YEASTBOOK
Topology and Control of the Cell-Cycle-Regulated
Transcriptional Circuitry
Steven B. Haase* and Curt Wittenberg†,1
*Department of Biology and Duke Center for Systems Biology, Duke University, Durham, North Carolina 27708, and yDepartment of Cell
and Molecular Biology, The Scripps Research Institute, La Jolla, California 92037
ABSTRACT Nearly 20% of the budding yeast genome is transcribed periodically during the cell division cycle. The precise temporal
execution of this large transcriptional program is controlled by a large interacting network of transcriptional regulators, kinases, and
ubiquitin ligases. Historically, this network has been viewed as a collection of four coregulated gene clusters that are associated with
each phase of the cell cycle. Although the broad outlines of these gene clusters were described nearly 20 years ago, new technologies
have enabled major advances in our understanding of the genes comprising those clusters, their regulation, and the complex
regulatory interplay between clusters. More recently, advances are being made in understanding the roles of chromatin in the control
of the transcriptional program. We are also beginning to discover important regulatory interactions between the cell-cycle
transcriptional program and other cell-cycle regulatory mechanisms such as checkpoints and metabolic networks. Here we review
recent advances and contemporary models of the transcriptional network and consider these models in the context of eukaryotic cellcycle controls.
TABLE OF CONTENTS
Abstract
65
Introduction
66
Evolution of Experimental Approaches and Models
The single-gene approach
Genomic approaches
Caveats associated with population-based approaches
Algorithmic approaches to population studies
Single-cell approaches
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Cell-Cycle-Regulated Gene Clusters: The Logic
and Strategy for Expression
The G1/S gene cluster
Activation of G1/S transcription factors
Repression of G1/S transcription factors
Temporal regulation within the G1/S gene cluster
Coupling of G1/S transcription to the rest of the transcriptional program
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Continued
Copyright © 2014 by the Genetics Society of America
doi: 10.1534/genetics.113.152595
Manuscript received May 9, 2013; accepted for publication September 16, 2013
1
Corresponding author: Department of Molecular Biology, MB-3, 10550 North Torrey Pines Road, La Jolla, CA 92037. E-mail: [email protected]
Genetics, Vol. 196, 65–90
January 2014
65
CONTENTS, continued
The
The
The
The
The
The
S phase gene cluster
G2/M gene cluster
M/G1 gene cluster
Mcm1 gene cluster
Sic1 gene cluster
MAT gene cluster
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Emerging Views of Cell-Cycle-Regulated Transcription
CDK’s role in cell-cycle-regulated transcriptional program
Checkpoint control of cell-cycle-regulated transcription
Yeast metabolic cycle control of cell cycle and transcription
The YMC-regulated transcriptional program:
YMC control of cell cycle:
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What is the role of chromatin in
cell-cycle-regulated transcription?
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Rationale for Cell-Cycle-Regulated Transcription: Need to Have or Nice to Have?
82
The Topology of Cell-Cycle-Regulated Transcriptional Circuitry is Conserved
83
T
HE cell cycle is the sum of the processes by which cells
replicate. That process, involving the replication and redistribution of all cellular components, requires the synthesis
of many proteins, including constituents of cellular structures and organelles, enzymes that catalyze the many anabolic and catabolic processes required for their replication
and distribution, as well as the regulatory proteins that govern those events. Although the genes encoding many of
those proteins are expressed constitutively, a large number
are expressed at or near the interval when they are needed.
Consequently, progression through the cell cycle is accompanied by dramatic reorganization of gene expression that
we refer to collectively as “cell-cycle-regulated transcription.” The nature and mechanism of that program of gene
expression is the subject of this review.
The first cell-cycle-regulated genes were observed in
yeast more than three decades ago (Hereford et al. 1981)
when the mRNA encoding histone genes was observed to
accumulate periodically during the course of the cell cycle in
a synchronized population of cells. Since then, the list of
cell-cycle-regulated genes has grown, slowly at first, one
gene at a time, and then very rapidly, largely as a consequence of genome-wide approaches, to encompass between
as much as 20% of the genome (Cho et al. 1998; Spellman
et al. 1998; Pramila 2002; Orlando et al. 2008; Guo et al.
2013). Despite the relatively large number of individual
genes that are periodically expressed, it has become clear
that they fall into a relatively small number of gene families
that are coregulated. Consequently, the entire program
appears to be controlled by a relatively small set of specific
transcriptional regulatory factors.
This general topic has been extensively reviewed (Bähler
2005; Wittenberg and Reed 2005; McInerny 2011) and in-
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depth reviews covering specific transcription factor families
and cell-cycle-regulated gene clusters have been presented
(Murakami et al. 2010; Cross et al. 2011; Eriksson et al.
2012). We will introduce the constituents and regulatory
logic of the cell-cycle transcriptional circuitry with discussion weighted toward more recent contributions.
A general understanding of both the pattern of gene
expression and the regulation of the cell-cycle transcriptional program is, in many cases, emerging. When viewed
in its entirety, the program appears as a continuum of
transcriptional activation and deactivation. However, we
now appreciate that waves of gene expression are coupled to
observable cell-cycle events, which, in most cases, depend
on the activity of the cyclin-dependent protein kinase Cdk1
(Cdc28, see below). The transcriptional program guides the
activity of Cdk1 by initiating the properly timed expression
of specific cyclin genes. In turn, cyclin/Cdk1 complexes
phosphorylate transcription factors and regulate their activity. Thus, there is a complex dynamic interplay between the
transcriptional program, CDK activity, and cell-cycle progression (Figure 1). Waves of gene expression are associated
with (i) cell-cycle initiation late in G1 phase prior to the
initiation of S phase (G1/S transcription), (ii) S phase
(S phase transcription), (iii) the transition from G2 phase
into M phase (G2/M transcription), and (iv) the transition of
cells from M phase back into G1 phase (M/G1 transcription). Genes within a coregulated cluster are not all activated at the same time but appear to be turned on in
a precise order during an interval that can span 20% of
the cell cycle (Eser et al. 2011; Guo et al. 2013). The consequence of this highly regulated pattern of transcription is
the sequential periodic expression of upwards of 1000
genes.
Figure 1 The cell-cycle transcriptional circuitry. This
transcriptional circuit depicts the major interactions between transcriptional activators and repressors and
their regulation by cyclin/CDK and APC discussed in
the course of this article. This circuit is not meant to
be exhaustive but rather to provide a reference for the
interaction between the cluster regulators that are
depicted in the subsequent figures. Many other interactions are discussed in the body of the article. Subunits in green are those with activating activity,
subunits in red are those with repressing or inhibitory
activity, and subunits in blue represent those that require a regulatory subunit. Arrows in green represent
activating activities, those in red represent repressing
activities, and those in black represent transitions in the
process.
The existence of this intricately orchestrated sequence of
transcriptional activity raises a number of important questions that form the basis for contemporary studies. Which
genes constitute the full cell-cycle-regulated set? How is the
transcriptional activation within a coregulated cluster ordered, and how is the temporal order of clusters maintained?
How is the cell-cycle-regulated transcriptional program influenced by regulatory pathways, including checkpoints and the
yeast metabolic cycle (YMC)? Recent findings that suggest
the transcriptional program can be uncoupled from cell-cycle
progression leave in question the mechanisms that maintain
synchrony between the transcriptional program and cell-cycle
events. Finally, why is the temporal order of this transcriptional program important for execution of the yeast cell cycle
and to what degree is this program conserved among
eukaryotes? In the sections that follow, we will review the
literature regarding these issues with a focus on the emerging
models.
Evolution of Experimental Approaches and Models
Models of biological phenomena are necessarily constrained
by the approaches we use to observe them. Views of cellcycle-regulated transcription have certainly evolved as new
technologies and approaches have been applied.
The single-gene approach
Most early studies identified cell-cycle-regulated genes one
at a time. Typically, populations of cells were synchronized
by arresting with mating pheromone, released from the
arrest, and sampled over time. RNAs were isolated from the
time series and subjected to Northern blot analysis using
radioactively labeled probes from the gene of interest. Taken
together, these early studies revealed that many of the genes
identified by genetic methods as important cell-cycle regulators were themselves transcriptionally controlled during
the cell cycle. Cell-cycle regulators, like the cyclins, were
generally found to be transcribed in the phase in which they
functioned (Wittenberg et al. 1990; Surana et al. 1991;
Richardson et al. 1992; Kuhne and Linder 1993; Schwob
and Nasmyth 1993). These findings pointed to a model in
which the cell cycle is driven, at least in part, by successive
waves of expression of regulatory proteins (Amon et al.
1993).
Genomic approaches
In 1998, the first global view of the cell-cycle-regulated
transcription program was revealed when several groups
examined synchronized populations of cells in time-series
experiments using microarrays (Cho et al. 1998; Spellman
et al. 1998). To date, five studies (using different types of
microarrays and different means of synchronizing populations) have reported global transcriptional expression profiles in wild-type yeast cells as they progress through the cell
cycle (Cho et al. 1998; Spellman et al. 1998; Pramila et al.
2006; Orlando et al. 2008; Granovskaia et al. 2010). These
studies, and comparable studies in other systems (Rustici
et al. 2004; Oliva et al. 2005; Peng et al. 2005), delineated
Cell-Cycle-Regulated Transcription
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the scope of the problem, identifying several hundred genes
whose transcript levels oscillate during the cell cycle. The
precise number of periodic genes is still a somewhat contentious issue, ranging from 600 to 1500 or between 10 and
20% of all yeast genes (discussed further below). This fraction is higher than expected, given the number of known
cell-cycle regulators identified by genetics, and suggests that
precise regulation of a large fraction of the genome might
contribute to proper progression through the cell cycle.
The emerging picture from these studies also seemed to
suggest genes are not transcribed in a small number of
discrete intervals, but are instead turned on and off in
a continuum as cells passed through the cycle. Nonetheless,
the program could be divided into large blocks of genes
(clusters) found to be transcriptionally active in specific cellcycle phases. The lists of periodic genes enabled searches to
identify promoter motifs governing cell-cycle-regulated
transcription. This effort benefited greatly from the genome
sequences of related Saccharomyces species, which allowed
for comparisons that highlighted conserved promoter elements (Cherry et al. 2004; Teixeira et al. 2006). Bioinformatic analyses determined that genes within clusters shared
common promoter elements and were likely coregulated by
specific subsets of transcription factors (Spellman et al.
1998; Pramila et al. 2006; Orlando et al. 2008).
At the turn of the century, high-throughput chromatin
immunoprecipitation-microarray chip (ChIP-chip) technologies enabled researchers to collectively identify the binding
sites of all known transcription factors across the budding
yeast genome (Ren et al. 2000; Simon et al. 2001; Lee et al.
2002; Harbison et al. 2004). These ChIP-chip experiments,
like other early generation high-throughput technologies,
likely produced false positives and false negatives at relatively high rates. Thus, functional experimental evidence is
needed to confidently assign regulatory roles to specific
transcription factor/gene combinations. Nonetheless, these
studies provided physical evidence that clusters of genes
identified by microarray experiments were indeed coregulated by specific transcriptional regulators expressed in discrete cell-cycle phases. It also became clear that the genes
encoding transcriptional regulators were themselves located in these clusters (Simon et al. 2001). When the
results of genome-wide transcription factor localization
were combined with transcript dynamics from microarray
experiments, it became apparent that the cell-cycle transcriptional program could be regulated by a relatively small
network of serially activated transcription factors (Simon
et al. 2001; Lee et al. 2002; Pramila et al. 2006). Furthermore, this limited network of factors can function independently or combinatorially to enhance the complexity of the
pattern of transcriptional regulation (Kato et al. 2004).
The idea that a transcription factor network could control
the cell-cycle transcriptional program was a paradigm shift
in the field (discussed below), and this shift was made
possible only through the development and application of
genomic technologies.
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S. B. Haase and C. Wittenberg
A qualitatively similar view of the cell-cycle transcriptional program emerged in Schizosaccharomyces pombe
when microarray approaches were applied to synchronous
populations of fission yeast (Rustici et al. 2004; Oliva et al.
2005; Peng et al. 2005). Cell-cycle-regulated transcripts in
S. pombe are expressed in waves that can be subdivided into
clusters of temporally coregulated genes. Many recognizable
orthologs in Saccharomyces cerevisiae and in S. pombe exhibit
similar transcript periodicity, suggesting that transcriptional
regulatory mechanisms are conserved (Oliva et al. 2005;
Orlando et al. 2007). Despite the fact that many of the transcription factors that mediate cell-cycle-regulated transcription and some specific genes regulated by those factors are
conserved, there is considerable divergence among cellcycle-regulated genes and programs, even among closely
related yeasts (Eser et al. 2011; Guan et al. 2013). Furthermore, computational models indicate some altered
behaviors that likely reflect unique aspects of cell-cycle
progression in these highly diverged yeasts (Orlando et al.
2007). Comparison with other yeasts and more highly
diverged metazoan systems has led to the concept of conservation of the general topology, but not the specifics, of
the cell-cycle-regulated transcriptional circuitry (discussed
below).
Caveats associated with population-based approaches
Taken together, the single-gene and genomic approaches
have been phenomenally successful in providing a global
view of how transcript levels are regulated during cell-cycle
progression. However, it seems likely that some of the more
subtle aspects of transcriptional regulation remain hidden,
due to the nature of the experiments. The approaches
described above rely on synchronous populations of cells
to produce enough RNA to be interrogated by Northern blot,
microarray, qPCR, or RNA-Seq. Because populations are
used, measurements of mRNA levels necessarily represent
the average level across the population. In an ideal case, all
of the cells in the population would be in the exact same
cell-cycle phase, and thus the measured values represent an
average over cell-to-cell variance (biological noise). Although budding yeast is well known for the ease of
synchronization using a variety of methods (mating pheromone, conditional mutants in cell-cycle genes, elutriation),
synchronized populations are never fully synchronous, and
they lose synchrony over time. Loss of synchrony is
especially severe in budding yeast where the division is
asymmetric (Hartwell and Unger 1977; Lord and Wheals
1980, 1981; Woldringh et al. 1993; Bean et al. 2006;
Orlando et al. 2007). Thus, the measured values from
time-series experiments using synchronized populations represent a convolution of values from cells distributed across
cell-cycle phases. The result is that transcript oscillations
appear to dampen over time. Convolution due to loss of
synchrony can obscure the true amplitude of transcript oscillations as well as the precise timing of the appearance and
disappearance of a specific transcript.
Algorithmic approaches to population studies
To better identify more subtle aspects of the cell-cycleregulated transcriptional program, several groups set out to
use algorithmic approaches to deconvolve the measured
values of transcript levels from microarray experiments
(Bar-Joseph et al. 2004; Qiu et al. 2006; Rowicka et al.
2007; Guo et al. 2013). The goal of these approaches was
to better approximate the dynamic behavior of transcripts in
the average cell in the population. The groups made various
assumptions about the underlying models that drive synchrony loss in time-series experiments, and each reported
an increased performance in accurately inferring both the
timing and amplitude of transcript levels during the cell
cycle. Additionally, Guo et al. (2013) explicitly modeled
asymmetric division using a branching process model that
allowed them to infer differences in expression dynamics
between mother and daughter cells. Thus, they were able
to identify 82 genes that were expressed exclusively in
daughter cells (see below). Taken together, these algorithmic approaches permit a more accurate view of the timing of
transcript dynamics and suggest that the amplitude of transcriptional responses is, in many cases, substantially higher
than previously recognized.
Although these algorithmic approaches likely provide
a sharper view of transcript dynamics during the cell cycle,
the values they return are still estimates rather than true
measurements. Of course, the estimates are only as good as
the underlying models for population asynchrony. Additionally, the algorithms report values for the average cell in the
population and do not tell us much about the distribution of
behaviors across cells in the population.
Single-cell approaches
While there are computational methods to address the
issues associated with approaches using synchronized populations, arguably the most direct way to address the issues
is to measure transcript abundance in single cells. Ideally,
one would be able to monitor the abundance of a variety of
transcripts in living cells as they progress through the cell
cycle. However, current protocols are limited to measuring
only a few transcripts per experiment in fixed cells or to
using the expression of destabilized proteins as a proxy for
mRNA abundance in living cells.
Single-cell fluorescence in situ hybdrization (FISH) studies (Femino et al. 1998) can be used in yeast to measure the
levels of a specific transcript. Because cells are fixed during
the hybridization procedure, measurements of transcript
fluctuations over time can only be made on a population
of synchronized cells, and thus this approach would suffer
from some of the same issues related to population synchrony. However, single-cell FISH was used effectively to
demonstrate that individual yeast cells in an unsynchronized
culture of slow growing yeast cells exhibit metabolic cycling
of gene expression (Silverman et al. 2010). This approach,
examining the coincidence of expression of several gene
pairs, enabled investigators to establish positive correlation
between genes expressed in the same phase of the metabolic
oscillation and negative correlation between those expressed in opposite phases. Earlier studies relied on sampling synchronized populations of cells from a chemostat
for microarray analysis (Klevecz et al. 2004; Tu et al.
2005). This approach also worked well when applied to
cell-cycle-regulated genes by Silverman et al. (2010).
Single-molecule fluorescence techniques, which have been
applied successfully to monitor the initiation and elongation
rates, as well as the stability of cell-cycle-regulated mRNAs
in living cells, may be applicable in studies of the dynamics
of cell-cycle-regulated transcription (Larson et al. 2011;
Trcek et al. 2011).
The dynamics of periodic transcripts can also be inferred
by time-lapse microscopy in live cells using strains with cellcycle-regulated promoters fused to unstable variants of
green or red fluorescent proteins (GFP/RFP) (Skotheim
et al. 2008; Eser et al. 2011). Time-lapse microscopy on
single cells does not suffer from population effects, and in
cases where other markers are available, coherence between
the expression of a fluorescent protein and other events can
be determined. However, this approach does not directly
measure transcript levels, and the kinetics of the appearance
and disappearance of transcripts can only be estimated from
the accumulation of the GFP protein. Precise estimation is
blurred by variations in the maturation time of the GFP/
RFP, and in the protein half-life, especially for short-lived
transcripts. Furthermore, detection of GFP/RFP fluorescence
is insufficiently sensitive to be useful for all but the most
powerful promoters. Despite the low-throughput nature of
the approach, transcriptional reporters with better signal to
noise (e.g., luciferase) and advancements in high-resolution
time-lapse microscopy should expand its utility (Howell
et al. 2012).
Cell-Cycle-Regulated Gene Clusters: The Logic
and Strategy for Expression
Despite the observation that transcripts appear and disappear in a continuum throughout the cycle, the cell-cycle
transcription program has historically been viewed as four
major waves of gene expression that define the dominant
cell-cycle-regulated gene clusters including the G1/S cluster,
the S phase cluster, the G2/M cluster, and the M/G1 cluster,
based upon the coincidence of the timing of expression
relative to the easily discernible cell-cycle events (reviewed
by Bähler 2005; Wittenberg and Reed 2005; McInerny
2011). Despite their apparently coherent expression, the
genes within a specific cluster are often coordinately regulated by distinct transcription factors or mechanisms. In addition to the prominent clusters, there are smaller gene
clusters or single genes with distinct patterns of cell-cycleregulated expression sometimes mediated by multiple transcription factors acting in concert. Our goal in this section is
to discuss the major clusters and their modes of regulation.
Cell-Cycle-Regulated Transcription
69
The G1/S gene cluster
The G1/S gene cluster (also called simply the G1 gene
cluster) comprises the largest and, arguably, the bestcharacterized family of cell-cycle-regulated genes in yeast.
This family of upwards of 200 genes is expressed largely in
G1 phase cells coincident with START and prior to the
initiation of DNA replication (reviewed by Bähler 2005;
Wittenberg and Reed 2005; McInerny 2011). These genes
are controlled by two heterodimeric transcription factors,
SCB-binding factor (SBF) and MCB-binding factor (MBF),
which bind to distinct but related DNA sequence motifs in
the promoters of their target genes (Figure 2). SBF binds to
SCB elements (SBF cell-cycle box) and MBF to MCB elements (MBF cell-cycle box), respectively (Lee et al. 2002;
Teixeira et al. 2006). The distinct DNA-binding properties of
those factors is attributable to their DNA-binding proteins,
Swi4 for SBF and Mbp1 for MBF. Both factors bind DNA via
a classical winged helix-turn-helix DNA-binding fold and
associate with the common subunit Swi6. Although both
DNA-binding proteins and Swi6 share a common domain
structure, Swi4 and Mbp1 bind distinct DNA sequences,
whereas Swi6 does not bind directly to DNA (McIntosh
et al. 2000; Xu et al. 1997; Nair et al. 2003; Taylor et al.
1997, 2000, 2010).
SBF and MBF bind and regulate partially overlapping
subsets of genes in the G1/S cluster because of the presence
of both consensus sequence motifs in some of the promoters
(Spellman et al. 1998; Iyer et al. 2001; Simon et al. 2001;
Teixeira et al. 2006; Ferrezuelo et al. 2010). The extent to
which the overlap is reflected in coordinate or differential
regulation is only beginning to be understood (see below)
(Bean et al. 2005; Eser et al. 2011; de Oliveira et al. 2012;
Smolka et al. 2012; Travesa and Wittenberg 2012; Travesa
et al. 2012). Finally, many potential and actual binding sites
have been identified in promoters that do not appear to
promote cell-cycle periodicity but may, in some cases, have
functional consequences (Iyer et al. 2001; Horak et al. 2002;
Bean et al. 2005; also see Haber 2012; Eriksson et al. 2012).
The G1/S gene cluster is rich in genes involved in
processes associated with progression from G1 into S phase.
Passage through START is particularly important for initiating the many events required for the new cell division cycle.
Those include dramatic changes in morphogenesis leading
to the formation of the bud, the nascent daughter cell with
its associated septin ring marking the future site of cytokinesis (reviewed in Bi and Park 2012; Howell and Lew
2012). Additionally, the duplication of the spindle pole body,
the first structures associated with assembly of the mitotic
spindle (reviewed by Winey and Bloom 2012), and the initiation of DNA replication are triggered following START
(reviewed by Remus and Diffley 2009). Although the separation of function is by no means absolute, in general, many
genes involved in morphogenesis, including enzymes required for cell wall biosynthesis and septin ring components,
are regulated by SBF, whereas genes for DNA replication
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S. B. Haase and C. Wittenberg
and repair, including nucleotide biosynthetic enzymes and
proteins acting at the replication fork, are regulated by MBF.
The rationale for segregating SBF and MBF targets based
upon function has, until recently, lacked explanation because
the two transcription factors are coordinately regulated during
the cell cycle. However, it has become apparent that segregation of targets into two groups can have dramatic consequences under specific physiological conditions (see below)
(Doncic et al. 2011; Eser et al. 2011; Smolka et al. 2012;
Travesa et al. 2012).
The hallmark of these two transcriptional regulators is
their capacity to abruptly activate their target genes during
G1 phase in response to the activation of G1 cyclinassociated Cdk1 (Figure 2) (Costanzo et al. 2004; Di Talia
et al. 2007; Taberner et al. 2009). That regulation is disrupted by cdc28-ts mutants (Marini and Reed 1992). We now
understand that the activation of G1/S genes is a consequence of the regulated accumulation of the G1 cyclin
Cln3 (Tyers et al. 1993; Dirick et al. 1995; Stuart and
Wittenberg 1995; Garí et al. 2001). The accumulation, activation, and localization of Cln3/CDK are critical determinants of the timing of START and are responsive to cell
growth and cell size (Nash et al. 1988; Tyers et al. 1993;
Baroni et al. 1994; Polymenis and Schmidt 1997; Miller and
Cross 2000; Edgington and Futcher 2001; Jorgensen et al.
2002; Di Talia et al. 2007; Taberner et al. 2009; Ferrezuelo
et al. 2012; Thorburn et al. 2013). Cln3, once accumulated
to a critical level in the nucleus, activates Cdk1, leading to
the phosphorylation of the SBF-bound transcriptional repressor, Whi5 (Costanzo et al. 2004; de Bruin et al. 2004;
Schaefer and Breeden 2004). Upon phosphorylation, Whi5,
which binds to the carboxy terminus of Swi6 through its
GTB-containing domain, dissociates from promoter-bound
SBF and is relocalized to the cytoplasm (Figure 2) (Costanzo
et al. 2004; de Bruin et al. 2004; Di Talia et al. 2007;
Skotheim et al. 2008; Travesa et al. 2013). Phosphorylation
of Swi6 by Cln/CDK may also contribute to the regulation of
Whi5 binding (Sidorova et al. 1995; Costanzo et al. 2004;
Wagner et al. 2009). Dissociation and nuclear export of
Whi5, in turn, leads to the activation of SBF and the expression of its target genes. Although Cln3/CDK-dependent
Whi5 phosphorylation appears to promote both dissociation
from promoters and export from the nucleus, those two
phenomena have not been separated in terms of their effect
on the activation of G1/S transcription (Costanzo et al.
2004; de Bruin et al. 2004; Di Talia et al. 2007; Skotheim
et al. 2008; Travesa et al. 2013). A recent study has established that release of the Whi5 repressor is the critical determinant of the event historically referred to as START by
Hartwell and colleagues (Doncic et al. 2011; Eser et al.
2011).
Activation of G1/S transcription factors
Our current understanding of SBF derives from its early
identification (by the Nasmyth and Herskowitz laboratories)
among the genes required for expression of the HO gene,
Figure 2 Transcriptional regulation of the G1/S gene cluster. The regulatory circuitry for the G1/S gene cluster is depicted. Two transcriptional
regulators, SBF and MBF, bind to SCB and MCB elements, respectively, in
the repressed promoters of G1/S genes as cells enter G1 phase. SBF
represses its targets due to the binding of Whi5. MBF appears to act, in
part, as a repressor, during early G1 phase. Once accumulated to appropriate levels, Cln3/Cdk1 phosphorylates Whi5, leading to Whi5 dissociation from SBF and activation of the G1/S gene promoters. Cln3/Cdk1 also
activates MBF-regulated promoters via an unknown mechanism. Activation of the CLN1 and CLN2 promoters by SBF leads to activation of the
Cln1/Cdk1 and Cln2/Cdk1, both of which feed back to further activate
SBF-regulated promoters, as well as activating MBF-regulated promoters.
The activation of MBF-regulated promoters leads to the accumulation of
Nrm1, a specific repressor of MBF that, when accumulated sufficiently,
feeds back to bind MBF, thereby repressing MBF targets. MBF also activates Swi4, thereby increasing the abundance and activity of SBF. Activation of SBF targets promotes cell-cycle progression, leading ultimately
to the accumulation of Clb2/Cdk1, which binds and phosphorylates Swi4,
leading to its dissociation from SBF-regulated promoters. Clb2/Cdk1 can
also promote repression of MBF-regulated promoters via an unknown
mechanism. The color of subunits and arrows is as in Figure 1.
which encodes the endonuclease that mediates switching of
mating types via DNA rearrangements at the MAT locus
(Breeden and Nasmyth 1987; Andrews and Herskowitz
1989; see Haber 2012). The known targets of SBF, which
now number 100, include those encoding the G1 cyclins,
Cln1 and Cln2, which, like Cln3, promote Cdk1-dependent
activation of G1/S transcription (Cross 1988; Wittenberg
et al. 1990; Nasmyth and Dirick 1991; Tyers et al. 1993;
Wijnen et al. 2002). Although earlier studies supported
the autonomy of Cln3/Cdk1 in the timely activation of
G1/S genes (Dirick et al. 1995; Stuart and Wittenberg
1995), the application of single-cell techniques to the analysis of START led to the realization that the rapid accumulation of Cln1 and Cln2, and their potent activation of Cdk1,
leads to a dramatic acceleration of nuclear export of Whi5.
This positive feedback greatly enhances the coherency of
activation of G1/S genes (Skotheim et al. 2008), confirming
a model advanced many years prior (Cross and Tinkelenberg 1991; Dirick and Nasmyth 1991). Thus, the combined
activity of the G1 cyclin-associated forms of Cdk1 is responsible for the switch-like behavior and irreversibility of the
START event.
The expression of MBF targets, like those regulated by
SBF, occurs during late G1 phase and depends upon Cln3/
Cdk1 activity (Tyers et al. 1993; Dirick et al. 1995; Stuart
and Wittenberg 1995; Wittenberg and Reed 2005). MBF
targets include genes encoding DNA replication and repair
proteins and the cyclin Clb5, which is critical for timely
activation of DNA replication. Interestingly, the SWI4 gene
was recently shown to be a target of MBF, which is sufficient
for much of its expression during G1 phase (Harris et al.
2013), suggesting there is crosstalk between the two major
G1/S transcription factors. The mechanism of activation of
MBF-regulated genes remains to be established. We do
know that, unlike SBF, MBF acts, in part, as a repressor of
its target genes outside of G1 phase, suggesting that Cdk1
acts to relieve its repressive activity (de Bruin et al. 2006).
However, as of yet, no protein playing an equivalent role to
Whi5 has been identified. This has led to the prevailing
hypothesis that the direct phosphorylation of the transcription factor by Cdk1 is the critical event in transcriptional
activation. Although both Swi6 and Mbp1 are known to be
substrates of Cdk1 in vitro and are phosphoproteins in vivo,
there is currently no evidence indicating that those phosphorylation events are required for activation of MBF targets
(Sidorova et al. 1995; Siegmund and Nasmyth 1996; Ubersax
et al. 2003; Geymonat et al. 2004; Holt et al. 2008).
Repression of G1/S transcription factors
Like SBF targets, MBF-regulated genes play important roles
in the circuitry regulating G1/S transcription (Figure 2).
One of its targets is the gene encoding the MBF-specific
transcriptional repressor, Nrm1 (de Bruin et al. 2006). Unlike Whi5, which represses transcription from M phase
through late G1 phase, Nrm1 acts only after G1/S transcription is strongly activated, leading to Nrm1 accumulation.
Nrm1 binds to the carboxy terminus of Swi6 at MBF target
promoters via interaction between its GTB-containing domain, a motif conserved with Whi5 and its fungal orthologs
(Ofir et al. 2012; Travesa et al. 2013). This leads to the
reestablishment of MBF as a transcriptional repressor such
that expression of MBF target genes, including that of NRM1
itself, is terminated by negative feedback. Subsequently, as
cells exit mitosis, the Nrm1 protein is targeted for degradation by the APCCdh1 ubiquitin ligase, preparing cells to reactivate MBF during the subsequent G1 phase (Ostapenko and
Solomon 2011). In addition to Nrm1, MBF also promotes
the expression of two B-type cyclins, Clb5 and Clb6.
Whereas Clb5 ultimately activates Cdk1 to drive entry into
S phase, Clb6/Cdk1 has been reported to phosphorylate
Swi6, promoting its accumulation in the cytoplasm (Geymonat
et al. 2004). It has been suggested that Swi6 is constitutively cycling between the cytoplasm and the nucleus such
that the phosphorylation-dependent inhibition of nuclear
uptake results in accumulation in the cytoplasm that is then
reversed upon dephosphorylation subsequent to mitosis by
the Cdc14 phosphatase (Queralt and Igual 2003; Geymonat
et al. 2004). Interestingly, cycling of Swi6 between the cytoplasm and nucleus appears to be important for SBFdependent transcriptional activity, although the molecular
basis for that requirement has not been established (Queralt
Cell-Cycle-Regulated Transcription
71
and Igual 2003). Like Swi6, nuclear localization of Whi5 is
promoted by Cdc14-dependent dephosphorylation upon M
phase exit. The capacity of Whi5 to relocalize to the nucleus
if CDK activity is reduced during other cell cycle phases
suggests that phosphatases other than Cdc14 must also be
competent to dephosphorylate it (Charvin et al. 2010). Nuclear exclusion of Whi5 is also promoted by CDK-dependent
phosphorylation, albeit earlier in the cell cycle than Swi6
(Costanzo et al. 2004; Di Talia et al. 2007; Taberner et al.
2009).
The repression of SBF does not seem to act through
Nrm1-like negative feedback. The primary mechanism terminating expression of SBF targets is the accumulation of
the B-type cyclin, Clb2 (Figure 2). Clb2/Cdk1 binds stably to
Swi4 through its conserved ankyrin repeat and phosphorylates Swi4, leading to its dissociation from DNA and export
to the cytoplasm, terminating SBF-dependent transcription
(Amon et al. 1993; Koch et al. 1993; Siegmund and Nasmyth
1996). SBF-dependent expression of the genes encoding the
Yox1 and Yhp1 transcriptional repressors may also contribute, albeit modestly, to the repression of SBF by repressing
SWI4 expression (Pramila 2002). Although a similar mechanism has not been demonstrated for MBF, it is clear that in
the absence of Nrm1, MBF can also be inactivated by a mechanism that depends upon B cyclin-Cdk1 (de Bruin et al. 2006).
proach that increases the temporal resolution of microarray
experiments (Guo et al. 2013). These findings suggests that
cells can adapt their sequential program of gene activation
in response to environmental or physiological conditions,
perhaps resulting in increased fitness.
Temporal regulation within the G1/S gene cluster
The S phase gene cluster
The application of algorithmic approaches to microarray
analysis of the cell-cycle transcriptome has revealed an
ordered timing in the activation of G1/S genes that had
previously appeared to be expressed simultaneously (Eser
et al. 2011; Guo et al. 2013). Although some hints of temporal resolution of gene expression within that cluster had
been observed in prior analyses, the lack of reproducibility
and the problems associated with differing amplitudes of
expression had obscured those differences. Using highthroughput analysis of multiple array experiments coupled
with single-cell fluorescence analyses, Eser and colleagues
found that CLN1 and CLN2 were among the earliest
expressed genes of the G1/S cluster, leading to the concept
of “feedback first,” wherein the first targets to be expressed
are those that promote further transcriptional activation of
G1/S targets. That mechanism ensures the irreversibility of
the process and is generally conserved in both closely related yeast species and in systems as divergent as humans.
Their findings also revealed that NRM1, encoding the repressor of MBF-regulated transcription, is among the last
genes of the cluster to be expressed, thereby ensuring that
MBF targets are not repressed before they are adequately
expressed. Different synchronization regimens can cause sequential waves of MBF-then-SBF or SBF-then-MBF activity
such that genes containing both SBF- and MBF-binding sites
are activated by whichever of the MBF or SBF factors is
activated first and inactivated by whichever transcription
factor is repressed first (Eser et al. 2011). Similar results
were obtained using a mathematical deconvolution ap-
The G1/S transition, characterized by the initiation of DNA
replication, is accompanied by the activation of two clusters
of S phase genes. One cluster is composed of the genes
encoding the histones, which make up the nucleosomes that
package newly replicated DNA. Histone genes are regulated
by an, as yet, poorly understood transcriptional regulatory
mechanism involving SBF, Spt10, histone chaperones, and
other factors. That S phase cluster has been recently
reviewed by Eriksson et al. (2012) and will not be discussed
further here. The second, much larger, cluster of genes
expressed during S phase is that regulated by Hcm1 (Figure
3). Although less thoroughly studied than transcription factors controlling the other major bursts of cell-cycle-regulated
gene expression, Hcm1 plays a central role in that regulatory
circuitry. It is one of four members of the forkhead family of
transcription factors found in budding yeast (Hcm1, Fkh1,
Fkh2, and Fhl1 (Forkhead-like) (reviewed in Murakami
et al. 2010). Like the other members of that family, which
include mammalian Fox transcription factors, Hcm1 binds
DNA directly via a winged helix DNA-binding motif. The
presence of putative CDK phosphorylation sites in Hcm1
and its capacity to be phosphorylated by Clb/Cdk1 in vitro
make it a likely target of the Cdk1 protein kinase (Ubersax
et al. 2003).
The Hcm1 transcription factor appears to regulate 180
genes based upon the timing of their expression and the
presence of the Hcm1-binding site in their promoters
(Pramila et al. 2006). However, only a small fraction of those
genes were found to bind Hcm1 by genome-wide chromatin
72
S. B. Haase and C. Wittenberg
Coupling of G1/S transcription to the rest of the
transcriptional program
The targets of the G1/S transcription factors are a large and
diverse set of genes involved in a broad spectrum of events,
many of which are initiated during the interval of G1/S gene
expression. Of particular interest, in the context of understanding cell-cycle-regulated gene expression, are genes
that act to limit G1/S transcription and those that induce
subsequent waves of transcription. Nrm1 is perhaps the best
example of a G1/S target that acts to limit G1/S transcription. Clb2 similarly limits SBF-regulated transcription, but
it is not itself a G1/S target (see below). Hcm1 is the transcriptional activator of a large group of S phase genes (discussed below), and is expressed during G1/S from a
promoter that is bound by both SBF and MBF (Iyer et al.
2001; Pramila 2002). Thus, Hcm1 targets are activated as
a consequence of the burst of G1/S transcription, thereby,
participating as a link in the cell-cycle-transcriptional circuit
(Figure 1).
Figure 3 Transcriptional regulation of the S phase and
G2/M gene clusters. The regulatory circuitry for the S
phase gene cluster (top) and the G2/M gene cluster
(bottom) is depicted. The Hcm1 transcription factor is
the major activator of S phase genes. It is uncertain
whether additional components are required to promote gene expression and little is known about the
activation or repression mechanisms. The G2/M gene
cluster is regulated by Mcm1 in complex with Fkh2,
which act as a repressor during S phase. Ndd1 binds
to Fkh2, and activates G2/M genes including those
encoding Clb2 and Cdc5. Clb2 and Cdc5 kinases act
via positive feedback to phosphorylate Ndd1 to promote binding to Fkh2 and transcriptional activation.
Clb2/Cdk1 activates the APCCdh1 ubiquitin ligase,
which targets Ndd1 for degradation, terminating the
expression of the G2/M gene cluster as cells complete
M phase. The color of subunits and arrows is as in
Figure 1.
immunoprecipitation (Horak et al. 2002), likely a consequence of the low reliability of the early studies using that
technique. Among the targets are 10 genes involved in
budding and morphogenesis and nearly 100 involved in
some aspect of chromosome segregation. Unsurprisingly,
these include protein components of chromosomes and elements of the chromosome segregation machinery, much of
which is assembled during S and G2 phases. However, a role
in the regulation of genes involved in budding was unexpected because bud morphogenesis is a well-established
function of many genes in the G1/S gene cluster. Nevertheless, under most circumstances, budding occurs coincident
with initiation of DNA replication so expression during S
phase under the control of Hcm1 is timely. That said,
Hcm1 is dispensable for budding but is required for the
proper timing and execution of mitosis (Pramila et al.
2006). The synthetic lethality of hcm1 mutations with mutations in a number of genes involved in chromosome segregation is consistent with its importance in the orchestration
of mitotic functions.
The relatively recent discovery of Hcm1 makes it something of a “missing link” in the transcriptional circuitry of the
cell cycle. Its targets include regulators of all of the other
major bursts of cell-cycle-regulated gene expression: the
transcriptional repressors Whi5 (G1/S transcription) and
Yhp1 (M/G1 transcription) as well as Fkh1, Fkh2, and
Ndd1, the central regulators of the G2/M gene cluster (see
below). Although these factors do not play a direct role in
limiting G1/S transcription via feedback, Hcm1 indirectly
activates Clb2, the feedback repressor of SBF genes, via
its role in the expression of Fkh2 and Ndd1 activators of
the G2/M gene cluster (Figure 1). Despite the central role
of Hcm1 in the regulation of genes involved in cell-cycleregulated transcription, hcm1 mutants are viable and retain
detectable periodicity of S phase gene expression (Pramila
et al. 2006; Simmons Kovacs et al. 2012). This suggests the
involvement of additional, as yet unknown, transcription
factors in the regulation of this gene family.
The G2/M gene cluster
Progression from S phase into G2 phase is accompanied by
activation of a family of 35 genes falling into the G2/M
cluster, also called the CLB2 cluster (Cho et al. 1998; Spellman
et al. 1998). As the latter name implies, this cluster
includes the genes encoding the B-type cyclin Clb2, along
with its paralog Clb1 and other important cell cycle regulatory factors including Cdc5, the yeast polo-like kinase,
Cdc20, a specificity subunit of the APC ubiquitin ligase,
and the Ace2 and Swi5 transcription factors. The primary
regulator of this gene cluster is the MADS box transcription factor Mcm1, in conjunction with the forkhead family
member Fkh2 and the coactivator Ndd1, both members of
the S phase gene cluster regulated by Hcm1 (Figure 3)
(Loy et al. 1999; Kumar et al. 2000).
Mcm1 plays diverse roles that derive from its capacity to
multimerize with a diverse set of coregulators. This allows it
to regulate distinct gene families including the M/G1 cluster
genes (see below), the MAT locus genes (Tuch et al. 2008),
and even to play a role in mating-type switching, which is
apparently transcription independent (Coïc et al. 2006).
The collaboration of Mcm1 with Fkh2 in the context of transcriptional regulation of the G2/M cluster is, in part, a
consequence of the occurrence of adjacent Mcm1- and
Fkh2-binding sites in G2/M cluster promoters (Lydall et al.
1991; Boros et al. 2003). The association of that heterodimer with the adjacent binding sites on those genes creates
a platform for association of the coactivator Ndd1 to activate
gene expression. Whereas other forkhead family members
possess the capacity to bind to the DNA via their conserved
winged helix motif, only Fkh2 can also associate directly
with Mcm1 and thereby exert its effect on gene expression
in that context. Interestingly, this binding interface has been
conserved sufficiently for Fkh2 to bind to the human serum
response factor (SRF), the homolog of Mcm1 (Boros et al.
2003). Fkh1, which lacks the Mcm1 interaction domain of
Fkh2, binds to the same DNA motif as Fkh2 at CLB2 cluster
Cell-Cycle-Regulated Transcription
73
promoters, but only in the absence of Mcm1 (Koranda et al.
2000; Hollenhorst et al. 2001).
The general view of regulation of the CLB2 cluster posits
that Mcm1 and Fkh2 remain bound to DNA, and to each
other, throughout the cell cycle. They repress transcription
during G1 and S phases, but subsequent recruitment of
Ndd1 activates those same genes as cells progress through
G2 and into M phase. FKH2 and NDD1 are coexpressed as
components of the S phase gene cluster. Ndd1 is destroyed
during late M phase, presumably as a consequence of ubiquitylation by the APC (Loy et al. 1999). In contrast, Fkh2 is
stable, persisting throughout the cell cycle (Koranda et al.
2000). Although association of Ndd1 with Fkh2 is the limiting event for activation of G2/M cluster expression, its
accumulation alone is reported to be insufficient for association and transcriptional activation (Reynolds et al. 2003).
Phosphorylation of Ndd1 by both Clb2/Cdk1 and Cdc5 pololike kinase appears necessary for efficient recruitment to the
Fkh2 FHA domain and robust activation of Clb2 cluster
genes (Reynolds et al. 2003; Pic-Taylor et al. 2004; Darieva
et al. 2006, 2012).
The requirement for Ndd1 phosphorylation constitutes
a positive feedback loop, wherein both Clb2 and Cdc5, when
expressed as G2/M cluster genes, feedback and activate
their own gene expression. This posed a problem because,
if these two protein kinases are required for activation of
their own expression, it is unclear how they would be activated to initiate the feedback loop. This is particularly vexing in the case of Cdc5, which, unlike Clb2, has no paralogs
in yeast and which appears to be the only kinase that phosphorylates S85 of Ndd1 (Darieva et al. 2006). However, this
problem was recently addressed by the report that at least
a subset of the Clb2 cluster genes (including CLB2) can be
partially activated, even in the complete absence of B-type
cyclins (Orlando et al. 2008). This finding suggests a mechanism by which the feedback loop can be activated in the
absence of phosphorylation by Clb2.
In addition, phosphorylation of Ndd1 by Pkc1 (protein
kinase C) has recently been shown to delay its association
with Mcm1/Fkh2 and, thereby, prevent activation of Clb2
cluster genes by Ndd1 until conditions are appropriate
(Darieva et al. 2012). Precisely what conditions are monitored via Pkc1 is unclear but it seems likely that it is related
to cell wall synthesis or stress (reviewed in Levin 2011).
Finally, it is believed that upon destruction of Ndd1 late in
M phase, Mcm1/Fkh2-bound CLB2 cluster promoters are
returned to their repressed state, prepared to initiate
a new cell cycle (Loy et al. 1999). It remains unclear
whether Ndd1 destruction alone is sufficient for repression
of CLB2 cluster genes or whether that repression also
involves a specific transcriptional repressor that acts either
via negative feedback or some other mechanism.
This view of regulation of G2/M cluster genes, although
sufficient to explain expression of CLB2 and other members,
does not explain the regulation of the entire gene cluster.
In fact, some genes within the cluster exhibit different
74
S. B. Haase and C. Wittenberg
behaviors in the mitotic cyclin mutants, suggesting that more
than one regulatory mechanism exists for triggering transcription (Orlando et al. 2008). It has recently been recognized
that other members of the family, including SPO12, despite
being Mcm1/Fkh2 targets, are subject to an additional level
of regulation by the homeodomain-containing repressor protein, Yox1 (Pramila 2002; Darieva et al. 2010). Yox1 was
previously recognized as an important repressor in the regulation of the M/G1 gene cluster (see below). However, in
addition to having a consensus DNA-binding site for Fkh2
downstream of that for Mcm1, some G2/M genes have
a Yox1-binding site upstream, suggesting that Yox1 also plays
a role in their transcriptional regulation (Darieva et al. 2010).
In fact, occupancy of that Yox1-binding site precludes the
binding of Fkh2. Consequently, Yox1 represses those genes
having a Yox1-binding site in a manner similar to its effect on
members of the M/G1 gene cluster. The basis for the mutual
exclusivity of binding sites for Yox1 and Fkh2 remains unclear
but presumably involves competition between Yox1 and Fkh2
for binding sites on Mcm1 and not the geometry of the DNAbinding sites themselves (Boros et al. 2003; Darieva et al.
2010). Regardless, this regulatory circuit is clearly effective
in repressing gene expression at those loci and shifting the
expression of those genes later in the cell cycle relative to the
other members of the G2/M gene cluster (Pramila 2002).
Importantly, this regulation does not affect Clb2 or Cdc5,
which act early during the transcriptional burst to promote
activation via positive feedback. This is similar to the
“feedback first” phenomenon observed earlier in the cycle,
wherein Cln1 and Cln2, encoded by the earliest expressed
genes of the G1/S cluster, feedback to promote SBF- and
MBF-regulated transcription.
The M/G1 gene cluster
As cells exit M phase and transition into G1 phase of a new
cell cycle, cells activate four families of genes, distinguishable based upon either the function of their products or the
transcription factors that regulate them. Together they
comprise the M/G1 gene cluster. That cluster includes genes
encoding the cyclin Cln3 and the Swi4 transcription factor,
two proteins responsible for promoting G1/S transcription
(SWI4 is also activated by MBF; see above). Components of
the prereplication complex that prepare cells to initiate DNA
replication during S phase, elements of the mating response
pathway, and genes of the PHO regulon that are required for
mobilization of phosphate are also members of that cluster
(Haber 2012; Ljungdahl and Daignan-Fornier 2012).
The Mcm1 gene cluster
The largest family of genes in the M/G1 gene cluster, like the
G2/M cluster, is regulated by Mcm1 but, in this case, without the involvement of Ndd1 (Zhu et al. 2000). Mcm1 is
bound to the promoters of target genes, including Cln3,
Swi4, and seven members of the prereplicative complex
(Cdc6 and Mcm2–7) (Spellman et al. 1998), via an element
referred to as an early cell-cycle box (ECB) (McInerny et al.
Figure 4 Transcriptional regulation of the M/G1 gene
clusters. The regulatory circuitry of the M/G1 gene
cluster is depicted, including the Mcm1 cluster (top)
and the Sic1 cluster (bottom). Mcm1 binds the
Mcm1 cluster promoters along with the repressors
Yox1 and Yhp1, repressing those genes prior to M
phase. As cells exit M phase, Yox1 and Yhp1 are targeted for degradation by the APCCdh1 ubiquitin ligase
and Mcm1 targets are activated. Expression of those
two repressors as products of G1/S genes restores repression of Mcm1 cluster genes as cells exit G1 phase.
Sic1 cluster genes are regulated by the Swi5 and Ace2
transcription factors, which share the same binding
sequence in target gene promoters. Swi5 and Ace2
are both excluded from the nucleus when Clb/Cdk1
is active. As cells progress through mitosis, Clb proteins
are degraded and the Cdc14 phosphatase is activated,
promoting accumulation of the transcription factors in
nuclei. Whereas Swi5 accumulates in both mother and
daughter nuclei, Ace2 accumulates only in daughter
nuclei. One class of genes characterized by SIC1 is
expressed in both mothers and daughters. At those
genes, either Swi5 or Ace2 can bind and activate. Another class of genes, characterized by CTS1, expressed
only in daughters, can bind both Swi5 and Ace2, but
Swi5 binds at those promoters along with Fkh1 or
Fkh2, which act as “antiactivators.” Consequently, in mother cells, the binding of Swi5 fails to activate transcription. In daughter nuclei, the same
promoters are activated by Ace2, despite the presence of Swi5. Whether Swi5/Fkh protein complexes also bind to those promoters but fail to activate is
not known. Swi5-specific and daughter-specific genes are not depicted. Finally, all of these genes are inactivated as cells pass START as a consequence
of the nuclear degradation of Swi5 and the nuclear export of Ace2. The color of subunits and arrows is as in Figure 1.
1997; MacKay et al. 2001). Mcm1 is regulated by one of two
transcriptional repressors, the homeodomain proteins Yox1
or Yhp1, that hold expression of this cluster of genes in
check outside of the M/G1 interval by binding directly to
Mcm1 and a DNA element that flanks the ECB (Mai et al.
2002; Pramila 2002) (Figure 4). Their interaction with both
Mcm1 and DNA is required to stabilize the repressive complex (Darieva et al. 2010). Consistent with the need to repress M/G1 cluster expression from G1 through M phase,
YOX1 is expressed as a G1/S cluster gene during G1 and
YHP1 somewhat later, perhaps as a consequence of the presence of Fkh2 sites in its promoter (Spellman et al. 1998;
Pramila 2002). Expression of Mcm1 target genes does not
occur until the Yox1 and Yhp1 repressor proteins are eliminated, presumably by APC-dependent ubiquitylation, during late M phase (Pramila 2002). Whereas Yox1 appears to
repress gene expression at some G2/M cluster promoters by
displacing Fkh2-Ndd1, no similar mechanism is known for
M/G1 cluster genes (Darieva et al. 2010). Nevertheless,
renewed accumulation of Yox1 and Yhp1 at the beginning
of the next cell cycle certainly contributes to M/G1 gene
repression.
The Sic1 gene cluster
The second family of M/G1 cluster genes, often referred to
as the Sic1 cluster, is regulated by the paralogous transcription factors Ace2 and Swi5 (Dohrmann et al. 1992, 1996;
McBride et al. 1999; Laabs et al. 2003) (Figure 4). These
factors bind identical DNA sequences via zinc finger domains
in vitro but regulate groups of genes in vivo that are only
partially overlapping. CTS1 is a classical Ace2-regulated
gene encoding chitinase, which is expressed only in daughter cells and mediates cell separation following mitosis
(reviewed in Bi and Park 2012). In contrast, the best-studied
Swi5 target, HO, is expressed only in mother cells (reviewed
in Haber 2012). Both factors participate in the activation of
SIC1, encoding a Clb-specific CDK inhibitor for which the
cluster is named. Sic1 is expressed very late in M phase,
where it contributes to the inhibition of Clb/CDK activity
during exit from mitosis. Recent analyses suggest that the
SIC1 transcription occurs primarily in daughter cells during
G1 phase (Guo et al. 2013), where it prevents premature
initiation of DNA replication (reviewed in Enserink and
Kolodner 2010). Consequently, regulating the activity of
Swi5 and Ace2 and their partitioning between mother and
daughter cells is critical for proper cell-cycle regulation.
The differential regulation of Sic1 cluster genes by paralogous transcription factors that not only bind to the same
DNA-binding site in vitro but, in many cases, bind to the
same genes in vivo poses a conundrum (Dohrmann et al.
1992, 1996). By evaluating the requirement for Ace2 and
Swi5 for expression of M/G1 genes and correlating that with
the binding of Ace2 and Swi5 to the promoters of those
genes, it became clear that the promoters of genes regulated
solely by Swi5 are bound only by Swi5, whereas those regulated by both factors or by Ace2 alone bind both factors
(Voth et al. 2007). Why, then, are some genes that bind both
factors only dependent upon Ace2 for expression, whereas
Cell-Cycle-Regulated Transcription
75
others can be activated by either one? This conundrum is, in
part, resolved by the finding that promoters regulated only
by Ace2 bind Swi5 along with Fkh1 and Fkh2, which act as
“antiactivators” for Swi5 at those promoters (Figure 4). In
contrast, genes that are activated by both factors or only by
Swi5, including SIC1, CDC6, and HO, lack the Fkh1/2 binding site. Consequently, it appears that Swi5 binds to the
promoters of all Sic1 cluster genes but only activates those
at which Fkh1 or Fkh2 is not bound. Whether the Swi5/
Fkh1/2 binds but fails to activate the same promoters in
daughters, as well as in mothers, is unclear.
Another aspect of the differential regulation of genes by
these transcription factors involves the asymmetric distribution of Ace2 to daughter nuclei following mitosis. That
asymmetry is a consequence of the failure to retain Ace2
in mother cell nuclei following mitosis (Colman-Lerner
et al. 2001; Weiss et al. 2002; Mazanka and Weiss 2010).
Daughter-specific retention of Ace2 is regulated, at least in
part, by the NDR/LATS family protein kinase Cbk1, a component of the so-called RAM complex (regulation of Ace2
and morphogenesis). A deconvolution algorithm developed
by Guo and colleagues revealed the existence of a large
group of genes subject to the daughter-specific transcription
program (Guo et al. 2013). In addition to those genes that
had been reported previously (Colman-Lerner et al. 2001; Di
Talia et al. 2009), the high temporal resolution afforded by
the deconvolution approach led to the identification of at
least four temporally distinct waves of daughter-specific
transcripts and identified several new daughter-specific
genes. Transcription factor enrichment analyses suggest
the earliest waves are regulated by Swi5 and Ace2, whereas
the later waves appear to be regulated by Mcm1. These
analyses also indicate a role for several other transcription
factors in that regulation, including Sok2, Phd1, Ste12,
Cin5, Yap6, and Tec1 (Guo et al. 2013).
One consequence of asymmetry is that chitinase, encoded
by the Ace2 target CTS1, is daughter specific. This pattern of
expression is responsible for the characteristic retention of
the bud scar on mother cells, but not daughter cells, subsequent to cytokinesis. Although Ace2 and Swi5 can both
bind to the CTS1 promoter, when Swi5 is bound, it fails to
activate due to the binding of Fkh1/2 (Voth et al. 2007)
(Figure 4). Only when Ace2 is bound, can it activate
CTS1. Another consequence of the daughter-specific localization of Ace2 is the characteristic delay in budding in
daughter cells relative to mother cells. This is, in part, a consequence of the Ace2-dependent delay in CLN3 expression
and, thereby, delayed activation of G1/S transcription in
daughter cells (Laabs et al. 2003; Di Talia et al. 2009). Although this impacts the range of cell size at which daughter
cells bud under various nutrient conditions, it does not explain the phenomenon of cell size control (reviewed by
Turner et al. 2012).
Finally, the Swi5 transcription factor exhibits specificity
for binding to some cell-cycle-regulated promoters (e.g.,
CDC6 and HO), a property that is apparently not shared
76
S. B. Haase and C. Wittenberg
by Ace2 (Voth et al. 2007). Swi5 binding to the HO promoter, which occurs late in M phase, licenses that promoter
for activation only in mother cells during the subsequent G1
phase. Similar activation does not occur in daughter cells
due to the daughter-specific localization of the transcriptional repressor Ash1, which prevents Swi5 binding (Bobola
et al. 1996; Sil and Herskowitz 1996; Cosma 2004; Shen
et al. 2009). Thus, the selective binding of Swi5 at promoters in the absence of the Fkh1 and Fkh2 transcription
factors defines genes that are uniquely regulated by Swi5.
Clb/CDK dependent phosphorylation is another critical
contributor to Ace2 and Swi5 regulation during the cell
cycle because it impacts their nuclear localization (Moll
et al. 1992) (Figure 4). Phosphorylation within the nuclear
localization sequences (NLSs) of Swi5 and Ace2 from S
phase through M phase, catalyzed by increasingly active
CDK, prevents transcriptional activation by preventing their
entry into the nucleus. Only when B-type cyclins are
destroyed during M phase and the Cdc14 phosphatase is
activated upon exit from mitosis, are their NLS motifs
dephosphorylated, allowing entry into the nucleus and promoter binding (Visintin et al. 1998). Thus, activation of
Swi5 and Ace2 targets in early G1 is dependent on exit from
mitosis. The asymmetric loss of Ace2 from mother cell nuclei
depends upon its nuclear export sequence (NES), a motif
that is not shared with Swi5 (Sbia et al. 2008). Inactivation
of Ace2 by nuclear exclusion is necessary because it is stable
throughout the cell cycle, whereas the bulk of Swi5 is apparently rapidly degraded upon localization to the nucleus
(Tebb et al. 1993). Together, these mechanisms account for
the selective inactivation of the M/G1 transcription factors
in mother and daughter cells.
The MAT gene cluster
Another cluster of M/G1 genes that is expressed differentially based upon cell type is the so-called MAT cluster of
haploid-specific genes regulated by the Ste12 transcription
factor (Oehlen et al. 1996; Spellman et al. 1998). Although
Ste12 participates in multiple transcription factor complexes, some as a heterodimer with Tec1 or Mcm1, regulation of the MAT gene cluster by mating pheromone occurs
via binding to pheromone response elements (PREs) in the
target promoters as a homodimer (Hwang-Shum et al.
1991). Genes in the MAT cluster encode many proteins involved in the response to mating pheromone, including
the Fus3 MAP kinase and Far1, both of which are critical
for induction of G1 phase cell-cycle arrest (Roberts et al.
2000). Whereas Ste12-regulated transcription is strongly
induced in response to mating pheromone signaling, its target genes are expressed at a basal level in haploid cells of
both mating types (Oehlen et al. 1996; Spellman et al.
1998).
Both basal and pheromone-induced expression of Ste12regulated genes are restricted to the pre-START interval
of G1 phase (reviewed by Dohlman and Thorner 2001;
Bardwell 2005). Confining mating to G1 phase prevents
inappropriate ploidy of the mating cells and proper organization of the apparatus for nuclear fusion. Transcriptional
and post-transcriptional controls contribute to the G1 phase
restriction. In particular, Far1, an inhibitor of G1 cyclinassociated CDK and regulator of cell polarity, is expressed
as a target of both Mcm1 and Ste12 and, thereby, accumulates from M phase through early G1 phase (Oehlen et al.
1996). Fus3 activates both Ste12 and Far1 by phosphorylation,
thereby, promoting gene expression and CDK inhibition.
Conversely, pheromone signaling is inhibited and Far1 is
targeted for destruction by ubiquitin-mediated proteolysis
by G1 cyclin/CDK, which accumulates as cells pass START
(McKinney et al. 1993; McKinney and Cross 1995; Henchoz
et al. 1997). Cln/CDK also attenuates mating pheromone
signal transduction by phosphorylating the MAPK cascade
scaffold protein Ste5 (Oehlen and Cross 1994; Strickfaden
et al. 2007; Bhaduri and Pryciak 2011). Because the substantial activation of Far1 by Fus3-dependent phosphorylation occurs only when the mating pheromone-signaling
pathway is induced by mating factor, stable arrest resulting
from Far1 inhibition of Cln/CDK does not occur in the absence
of pheromone. Instead, under those conditions, Cln/CDK
attenuates Ste12-dependent gene expression and promotes
cell proliferation. Consequently, cycling cells are only sensitive
to mating pheromone-induced cell-cycle arrest during early
G1 phase.
The mechanism of periodic expression of Ste12-dependent
genes in cycling cells is associated with basal signaling
through the pheromone response pathway leading to basal
activation of Ste12. Like pheromone-inducible expression,
basal expression of the MAT cluster genes occurs only during
G1 phase because the pathway is repressed by CDK, which
remains active from START until the Clb/CDK inhibitor Sic1
is expressed as cells exit M phase. Although basal pheromone
signaling is insufficient to cause cell cycle arrest, it is likely
responsible for the enhanced rate of cell proliferation observed in pheromone signaling mutants even in the absence
of pheromone (Lang et al. 2009).
The stability of both the START transition and pheromone arrest creates a tension between the mating pathway
and cell proliferation. Both the stability and reversibility of
those pathways is a consequence of the architecture of the
cell-cycle- and pheromone-regulated transcriptional networks (Doncic et al. 2011; Doncic and Skotheim 2013).
Pheromone arrest is stable due to pheromone-induced expression of Far1, Fus3, and other factors in the pathway. A
drop in pheromone leads rapidly to mitotic growth due both
to the rapid decay of signaling through the pathway and the
associated activation of Cln/CDK activity resulting from the
decreased expression of Far1. Cln/CDK activity promotes
proliferation by attenuating the basal activity of the pheromone pathway and thereby Ste12-dependent transcription
by promoting Far1 proteolysis and by activating G1/S transcription via positive feedback. Together, these responses flip
the bistable switch governing START (reviewed by Ferrell
2011).
Emerging Views of Cell-Cycle-Regulated
Transcription
The broad outlines of cell-cycle-regulated gene expression
have been worked out for almost a decade. Yet, as can be
seen from our discussion of the regulation of the cell-cycle
gene clusters, substantial fleshing out and numerous revisions of that view have led to a much greater understanding.
In the following vignettes, we introduce some of the recent
developments in the general area of cell-cycle-regulated
gene expression and a number of specific issues that are just
beginning to influence our understanding of this important
problem. Important topics impacting our understanding of
cell-cycle-regulated gene expression, including the influence
of nutrients, pheromones, stress responses, cell size, and
other factors, are discussed in other Reviews and Yeastbook
articles previously published and to come.
CDK’s role in cell-cycle-regulated transcriptional program
CDKs are widely believed to be the core mechanism
controlling cell-cycle-regulated transcription. Support for
the idea that periodic transcription occurs as a consequence
of periodic cyclin/Cdk activity came from experiments in
which specific groups of transcripts were found to be
misregulated in cells mutated for specific cyclins (Dirick
and Nasmyth 1991; Amon et al. 1993; Stuart and Wittenberg
1994; Koch et al. 1996). An attractive model was proposed by
Nasmyth and colleagues in which successive waves of cyclin
expression drive progression through the cell cycle as well as
control periodic transcription (Amon et al. 1993; Koch et al.
1996). When cells commit to a new cell cycle, Cln3 activates
a wave of transcription peaking in late G1. Clb2 then downregulates a subset of those transcripts and simultaneously
promotes the accumulation of a later wave of transcripts
peaking in G2/M. A subsequent wave of transcripts peaking
in late M/G1 is triggered by the destruction of Clb2 as cells
exit mitosis.
Textbook models suggest that the temporal program of
transcription is regulated by an interconnected regulatory
network containing both CDKs and transcription factors. In
these models, CDKs act as the key regulatory components
that control the activity of cell-cycle transcription factors
(Morgan 2007). There is good motivation for proposing this
regulatory relationship, as many of the 10–15 transcription
factors that are commonly placed in the cell-cycle network
are phosphorylated by CDKs. CDK-mediated phosphorylation regulates both nuclear localization and the formation
of transcription factor complexes at promoters (Ho et al.
1999; Costanzo et al. 2004; de Bruin et al. 2004). Consequently, they may serve as critical molecular switches for
the interconnected network. In particular, G1-cyclin/Cdks
activate SBF and MBF through phosphorylation of Whi5
(Costanzo et al. 2004; de Bruin et al. 2004) and, perhaps,
Swi6 (Sidorova et al. 1995), triggering accumulation of
many late G1 transcripts; SBF is then inhibited by the mitotic B-cyclin/Cdk complexes (Amon et al. 1993; Koch et al.
Cell-Cycle-Regulated Transcription
77
1996); B-cyclin/Cdks also phosphorylate and activate Ndd1
and Fkh2 to promote complex formation and trigger accumulation of many G2/M transcripts (Pic-Taylor et al. 2004);
B-cyclins inhibit nuclear localization of Swi5 (Moll et al.
1991) and Ace2 (O’Conallain et al. 1999) during mitosis
and, it is likely that B-cyclin destruction at the end of mitosis
triggers late M/early G1 transcript accumulation (Breeden
and Nasmyth 1987; Knapp et al. 1996; Kovacech et al. 1996;
Toyn et al. 1997). Thus, successive waves of transcription
could arise due to changing cyclin/CDK activities that act
post-translationally to turn key factors on and off.
By contrast, in network models arising from genomic
approaches, the temporal program of transcription can arise
largely independent of CDK regulation (Simon et al. 2001;
Lee et al. 2002; Pramila et al. 2006). In these models, successive waves of transcription emerge from oscillations in
transcription factor abundance, rather than regulation of
transcription factor activity. In a simplified model, transcription factors activate the expression of gene clusters that include genes encoding other transcription factors. Thus, the
program could be controlled intrinsically by a network of
serially activated transcription factors.
At the time when the first network models were proposed, the CDK and network models seemed incompatible.
Both existing models had widely acknowledged gaps. The
network models did not incorporate much of the posttranslational regulation of transcription factors by CDK
complexes that had been established by directed studies of
specific transcription factors. However, although the CDK
model was supported by rigorous experiments interrogating
small subsets of target genes, this model had not been tested
on a genome-wide scale.
The extent to which the cell-cycle-regulated transcriptional program was controlled by oscillations of specific
cyclin/CDK complexes or by a network of serially activated
transcription factors was examined directly by Orlando et al.
(2008). Using budding yeast cells in which all six B-type
cyclin genes had been inactivated (clb1,2,3,4,5,6Δ), they
showed that nearly 70% of the cell-cycle-regulated genes
continued to be expressed at the proper time. In addition,
the program of transcription repeated in a second cycle, despite the fact that the cells were arrested at the G1/S border
by all conventional cell cycle measures. In a later study,
similar results were observed in cdc28-4 cells at restrictive
temperature, although the number and amplitude of the
periodic genes was substantially reduced (Simmons Kovacs
et al. 2012).
Taken together, these results suggested that interconnected networks of transcription factors could drive most of
the periodic transcriptional program in the absence of Clb/
CDK activity and function as an autonomous oscillator
(Orlando et al. 2008; Simmons Kovacs et al. 2008, 2012).
Nonetheless, while many transcripts continue to oscillate in
the cells depleted for CDK activities, there were clear alterations in the behaviors of others. Thus, post-transcriptional
regulation by CDKs may act more like a rheostat in the
78
S. B. Haase and C. Wittenberg
control of periodic transcription rather than simply functioning as an “on–off” switch. In general, as CDK activities decreased, the number of identifiable periodic genes also
decreased (Simmons Kovacs et al. 2012). Furthermore, decreasing CDK activity was associated with diminished transcript amplitudes as well as increasing oscillatory periods.
The regulatory mechanisms responsible for the loss of amplitude and increased period may be positive feedback loops
in which cyclin expression is promoted by a transcription
factor that is then phosphorylated by the cyclin/CDK complex,
which increases the transcription factor’s activity (Simmons
Kovacs et al. 2012). This regulatory network motif affects both
G1 (SBF and Cln2; see section titled “Activation of G1/S transcription factors”) (Cross and Tinkelenberg 1991; Dirick and
Nasmyth 1991; Skotheim et al. 2008) and G2/M gene clusters
(Pic-Taylor et al. 2004).
Although CDKs may not be required as a core mechanism
for generating the temporally ordered program of transcription during the cell cycle, they clearly play a role in
maintaining the robust character of the program. Through
positive feedback loops, CDKs provide the basis for important switch-like behavior observed at G1/S and G2/M
transitions. Furthermore, the inhibition of Swi5/Ace2-regulated
genes by mitotic cyclin/CDK ensures that the initiation of a new
program of G1 transcription is dependent on exit from mitosis
and the subsequent loss of Clb2/CDK activity. Thus, transcription factor networks and CDKs collaborate to coordinate the
execution of the transcriptional program with the cell cycle
(Simmons Kovacs et al. 2012).
Checkpoint control of cell-cycle-regulated transcription
Cell-cycle checkpoints maintain the appropriate order of
cell-cycle progression by placing prerequisites on progression into a particular cell-cycle phase or on the execution of
phase-specific events (Hartwell et al. 1994). For example,
a checkpoint mechanism restricts progress into M phase
when replication is incomplete, either due to ongoing replication or stalling of DNA replication forks. Similarly, another
checkpoint mechanism restricts mitotic spindle elongation
when chromosomes are not appropriately attached to the
mitotic spindle. To elicit such responses, checkpoint pathways must impinge upon cell-cycle-regulatory mechanisms
(Elledge 1996). Furthermore, they must allow the cell time
to correct the problem yet retain the capacity to proceed
once that problem is corrected.
The observation that the cell-cycle-regulated transcriptional program can proceed (albeit less robustly) when CDK
activity is lost suggests that checkpoint arrest, which blocks
CDK cycling, might allow the transcription program to be
decoupled from cell-cycle events. This problem would be
resolved if checkpoint pathways directly regulate the transcription network to maintain synchrony between transcription and cell-cycle progression. Some evidence suggests that
checkpoints do, in fact, regulate transcription to enforce
robust cell-cycle arrest (Gardner et al. 1999; Chu et al.
2007). Several global transcript-profiling studies indicate
that the DNA replication and DNA damage checkpoint pathways trigger a checkpoint-dependent transcription program
controlled by Dun1 kinase (Figure 5) (reviewed in Fu et al.
2008). The Dun1 kinase induces a well-described transcriptional response involving genes (such as RNR2 and RNR3)
that mediate recovery from DNA damage or blocks to DNA
replication (Elledge et al. 1993; Zhou and Elledge 1993;
Allen et al. 1994; Gasch et al. 2001). Dun1 activity is stimulated by Rad53. Crt1, a repressor of DNA damage response
genes, is phosphorylated by Dun1 upon checkpoint activation and released from the DNA, thereby, allowing transcriptional activators access to promoter regions of Crt1-target
genes (Huang et al. 1998). In addition to its role in recovery
from checkpoint stresses, genetic evidence suggests that
Dun1 also plays a role in enforcing a robust checkpoint
arrest (Gardner et al. 1999). Although some of these studies
hint at regulation of cell-cycle genes (Gasch et al. 2001),
a specific role for Dun1-regulated transcription in cell-cycle
arrest has yet to be described.
In addition to activating the Dun1 pathway, Rad53
also directly phosphorylates Nrm1, the repressor of MBFregulated transcription, preventing its association with MBF
at target promoters (Figure 5) (de Oliveira et al. 2012; Travesa
et al. 2012). Consequently, MBF targets, which would normally be repressed when DNA replication is underway, remain active, leading to the accumulation of their products.
Because many of those targets are required for DNA replication and repair, this positions cells to correct the problems
leading to induction of the checkpoint and to progress
through S phase once that problem has been eliminated.
A similar mechanism is observed in fission yeast where the
Rad53 ortholog, Cds1, phosphorylates both Nrm1 and Cdc10,
the S. pombe ortholog of Swi6. The result is persistent activation of transcription by MBF, the sole G1/S transcription
factor in that organism (de Bruin et al. 2008b; Dutta et al.
2008; Aligianni et al. 2009; Gómez-Escoda et al. 2011). Finally,
Rad53 also phosphorylates Swi6 in S. cerevisiae (Sidorova and
Breeden 1997). Although the importance of that phosphorylation has not been established, it has been suggested that it
delays expression of CLN1 and CLN2 in G1 cells responding to
DNA damage.
The phosphorylation of Nrm1 by Rad53 specifically
affects MBF, allowing SBF to be repressed, like it is in untreated cells (Travesa et al. 2012). Consequently, repression
of SBF targets by Clb2/CDK occurs normally because Clb2 is
stabilized in response to down-regulation of APCCdc20 activity by the checkpoint. Thus, inappropriate accumulation
transcripts associated with morphogenesis and other processes promoted by targets of SBF is prevented. The differential regulation of the SBF and MBF transcription factors,
and thereby their specific targets by the DNA replication
checkpoint, provides one clear rationale for the existence
of independent transcription factors regulating the two
groups of G1/S cluster genes (de Oliveira et al. 2012;
Smolka et al. 2012; Travesa et al. 2012). This dual regulation is dispensable in fission yeast where MBF, the sole G1/S
Figure 5 Control of MBF-regulated transcription by the DNA replication
checkpoint. The influence of the DNA replication checkpoint on the MBF
transcriptional regulatory circuit is depicted. MBF-regulated promoters are
normally repressed as cells progress into S phase as a consequence of the
expression of the NRM1 gene and the accumulation of its product, the
MBF-specific repressor, Nrm1 (dashed line, graph). When the DNA replication checkpoint is activated as a consequence of stalling of DNA replication forks, the DNA replication checkpoint pathway leads ultimately to
the activation of the Rad53 protein kinase, which can phosphorylate the
downstream protein kinase Dun1 and the Nrm1 protein. Phosphorylation
activates Dun1, which can then phosphorylate the transcriptional repressor Crt1, leading to the activation of DNA damage response (DDR) genes.
Phosphorylation of Nrm1 inactivates it as a transcriptional repressor, leading to persistent activation of MBF target genes (solid line, graph) to
support repair of DNA and restoration of DNA replication. The color of
subunits and arrows is as in Figure 1.
transcription factor, is also subject to checkpoint regulation
via phosphorylation of the transcriptional repressor Nrm1
(de Bruin et al. 2006, 2008b; Dutta et al. 2008). In that
system, proliferation does not occur via budding and, consequently, morphogenesis is not regulated during G1 phase.
In addition to providing a mechanism by which the
checkpoint can activate only a subset of G1/S genes, the
differential regulation of SBF and MBF by the DNA replication checkpoint also poses a conundrum. SBF- and
MBF-regulated genes can be repressed by the accumulation
of Clb/CDK activity during the unperturbed cell cycle (Amon
et al. 1993; de Bruin et al. 2006). Why, then, is SBF repressed normally during S phase in cells responding to the
checkpoint signal, whereas MBF remains unaffected? This
observation suggests that a mechanism invoked by the
checkpoint signal allows MBF to evade the repressive activity of Clb/CDK. Although the mechanism of that evasion is
unknown, it is possible that the checkpoint-induced phosphorylation of Swi6 by Rad53 makes MBF refractile to inhibition by Clb/CDK (Sidorova and Breeden 1997). Whatever
the mechanism, this difference in their response to Clb/CDK
provides another example of differential regulation of SBF
and MBF during the cell cycle.
Although, the differential regulation of SBF and MBF
by the DNA replication checkpoint provides a seemingly
Cell-Cycle-Regulated Transcription
79
straightforward rationale for the involvement of two transcription factors in G1/S transcription, several observations
suggest that the situation is really much more complex. First,
many G1/S genes have multiple binding sites for both
transcription factors that appear to be functional in vivo (Iyer
et al. 2001; Horak et al. 2002; Bean et al. 2005; de Bruin
et al. 2006). Second, the expression of many G1/S genes is
impacted by inactivation of either of the two transcription
factors, suggesting that both factors can contribute to their
regulation (Bean et al. 2005; de Bruin et al. 2006). Finally,
some genes undergo a switch from binding SBF to binding
MBF during the course of the G1/S transcriptional burst as
a consequence of the presence overlapping binding sites for
the two transcription factors (de Oliveira et al. 2012). Those
genes seem to be activated by SBF, dependent upon inactivation of Whi5, and repressed by MBF, via the activity of
Nrm1. One rationale that has been proposed for this regulatory switch is to enable inducibility of those genes by the
DNA replication checkpoint and to ensure that the genes
are not expressed inappropriately if MBF is inactivated by
mutation or other insult (reviewed by Smolka et al. 2012).
Regardless of the reason, it is clear that the regulation of G1/S
genes by the combined activities of SBF and MBF provides
opportunities for more diverse modes of regulation than does
the involvement of either transcription factor on its own.
We are just beginning to understand the complex mechanisms by which checkpoint effector pathways control cellcycle-regulated transcription of the G1/S cluster. It also
seems likely that checkpoints may regulate other clusters of
periodic genes. Thus, it is likely that checkpoint control of cellcycle-regulated transcription will be a rich area of future
investigation.
Yeast metabolic cycle control of cell cycle and transcription
It has been recognized for some time that metabolic
processes in yeast are periodically regulated (Futcher 2006
and references within) and may be coordinated with the cell
cycle (e.g., Creanor 1978; Novak and Mitchison 1986,
1990). When budding yeast cells are grown in continuous
culture conditions at appropriate densities and growth rates,
the cells synchronize in robust metabolic cycles that can be
monitored by periodic changes in dissolved oxygen. The
reported periods of the YMC can vary substantially (Klevecz
et al. 2004; Tu et al. 2005; Slavov and Botstein 2011), and
seem to be linked to growth rate (Brauer et al. 2008; Slavov
and Botstein 2011). The YMC regulates a large transcriptional program and appears to be coordinated with the cell
cycle under slow growth conditions.
The YMC-regulated transcriptional program: By sampling
continuous cultures of metabolically synchronous populations of budding yeast over time, investigators have identified oscillations in gene expression that are coincident with
the periodicity of the YMC. In one of the initial reports, cells
were sampled over the course of three metabolic cycles and
by microarray analyses found that the bulk of the genome
80
S. B. Haase and C. Wittenberg
exhibited oscillations in transcript abundance (Klevecz et al.
2004). The amplitudes of these oscillations were modest,
with peak-to-trough ratios of 2. In a subsequent study,
the McKnight group (Tu et al. 2005) also sampled metabolically synchronous cells growing in a chemostat over three
metabolic cycles. Their findings suggested approximately
half of the genes in the genome exhibited transcript oscillations coincident with the YMC. The amplitudes of these
transcript oscillations were in the range of 10-fold (Tu
et al. 2005), substantially higher than observed by Klevecz
and colleagues.
Using clustering approaches, both groups were able to
identify sets of transcripts that appeared to be coregulated,
and many of the coregulated genes shared related functions
(Klevecz et al. 2004; Tu et al. 2005). Analysis of the data
from Tu and colleagues revealed three “super clusters” of
genes that were expressed in relatively discrete temporal
intervals that correspond to three phases of the YMC: oxidative (Ox), reductive building (R/B), and reductive charging
(R/C) (Tu et al. 2005). Within these clusters of oscillating
transcripts were several cell-cycle-regulated genes, suggesting
that the YMC transcriptional program and the cell-cycle transcriptional program are interconnected (Klevecz et al. 2004;
Tu et al. 2005).
Botstein and colleagues approached YMC control of
transcription from a different angle. They used continuous
culture conditions to investigate the transcriptional response
of yeast to a wide variety of growth limiting conditions in
which cells were starved for a variety of different nutrients
(Brauer et al. 2008). Under this broad range of growth conditions, they found that the expression of approximately
one-fourth of the genes in the genome is linearly correlated
(either positively or negatively) with growth rate. Surprisingly, this set of growth rate responsive (GRR) genes
exhibited substantial overlap with a set of genes previously
identified as environmental stress response (ESR) genes
(Gasch et al. 2000), suggesting a functional relationship
between the gene sets. Genes that exhibited positive correlation with growth rate tended to be genes inhibited by
stress, while genes that were down-regulated in fast growth
conditions were often activated by stress (Brauer et al.
2008). As well, there was considerable overlap between
the GRR gene set and the YMC responsive gene sets identified by Tu et al. (2005) (Brauer et al. 2008). In a recent
study using continuous culture conditions and a variety of
nutritional limitations, Slavov and Botstein (2011) demonstrated that all of the GRR transcripts were, in fact, periodic during the YMC. Strikingly, negatively and positively
correlated GRR transcripts oscillate in opposing YMC
phases. Taken together, these findings suggest that the
mechanisms controlling oscillation in transcript abundance
may be integrating signals from stress, growth rate, YMC,
and cell cycle.
YMC control of cell cycle: The identification of cell-cycleregulated genes as part of the YMC transcriptional program
suggested that these genes were regulated by multiple
conditions or that the YMC and the cell cycle may be
coordinated (Klevecz et al. 2004; Tu et al. 2005). DNA content and budding analyses both suggest that the YMC and
the cell cycle are indeed synchronized in continuous culture
conditions (Klevecz et al. 2004; Tu et al. 2005). Both groups
suggest that only a fraction of cells in the population [Tu
et al. (2005) estimates 50%] enter the cell cycle during each
metabolic cycle. In addition, it appeared that the YMC
phases might be coherent with cell-cycle phases, with S
phase being restricted from occurring during the Ox phase
of the YMC (Klevecz et al. 2004; Tu et al. 2005).
Both groups proposed that the YMC gates the cell cycle so
that the execution of S phase is restricted to the reductive
phase and that this restriction is important for preventing
oxidative damage of DNA. Supporting evidence for this
hypothesis was reported in a subsequent study from the
McKnight group (Chen et al. 2007). In mutant cells where S
phase was allowed to proceed during the Ox phase of the
YMC, they observed an increase in the spontaneous mutation rates. However, the mutant genes found to disrupt the
phase coherence of the YMC and the cell cycle were themselves genes controlling the cell division cycle (Chen et al.
2007), suggesting the increase in mutation rate observed in
these mutants could be related to improper execution of
cell-cycle functions rather than replicating DNA during the
oxidative phase of YMC.
In a recent study, the connection between the cell cycle
and the YMC was directly tested (Slavov and Botstein
2011). By varying nutrient availability, and thereby altering
the growth rate of continuous cultures, it was demonstrated
that the restriction of S phase from the Ox phase of the YMC
was dependent on the growth rate. Under specific growth
rate conditions, synchrony between the cell cycle and YMC
could be maintained, but execution of S phase occurred
during the Ox phase of the YMC (Slavov and Botstein
2011). Interestingly, the cell cycle mutants that disrupted
the coordination between S phase with the reductive phases
of the YMC were also the mutants that exhibited the most
substantial alterations in the growth rate and YMC period
(Chen et al. 2007).
One interpretation of these data is that the duration of
the cell division cycle influences the YMC period. Consistent
with this possibility, it has been proposed that the coherence
of cell-cycle phases with YMC phases may reflect the storage
of carbohydrates under slow-growth conditions and a sudden “burn” of these carbohydrates during G1 phase (Futcher
2006). This rapid utilization of carbohydrates provides energy and drives rapid protein synthetic rates, providing
a “finishing kick” for cells to pass through START and enter
S phase (Futcher 2006). This coupling of cell-cycle and YMC
programs seems to be restricted to conditions of slow
growth, and oscillation of YMC or GRR transcripts has not
been observed in microarray studies performed using rapidly growing cell populations synchronized in the cell cycle
(Cho et al. 1998; Spellman et al. 1998; Pramila et al. 2006;
Brauer et al. 2008; Orlando et al. 2008). This may be because the rapidly growing cells have abundant glucose and
do not store substantial amounts of carbohydrates.
Many open questions remain regarding how and why
budding yeast cells coordinate the YMC with the cell-cycle
program. The interaction between the YMC and the cell
cycle is likely to be more complex than a simple gating of the
cell cycle by a metabolic oscillator. Although the mechanisms driving the YMC remain undiscovered, like the cell
cycle, it is clearly capable of producing oscillations in an
abundance of transcripts on a genome-level scale. The
proposal that the cell-cycle period may be controlled by
a transcriptional network oscillator suggests that the YMC
and the cell cycle may be coupled via coordinated control
of key transcriptional regulators (Orlando et al. 2008;
Simmons Kovacs et al. 2008, 2012).
What is the role of chromatin in
cell-cycle-regulated transcription?
Transcription takes place in the context of chromatin and, as
a consequence, is influenced by it (Lenstra et al. 2011;
Rando and Winston 2012). Cell-cycle-regulated genes have
proven useful for understanding both general, and genespecific contributions of chromatin to the regulation of expression. The specific roles of chromatin in the control of
cell-cycle-regulated gene expression and the relationship between chromatin and the transcription factors that control
the cell-cycle gene clusters are only beginning to be understood. In many cases, the specific vs. general roles of chromatin in the control of cell-cycle-regulated gene expression
have yet to be established.
The earliest studies of the contribution of chromatin
modification and remodeling to gene expression and, more
specifically, cell-cycle-regulated gene expression were based,
in large part, upon the pioneering studies of the G1/S gene,
HO, by the Nasmyth and Herskowitz laboratories (reviewed
by Stillman 2013; see Haber 2012). Most of the studies of
the role of chromatin in the regulation of HO expression
focused upon licensing for differential expression in mother
cells, mediated by an upstream regulatory region known as
URS1. However, the balance of the regulatory functions,
mediated by URS2, are largely indistinguishable from expression of other SBF target genes (Peterson and Herskowitz
1992; Cosma et al. 1999, 2001). Consequently, most of the
observations regarding URS2 can be generalized to CLN2
and other G1/S promoters.
Recent study of the CLN2 and HO promoters has focused,
in part, upon the role for histone deacetylation by the Rpd3
(L) HDAC complex in repression of SBF and MBF genes (de
Bruin et al. 2008a; Takahata et al. 2009a,b, 2011; Wang
et al. 2009). Rpd3(L) recruitment to SBF promoters depends
upon the binding of the Sin3-associated protein Stb1 to promoters in a manner that depends upon the SBF-specific
transcriptional repressor Whi5 (Wang et al. 2009; Takahata
et al. 2011). At MBF genes, Stb1 binding occurs, but is, by
necessity, Whi5 independent (Costanzo et al. 2003; de Bruin
Cell-Cycle-Regulated Transcription
81
et al. 2008a; Takahata et al. 2009a). Stb1 binding is accompanied by the binding of Cdk1, which, via some combination
of phosphorylation of Whi5, Stb1, and the transcription
factors, promotes activation of the promoters. Cln/Cdk1dependent phosphorylation of Whi5 and Stb1 leads to the
dissociation and export of Whi5 from the nucleus and the
dissociation of Rpd3(L) from promoters. Promoter activation occurs upon the loss of Rpd3(L) binding, presumably
accompanied by the restoration of histone acetylation. Similar changes in histone acetylation during G1 and S phases
are observed at both SBF and MBF target promoters. Although Rpd3(L) is an important factor in the regulation of
histone acetylation state and involved in the repression of
many genes, Stb1 may affect only a small subset of genes,
including those in the G1/S gene cluster.
Activation of both CLN2 and HO expression, and likely
the entire G1/S gene cluster, also involves the FACT complex, which promotes nucleosome eviction at those promoters (Takahata et al. 2009a,b; Wang et al. 2009). FACT
acts just prior to, or at the time of, transcriptional activation
and is required for G1/S expression. The binding of FACT to
both CLN2 and HO promoters depends upon the Swi6 subunit of SBF along with Whi5 and Stb1. The G1/S genes
provide the only example of a promoter-associated role for
FACT, which is most often associated with transcriptional
elongation (Rando and Winston 2012). FACT has also been
implicated in CLN3 transcription, although whether that
occurs via a direct or indirect mechanism remains obscure
(Morillo-Huesca et al. 2010).
At least some of these factors play well-established roles
at genes that are expressed independent of cell-cycle
position, but the extent to which these transcription-factordependent mechanisms for chromatin modification and
remodeling play specific roles at cell-cycle genes is not yet
clear. However, those factors are clearly distinct from the
more general role of nucleosome-depleted regions (NDRs)
that encompass the SBF-binding sites at the CLN2 promoter
and are required for the reliability of cell-cycle-dependent
transcriptional activation (Bai et al. 2010, 2011). Consistent
with that, the formation of NDRs at the CLN2 promoter does
not require any of the known G1/S-specific transcriptional
regulators. Additionally, histone H3 K79 methylation has
also been associated with the function of SBF, Whi5, and
Nrm1, although the relevance of that association to the regulation of G1/S promoters is not clear (Schulze et al. 2009).
There are many studies suggesting a role for chromatin in
the regulation of other cell-cycle-regulated gene clusters,
although most are less well studied. For example, Fkh2/
Mcm1 promotes recruitment of Rpd3(L) leading to repression of G2/M cluster genes (Veis et al. 2007). Interestingly,
in a two-step process, Rpd3(L) dissociates from G2/M gene
promoters at the onset of S phase in a manner that depends
upon Cln/CDK and, only then, does the combined activation
of Clb2/CDK and Cdc5 promote eviction of a nucleosome
from those promoters and binding of Ndd1 to Fkh2. Recent
studies describing a role for histone chaperones and chro-
82
S. B. Haase and C. Wittenberg
matin boundary proteins in chromatin remodeling at the
histone gene cluster are reviewed elsewhere (Eriksson et al.
2012). Together, these studies suggest both direct and indirect
roles for cluster-specific transcription factors in the recruitment and regulation of chromatin modifying and remodeling
proteins. Nevertheless, the degree of uncertainty regarding
the specificity of these processes makes this a fertile ground
for future research.
Rationale for Cell-Cycle-Regulated Transcription:
Need to Have or Nice to Have?
To understand the rationale for regulating the expression of
genes during the cell cycle, it is helpful to examine which
genes are controlled. Deciding whether or not a gene is cellcycle-regulated is not as trivial as it might seem. Many
definitions of “cell-cycle regulated” can and have been applied, yielding a variety of estimates of the number and
identities of cell-cycle-regulated genes in S. cerevisiae (Cho
et al. 1998; Spellman et al. 1998; de Lichtenberg et al. 2005;
Pramila et al. 2006; Orlando et al. 2008; Granovskaia et al.
2010; Guo et al. 2013). The number of transcripts reported
as cell-cycle regulated has varied from 416 to 1270. A consensus set of 440 genes (Orlando et al. 2008) was identified
by each of three distinct studies (Spellman et al. 1998;
Pramila et al. 2006; Orlando et al. 2008). The remaining
genes, which are unique to each of the studies, could result
from differences in strain backgrounds or experimental conditions including synchrony procedures, growth medium,
microarray platform, or the approach used to identify periodic genes.
So which genes are cell-cycle regulated? The best answer
is: There is no single, identifiable set of cell-cycle-regulated
genes. Because there is no precise and universally accepted
definition of cell-cycle regulated, there is no single algorithmic method available for identifying genes with periodic
behaviors over the cell cycle. In addition, each of the algorithms returns a rank-ordered list of genes with no obvious
gap in the distribution that distinguishes periodic from nonperiodic genes. Thus, all cut-offs chosen by investigators are
somewhat arbitrary, and it is likely more accurate to say that
there is a continuum of cell-cycle-regulated genes.
Despite the difficulty in precisely defining a single set of
cell-cycle-regulated genes, it is likely that 20% of genes in
the yeast genome are expressed periodically during the cell
cycle at a substantial energy cost to the cell. This observation
suggests that coordination of gene expression with cell-cycle
events is important for the well being of cells. Several distinct though potentially overlapping rationales have been
advanced as reasons for phase-specific expression of various
genes.
First, some genes are expressed at the time they are
required during the cell cycle. This is clearly true for a large
number of genes whose function is required for, or is related
to, morphogenesis, DNA replication, and chromosome
segregation. The logic for this pattern of expression is
referred to as “just in time.” Whether genes are actually
expressed just in time is not always so clear. For example,
because cytokinesis occurs subsequent to M phase, one
might expect the expression of genes encoding the structural
elements of the bud neck, including septins, to occur in M
phase rather than G1. However, those genes are, in fact,
expressed just in time because the structures at the bud
neck, which play roles from G1 through M phases, are assembled during G1 phase prior to budding, precisely when
G1/S gene expression is maximal (reviewed in Bi and Park
2012; Howell and Lew 2012). Similar scenarios may explain
the periodic expression of some of the genes for which no
rationale is currently apparent.
Many genes that fall into the just-in-time category,
despite being expressed at the time of their essential
function, need not be expressed precisely at that time and,
in some cases, need not be expressed periodically at all. In
fact, several genome-wide studies have revealed the vast
majority of yeast genes cause no significant growth disadvantage, at least when constitutively overexpressed in
cycling cells from the inducible GAL1 promoter (Liu et al.
1992; Sopko et al. 2006; Douglas et al. 2012).
In addition to those genes that are expressed just in time,
there are a large number of genes expressed at a specific
cell-cycle interval during which they have no apparent
function. In some cases, the apparent lack of phase-specific
function may be a consequence of our lack of knowledge,
but in other cases the genes may just “tag along” with other
genes expressed during the same phase. That is, the pattern
of expression may be adequate but not necessarily advantageous. However, some of those genes might be better described as just in case, because their expression may prepare
the cell to deal with a sporadic event that does not occur in
all cell cycles. For example, whereas many DNA repair proteins are expressed prior to every S phase, damage may
occur only in a subset of cells during any round of DNA
replication.
Finally, some genes that are expressed just in time are
deleterious when expressed outside of the interval during
which they function. Those genes often encode proteins
with regulatory functions that are temporally coupled to
their accumulation. For example, the genes encoding the
B-type cyclins Clb2 and Clb5 and others encoding the anaphase regulators Cdc20 and Pds1 perturb cell-cycle progression when they are expressed continuously throughout the
cycle, with the caveat that the studies in question involve
overexpression (Nasmyth 1993; Sopko et al. 2006). However, even in the context of overexpression, such genes are
surprisingly rare.
So, why express a gene just in time when that pattern
of expression is unnecessary? One possibility is that justin-time expression is simply a matter of economy. Often,
proteins expressed outside of the interval during which they
function are disposed of by proteolysis or other mechanisms
(e.g., those encoding the G1 cyclins Cln1–3 and the CDK
inhibitor Sic1) (Lanker et al. 1996; Willems et al. 1996;
Verma et al. 1997). Consequently, there is a large energy
cost to the cells of expressing those genes outside of their
functional interval. Although expressing those proteins
throughout the cycle may not produce a noticeable or debilitating phenotype, it may result in a slight competitive
disadvantage under optimal or suboptimal conditions. Consequently, such changes would not be noticed in typical laboratory growth conditions. A definitive answer to these
questions is lacking because, except for studies involving
overexpression, the impact of continuous expression of
cell-cycle-regulated genes on overall fitness has not been
systematically investigated.
The Topology of Cell-Cycle-Regulated Transcriptional
Circuitry is Conserved
One of the overarching questions arising in the study of
model systems is the degree to which their regulatory
circuitry is conserved with humans. Comparative analysis
of cell-cycle transcriptional programs across the Eukaryota
has provided one window through which to examine that
question. The general organization and structure of the cellcycle apparatus is conserved among the organisms so far
examined. Finally, all of those organisms express a portion of
their genes in a cell-cycle-dependent manner. Consequently,
it has been anticipated that the conservation of the cellcycle-dependent transcriptional machinery will be substantial. This supposition has been met with both confirmation
and contradiction.
The cell-cycle transcription program has now been
examined on a genomic scale in plants, a variety of yeasts
including budding and fission yeast, and in humans (Spellman
et al. 1998; Whitfield et al. 2002; Menges et al. 2003; BarJoseph et al. 2004; Rustici et al. 2004; Pramila et al. 2006;
Gauthier et al. 2008). Although it is often difficult to assess
in light of the evolution of both sequence and function,
comparative analysis of cell-cycle-regulated gene expression
has led to the conclusion that the ontology of genes under
cell-cycle control is often conserved (CDK regulators, chromatin proteins, transcription factors, DNA replication and
repair proteins, mitotic proteins, etc.), but the expression
pattern of orthologs is not always conserved (Lu et al.
2007; Orlando et al. 2007). The degree of divergence seems
to increase with the evolutionary distance between the species and is even obvious in closely related Saccharomyces
species, cerevisiae and bayanus (Guan et al. 2010, 2013; Eser
et al. 2011). This seems surprising, considering the similarities in cell-cycle organization between those organisms.
However, the observation that the timing of expression of
cell-cycle-regulated genes in yeast is flexible, and, in some
cases, completely dispensable (see above) suggests that such
changes in expression might be easily accommodated. Of
course, the differences in periodicity at the transcription
level do not account for the loss or acquisition of the broad
array of post-transcriptional mechanisms that might restore
periodic regulation of many of those gene products. In fact,
Cell-Cycle-Regulated Transcription
83
Figure 6 Conservation of the G1/S regulatory circuitry
in yeast and human. Comparison of the topology of the
G1/S transcriptional circuitry between yeast and human
reveals a high degree of conservation. Both SBF and E2F
are heterodimeric transcription factors that bind to promoters in quiescent cells and repress transcription by
virtue of their interaction with repressors, Whi5 and
Rb, respectively. Both Whi5 and Rb, the retinoblastoma
tumor suppressor, are phosphorylated by a G1 cyclinassociated CDK, leading to their dissociation from the
transcription factor and activation of G1/S cluster promoters. Those genes are then repressed in S phase via
mechanisms that promote the loss of the activating
transcription factor from their target promoters. Despite
the high degree of conservation of the topology of
these regulatory circuits, the transcription factors exhibit
little or no relatedness in terms of either amino acid
sequence or structure. The color of subunits and arrows
is as in Figure 1.
differences in the periodicity of transcription that have
coevolved with changes in post-transcriptional regulation
have been documented (de Lichtenberg et al. 2005; Jensen
et al. 2006). Another indication of the evolutionary flexibility
of cell-cycle-regulatory phenomenon is the finding that within
protein complexes where one or more subunits is periodically
expressed, the identity of the periodically expressed subunits
may differ between species (Jensen et al. 2006). In fact, it
appears that just-in-time assembly may be important and
conserved, whereas just-in-time expression may be dispensable (de Lichtenberg et al. 2005). This notion is consistent
with the finding that within closely related yeast species,
there is extensive repurposing of conserved transcription factor subunits to form novel complexes with other factors and
produce entirely new transcriptional outcomes (Tuch et al.
2008).
Like cell-cycle-regulated transcripts, comparison of the
regulators of periodic gene expression reveals strong conservation of the general topology of the systems and
dramatic divergence of the details. This is well illustrated
by analysis of the transcriptional machinery regulating the
G1/S gene clusters of fungi and mammals (reviewed by
Wittenberg and Reed 2005; Cross et al. 2011; Bertoli et al.
2013b). The activating members of the E2F family of transcriptional regulators, E2F1 being the best studied, exhibit
many parallels with SBF (Figure 6) (Dimova and Dyson
2005). E2F1/DP1 binds to promoters in quiescent cells in
complex with a transcription repressor, in this case Rb, the
retinoblastoma tumor suppressor protein, a functional homolog of Whi5. Upon entry into the cell cycle, Rb is phosphorylated by a G1 cyclin/CDK, in this case CycD/Cdk4/6,
functional analogs of Cln3/Cdk1, leading to activation of the
E2F1/DP1 targets. Finally, the G1/S genes are repressed by
the binding of the E2F6, a repressive form of E2F, as cells
progress into S phase, leading to the dissociation of E2F1/
DP1 from promoters (Bertoli et al. 2013a). This precisely
parallels the regulation of SBF in budding yeast. However,
examination of E2F1, DP1, and Rb, reveals no recognizable
84
S. B. Haase and C. Wittenberg
protein sequence conservation, or even structural conservation, with Swi4, Swi6, or Whi5 (Cross et al. 2011).
Whether this is an example of rapid evolution between
fungi and humans or convergent evolution directed at
a redundant function remains to be established. However,
it is clear that the E2F/Rb family and not the SBF/Whi5
family of regulators exists in the plant kingdom, which is
evolutionarily distant from both fungi and mammals (Cross
et al. 2011). Furthermore, despite the lack of relationship
between the transcriptional regulators, all of these organisms use a conserved, albeit diversified, family of CDK
kinases to coordinate gene expression with the cell cycle
(Enserink and Kolodner 2010).
In contrast to the lack of conservation of SBF, E2F, and
their regulators, other cell-cycle-associated transcriptional
regulators from yeast exhibit strong sequence and structural
conservation with other eukaryotes. For example, both the
MADS box transcription factor, Mcm1, and Forkhead transcription factors, Fkh1 and Fkh2, are relatively well conserved among eukaryotes. However, at least in the case
of the MADS box transcription factors, their role in cellcycle regulation appears not to be conserved. SRF is a
well-established regulator of growth-associated genes, but
not those expressed periodically during the cell cycle (Shore
and Sharrocks 1995). In plants, MADS box transcription
factors are prominently associated with developmentally
regulated gene expression (reviewed by Masiero et al.
2011). Interestingly, although yeast Mcm1 plays a prominent role in the regulation of G2/M and M/G1 regulated
genes, it is also involved in a number of functions that are
not related to the cell cycle.
In contrast, the human Forkhead protein, FoxM1, shares
many regulatory properties with Fkh2. That regulation
appears not to involve either a MADS box transcription factor or an Ndd1 ortholog, as it does in yeast. However, like in
yeast, the activity of FoxM1 is required for the proper expression of a number of mitotic genes during G2/M and, like
the Mcm1/Fkh2/Ndd1 complex, it is subject to regulation by
both Cdk1 and the human Polo kinase, Plk1 (reviewed by
Murakami et al. 2010).
Together these observations suggest a complex evolutionary relationship between the factors mediating cellcycle-regulated gene expression and their targets. Perhaps unsurprisingly, it is the final product, the overall
execution of cell-cycle events, that is best conserved rather
than the details of the regulatory pathway that controls
that outcome.
Acknowledgments
The authors thank Anna Travesa, Danny Lew, Nick Buchler,
Sara Bristow, Adam Leman, Christina Kelliher, Mark Chee,
and anonymous reviewers for helpful discussion and comments on the manuscript. S.B.H. is supported by grants from
the Defense Advanced Research Projects Agency and the
National Institutes of Health (NIH) P50-GM081883. C.W. is
supported in part by grants R01 GM059441 and R01
GM100354 from the National Institutes of Health.
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Communicating editor: M. Tyers