REVIEW ARTICLE
DNA damage-induced gene expression in Saccharomyces cerevisiae
Yu Fu, Landon Pastushok & Wei Xiao
Department of Microbiology and Immunology, University of Saskatchewan, Saskatoon, SK, Canada
Correspondence: Wei Xiao, Department of
Microbiology and Immunology, University of
Saskatchewan, 107 Wiggins Road,
Saskatoon, SK, Canada S7N 5E5. Tel.: 11 306
966 4308; fax: 11 306 966 4311;
e-mail: [email protected]
Received 15 February 2008; revised 27 May
2008; accepted 28 May 2008.
First published online 8 July 2008.
DOI:10.1111/j.1574-6976.2008.00126.x
Editor: Martin Kupiec
Keywords
Saccharomyces cerevisiae ; DNA damage;
transcriptional regulation; SOS response;
cell-cycle checkpoint; signal transduction.
Abstract
After exposure to DNA-damaging agents, both prokaryotic and eukaryotic cells
activate stress responses that result in specific alterations in patterns of gene
expression. Bacteria such as Escherichia coli possess both lesion-specific responses
as well as an SOS response to general DNA damage, and the molecular mechanisms
of these responses are well studied. Mechanisms of DNA damage response in lower
eukaryotes such as Saccharomyces cerevisiae are apparently different from those in
bacteria. It becomes clear that many DNA damage-inducible genes are coregulated
by the cell-cycle checkpoint, a signal transduction cascade that coordinates
replication, repair, transcription and cell-cycle progression. On the other hand,
among several well-characterized yeast DNA damage-inducible genes, their
effectors and mechanisms of transcriptional regulation are rather different. This
review attempts to summarize the current state of knowledge on the molecular
mechanisms of DNA damage-induced transcriptional regulation in this model
lower eukaryotic microorganism.
Introduction
DNA is the carrier of genetic information in most organisms.
Any damage to the molecular structure of DNA has the
potential to cause genomic instability, mutagenesis or even cell
death. Unfortunately, DNA damage is unavoidable; DNA is
continually exposed to insults resulting from exogenous and
endogenous DNA-damaging agents as well as challenges posed
by DNA replication. Therefore, it is not surprising that living
organisms have developed numerous pathways to deal with
DNA damage. In response to DNA damage, cells can alter many
of their intracellular processes, including DNA repair, metabolism and cell-cycle progression, for cell survival and maintenance of genomic stability (Friedberg et al., 2006). Such
dynamic responses can be achieved through a variety of
molecular mechanisms such as protein localization, protein
degradation, posttranslational modification and genetic regulation. Generally speaking, altering gene expression is perhaps
one of the most fundamental responses and it is reasonable to
suggest that most, if not all, cellular processes are affected at the
level of transcriptional regulation. In the field of DNA repair
and mutagenesis, the notion that DNA damage causes an
alteration in the expression profile of damage responsive genes
has been an important area of research for many years. It is
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c
expected that the expression profile of a cell following DNA
damage will provide us with critical information on how a cell
protects itself from such stress.
Transcriptional regulation in response to DNA damage has
been studied extensively in model bacteria such as Escherichia
coli. In contrast, such information is relatively scarce in
eukaryotic organisms, from simple lower unicellular eukaryotes
such as Saccharomyces cerevisiae to humans. In this review, we
attempt to summarize recent findings related to gene regulation
in response to DNA damage in lower eukaryotes, particularly
the budding yeast S. cerevisiae, and to compare this response
with the well-studied bacterial SOS response.
Bacterial transcriptional responses to DNA
damage
The SOS response in E. coli
When E. coli cells are subjected to DNA damage, about 48
unlinked genes are coordinately induced through a complex
SOS regulatory network (Courcelle et al., 2001). The increased expression of these SOS regulon genes results in the
elaboration of a set of physiological responses such as an
enhanced capacity for recombination repair and excision
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DNA damage-induced transcription in budding yeast
repair, enhanced mutagenesis (due to error-prone translesion DNA synthesis mediated by PolIV and PolV) and
inhibition of cell division. These responses have been
collectively termed the SOS response (Radman, 1975;
Witkin, 1976; Gottesman, 1981; Little & Mount, 1982).
Current model for transcriptional control of the
SOS response
The SOS regulatory network is mainly controlled by two
proteins: RecA and LexA (Radman, 1975; Little & Mount,
1982; Walker, 1984). LexA is a transcriptional repressor that
binds to SOS boxes located near or inside the operator site of
the SOS-induced genes. SOS boxes are often palindromic
structures with a high degree of homology in nucleotide
sequence. An ideal symmetrical consensus sequence 5 0 TACTGTATATATATACAGTA-3 0 was derived from the analysis of a pool of SOS boxes (Berg, 1988). The sequence
distinction in SOS boxes allows the LexA repressor to bind
to operators with different strengths (Lewis et al., 1994). The
LexA occupancy prevents accessibility to RNA polymerase
so as to inhibit the initiation of transcription. LexA is likely
to interact with DNA via its N-terminal domain, and the
C-terminus of LexA is required for its dimerization. The
dimerization of LexA is essential for its ability to repress
SOS-regulated genes in vivo. Meanwhile, LexA is able to
undergo a slow intramolecular self-cleavage termed autodigestion, whose rate significantly increases upon interaction with RecA (Little, 1984, 1991, 1993).
The RecA protein of E. coli has at least three functions
in the SOS response. It not only plays a role in the
transcriptional regulation in response to DNA damage but
also directly participates in translesion synthesis and homologous recombination. Following DNA damage, DNA
synthesis becomes discontinuous and single-strand DNA
(ssDNA) is produced by failed attempts to replicate damaged DNA. In the presence of ATP, RecA binds to the
ssDNA region and forms helical RecA-ssDNA nucleoprotein filaments. LexA then diffuses to deep grooves in the
RecA-ssDNA filaments and interacts with them in a manner that results in autocatalytic cleavage of LexA at a scissile
peptide bond located between Ala84 and Gly85. Cleavage
of LexA inactivates its ability as a repressor, thus releasing
the SOS regulon genes from transcriptional repression. As
cells begin to recover from the inducing treatment by
various DNA repair and tolerance processes, the regions of
ssDNA disappear, and thus the inducing signal is diminished. Without the cleavage stimulated by RecA-ssDNA
filaments, the pool of LexA is boosted, which leads to
repression of the transcription of SOS regulon genes and a
return to the uninduced state (Walker, 1984; Friedberg
et al., 2006).
FEMS Microbiol Rev 32 (2008) 908–926
Fine tuning in the induction of the SOS response
The SOS regulatory system provides E. coli with a rapid
transcriptional response to the presence of DNA damage.
Furthermore, during SOS induction, the timing, the duration and the level of induction are diverse for different LexAregulated genes, suggesting a fine-tuning mechanism in the
SOS induction. The fine tuning is possibly determined by
at least four parameters: (1) the binding affinity of LexA
for the SOS box in the operator region; (2) the number of
SOS boxes in the operator region; (3) the location of the
SOS box relative to the promoter; and (4) the strength of
the promoter.
After exposing E. coli cells to DNA-damaging agents,
genes with operators that bind LexA relatively weakly are
the first to turn on fully. For example, uvrA, uvrB, ruvA,
ruvB, recN and sulA are induced within 5 min after 40 J m2
UV radiation (Courcelle et al., 2001). uvrA and uvrB encode
proteins involved in nucleotide excision repair (NER); recN,
ruvA and ruvB encode proteins used for recombination
repair; while the protein product of the sulA gene can
temporarily arrest cell division to allow bacteria time to
complete the repair of damaged DNA. After the activation of
loosely controlled SOS genes, if the damage cannot be fully
repaired by NER and homologous recombination, genes
with operators that are tightly controlled by LexA will be
turned on. For example, the full induction of umuC and
umuD is not observed until 20 min after 40 J m2 UV
irradiation (Courcelle et al., 2001). Similar to LexA, the
protein encoded by umuD also has a latent ability to autodigest, and the auto-digestion is strongly stimulated by the
interaction between the RecA/ssDNA nucleoprotein filament and UmuD (Nohmi et al., 1988). UmuC and a
posttranslationally processed form of UmuD (UmuD 0 ) serve
as a mutagenic lesion-bypass DNA polymerase (PolV). This
last response allows the survival of E. coli after severe DNA
damage, but at the expense of introducing errors into the
genome.
In the SOS regulatory network, the transcript level of key
regulatory genes recA and lexA is also regulated by LexA,
thus forming a delicately controlled circuit (Brent &
Ptashne, 1980; Little et al., 1981; Brent, 1982). recA has one
SOS box positioned between the 35 and 10 regions of
the promoter, and a tandem pair of SOS boxes is located in
the lexA promoter region (Brent & Ptashne, 1981; Little
et al., 1981; Schnarr et al., 1991). LexA can bind to these SOS
boxes to prevent the initiation of transcription. Relative
binding affinity experiments reveal that LexA binds to the
recA operator more strongly than to the lexA operator and
many other operators of SOS regulon genes such as uvrA,
uvrB and uvrD (Brent & Ptashne, 1981; Peterson & Mount,
1987; Schnarr et al., 1991). Accordingly, it allows an intermediate inducible state, bridging an uninduced state and a
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910
fully induced state (Little, 1983). A low amount of inducing
signal can thus lead to the activation of some of the SOS
functions, such as uvr1-dependent NER, without substantial amplification of the RecA protein. When the inducing
signal continues to accumulate, recA will be induced, resulting in a full induction of SOS regulon genes. Meanwhile, due
to the repression of lexA by LexA itself, the SOS response
system is robust to prevent substantial induction caused by
very small amounts of inducing signal. Furthermore, the
strong repression of recA by LexA and the constant expression of LexA in SOS response ensure a fast return to the
uninduced state once the level of the inducing signal begins
to decrease (Walker, 1984).
In addition to the two central regulators RecA and LexA,
recent studies reveal that some other proteins are also
involved in the subtleties of SOS induction. All these
proteins appear to affect SOS regulation by modulating
RecA-ssDNA stability. RecX can block the assembly of the
RecA-ssDNA filament while not affecting the disassembly
through capping the assembly ends (Drees et al., 2004). As a
result, it strongly inhibits RecA-mediated DNA strand
exchange, ATPase and coprotease activities. The recX gene
is located 76-bp downstream of recA, and these two genes
belong to the same operon. Although recX is cotranscribed
with recA, recX transcription is downregulated with respect
to recA by an intrinsic transcription terminator that is
located between the recA and recX coding sequences. Despite
the presence of this terminator, a recA–recX message resulting from transcriptional read-through is detected at a level
of 5–10% of the recA message (Pages et al., 2003). RecX is
barely detected during vegetative growth, but robust expression of recX is observed after treating cells with DNAdamaging agents (Stohl et al., 2003). The maximal recX
expression is observed at a later time than maximal expression of RecA after UV irradiation (Courcelle et al., 2001). All
these observations suggest that RecX is likely involved in the
subtle regulation that helps shut off the SOS response.
Early studies have reported that DinI can destabilize the
RecA-ssDNA filament when its concentration is 50–100-fold
above the natural RecA concentration, and it inhibits all
activities of RecA (Yasuda et al., 1998; Voloshin et al., 2001).
Therefore, DinI was initially thought to aid the return of
SOS-induced cells to a steady state. In contrast, recent
research has shown that DinI is also able to stabilize the
RecA-ssDNA filament to prevent its disassembly when
present at concentrations that are stoichiometric with or
somewhat greater than those of RecA (Lusetti et al., 2004).
Furthermore, DinI-mediated stabilization affects RecAmediated UmuD cleavage rather than RecA-mediated ATP
hydrolysis and LexA coprotease activities (Lusetti et al.,
2004), indicating that DinI could have a biological role in
fine-tuning the activity of UmuD in order to limit SOS
mutagenesis.
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Y. Fu et al.
Role of DNA helicases and nucleases in SOS
induction
A critical step in the SOS response is the production of a
RecA-ssDNA filament, and this step requires DNA helicases
and nucleases. Either the RecFOR or the RecBCD pathway is
necessary for the SOS response after UV irradiation
(Ivancic-Bace et al., 2006). In E. coli, SOS induction immediately after UV irradiation is dependent on the RecFOR
pathway. The RecFOR pathway includes the DNA helicase
RecQ, the nuclease RecJ and the RecFOR complex, which
facilitates RecA loading (Lusetti et al., 2006). It has been
suggested that the pathway may aid RecA binding to ssDNA
gaps. RecQ appears to be needed for fast degradation of the
LexA repressor (Hishida et al., 2004). This observation leads
to a model in which RecQ unwinds the template duplex in
front of a stalled fork on the leading strand, and then
switches over to the lagging strand to generate ssDNA on
the leading strand template, allowing formation of the RecA
filament in the 5 0 –3 0 direction for SOS induction (Heyer,
2004; Hishida et al., 2004).
RecBCD, also known as Exonuclease V, contains helicase,
5 0 –3 0 exonuclease, and RecA-loading activities. It can directly load RecA on the processed double-strand break
(DSB) ends (Singleton et al., 2004). SOS induction after
UV irradiation in recFOR mutants is not completely eliminated but is delayed, and this induction is dependent on the
RecBCD enzyme (Thoms & Wackernagel, 1987; Hegde et al.,
1995; Whitby & Lloyd, 1995; Renzette et al., 2005). Accordingly, it has been proposed that SOS induction requires
RecBCD when the DSB ends appear later because of NER
and replication fork collapse after UV irradiation (IvancicBace et al., 2006).
SOS-independent DNA damage induction in
bacteria
Bacteria also possess damage-inducible systems that are
independent of the SOS response. For example, alkylating
or oxidative agents induce not only the SOS response but
also other more specific DNA damage responses.
Following exposure to a sublethal dose of an alkylating
agent (e.g. N-methyl-N 0 -nitro-N-nitrosoguanidine, MNNG),
bacteria such as E. coli manifest a pronounced resistance to
both lethal and mutagenic effects caused by a much higher
dose of the same or a similar alkylating agent (Jeggo et al.,
1977). The resistance is due to the induction of one or more
genes in response to low levels of the alkylating agents; the
protein products of these genes can directly remove the
alkylated adducts in DNA (Samson & Cairns, 1977). Therefore, this regulatory pathway is called the adaptive response
to alkylation damage.
In E. coli, the Ada protein possesses dual functions and
plays a pivotal role in the adaptive response: it is a
FEMS Microbiol Rev 32 (2008) 908–926
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DNA damage-induced transcription in budding yeast
methyltransferase that removes alkyl groups from the
methylated base and a strong transcriptional activator of
several genes. The 39-kDa Ada protein is comprised of
two functional domains linked by a hinge region. The
C-terminal 19-kDa domain is capable of acquiring the
methyl group from a mutagenic O6-methylguanine or
O4-methylthymine to its Cys321 residue. The N-terminal
20-kDa domain specifically catalyzes the transfer of methyl
groups from methyl phosphotriester lesions to its Cys38
residue in a direct and irreversible way (Demple et al., 1985;
Nakabeppu & Sekiguchi, 1986; Teo et al., 1986; Lindahl
et al., 1988; Myers et al., 1993b; Takinowaki et al., 2006).
Methylation of the Cys38 triggers a conformational change
in the Ada protein, thereby dramatically (up to 1000-fold)
enhancing the sequence-specific DNA-binding affinity of
the Ada protein to the promoter regions of its own gene,
ada, and other alkylation resistance genes including alkA,
aidB and alkB (Teo et al., 1986; Sakumi & Sekiguchi, 1989;
Akimaru et al., 1990; Myers et al., 1993a; Sakashita et al.,
1993; Takinowaki et al., 2006). The transcriptional regulatory element (Ada box) in the ada gene to which methylated
Ada binds has been defined precisely, with the sequence 5 0 AAAGCGCA-3 0 . Methylated Ada also displays binding affinity for the sequences 5 0 -AAANNAAAGCGCA-3 0 and 5 0 AAT(N)6GCAA-3 0 in the alkA and aidB promoter regions,
respectively (Teo et al., 1986; Nakamura et al., 1988; Landini
& Volkert, 1995).
Because the alkylation of Ada is irreversible, how is the
adaptive response switched off? Several mechanisms have
been proposed. One suggests that in the absence of alkylation adducts, the activated Ada is simply diluted by cell
divisions (Lindahl et al., 1988). Research has revealed that
physiologically relevant high concentrations of unmethylated Ada are able to inhibit the activation of ada transcription by methylated Ada, both in vitro and in vivo (Saget &
Walker, 1994). It is also possible that activated Ada is
proteolytically degraded. The proteolytic cleavage of activated Ada in the hinge region linking the N-terminal to
C-terminal domain of the protein might downregulate the
expression of ada. Consistent with this model, a methylated
20 kDa Ada can bind to the ada promoter; however, it does
not facilitate further binding of RNA polymerase to the
promoter nor does it promote ada transcription in vitro
(Akimaru et al., 1990).
The expression of genes responding to oxidative DNA
damage can be induced by the presence of reactive oxygen
species (ROS). Two regulatory responses induced by ROS
have been identified in E. coli – one controlled by soxRS and
the other by oxyR. In the soxRS regulatory system, SoxR
serves both as a sensor and as an activator. In the presence of
ROS, SoxR forms 2Fe–2S centers, which convert SoxR to an
active state. The activated SoxR induces transcription of
soxS, a positive regulator that stimulates transcription of
FEMS Microbiol Rev 32 (2008) 908–926
superoxide-responsive genes. Upon relief of oxidative stress,
SoxR is rapidly converted to its transcriptionally inactive
form, thus turning off the response (Wu & Weiss, 1992;
Hidalgo & Demple, 1994; Hidalgo et al., 1995; Ding et al.,
1996; Gaudu et al., 1997; Volkert & Landini, 2001). The oxyR
regulatory system is in response to hydrogen peroxide
(H2O2). H2O2 activates the transcriptional activity of OxyR
through formation of a disulfide bond between two of its
cysteine residues. Activated OxyR induces the transcription
of several oxidative stress genes (Zheng et al., 1998; Aslund
et al., 1999; Volkert & Landini, 2001).
Transcriptional responses to DNA damage
in S. cerevisiae
From the above analyses, it appears that bacteria possess
transcriptional responses to specific types of DNA lesions as
well as general DNA damage, and that these regulatory
pathways are fairly well understood. In contrast, DNA
damage response in eukaryotes is apparently more complex
than in prokaryotes. Rather than direct cleavage of transcription repressors, eukaryotes most likely use posttranslational modifications, such as phosphorylation and
ubiquitination, and a sophisticated signal transduction
cascade to achieve gene regulation in response to DNA
damage. Here, we attempt to summarize the current knowledge of mechanisms of DNA damage response in S. cerevisiae, a unicellular lower eukaryotic model organism.
Following a historic overview of identification and characterization of genes whose transcript levels are altered following treatment of cells with DNA-damaging agents, this
review focuses on regulatory responses that are either
common or unique to some well-studied genes.
Genome-wide DNA damage-induced expression
studies in yeast
Following work in prokaryotes revealing that DNA damage
causes alterations in the transcriptional program of a cell, an
important initiative was to identify a comparable response
in the lower model eukaryote, S. cerevisiae. Although some
of these experiments were performed one gene at a time, a
precedent for systemic studies of transcription in bacteria
(Kenyon & Walker, 1980) and the emergence of highthroughput technologies spurred the use of large-scale
screens for such studies in yeast. To originally identify some
of the genes that might be induced in response to DNA
damage in S. cerevisiae, two early studies utilizing different
genomic screens for DNA damage-induced genes are most
notable. The first was based on a transcriptional fusion
technique that is still widely used today for routine expression analysis. Using an experimental approach used previously in E. coli to screen for DNA damage-inducible genes
(Kenyon & Walker, 1980), Szostak and colleagues tested
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912
yeast genes as a random pool of lacZ fusions and screened
for coordinately regulated DNA damage-inducible genes
over various DNA damage treatments. From c. 8000 independent clones, a total of six damage-inducible (DIN) genes
were identified, including one gene from a previous screen
(Ruby et al., 1983; Ruby & Szostak, 1985). Based on different
expression profiles of DIN genes to specific DNA-damaging
agents, the authors proposed at least two DIN classes. Such
grouping of inducible genes is analogous to the clustering of
large-scale data sets that is often necessary to manage
and interpret modern DNA microarray experiments. Meanwhile, McClanahan & McEntee (1984) used a differential
plaque hybridization strategy to identify genes whose transcript levels are altered after DNA damage treatment.
Radiolabeled cDNA probes were generated from poly(A)
mRNA harvested from control and treated cells, and
subsequently used for differential plaque hybridization.
By comparing the intensity of labeled probe bound between
the control and the treated samples, differential expression
of genes could be evaluated. Altogether, c. 9000
genomic clones were screened and four DNA damageresponsive (DDR) genes were identified. The experimental
approach also enabled the identification of another set of
genes, with decreased expression in response to DNA
damage treatment. This latter finding testifies to the
complexity of the DNA damage response in eukaryotes,
because no such deactivation of genes was ever found in the
E. coli SOS response (Walker, 1984).
These two early screens were significant to the field of
transcriptional response to DNA damage in yeast. Not only
were they among the first to demonstrate altered levels of
transcripts in S. cerevisiae after DNA damage insult but also
they were carried out in a genome-wide manner and
foreshadowed some of the benefits and complications
associated with analogous DNA microarray experiments
that would follow. The powerful yet straightforward experimental global approaches also paid dividends by considerably expanding the number of genes known to be induced by
DNA damage at the time. Together, 10 new DNA damageinducible (DIN/DDR) and four DNA damage-repressive
genes were identified. These studies allowed estimation of
total DNA damage-responsive genes. Because the genomic
library of Ruby and Szostak likely contained only c. 500 yeast
genes representing about 8% of the genome, extrapolation
led to an estimation of roughly 80 DNA damage-inducible
genes in yeast. Significantly, the two independent screens
yielded nonoverlapping data sets, indicating that neither
screen was saturated and that the number of DNA damageinducible genes was probably underestimated. Thus, even in
the earliest stages of the studies of yeast DNA damageinduced gene expression, it was obvious that S. cerevisiae
would have a much larger repertoire of DNA damageinduced genes than E. coli.
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Y. Fu et al.
With the exception of these studies, the identification
of DNA damage-inducible genes in the premicroarray era
was performed on a gene-by-gene basis, and the entire
complement of DNA damage-inducible genes grew very
slowly. A comprehensive list of these genes, including
relevant references, has been compiled and tabulated
(Friedberg et al., 1995) with references herein (McDonald
et al., 1997; Basrai et al., 1999; Bennett, 1999; Brusky
et al., 2000; Fry et al., 2003; Zaim et al., 2005; Fu & Xiao,
2006).
DNA microarray analyses in S. cerevisiae
Despite the above large-scale experimental approaches, a
plausible high-throughput and truly global approach was
not available until the emergence of genomic technologies.
In particular, DNA microarrays have provided the majority
of damage-induced gene expression data in budding yeast to
date. This wealth of expression data provided by DNA
microarrays has revealed a new realm for conceptualizing
regulatory networks in a global manner. In the following
paragraphs, we highlight only the first DNA microarray
studies of DNA damage-induced expression in yeast, and
note that there are numerous other significant studies that
cannot be feasibly covered in this review. Table 1 is provided
for an overview of genomic expression studies in S. cerevisiae focusing on DNA damage treatments or related gene
mutants. We refer the reader to the Yeast Functional
Genomics Database (http://yfgdb.princeton.edu/) for a navigable compilation of these and related studies, 93 of which
are currently listed as genomic expression studies involving
stress.
The premiere microarray study on DNA damage-induced
expression in S. cerevisiae was published in 1999 (Jelinsky &
Samson, 1999). Representative sequences of 6218 yeast ORFs
were arrayed and cRNA probes were generated from untreated
cultures or cultures exposed to the DNA-alkylating agent
methyl methanesulfonate (MMS). As evidenced by the overlap
with the previous independent data mentioned above, the
results validated the use of DNA microarrays for damageinduced expression analysis. Of the genes already known to be
induced by DNA-damaging agents by all other methods
combined, 86% (18 out of 21) were identified in the DNA
microarray experiment. The use of a single MMS dose and a
fourfold minimum cut-off likely explain the omission of the
three other previously identified inducible genes. Altogether, an
astonishing 325 yeast genes (c. 5% of entire yeast genes) were
found to be induced by MMS treatment, with a significant
portion (112) representing uncharacterized ORFs. In addition,
76 ORFs had reduced expression upon MMS exposure. The
reduced expression of some genes after DNA damage treatment
is reminiscent of the early DDR screen (McClanahan &
McEntee, 1984). Altogether, a single DNA microarray
FEMS Microbiol Rev 32 (2008) 908–926
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DNA damage-induced transcription in budding yeast
Table 1. Genomic DNA microarray studies of transcriptional response to
DNA-damaging agents and DNA repair-associated mutations in Saccharomyces cerevisiae
References
Cell type
Jelinsky & Samson (1999)
Jelinsky et al. (2000)
Haploid
Haploid
Gasch et al. (2000)
Gasch et al. (2001)
De Sanctis et al. (2001)
Haploid
Haploid
Haploid
Haploid
Haploid
Haploid
Haploid
Oshiro et al. (2002)
Ostapenko & Solomon
(2003)
Green & Johnson (2004)
van Attikum et al. (2004)
Haploid
Haploid
Keller-Seitz et al. (2004)
Diploid
Mercier et al. (2005)
John et al. (2005)
All types
Haploid
Benton et al. (2006)
Guo et al. (2006)
Kelly et al. (2006)
Haploid
Diploid
Haploid
Haploid
Haploid
Marques et al. (2006)
Shenton et al. (2006)
Workman et al. (2006)
Kugou et al. (2007)
Haploid
Haploid
Haploid
Diploid
Fu et al. (2008)
Haploid
Mutant
rpn4
dun1
mec1
crt1
rad53,
rad6
ctk1
tup1
ino80,
arp8
mec1
30 TFs
mre11,
rad50
rad6,
rad18
Treatment
MMS
MMS, MNNG,
BCNU, t-BuOOH,
4-NQO, IR, MMC
MMS
H2O2
MMS, IR, H2O2
MMS
MMS, IR
NA
IR
MMS, HU, UV,
HU
NA
MMS
Aflatoxin B1
(N7-guanine
DNA adducts)
IR
Cigarette smoke
extract
MMS, IR
Aflatoxin B1
Azinomycin B
(interstrand
crosslinks and
alkylating agent)
H2O2
H2O2
MMS
NA
MMS
MMS, methyl methanesulfonate; MNNG, N-methyl-N 0 -nitro-N-nitrosoguanidine; BCNU, 1,3-bis(2-chlorothyl)-1-nitrosourea; t-BuOOH, tertbutyl hydroperoxide; 4-NQO, 4-nitroquinoline-n-oxide; IR, ionizing radiations; MMC, mitomycin C; HU, hydroxyurea; UV, ultraviolet irradiation.
NA, not applicable.
experiment augmented the complement of known DNA damage-induced genes in S. cerevisiae by over 20-fold. Despite this
wealth of new data, however, meaningful conclusions regarding
transcriptional regulation were not possible with such an
abundance of raw, unorganized data.
A follow-up study (Jelinsky et al., 2000) provided a more
comprehensive analysis of transcriptional response to DNA
damage and demonstrated how large amounts of microarray
data can be manageably interpreted. This enabled the use of
a large dataset to address a specific hypothesis, namely that
FEMS Microbiol Rev 32 (2008) 908–926
MAG1 belongs to a novel regulatory network. The DNA
microarray data were computationally assembled into selforganizing maps (SOMs) (Tamayo et al., 1999) that group
genes with similar expression profiles. Using SOMs, 18
different groups of genes were coregulated across various
experimental conditions. Of the genes in the SOM containing MAG1, it was noted that most contained an upstream
repressor sequence 2 element (URS2) previously identified
upstream of several genes involved in DNA repair and
metabolism (Xiao et al., 1993; Singh & Samson, 1995). It
was thus hypothesized and resolved that the genes within the
MAG1 SOM were regulated at the transcriptional level by
the same repressor element and constitute a regulatory
network. This study identified Rpn4 as a putative transcriptional regulator of a large number of genes in response to
DNA damage. The fact that Rpn4 serves as a 26S proteasome-associated protein as well as a transcriptional factor
that regulates at least 26 out of 32 proteasomal genes
(Mannhaupt et al., 1999) points to a possibility that
ubiquitin-mediated proteolysis may play a role in the
transcriptional response to DNA damage. Other studies
on the transcriptional response to DNA damage in yeast
using DNA microarray experiments have successfully applied similar methods of data management to test certain
hypotheses (see Table 1).
DNA damage checkpoint pathways and
transcriptional regulation
Both individual gene analyses and global transcriptional
studies in budding yeast indicate that many genes are
coordinately regulated in response to DNA damage.
Although transcriptional responses to specific damaging
agents result in distinct and signature profiles, the microarray data point to a general stress response pathway that
controls transcription. These observations are consistent
with previous reports based on individual gene analyses that
DNA damage and replication checkpoints are involved in a
transcriptional response to DNA damage.
The DNA damage checkpoint mutants were first isolated
by their failure to delay cell-cycle progression into mitosis
after irradiation with X-rays (Weinert & Hartwell, 1988;
Rowley et al., 1992; Weinert et al., 1994). These mutants
displayed increased radiation sensitivity, which could be
largely negated through reimposing an artificial arrest before
the M phase by an antitubulin agent. Hence, DNA damage
checkpoint was initially defined as a nonessential and
reversible response that slows down or arrests cell-cycle
progression in response to DNA damage, allowing time for
DNA repair (Hartwell & Weinert, 1989). At present, about
20 genes in budding yeast have been identified or anticipated to be involved in DNA damage checkpoints (Table 2)
(Elledge, 1996; Nyberg et al., 2002; Friedberg et al., 2006).
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Y. Fu et al.
Table 2. DNA damage sensors and signal transducers in Saccharomyces cerevisiae and their Schizosaccharomyces pombe and mammalian orthologs
Category
Protein function
S. cerevisiae
S. pombe
Mammals
Damage sensors
RFC-like clamp loader
Clamp loader
PCNA-like clamp
S-phase sensors
Pole
Rad24
Rfc2-5
Ddc1
Rad17
Mec3
Pol2/Dun2
Dpb11
Drc1/Sld2
Sgs1
Mec1
Lcd1/Ddc2
Tel1
Rad9
Mrc1
Tof1
Csm3
Rad53
Chk1
Pds1
Dun1
Rad17
Rfc2-5
Rad9
Rad1
Hus1
Cdc20
Cut5/Rad4
Drc1
Rqh1
Rad3
Rad26
Tel1
Crb2
Mrc1
Swi1
Swi3
Cds1
Chk1
Cut2
Unknown
RAD17
RFC2-5
RAD9
RAD1
HUS1
Pole
TopBP1
Unknown
WRN, BLM, RTS
ATR
ATRIP
ATM
BRCA1, MDC1, 53BP1
Claspin
Unknown
Unknown
CHK2
CHK1
PTTG
Unknown
Sensor/transducer
Transducers
DNA helicase
PI3K-like kinases
ATR partner
PI3K-like kinases
Damage mediator
S-phase mediator
Kinase
Kinase
Effector kinase
These genes are required for different phases of the cell cycle
and for different types of lesions. Because inactivation of
damage checkpoint genes affects the transcriptional response to DNA damage as well as other cell fates, it has been
well accepted that checkpoint pathways play critical roles in
addition to cell-cycle arrest (Zhou & Elledge, 2000).
It is now clear that the checkpoint pathway comprises
a subroutine integrated into the larger DNA damage
response that regulates multifaceted responses (Zhou &
Elledge, 2000). Besides arresting cell-cycle progression, the
damage checkpoint pathway has been shown to promote
DNA repair (Mills et al., 1999), control telomere length
(Ritchie et al., 1999), activate transcription (Elledge, 1996)
and trigger apoptosis in metazoan cells (Roos & Kaina,
2006). In a broad sense, the damage checkpoint coordinately
regulates DNA damage responses including transcriptional
regulation.
Based on the data from previous research, a putative
model is proposed for the signal transduction pathways in
response to DNA damage, consisting of sensors, transducers
(including mediators) and effectors (Bachant & Elledge,
1998). The sensors are proteins that initially sense the
damaged DNA and initiate the signaling response. Transducers can be activated by the DNA damage signal passed
down from the sensor’s and then amplify and relay the signal
to the downstream effectors. The ultimate effectors are
defined as proteins that execute the cellular response. In the
case of transcriptional regulation, the effectors are likely to
be transcription factors that recognize cis-acting elements in
the promoter of DNA damage-inducible genes.
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DNA damage sensors in S. cerevisiae
Because of the complexity of the source and type of DNA
damage, as well as the different cell-cycle stages at which
damage may occur, DNA damage sensors and their intrinsic
interactions have not been well established. Much of our
knowledge comes from studies with damage and replication
checkpoints.
The budding yeast proliferating cell nuclear antigen
(PCNA)-like 9-1-1 complex composed of Rad17, Ddc1 and
Mec3 is loaded on the DNA damage site with the assistance
of the clamp loader Rad24–Rfc complex, which leads to
activation of the main damage response kinase Mec1
(Kondo et al., 1999; Rouse & Jackson, 2002; Majka &
Burgers, 2003; Majka et al., 2006). Therefore, Rad24–Rfc
and the 9-1-1 complexes together have been speculated to be
sensors. Mec1 forms a stable complex with Lcd1/Ddc2, and
this complex itself has been anticipated to be a sensor due to
the ability of Ddc2 to bind to ends of single- and doublestranded oligonucleotides in vitro and to recruit Mec1 to
double-strand break (DSB) sites in vivo in a manner
independent of the 9-1-1 complex (Kondo et al., 2001; Melo
et al., 2001; Rouse & Jackson, 2002; Niida & Nakanishi,
2006). Another sensor candidate is Rad9. Rad9 is required
for the activation of DNA damage checkpoint pathways in
budding yeast, is phosphorylated after DNA damage in a
Mec1- and Tel1-dependent manner and subsequently interacts with the downstream kinase Rad53 (Naiki et al., 2004).
Rad9 displays the ability to associate with DSBs and controls
the DNA damage-specific induction of some repair,
FEMS Microbiol Rev 32 (2008) 908–926
915
DNA damage-induced transcription in budding yeast
replication and recombination genes (Aboussekhra et al.,
1996; Naiki et al., 2004). Furthermore, Rad9 is thought to
act in a pathway distinct from that of Rad24/9-1-1 for
damage checkpoint (Lydall & Weinert, 1995) and transcriptional response (de la Torre-Ruiz et al., 1998), suggesting
that Rad9 may serve as an alternative DNA damage sensor.
The above candidate sensors primarily respond to DNA
damage caused by UV, ionizing radiations and endogenous
DSBs that induce G1/S and G2/M checkpoints. They also
respond to excessive telomere ssDNA in the cdc13-1 mutant
(Lydall & Weinert, 1995) and the accumulation of DSBs due
to a defective Cdc9 DNA ligase (Weinert & Hartwell, 1993),
both inducing the G2/M checkpoint.
The intra-S checkpoint is experimentally activated by
modest doses of alkylating agents, such as MMS, that cause
replication–blocking lesions. In addition, inhibiting replication by means other than DNA damage, such as treatment
with the ribonucleotide reductase (Rnr) inhibitor hydroxyurea, can activate similar cellular responses. Allele-specific
mutants in the catalytic subunit of DNA polymerase e (Pol2)
are defective in MMS- and hydroxyurea-induced RNR3
(Navas et al., 1995) and MAG1 (Zhu & Xiao, 1998) expression. Hence, Pol2 and its interacting partners Dpb11 and
Drc1 may serve as S-phase sensors. It is of great interest to
note that Dpb11 has been reported to interact with the Ddc1
subunit of 9-1-1 (Wang & Elledge, 2002), and that physcial
interactions between Pole and the human homologs of
Rad24/9-1-1 were also reported (Makiniemi et al., 2001;
Post et al., 2003), indicating that the damage signal may be
relayed between sensors. Recently, the helicase Sgs1, which is
a member of the E. coli RecQ helicase subfamily that includes
mammalian homologs of human BLM, WRN and RTS,
responsible for the Bloom, Werner and a subset of Rothmund–Thomson syndromes, respectively, was proposed as a
candidate sensor. Sgs1 possesses ATPase activity and preferentially binds to branched DNA substrates (Bennett et al.,
1998, 1999). Sgs1 might be involved in the creation of the
signal for checkpoint activation, perhaps by resolving aberrantly paired double helices (Cobb et al., 2002). It functions
in the same epistasis group as Pole to activate Rad53 in the
presence of hydroxyurea, and this signaling pathway acts in
parallel to that of Rad17 and Rad24 (Navas et al., 1995; Frei
& Gasser, 2000). However, it remains unclear whether
putative S-phase-specific sensors like Pole and Sgs1 recognize a subset of DNA damage or their activities lead to
common DNA damage intermediates such as ssDNA (like
RecA in the SOS response).
The above checkpoint proteins are placed in the sensor class
primarily based on genetic and biochemical properties or
inference, as little direct biochemical evidence yet exists to
support their sensor role. In principle, checkpoint sensors are
capable of directly recognizing DNA lesions or interacting with
pre-existing DNA damage repair proteins specific for indiviFEMS Microbiol Rev 32 (2008) 908–926
dual lesions. In this context, DNA repair proteins may serve as
damage sensors for both checkpoint and/or transcriptional
responses. A good example is the interaction between NER and
DNA damage checkpoint in response to UV treatment. A
genetic screen for mutants defective in the activation of
checkpoint following UV lesions but not other types of damage
resulted in the identification of RAD14, which is absolutely
required for the UV-induced damage checkpoint. Interestingly,
the checkpoint activation requires processing of UV lesions by
the NER complex instead of mere lesion recognition by Rad14,
and Rad14 physically interacts with Ddc1 (Giannattasio et al.,
2004). Similarly, the Mre11/Rad50/Nbs1 (MRN) DSB-binding
complex is required for the activation of ATM kinase in
mammalian cells (Lee & Paull, 2005), although a similar
function has not been reported for the corresponding Mre11/
Rad50/Xrs2 (MRX) complex in budding yeast. In both the
above cases, processing of UV-induced lesions by NER and
DSBs by MRN/MRX results in ssDNA regions that may serve
as an ultimate damage signal.
DNA damage transducers in S. cerevisiae
Considerable efforts have been made to understand signal
transducers in the damage and replication checkpoint pathways, and the same signal transduction cascade likely functions in the transcriptional response to DNA damage. Most
of the transducers are protein kinases that, once activated by
the DNA damage signal passed down from the sensors,
amplify and relay the signal to the downstream effectors. In
S. cerevisiae, checkpoint protein kinases Mec1, Rad53 and
Dun1 are necessary for the transcriptional response of most,
if not all, genes to DNA damage, and they appear to be
central transducers in the regulation network (Zhou &
Elledge, 1993; Allen et al., 1994; Kiser & Weinert, 1996;
Gasch et al., 2001). The initiator of this signal transduction
kinase cascade in budding yeast appears to be Mec1 and its
regulatory subunit Lcd1/Ddc2, which is a serine/threonine
protein kinase belonging to the phosphatidylinositol-3kinase (PI3K) family. Mec1 is required for the activation of
Rad53 through phosphorylation of consensus PI3K sites
within Rad53 (Pellicioli & Foiani, 2005; Ma et al., 2006).
However, efficient and direct phosphorylation of Rad53
by Mec1 is only observed in the presence of Rad9 in vitro,
and the stimulatory activity of Rad9 requires both phospho- and FHA-dependent interaction with Rad53, which
allows Rad53 to be recognized as a substrate for Mec1
(Sweeney et al., 2005). Hence, Rad9 is defined as a
checkpoint mediator (or adaptor) that serves to facilitate
and amplify signals. Rad9 functions primarily at G1/S and
G2/M, while Mrc1 and its mammalian homolog claspin
has been proposed to function as an S-phase or a replication checkpoint mediator. Mrc1 and Tof1 form a stable
replication-pausing complex at the stalled replication fork,
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which is required to anchor subsequent DNA repair events
(Katou et al., 2003).
Activation of Rad53 is an essential intermediary step in
yeast DNA damage responses that include delaying cell-cycle
progression, promoting repair processes, stabilizing stalled
replication forks and regulating transcription (Branzei &
Foiani, 2006). The kinase Dun1 is one of the identified
targets of Rad53, and the activation of Dun1 requires
phosphorylation by Rad53 (Chen et al., 2007). Dun1 was
originally identified as a DNA damage-uninducible (dun)
mutant defective in the transcriptional activation of genes
encoding Rnr in response to DNA damage (Zhou & Elledge,
1993), and was subsequently shown to be required for the
induction of other DNA damage-inducible genes (Basrai
et al., 1999; Zhu & Xiao, 2001; Fu & Xiao, 2006). Genomic
expression research showed that deletion of DUN1 affected
the expression of 4 1000 genes in response to MMS, and
the response in the dun1D mutant is largely the same as the
response seen in the mec1 mutant (Gasch et al., 2001),
suggesting that most of the Mec1-dependent effects on
genomic expression are mediated by the downstream Dun1
kinase. The mechanism used by Dun1 for DNA damage
induction appears to be rather diverse and requires further
investigation.
In response to DNA damage, the primary mechanism for
transcriptional regulation is likely to be at the stage of
transcription initiation. The downstream effectors are therefore expected to be transcription factors that directly influence transcription initiation. However, unlike damage
checkpoint transducers that appear to be aligned in a linear
cascade and control most if not all DNA damage-inducible
genes, the downstream effectors are clearly diverse and
dedicated to a particular gene or a group of genes (Fig. 1).
In the next section, selected examples are presented to
provide insights into the molecular mechanisms of DNA
damage-induced transcriptional response and their interaction with the checkpoint cascade.
Mechanisms of transcriptional regulation
of yeast genes in response to DNA
damage
RNR genes
Regulation of RNR genes is the best-known example of a
eukaryotic transcriptional response to DNA damage to date.
Rnr is an enzyme that converts nucleoside diphosphates
(NDPs) into deoxynucleoside diphosphates (dNDPs),
which represents the rate-limiting step in the production of
the four dNTPs for DNA synthesis and repair (Reichard,
1988; Elledge et al., 1993; Jordan & Reichard, 1998). Altered
levels or an imbalance of dNTP pools can lead to a higher
rate of spontaneous mutagenesis or cell death. Rnr is an
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Y. Fu et al.
Fig. 1. A working model of DNA damage-induced transcription in
budding yeast. In budding yeast, DNA damage-induced transcription is
controlled by signal transduction pathways including sensors, transducers and effectors. Mec1, Rad53 and Dun1 form the central kinase
cascade to regulate the DNA damage-induced transcription of most
genes examined. The phosphorylation of trans-acting factors (effectors)
may change their affinity for the corresponding cis-acting elements in
the promoter region of target genes, thus activating their transcription.
Only three sets of well-characterized genes are depicted. Solid ovals
indicate protein kinases. Dotted lines and question mark indicate that the
molecular mechanism of this step(s) remains unknown. Note that the
signal-sensor-transducer cascades are based on studies with cell-cycle
checkpoints and have not been extensively examined with transcriptional
regulation.
a2b2 tetramer, and four genes, RNR1-4, encode the subunits
of budding yeast Rnr: RNR1 and RNR3 encode a (large)
subunits, while RNR2 and RNR4 encode b (small) subunits.
Expression of all four genes is inducible at the transcriptional level by a variety of DNA-damaging agents (Elledge
& Davis, 1987, 1989, 1990; Huang & Elledge, 1997), among
which RNR3 exhibits the highest level of induction, and
its transcriptional regulation has been examined most
extensively.
The transcriptional level of the RNR3 gene is very low
under normal conditions. However, when treating yeast cells
with DNA-damaging agents such as UV, MMS or 4-nitroquinoline-1-oxide (4-NQO), the transcript level of RNR3
can be increased 100–500-fold (Elledge & Davis, 1990). In
order to determine the mechanisms of induction, a series of
mutants have been isolated that cause constitutive expression of RNR3 (crt mutants) (Zhou & Elledge, 1992). These
negative regulators of RNR3 expression are divided into two
groups: indirect regulators that result in endogenous DNA
damage or a state of metabolic stress such as nucleotide
depletion that results in the upregulation of RNR3, and
FEMS Microbiol Rev 32 (2008) 908–926
917
DNA damage-induced transcription in budding yeast
Pol
DNA damage
Stalled replication
Mec1
P
P
Rad53
P
Crt10
Dun1
P
P
Crt1
Crt1
CRT1 mRNA
Ccr4
Crt1
X-box
RNR3
X-box
Wtm2
CRT1
Rpd3
X-box
Hos2
RNR3
Crt1
Fig. 2. Summary of the transcriptional regulation of RNR3. In response to DNA damage or stalled replication, activation of the Mec1-Rad53-Dun1
kinase cascade results in phosphorylation of the repressor protein Crt1, and phosphorylated Crt1 loses its binding affinity for the RNR3 promoter. With
the assistance of Wtm1, Rpd3 and Hos2, the transcription of RNR3 is highly activated. The transcription of CRT1 is negatively regulated by itself and
positively regulated by Crt10. Ccr4 influences Crt1 protein abundance by controlling the CRT1 mRNA stability.
direct regulators involved in the regulatory pathway, including transcription factors. The CRT1, TUP1 (CRT4) and
SSN6 (CRT8) genes encode negative regulators that bind to
the RNR3 promoter. A second screen was carried out for the
mutations that disrupt the ability of DNA damage to induce
transcription of RNR3, and the corresponding genes were
designated DUN for DNA damage uninducible (Zhou &
Elledge, 1993). The nonessential serine/threonine protein
kinase Dun1 was isolated through this screen. Genetic
analysis of crt1, tup1 and ssn6 showed that these mutations
were epistatic to dun1, providing a strong genetic verification that CRT1, TUP1 and SSN6 function downstream of
DUN1 (Huang et al., 1998). Combined with observations
that the Dun1 upstream kinases Mec1 and Rad53 are also
essential for the DNA damage-induced transcription of
RNR3 (Huang et al., 1998), the signal transduction pathway
for RNR3 becomes conceivable (Fig. 2).
It is now clear that in response to DNA damage or
replication blocks, the Rad53 protein kinase is activated in
a Mec1-dependent manner, and activated Rad53 further
phosphorylates the protein kinase Dun1. The Mec1-Rad53Dun1 kinase cascade culminates in the phosphorylation of
Crt1. Crt1 is a DNA-binding protein that recognizes a 13-bp
consensus sequence termed the X box. Multiple X boxrelated sequences of different strength can be found in the
FEMS Microbiol Rev 32 (2008) 908–926
promoter region of all four RNR genes. Crt1 is able to
recruit the Tup1–Ssn6 general repressor complex to suppress
the transcription of RNR genes. In the presence of DNA
damage or replication blocks, Crt1 is phosphorylated in a
Dun1-dependent manner and this phosphorylation
abolishes its ability to bind X boxes, leading to the transcriptional induction of RNR genes (Huang et al., 1998).
Surprisingly, multiple X boxes were also identified in the
CRT1 promoter. Data from mutated X boxes showed that
they confer CRT1-dependent repression on CRT1 itself
(Huang et al., 1998). The expression level of CRT1 is very
low under normal growth conditions but is inducible by
DNA-damaging agents. Interestingly, the X boxes in the
CRT1 promoter consist of one with a weak affinity for Crt1
and one with a strong affinity. Thus, it is speculated that a
weak induction of CRT1 occurs immediately following DNA
damage, which provides a buffer against spurious transcriptional activation of the pathway. Delaying full activation
may ensure a rapid restoration of the basal repressed state
when the DNA damage is repaired (Huang et al., 1998),
which may serve as an autonomous regulatory circuit (Fig.
2). This mechanism is reminiscent of the autonomous
regulation of the LexA repressor in the E. coli SOS response.
Crt1 mediates repression by recruiting the general repressor Tup1–Ssn6 complex to RNR promoters via its
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918
N-terminus (Huang et al., 1998; Li & Reese, 2000). Tup1Ssn6 recruitment establishes a nucleosomal array over the
promoter of RNR3 with a positioned nucleosome occupying
the TATA box to block access by the general transcriptional
machinery (Li & Reese, 2000, 2001; Sharma et al., 2003). The
derepression of RNR3 correlates with the disruption of the
nucleosome position. In response to DNA damage signals,
hyperphosphorylated Crt1 loses the ability to bind to the
RNR3 promoter, and the Tup1–Ssn6 complex is not recruited to the promoter region. Consequently, the chromatin
structure is remodeled and thus increases the accessibility of
DNA to transcription factors. The chromatin remodeling at
the RNR3 promoter requires a number of general transcriptional factors, such as TBP-associated factors (TAFIIS), and
RNA polymerase II. Furthermore, the remodeling is also
dependent on the SWI/SNF complex, which possesses a
DNA-stimulated ATPase activity and can destabilize histone–DNA interactions in an ATP-dependent manner (Sharma
et al., 2003). The preinitiation complex components TFIID
and RNA polymerase II aid in recruitment and retention of
the SWI/SNF complex to the RNR3 promoter.
Recently, it was reported that the expression of CRT1 is
also controlled by the novel regulator Crt10 (Fu & Xiao,
2006) (Fig. 2). CRT10 was identified through screening of
the S. cerevisiae deletion strains for hydroxyurea resistance.
Epistatic analysis indicates that CRT10 belongs to the CRT1
pathway. Deletion of CRT10 does not affect the transcript
level of TUP1 or SSN6 regardless of hydroxyurea treatment,
but significantly reduces the basal level as well as hydroxyurea-induced expression of CRT1; thus, CRT10 appears to be a
positive regulator of CRT1 transcription (Fu & Xiao, 2006).
Furthermore, the dun1 mutation is epistatic to crt10 with
respect to both hydroxyurea sensitivity and RNR gene
expression. Interestingly, the expression of CRT10 itself is
induced by DNA-damaging agents and this induction
requires DUN1 (Fu & Xiao, 2006). The increased Crt10
activity may be required to bring Rnr activity back to a
normal level through increasing the expression of repressor
Crt1 once the DNA damage is repaired. Like CRT1, the
induction of CRT10 itself depends on DUN1, suggesting
that Crt10 functions downstream of Dun1 and forms
another component of the autoregulatory circuit to regulate
the expression of RNR genes (Fu & Xiao, 2006).
CCR4 encodes a component of the major cytoplasmic
deadenylase, which is involved in mRNA poly(A) tail shortening (Tucker et al., 2001). Cells defective in CCR4 display
particular sensitivity to the Rnr inhibiter hydroxyurea
(Woolstencroft et al., 2006). The ccr4 dun1 double mutants
exhibit synergistic sensitivity to hydroxyurea, and simultaneous overexpression of RNR2, RNR3 and RNR4 partially
rescues the hydroxyurea hypersensitivity of a ccr4 dun1
strain, implying that CCR4 and DUN1 function in different
pathways to regulate the activity of Rnr. Deletion of CRT1
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Y. Fu et al.
suppresses hydroxyurea sensitivity of the ccr4 mutant, and
overexpression of CRT1 hypersensitizes ccr4 to hydroxyurea.
These observations lead to the conclusion that Ccr4 regulates CRT1 mRNA poly(A) tail length and may thus subtly
influence the Crt1 protein abundance (Woolstencroft et al.,
2006) (Fig. 2).
More regulatory factors were found to be involved in the
transcriptional regulation of RNR genes. Wtm1 and Wtm2
have been reported to modulate the expression of RNR3
(Tringe et al., 2006). Moderate overexpression of both genes
or high-level expression of WTM2 alone upregulates RNR3lacZ in the absence of DNA damage. In response to hydroxyurea and g-rays, the expression level of RNR3 attenuated
45% in wtm2D mutants, but not in wtm1 mutants. Wtm2
was found to associate directly with the RNR3 promoter,
and the association correlates with its ability to increase
constitutive RNR3 expression. It remains unknown how
Wtm2 increases RNR3 transcription, although some observations hint that Wtm2 might enhance RNR3 transcription
by participating in chromatin remodeling (Tringe et al.,
2006). Furthermore, Wtm1 associates with and anchors the
Rnr2/Rnr4 complex in the nucleus in untreated cells. In
response to DNA damage or replication inhibition, the
interaction between Wtm1 and Rnr2/Rnr4 is disrupted and
this complex is released from the nucleus to the cytoplasm
(Lee & Elledge, 2006; Zhang et al., 2006). A recent report
shows that two histone deacetylases (HDAC), Rpd3 and
Hos2, are required for the transcriptional activation of
RNR3 in response to DNA damage (Sharma et al., 2007)
(Fig. 2). Although a direct association between Hos2 and the
RNR3 promoter has not been observed, Rpd3 was found to
specifically bind to X boxes in the RNR3 promoter region in
a Tup1- or a Crt1-independent manner. The HDAC activity
of Rpd3 and Hos2 is essential for the activation of RNR3.
The observation that the recruitment of RNA polymerase II
is dramatically reduced in the rpd3 hos2 mutant indicates
that Rpd3 and Hos2 activate the transcription by regulating
the assembly of the preinitiation complex or inducing
multiple rounds of RNA polymerase recruitment (Sharma
et al., 2007).
PHR1
PHR1 encodes a photolyase that specifically and exclusively
repairs pyrimidine dimers, which are the most abundant
lesions found in DNA following UV irradiation (Sancar,
1985). Interestingly, the transcription of PHR1 is induced
by various DNA-damaging agents such as MMS, MNNG,
UV irradiation, 4-NQO or g-ray (Robinson et al., 1986;
Sebastian et al., 1990) despite the fact that Phr1 only reverses
UV-induced pyrimidine dimers but not lesions induced by
other DNA-damaging agents. Three transcriptional regulatory elements have been defined within the PHR1 promoter
FEMS Microbiol Rev 32 (2008) 908–926
919
DNA damage-induced transcription in budding yeast
region: an upstream activation sequence (UAS), an upstream repression sequence (URS) and an upstream essential sequence (UES) (Sancar et al., 1995). A 22-bp
interrupted palindrome comprises UASPHRI, and it is responsible for 80–90% of basal and induced expression. It
alone can activate transcription of a CYC1 minimal promoter but does not confer damage responsiveness (Sancar et al.,
1995). URSPHR1 is defined to a 39-bp region that includes a
22-bp palindrome. Deletions or specific mutations within
URSPHR1 increase basal-level expression but decrease the
induction ratio. It functions as a strong URS and confers a
low-level damage inducibility when placed in the context of
a heterologous gene (Sancar et al., 1995). The UESPHR1 is
required for efficient derepression when URSPHR1 is present.
Deletion of URSPHR1 also eliminates the requirement for
UESPHR1 for transcriptional activation (Sancar et al., 1995).
Three proteins have been identified that regulate the
expression of PHR1 by binding to the upstream regulatory
elements. Ume6 is a bifunctional transcriptional regulator
involved in several metabolic pathways (Strich et al., 1994).
It is a positive regulator of PHR1 transcription and binds
specifically to the UASPHR1 (Sweet et al., 1997). Multiple
copies of Ume6 enhance the expression of PHR1; however,
deletion of UME6 reduces the expression of PHR1 during
vegetative growth, but only at a distinct cell-cycle phase
(Sweet et al., 1997; Sancar, 2000). Rph1 and Gis1, which are
35% identical to each other at the amino acid sequence level,
are two DNA damage-responsive repressors of PHR1 transcription (Jang et al., 1999; Sancar, 2000). Both Rph1 and
Gis1 contain two putative zinc fingers that are 4 90%
identical overall and identical in the DNA-binding loop,
and they regulate the transcriptional response of PHR1
through binding to URSPHR1. Deletion of both RPH1 and
GIS1 is required to fully derepress PHR1 in the absence of
damage, suggesting that they are functionally redundant. In
vitro footprinting and binding competition studies indicate
that the sequence AG4 (C4T) within the URSPHR1 is the
binding site for Rph1 and Gis1 (Jang et al., 1999).
Induction of PHR1 is controlled by the DNA damage
signal transduction pathway. The serine and threonine
residues of Rph1 can be phosphorylated, and the phosphorylation of Rph1 is increased in response to DNA damage.
The DNA damage-induced Rph1 phosphorylation requires
DNA damage checkpoint proteins Rad9, Rad17, Mec1 and
Rad53, indicating that the phosphorylation of Rph1 is under
the control of the Mec1-Rad53 DNA damage checkpoint
pathway (Kim et al., 2002). On the other hand, deletion of
DUN1, TEL1 or CHK1 does not affect the phosphorylation
of Rph1, indicating that PHR1 is regulated by a potentially
novel damage checkpoint that is distinct from the Mec1Rad53-Dun1 protein kinase cascade. Based on the results of
a coimmunoprecipitation assay, Rad53 does not appear to
interact physically with Rph1, indicating the existence of a
FEMS Microbiol Rev 32 (2008) 908–926
yet unknown kinase(s) in the Mec1-Rad53-Rph1 pathway
(Kim et al., 2002).
MAG1 and DDI1
MAG1 encodes a 3-methyladenine DNA glycosylase that
initiates the base excision repair pathway by removing lethal
lesions such as 3-methyladenine (Chen et al., 1989). MAG1
is not only induced by DNA-alkylating agents such as MMS
but also by UV, 4-NQO or hydroxyurea (Chen et al., 1990;
Chen & Samson, 1991; Xiao et al., 1993). By analyzing the
DNA sequence immediately upstream of MAG1, another
DNA damage-inducible gene, named DDI1 for DNA Damage Inducible, was identified (Liu & Xiao, 1997). Similar
to MAG1, DDI1 is also induced by MMS, UV, 4-NQO and
hydroxyurea. Furthermore, both genes require a similar
dosage for maximum induction, and the induction profile
is similar (Liu & Xiao, 1997).
MAG1 and DDI1 lie in a head-to-head configuration and
are transcribed divergently. The expression of both MAG1
and DDI1 is controlled by common as well as distinct UAS
and URS elements, possibly through antagonistic mechanisms (Liu & Xiao, 1997). A UASMAG1 and a URSMAG1 have
been identified in the promoter region of MAG1 (Xiao et al.,
1993). Interestingly, fine mapping of the UASMAG1 sequence
reveals that it is located within the DDI1 protein coding
region, and the electrophoretic mobility shift assay using
labeled UASMAG1 as a probe detected sequence-specific
binding protein(s) (Liu & Xiao, 1997), although the identity
of the UASMAG1-binding protein has not been reported. The
expression of DDI1 is negatively regulated by a URSDDI1 in
its promoter region (Liu & Xiao, 1997). The intergenic
region between MAG1 and DDI1 also contains a cis-acting
element that coregulates the expression of both genes. In this
shared promoter region, UASDM, which contains two 8-bp
tandem repeat sequences 5 0 -GGTGGCGA-3 0 , is required for
the bidirectional expression of MAG1 and DDI1; deletion or
point mutations of this tandem repeat result in a reduced
basal level expression and significant reduction of damageinduced expression (Liu & Xiao, 1997). Furthermore,
UASDM alone is able to confer a DNA damage response
when fused to a CYC1 promoter (Liu & Xiao, 1997),
indicating that this cis-acting element independently interacts with a transcriptional factor(s). With a yeast one-hybrid
screen using UASDM as a bait, a transcriptional activator
called Pdr3 was isolated (Zhu & Xiao, 2004). Pdr3 binds to
UASDM in vivo and in vitro, and deletion of PDR3 reduced
both the basal level and the DNA damage-induced expression of MAG1 and DDI1. In addition, deletion of PDR3 does
not further affect MAG1 and DDI1 expression if UASDM is
deleted, indicating that UASDM is indeed the target for Pdr3
activation (Zhu & Xiao, 2004). Another transcriptional
activator, Rpn4, was shown to be required for MAG1
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920
(Jelinsky et al., 2000) and DDI1 (Zhu & Xiao, 2004)
expression; however, Rpn4 does not appear to bind UASDM.
Moreover, deletion of RPN4 does not alter MAG1 and DDI1
expression in the pdr3 mutant cells, suggesting that RPN4
acts upstream of PDR3 (Zhu & Xiao, 2004). Deletion of
PDR3 and RPN4 has no effect on the basal level or the DNA
damage-induced expression of PHR1, RNR2 or RNR3.
Meanwhile, Crt1 and Tup1/Ssn6, the repressors of the RNR
genes, and Rph1 and Gis1, the repressors of PHR1, are not
involved in the control of MAG1 expression (Zhu & Xiao,
2001). Hence, it becomes apparent that all three sets of wellstudied yeast damage-inducible genes (RNR, PHR1 and
MAG1-DDI1) have distinct regulators and that the regulatory mechanisms are also different from each other.
The expression of MAG1 and DDI1 is also controlled by
the DNA damage checkpoint. Mutation of POL2, MEC1 or
DUN1 reduces the DNA damage response of MAG1 (Zhu &
Xiao, 1998, 2001), suggesting that MAG1 is regulated by the
POL2-MEC1-RAD53-DUN1 checkpoint pathway. In contrast, DDI1 remains inducible in sad1-1 (rad53), dun1 or
mec1 mutants, but its induction is diminished in pds1 sad1-1
(rad53) or pds1 dun1 double mutants (Zhu & Xiao, 2001).
PDS1 encodes an anaphase inhibitor and functions downstream of CHK1; CHK1-PDS1 and RAD53-DUN1 may form
two parallel branches in the DNA damage checkpoints (Fig.
1) (Gardner et al., 1999; Schollaert et al., 2004). This
suggests that the CHK1-PDS1 and MEC1-RAD53-DUN1
checkpoint pathways may function redundantly in the
control of DDI1 expression (Zhu & Xiao, 2001).
A putative eukaryotic SOS response in budding
yeast?
Although cell-cycle checkpoints have been regarded as a
eukaryotic SOS response, their similarity to the bacterial
SOS response is limited. In budding yeast, the Rad51 protein
is a sequence homolog of E. coli RecA and possesses ssDNAbinding and ATPase activities (Shinohara et al., 1992).
However, the only Rad51 enzymatic activity reported to date
is ATP- and ssDNA-dependent recombinase, and it is involved in homologous recombination and homologymediated DNA repair and damage tolerance pathways (Sung,
1994; Paques & Haber, 1999; Gangavarapu et al., 2007).
Although RAD51 itself is DNA damage inducible (Shinohara
et al., 1992), deletion of RAD51 does not affect the expression
of the DNA damage-inducible genes examined (Y. Fu &
W. Xiao, unpublished results). Nevertheless, it remains
possible that there is a functional homolog of RecA in yeast
with respect to a transcriptional response but it shares no
sequence homology and enzymatic activity with RecA.
In a broad sense, E. coli RecA controls three important
cellular responses to DNA damage, namely homologous
recombination via RecBCD and RecFOR, TLS by working
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c
Y. Fu et al.
with PolIV and PolV and the SOS response, which collectively provides a survival mechanism when cells encounter
replication blocks and/or contain excessive ssDNA. The
above functions are reminiscent of the Rad6–Rad18 complex in budding yeast. Rad6–Rad18 forms a stable complex
that is essential for the DNA postreplication repair (PRR)
pathway (Broomfield et al., 2001). The Rad6–Rad18 complex possesses ssDNA-binding and ATPase activities (Bailly
et al., 1997) and assumes most if not all RecA functions to
coordinate broad cellular responses to DNA damage. Firstly,
Rad6–Rad18 as an E2–E3 ubiquitination complex monoubiquitinates PCNA to promote Polz- and PolZ-mediated
TLS (Hoege et al., 2002; Stelter & Ulrich, 2003). Secondly,
this mono-Ub-PCNA is required for PCNA poly-ubiquitination via a Lys63 chain linkage for an error-free mode of
DNA damage tolerance (Hoege et al., 2002) reminiscent of
the RecFOR activity in E. coli (Chow & Courcelle, 2004).
Finally, although Rad6–Rad18 does not have homologous
recombinase activity like RecA, it may compete with the
Ubc9–Siz1 complex that sumorylates PCNA at the same
K164 residue; sumorylated PCNA recruits the DNA helicase
Srs2, which inhibits the recombinase activity of the yeast
RecA homolog Rad51 (Papouli et al., 2005; Pfander et al.,
2005). Hence, it would be of great interest to investigate
whether the Rad6–Rad18 complex also plays a role in an
SOS response and, if so, how it interacts with the damage
checkpoints.
Conclusions
In summary, transcriptional response to DNA damage is an
important process for cell survival of microorganisms under
such life-threatening stress. In bacteria such as E. coli,
the DNA damage-induced transcriptional regulation is
centrally controlled by an SOS response. All SOS-inducible
genes contain SOS boxes in their operator regions, and
their expression is repressed by the interaction between the
transcriptional repressor LexA and SOS boxes under
noninduced conditions. The DNA damage induction is
achieved through enhanced self proteolysis of LexA
assisted by ssDNA-activated RecA. In addition, bacteria also
possess regulatory mechanisms in response to specific DNA
damage.
DNA damage-induced transcriptional response in eukaryotic microorganisms is apparently more complicated and
less understood. Nevertheless, several conclusions can be
drawn from data analyses as presented in this review. Firstly,
the idea that eukaryotic microorganisms possess an SOS
response structurally conserved from bacteria has been
effectively ruled out. Secondly, in budding yeast, a large
number of yeast genes (more than 5%) are induced after
DNA damage. Almost all DNA damage-induced genes
individually investigated to date respond to a broad range
FEMS Microbiol Rev 32 (2008) 908–926
921
DNA damage-induced transcription in budding yeast
of DNA-damaging agents regardless of whether the gene
function provides a survival value to the particular damage
(Birrell et al., 2002). This observation supports the hypothesis that there may exist a coordinated signal transduction
pathway to regulate transcriptional response to DNA damage. Thirdly, DNA damage checkpoint pathways variably
affect transcriptional response to DNA damage in budding
yeast and have been regarded as a functional analog of the
bacterial SOS response. Fourthly, unlike the SOS-inducible
genes in bacteria, few DNA damage-inducible genes in
budding yeast share a common promoter sequence, suggesting that the downstream effectors are diversified. This is
consistent with reports that all three well-characterized sets
of damage-inducible genes in budding yeast are controlled
by different transcriptional regulators and furthermore,
their modes of regulation are different: while RNRs and
PHR1 are induced by derepression, MAG1 and DDI1 are
induced by activation. Fifthly, the search for a novel
mechanism(s) in eukaryotic microorganisms that may better resume the RecA-mediated SOS response is underway
and may result in exciting insights into the eukaryotic SOS
response. Finally, although transcriptional response to DNA
damage is a convenient and useful indicator for the gene
function, transcriptional levels do not necessarily reflect
cellular protein levels, and this appears to be the case for
DNA damage response in budding yeast (Lee et al., 2007).
Hence, other posttranscriptional regulatory mechanisms
have to be taken into account when examining a particular
gene function and its product activity.
Acknowledgements
The authors wish to thank the laboratory members for
helpful discussion and Michelle Hanna for proofreading
the manuscript. This work is supported by the Natural
Sciences and Engineering Research Council of Canada
discovery grant OGP0138338 to W.X. Y.F. is a recipient of
the Arthur Smyth Memorial Scholarship from College of
Medicine, University of Saskatchewan.
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