hurt MM, Brown PO and Botstein D. (2002). Mol. Biol. Cell, 13, 1977–2000. Wijnen H and Futcher B. (1999). Genetics, 153, 1131–1143. Wijnen H, Landman A and Futcher B. (2002). Mol. Cell. Biol., 22, 4402–4418. Withee JL, Sen R and Cyert MS. (1998). Genetics, 149, 865– 878. Wittenberg C and Reed SI. (1991). Crit. Rev. Eukaryot. Gene Expr., 1, 189–205. Zhang J, Schneider C, Ottmers L, Rodriguez R, Day A, Markwardt J and Schneider BL. (2002). Curr. Biol., 12, 1992–2001. Zhu G, Spellman PT, Volpe T, Brown PO, Botstein D, Davis TN and Futcher B. (2000). Nature, 406, 90–94.

Oncogene