DNA Damage Tolerance and Mutagenesis: The Regulation of S. cerevisiae Revl
by
Mary Ellen Wiltrout
Submitted to the Department of Biology in Partial
Fulfillment of the Requirements for the Degree of
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DNA Damage Tolerance and Mutagenesis: The Regulation of S. cerevisiae Rev1
by
Mary Ellen Wiltrout
Submitted to the Department of Biology on May 22, 2009 in partial fulfillment of the
requirements for the Degree of Doctor of Philosophy in Biology
ABSTRACT
DNA damage constantly challenges the integrity of genetic material during the
lifetime of every cell. Accurate duplication of DNA and its proper transmission to a new
cell are critical to avoid mutations or loss of genetic information that ultimately may
cause altered cellular functions, cell death, or uncontrolled growth as in the case of tumor
cells. Fortunately, cells possess a multitude of mechanisms to ensure the fidelity of DNA
replication and protect against permanent changes to DNA.
These mechanisms divide into the categories of DNA repair and DNA damage
tolerance, although some of the proteins involved overlap between both mechanisms.
DNA repair restores damaged DNA back to the original, unmodified state. Alternatively,
the cell may require DNA damage tolerance to temporarily deal with DNA damage
during replication. The altered DNA remains as a result of tolerance and is later a
candidate for repair.
This work focuses on the DNA damage tolerance pathway of tranlesion synthesis
(TLS). TLS involves specialized DNA polymerases with the capacity to bypass DNA
lesions that are otherwise inhibitory to replicative polymerases. Specifically, the TLS
polymerases, Revl, Pol , and Pol r, perform TLS in Saccharomyces cerevisiae. In
addition to their contribution to cellular survival after DNA damage, Revl and Pol are
responsible for the majority of spontaneous and damage-induced mutagenesis. Thus
mutagenesis, at least for Pol , is a consequence of a catalytic activity with an increased
error rate relative to replicative polymerases like Pols 6 and E.
Due to the mutagenic nature of Revl, the cell must regulate the employment of
TLS to optimize the benefits of tolerance. One manner this is accomplished in S.
cerevisiae is through the cell cycle regulation of Rev1's protein levels. I begin with the
study of proteasome-dependent degradation as a means to govern the cell cycle regulation
of Revi and the non-cell cycle regulated levels of the Rev3 subunit of Pol . Next, I
describe another layer of regulation involving Rev 's lesion-specific catalytic activity
that is otherwise largely ignored in vivo and how the error-free tolerance pathway masks
the loss of this function.
Thesis Advisor: Graham C. Walker
Title: Professor of Biology
ACKNOWLEDGEMENTS
I will begin by thanking my thesis advisor, Graham C. Walker. Graham's style of
mentoring has allowed me to develop independence in research. His emphasis on
creativity exposed me to many types of science in our diverse lab. I appreciate Graham's
very generous, caring personality, his tremendous support, and the respect that he gives
me as a colleague during our scientific discussions. His encouragement for his lab
members to have a well-rounded life including family, friends, and hobbies created an
interactive lab environment that led to an overall intellectually stimulating and fun
graduate career.
I am very grateful to my committee members for reading this thesis and for their
constructive input. Steve Bell has provided many insightful experimental suggestions and
expertise in replication. Frank Solomon goes above and beyond the duties as a committee
member. I cherish his guidance and time that he took for the emails and talks in his
office. My more recent additions to my committee, Michael Hemann and Alan D'Andrea,
offer their valuable perspectives from mammalian and disease research and always show
enthusiasm for my work.
I would not be here today without my amazing undergraduate experience at
Carnegie Mellon University. I have to thank my research advisor, Chien Ho, for always
challenging me to reach my fullest potential while treating me like a family member in
his lab. Carrie Doonan made a huge impact on my career goals by allowing me to
discover my love for teaching during my year as her teaching assistant. Elizabeth (Beth)
Jones had the greatest overall influence on me from Carnegie Mellon. She truly cared and
performed her responsibilities at the highest level as an educator, scientist, leader, and
mentor. She always believed in me and managed to secretly know a lot about me. I
trusted her opinion on every major decision including my choice to attend MIT for
graduate school. I will always remember her as my role model and miss her as I continue
in academics.
I have to acknowledge all of the people that made my life at MIT better in every
way. I have to thank Daniel Jarosz, Laurie Waters, and Rachel Woodruff for their advice,
technical guidance, and enthusiasm for my preliminary data during my early years in the
Walker lab. Michael Onwugbufor, my summer MSRP student, helped to significantly
expand my catalytic project. Susan Cohen has been there for all of our lab and
departmental activities and was a partner in the thesis writing process. Nicole De Nisco is
an excellent baymate and friend. Lyle Simmons advised me as his "practice" graduate
student and occasionally acted as my protective big brother. Hajime Kobayashi provided
lasting friendship and entertainment. Brenda Minesinger shares my excitement for TLS
polymerases and was always willing to discuss data. Katherine Gibson, Asha Jacobs,
Michi Taga, other members of the Walker lab, and friends in Building 68 made going to
lab everyday enjoyable. Dan Pagano contributed to my thesis project during his rotation
but more notably is great for much needed trips to the Tavern after late nights in lab. My
classmates, Emily Miller, Josh Wolf, and Vineet Prabhu, have been through everything
with me since first year and will continue to do so as we move on to our next steps in life.
Giselle Roman Hernandez and I initially bonded over our love of salsa and became great
friends. My non-MIT friends including Yogesh Oka and the salsa community keep me
stress free and in check with reality. I learned a great deal about teaching from Michelle
Mischke. Betsey Walsh and Judy Carlin always knew the answer and would drop
everything to help me out.
Lastly, but most importantly, I have to thank my family. My parents constantly
give their love and support. My sister, Elizabeth, and I understand each other on a level
only comparable to twins. Aside from being the only relative that discusses scientific
research with me, I turn to her for honest criticism, editor-like proofreading, and opinions
on tough decisions. My grandparents, aunts, uncles, and cousins make up the large family
that I come from and love.
Table of Contents
Title
1
Abstract
3
Acknowledgements
5
Table of Contents
7
Chapter 1:
Introduction - Eukaryotic Translesion Polymerases and Their Roles
and Regulation in DNA Damage Tolerance
Introduction
DNA Repair and Tolerance
Translesion Synthesis
Eukaryotic Translesion Polymerases
Global Models for the Mechanism of Lesion Bypass by
TLS Polymerases
The Evolutionary Significance of TLS Polymerases
Summary
Table and Figures
References
9
10
11
13
20
46
47
48
51
58
Chapter 2:
Proteasomal Degradation of the Mutagenic Translesion DNA
Polymerases, Saccharomyces cerevisiae Rev1 and Pol
Abstract
Introduction
Materials and Methods
Results
Discussion
Table and Figures
References
77
78
79
84
87
93
97
110
Chapter 3:
The DNA Polymerase Activity of Saccharomyces cerevisiae Revl is
Biologically Significant in a Lesion-Specific Manner
Abstract
115
116
Introduction
Materials and Methods
Results
Discussion
Tables and Figures
References
117
122
126
137
143
170
Conclusions and Future Directions
175
Chapter 4:
Summary of Results
The molecular mechanisms governing Rev l's cell
cycle regulation
The lesion specificity of Revl's catalytic activity
The genetic and physical interactions with the
error-free tolerance pathway
Table
References
176
177
179
181
184
185
Appendix A:
The Role of Saccharomyces cerevisiae Rev1 in the Cellular
Resistance to Cisplatin
Introduction
Materials and Methods
Results
Discussion
Table and Figures
References
187
188
190
192
195
198
207
Chapter 1
Introduction
Eukaryotic Translesion DNA Polymerases and Their Roles and
Regulation in DNA Damage Tolerance
This chapter is a modified form of that previously published in Microbiology and
Molecular Biology Reviews, Volume 73, pages 134-154 in March 2009. The authors
were Lauren S. Waters, Brenda K. Minesinger, Mary Ellen Wiltrout, Sanjay D'Souza,
Rachel V. Woodruff, and Graham C. Walker. I was a co-first author.
9
INTRODUCTION
The faithful replication of DNA and proper transmission of chromosomes is
essential to inherit an accurate and complete genome, which encodes the information
necessary for life. Ironically, the process of living itself generates reactive metabolites
that can cause DNA damage. Cells are also exposed to a vast array of exogenous stresses
that can directly or indirectly lead to DNA damage. Although cells contain multiple,
highly complex systems to faithfully restore DNA to its original sequence and structure,
at times distinct mechanisms are required to temporarily tolerate DNA damage without
mediating repair of a lesion. These DNA damage tolerance processes contribute to
survival after DNA damage and, in some situations, also actively promote the generation
of mutations. The factors responsible for spontaneous and damage-induced mutagenesis
are now known to include specialized DNA polymerases, termed translesion
polymerases, found in all domains of life. Understanding of these potentially mutagenic,
yet highly conserved polymerases is critical to a complete knowledge of cell stress
responses, mechanisms of genomic integrity, cell death after DNA damage, induction of
mutations, disease development, and the processes of adaptation and evolution.
Here I will briefly introduce the many strategies a cell may employ to allow
survival in the face of DNA damage before turning to the contribution of damage
tolerance mechanisms, in particular translesion synthesis. I describe the DNA
polymerases that mediate translesion synthesis and highlight the unique properties of the
Rev1 and Pol
families, which are together responsible for the majority of mutations in
eukaryotes from yeast to humans. I review our current understanding of the eukaryotic
translesion polymerases and emphasize the complex regulation that utilizes mutagenesis
for a cell's benefit while preventing rampant mutations under normal conditions. I
conclude with a brief description of the two major models for the regulation of
mutagenesis resulting from translesion synthesis.
DNA REPAIR AND TOLERANCE
DNA damage is a highly complex cellular insult and represents a major obstacle
to proper cellular functions. DNA damage can lead to cell death or alternatively, diseases
in which damaged cells fail to die, such as cancer. DNA lesions and strand breaks
interfere with replication, potentially causing mutations, and also hinder transcription,
affecting gene expression and cellular physiology. Compounding the challenge for the
cell, DNA damage is also extremely prevalent. Approximately -30,000 lesions are
generated spontaneously in a mammalian cell per day (127). Major sources of
spontaneous DNA damage include reactive oxygen species produced primarily during
aerobic metabolism; base deamination, especially of cytosine to uracil; and the inherent
susceptibility of DNA to depurinations and depyrimidinations (47, 127). Additionally,
many environmental factors can cause DNA damage, such as ionizing or ultraviolet (UV)
radiation and chemical agents including methyl methanesulfonate (MMS), cisplatin, and
benzo[a]pyrene (47). These agents can cause modifications of the nitrogenous bases or
breaks in the sugar-phosphate backbone.
The wide variety of DNA lesions that result from diverse DNA damaging agents
has necessitated the evolution of a multitude of cellular responses to DNA damage
(Figure 1A). These DNA repair pathways consist of systems that directly reverse the
damage and several types of excision repair: nucleotide excision repair, base excision
repair and mismatch repair. Additional mechanisms of DNA repair include single strand
break repair and the repair of double strand breaks by non-homologous end joining,
homologous recombination, or single-strand annealing. The reader is referred to
Friedberg et al. (47) and the many excellent reviews that are available for further
information on DNA repair.
Additionally, cells possess mechanisms to temporarily tolerate DNA damage until
DNA repair processes can remove the damage (Figure 1A). In eukaryotes, tolerance
includes an error-free pathway and a parallel more mutagenic pathway as reviewed in
Andersen et al. (7). The type of posttranslational modification on the processivity clamp
PCNA (proliferating cell nuclear antigen) plays a major role in determining the tolerance
pathway utilized. Part of the tolerance to DNA damage lies in the ability of cells to
replicate across damaged DNA, a process called translesion synthesis (TLS) that is a
major component of the more mutagenic branch of tolerance. Without DNA damage
tolerance, cells face the risk of replication fork collapse, translocations, chromosome
aberrations, and cell death.
Conceptually, DNA damage tolerance is quite different from DNA repair in that,
rather than restoring DNA to its proper sequence and structure, the lesion is still present
in the DNA after DNA damage tolerance pathways act (e.g. Figure 1B) (47). Since the
function of damage tolerance is to temporarily bypass a DNA lesion rather than to
regenerate the original sequence, damage tolerance mechanisms are optimized to allow
survival by promoting the completion of DNA replication rather than protecting the
accuracy of the genomic information. Therefore, it is not surprising that DNA damage
tolerance often operates in a mutagenic manner. In this introduction, I focus on the
molecular mechanisms behind the generation of mutations by DNA damage tolerance
and how these potentially mutagenic pathways are exquisitely regulated to promote
survival while restricting the introduction of mutations. Another consequence of these
complex regulatory pathways is that the mutations which do arise, occur under conditions
when they might be evolutionarily advantageous through increasing genetic variability,
as demonstrated in somatic hypermutation (SHM) of immunoglobulin genes.
TRANSLESION SYNTHESIS
Translesion synthesis is the process by which a DNA lesion is bypassed by the
incorporation of a nucleotide opposite to the lesion (47) (Figure 1B). Many DNA lesions
cannot be used as a template by the highly stringent replicative DNA polymerases, which
are optimized to replicate the entire genome with high accuracy and efficiency (13, 47).
However, a class of DNA polymerases with particular characteristics, termed TLS
polymerases, can use damaged DNA as templates and insert nucleotides opposite lesions
despite the conformational constraints many modified bases may impose (47, 60, 189).
TLS polymerases are found in organisms throughout all three domains of life. Most TLS
polymerases are members of the Y family of DNA polymerases (173), a unique class of
DNA polymerases with specialized structures optimized to allow replication on damaged
DNA substrates and, in some cases, to promote mutagenic DNA synthesis. Additionally,
other classes of DNA polymerases, such as the A and X families, can exhibit TLS
activity. Since this activity is often weak or not the primary function of these polymerases
(see below), I focus on the Y family of DNA polymerases, which are uniquely adapted
for translesion synthesis. The Y family members include: Revl, Pol K (DNA Pol IV in
bacteria), Pol r, and Pol L,and bacterial DNA Pol V (UmuD' 2C). For historical reasons,
each polymerase has multiple names, resulting from genetic or biochemical
characterizations carried out over many years using different organisms. Since these
names are used interchangeably in the literature, additional names for polymerases are
indicated in the section titles and in Table 1. Another eukaryotic DNA polymerase, Pol ,
is a member of the B family of DNA polymerases, which includes replicative DNA
polymerases, yet is capable of TLS and has a specialized ability to extend from
mismatched and/or distorted primer-template pairs, including those opposite to DNA
lesions, with remarkably high efficiency compared to most other polymerases.
Discovery/history of translesion polymerases:
Genes encoding translesion polymerases have been known for decades, however
their function remained mysterious until relatively recently. In 1971, Jeffrey Lemontt
isolated genes actively involved in the process of mutagenesis by screening for
reversionless mutants of S. cerevisiae that were unable to revert an auxotrophic marker to
the wild-type allele after UV irradiation (119). Using this approach, REVI (encoding the
Rev 1 DNA polymerase) and REV3 (encoding the catalytic subunit of Pol ) were
identified as genes that, when mutated, conferred a strikingly lower frequency of
mutations than the wild-type strain. A conceptually analogous screen for unmutable
genes in E. coli led to the identification of umuC (encoding the catalytic subunit of
UmuD' 2 C, DNA Pol V) (93).
Although the REV1, REV3, and umuC genes were identified by their profound
contributions to damage-induced mutagenesis, other translesion polymerase genes have
14
more subtle effects on mutagenesis and were first identified primarily by homology
searches with other TLS polymerases. For example, RAD30 and Pol L(hRAD30B), were
first identified solely by their homology to the TLS polymerase genes REV1, umuC and
dinB (initially identified in E. coli) (96, 144, 145, 196, 219). It was not until 1996 that the
first biochemical description of a specialized translesion polymerase appeared [Pol ; a B
family DNA polymerase (162)], followed rapidly by the demonstration that the Revl
protein had a restricted DNA polymerase activity and was able to insert Cs opposite an
abasic site (161). Even then, it was not until 1999, with the characterization of the
translesion polymerase activity of DNA polymerase i. (84, 136, 137), that it was
recognized that all of the genes which shared homology with the eukaryotic REV1 and
bacterial umuC were in fact DNA polymerases with the unique ability to replicate over
DNA lesions (extensively reviewed in (60, 189). This realization took so long in part
because these novel translesion polymerases share almost no primary sequence homology
with classical replicative DNA polymerases and some have proved to be particularly
difficult to purify. This new family of translesion DNA polymerases was named the Y
family of DNA polymerases (173).
Physical features of TLS polymerases:
Several seminal crystal structures provided initial insights into the architectural
features that confer unique catalytic properties to the Y family members (128, 129, 157,
158, 202, 213, 214, 247) and recently many additional structures have been elucidated
which refine our understanding. Despite a nearly complete lack of primary sequence
homology with all other known DNA polymerases, Y family members share the classic
"right-hand" DNA polymerase fold (Figure 2) (13, 189, 235). Like replicative
polymerases, the catalytic aspartate and glutamate residues, which coordinate the divalent
magnesium ions that stabilize the triphosphate group of the incoming dNTP, are located
in the central palm region (13, 189, 234). The thumb and fingers domains of Y family
polymerases, analogous to those in replicative polymerases despite secondary structure
variations, grip the DNA and make specific contacts with the primer and template
strands, respectively (Figure 2) (13, 189, 234).
Although they share a common overall architecture, Y family polymerases differ
from replicative polymerases in certain key ways that allow them to perform translesion
synthesis. At the domain level, Y family polymerases lack the intrinsic 3' to 5'
exonuclease domain of replicative DNA polymerases that functions to proofread the
newly replicated strand (60, 235). A novel little finger domain, also called the
polymerase associated domain (PAD) (129, 202) or wrist (213), is present only in Y
family polymerases and extends from the classical fingers domain to make extra contacts
with the DNA (Figure 2) (234, 235). Certain TLS polymerases also contain additional
regions, such as the "N-clasp" of DNA Pol i, that further contribute to DNA binding
(131). These additional DNA binding regions provide important stability for the ternary
complex, since Y family members have short, stubby thumb and fingers domains that
make few contacts with the DNA backbone (189, 234). Y family polymerases generally
have an open grip on the DNA (Figure 2) and a greatly reduced processivity relative to
replicative DNA polymerases (48, 189); truncations of the little fingers domain or Nclasp reduces DNA binding and processivity even further (129, 131, 189). Intriguingly,
the little finger domain (the least conserved region of the TLS polymerase domain)
appears to contact the region of the template containing the lesion (Figure 2) (140, 142)
and has been implicated in lesion specificity (21).
Closer inspection of the active sites of Y family and replicative polymerases also
reveals significant differences. Particularly for the archaeal and bacterial Y family
polymerases, the active site is larger and more open than in other DNA polymerases (189,
234, 235). A more spacious active site allows accommodation of large bulky adducts
(130, 157), and even two covalently linked bases in a thymine-thymine dimer (128).
Other Y family members appear to have more constrained active sites that nonetheless
are specialized to accommodate particular classes of DNA lesions (131). Further
promoting their ability to use modified DNA templates, Y family polymerases make
fewer contacts with the forming base pair (234, 235) and, in particular, lack the O-helix
of replicative DNA polymerases which, upon binding of a dNTP, rotates -40' to
sterically check the forming base pair (180). Based on crystallographic analysis, it has
been proposed that some Y family polymerases may not exhibit an induced fit upon
binding of the incoming dNTP, which contributes to the replicative fidelity of replicative
polymerases (235), however, there is evidence to suggest a conformational change during
catalysis by archaeal Dpo4, S. cerevisiae Pol rl, and perhaps human Pol
Yr
as well (43, 86,
223).
As of the writing of this introduction, four eukaryotic Y family polymerases have
been co-crystallized with DNA: human Pol L(158), S. cerevisiae Pol r with a cisplatin
adduct (Figure 2A) (6), human Pol K (131), and S. cerevisiae Revl (Figure 2B) (156,
157). Each polymerase displays distinct and often unusual interactions with the DNA
lesion and incoming nucleotide. For example, Revi uses an amino acid in the catalytic
domain to base-pair with an incoming dCTP in lieu of pairing with the DNA template
(Figure 2B) (157), whereas Pol Lappears to use an unusual Hoogsteen base pairing
mechanism (158). In contrast, Pol
iKuses
Watson-Crick base-pairing to incorporate
nucleotides and has a relatively constrained active site (131). Thus, although we have
gained substantial insight at the molecular level into how Y family members are
specialized to bypass DNA lesions, we still have much to learn about how the molecular
architecture of each TLS polymerase active site helps it to achieve its bypass specificity.
TLS polymerases have reduced fidelity relative to replicative DNA polymerases:
The novel features of translesion polymerases that allow them to use an increased
variety of altered DNAs as templates also confer decreased replication fidelity.
Compared with replicative DNA polymerases, which utilize proofreading and exhibit
error rates in the range of incorporating one incorrect nucleotide for every 106 to 108
bases replicated, TLS polymerases display error rates that can range from approximately
one incorrect nucleotide for every 10 bases to one for every 10,000 bases when
replicating undamaged DNA (60, 103, 189). Therefore, TLS polymerases have a
potentially mutagenic activity inside the cell (46). The lack of a 3' to 5' proofreading
domain reduces the fidelity of TLS polymerases operating on undamaged DNA
_10-2
compared to replicative DNA polymerases (60, 103, 235), and the limited number of
contacts made with the template base and incoming nucleotide further decrease accuracy.
Additionally, as mentioned above, it has been suggested that some TLS polymerases are
less accurate because they do not undergo an induced fit upon nucleotide binding (234,
235) and certain TLS polymerases, like DNA Pol Land Revl, do not use canonical
Watson-Crick base pairing (157, 189). Thus, as a consequence of their unusual
polymerization mechanisms, TLS polymerases exhibit a markedly lower accuracy of base
pair insertion on undamaged DNA templates relative to the replicative DNA
polymerases.
Some TLS polymerases are specialized for replicating cognate DNA lesions or
particular DNA substrates:
Despite their relatively low fidelity on undamaged DNA, a paradigm shift has
reclassified TLS polymerases from generally being considered "error-prone" polymerases
(60), as often initially described, to a more nuanced understanding of their role as lesionspecific bypass polymerases (46). It is now appreciated that certain TLS polymerases are
optimized to efficiently replicate over particular DNA lesions, referred to as their cognate
lesions, in a relatively accurate manner (81, 84, 221). Cognate lesions have been defined
for several TLS polymerases by showing that the polymerase is able to bypass the lesion
accurately in vitro and in vivo and that the efficiency of nucleotide insertion opposite to
the lesion occurs with equal or higher efficiency than on undamaged DNA (81, 84, 142,
221). This is strikingly seen in the case of DNA Pol r~, which is specialized to bypass
cis-syn TT dimers caused by UV irradiation (83, 84, 137). Although DNA Pol rl exhibits
among the lowest fidelities of any TLS polymerase on undamaged DNA (-101 )(138), it
is highly accurate in bypassing this UV-induced lesion. This property makes Pol
,r critical for the avoidance of sunlight-induced skin cancers in humans (discussed further
below) (114). Additionally, Pol K,and its archeal and bacterial homologs, can bypass
certain N2 -dG adducts accurately and efficiently (81).
EUKARYOTIC TRANSLESION POLYMERASES
Phylogenetic analysis and extensive biochemical characterization has revealed
that there are five subfamilies within the Y family of DNA polymerases: Rev 1, UmuC,
DinB/Pol K, Pol U,and Pol . (173), each with their own unique enzymatic and
physiological properties. Additionally, one non-Y family polymerase is required for the
mutagenic bypass of DNA lesions in eukaryotes, DNA Pol ( (Rev3 /Rev7) (106). For this
introduction, I will focus on the eukaryotic DNA polymerases involved in TLS. After
highlighting the differing lesion bypass capabilities for each polymerase subfamily,
including the accurate and efficient bypass of cognate lesions by some TLS polymerases,
the mechanisms of regulation for expression and activity of each polymerase are
reviewed.
Before fully discussing each eukaryotic TLS polymerase, one recurring theme for
the regulation of eukaryotic TLS polymerases needs to be introduced involving the
physical and genetic interactions with PCNA and the proteins involved in modifying
PCNA. The homotrimeric sliding clamp, PCNA, serves as the processivity factor for Pol
6 and Eduring replication (23). For its role in DNA damage tolerance, particular ubiquitin
modifications of PCNA are involved. Specifically, the Rad6-Radl 8 complex catalyzes
the monoubiquitination of PCNA at K164, a modification that stimulates TLS, the more
mutagenic branch of tolerance (207). Monoubiquitination of PCNA can be later extended
to polyubiquitination by the Mms2-Ubcl3-Rad5 complex which elicits an error-free
mode of tolerance (7). In addition, attachment of SUMO (small ubiquitin-related
modifier) at positions K164 and K127 of PCNA in S. cerevisiae has been found to affect
phenotypes for DNA damage-induced survival as well as TLS-dependent mutagenesis in
the absence of exogenous DNA damage, as reviewed by Ulrich (215). Therefore, the
interaction(s) between TLS polymerases and PCNA in its modified forms are key in
comprehending TLS activity and regulation, as discussed frequently in the sections
below.
Rev1:
Uniquely among eukaryotic Y family polymerases, Rev actively promotes the
introduction of mutations in organisms ranging from unicellular yeast to multicellular
organisms, including humans (57, 119). Cells bearing rev] mutations display a drastic
reduction in spontaneous and induced mutagenesis by a wide variety of DNA damaging
agents (47, 106). In multiple genetic backgrounds and in response to different types of
DNA lesions, mutants of REV1 abolish most mutagenesis, indicating its fundamental
importance to this biologically important process (106). For example, Revl is required
for -95% of all UV-induced base-pair substitutions (107). Although only marginally
correlated with the onset of cancer to date (73, 200), REV1 has been shown to modulate
the frequencies with which cisplatin resistant cells are generated from an ovarian
carcinoma cell line (125, 175). Therefore, REV] may contribute to cancer progression
and could be an important target of cancer therapy.
The catalytic activity of Revl.
Revl was the first member of the Y family to be shown to have the capability of
catalyzing the formation of a phosphodiester bond (161). However, because its activity
was limited, Rev 1 was only described as a DNA polymerase after the discovery of the
DNA polymerase activity of other Y family members (173). Revl has a polymerase
activity that is restricted primarily to inserting dCMP nucleotides opposite template Gs
and across from certain DNA lesions, such as abasic sites and adducted G residues (106,
161, 222). To accomplish this specificity, Rev1 uses a novel mechanism that selects
dCTP as the incoming nucleotide by forming hydrogen bonds with a conserved arginine
residue in the catalytic domain rather than by base pairing with the template base, as for
all other known DNA polymerases (Figure 2B) (157). Contacts are made between Rev1
and the template base to ensure its identity as a G, but the template base is flipped out of
the active site by interactions with other conserved residues, allowing bypass of bulky G
adducts (Figure 2B) (157).
In contrast, a catalytically inactive mutant of Revi displays no reduction in levels
of mutagenesis induced by a wide range of DNA damaging agents including UV light
(69, 195), although a change in the mutation spectrum is observed (176, 195).
Interestingly, DNA damage sensitivity and mutagenesis phenotypes are observable for
the catalytic dead mutant after cells are exposed to 4-NQO, an alkylating agent that
produces, among other lesions, N 2-dG adducts. The phenotypes are even more dramatic
in the absence of error-free tolerance in S. cerevisiae, suggesting that the DNA
polymerase activity is indeed important for the bypass of certain N 2-dG adducts (Chapter
3).
The noncatalytic function(s) of Rev1 and its protein interactions.
REVI is required for bypass of a 6-4 TT dimer in vivo, even though purified Rev1
is unable to insert a nucleotide opposite to UV photoproducts in vitro (160, 176, 244).
Therefore, although Revl's unique and highly specialized dCMP transferase catalytic
activity is conserved from yeast to humans (123, 161), its DNA polymerase activity does
not seem to be required for bypass of many lesions for which Rev 1 function is required in
vivo. Instead, the ability of Rev 1 to confer resistance to DNA damaging agents and
promote mutagenesis results mainly from its interactions with other proteins, particularly
other translesion DNA polymerases. Rev1 is notable among TLS polymerases for its
multiple binding partners and possesses several protein-protein interaction modules, all of
which are individually required for its function in vivo. These are: the BRCT domain,
the C-terminal -100 amino acids, the PAD, and the UBMs (Figure 3).
The N-terminal BRCT domain of Revl was the first to be characterized since the
original loss-of-function revl-] mutation in S. cerevisiae, whereby the REV] gene was
identified, is located in the BRCT domain (57, 105, 119). The BRCT (BRCA1 Cterminus) domain was initially characterized as an important motif in the BRCA1 breast
cancer susceptibility protein and has subsequently been identified in a variety of proteins
associated with cell cycle regulation and cellular responses to DNA damage (26, 218). In
striking contrast to mutations in the catalytic active site, mutations affecting the BRCT
domain largely inactivate Revl in vivo. In yeast, BRCT mutants exhibit a severe defect
in survival and mutagenesis after DNA damage (119). More recently, mutations
affecting the BRCT domain have also been shown to reduce REV] function in higher
eukaryotes, however, the extent of the defect varies between studies (62, 80, 195). It was
this finding that REV1 function could be inactivated by a mutation which left the
polymerase activity of Rev1 intact that first led Lawrence to propose a "second function"
for Rev 1 (160). This idea was supported by the recognition that BRCT domains can
mediate protein-protein or protein-DNA interactions (26, 58, 99, 228). A model has
subsequently been developed in which Rev1 mediates its function in survival and
mutagenesis by recruiting and coordinating other DNA damage tolerance factors at the
site of lesions rather than by bypassing DNA lesions directly (61, 69, 212).
Recently, the BRCT domain of Revl has been shown to interact with two DNAbinding proteins-the replicative clamp PCNA (62) and the Rev7 subunit of DNA Pol
(35). Since a BRCT domain is not found in other TLS polymerases, these interactions
are likely to confer unique properties upon Revl. It is probable that the BRCT domain
promotes a specialized interaction with damaged DNA to mark sites of incomplete
replication that require TLS. Indeed, in sub-cellular localization studies, the BRCT
domain is sufficient for nuclear localization (212) and is required for constitutive
localization of REV1 to replication foci (62). This may be because mouse Rev1 interacts
with PCNA via the BRCT domain of Rev1 (62). Interestingly, Rev1 lacks the conserved
PIP motif through which most proteins, including all of the other eukaryotic Y family
polymerases, bind to PCNA (7). Thus, Revl likely interacts with a different surface of
PCNA than other TLS polymerases. Despite Revl interacting with unmodified PCNA,
most of the functional studies have involved monoubiquitinated PCNA. As
monoubiquitinated PCNA accumulates in response to agents that block replication forks
(91), this unique interaction may have functional consequences for differential
recruitment to lesions and/or, as one study indicates, stimulation of catalysis by Revl
relative to Pol
(51). The biological relevance of stimulation of Revl catalysis by
monoubiquitinated PCNA remains unresolved though, since conflicting in vitro results
have been reported (51, 71).
The BRCT of Revi may also be directly involved in localizing Rev1 to aberrant
DNA structures. BRCT domains in other proteins also have been found to interact with
DNA at single-stranded regions or double strand breaks (99, 228). Supporting this
suggestion is a study demonstrating that both S. cerevisiae Pol
and Rev1 are recruited
to the vicinity of an endonuclease induced double-strand break (75). This property was
found to require the Revl BRCT domain but not its catalytic activity.
In an activity unrelated to DNA binding, the BRCT domain of Revl may enable
specific interactions with proteins phosphorylated by the DNA damage checkpoint kinase
cascade. Tandem BRCT domains have been shown to interact preferentially with
phosphorylated targets (58, 134, 238). Although Rev1 only has a single BRCT domain, it
has also been implicated in phosphopeptide binding in vitro (238). To date, an
interaction between the Revl BRCT domain and phosphorylated target proteins has not
been demonstrated in vivo. In principle, however, the ability to bind phosphorylated
target proteins would allow for the regulation (or activity) of Revl or by Revl relative to
other TLS polymerases.
In addition to probable localization to DNA through interactions involving its
BRCT domain, Rev1 interacts with, and may regulate, the activity of other TLS
polymerases through its C-terminus and PAD domain. The last -100 amino acids of
mammalian Revl interact with the TLS Pols n, t, K and
(61, 154, 172, 212). Initially,
the polymerase interaction region at the C-terminus of Revl did not seem to be conserved
between higher eukaryotes and yeast (91, 154, 212). However, extensive sequence
alignment and functional studies have revealed that S. cerevisiae Rev1 does interact with
another TLS polymerase, DNA Pol , through its C-terminus (2, 4, 35, 36, 101, 195).
Beyond potentially regulating localization of TLS polymerases to DNA lesions, Rev1 can
also affect the catalytic activity of other TLS polymerases in vitro, as in the case for the
interaction with Rev3 that stimulates extension from a mismatch or opposite a DNA
lesion (4). Importantly, mammalian and S. cerevisiae revl mutants lacking this Cterminal polymerase interaction region are unable to complement a revlA strain for
survival or mutagenesis after DNA damage (4, 36, 101, 105, 195), showing that Rev
functions in vivo through interactions with other TLS polymerases. Additionally, in
vitro, the PAD region of S. cerevisiae Rev 1 interacts with the Rev7 subunit of DNA Pol (
(2), as well as DNA Pol T, an interaction that stimulates the polymerase activity of Revl
(3).
Finally, mouse Rev1 binds ubiquitin (Ub) via a non-canonical ubiquitin-binding
motif (UBM) (64). Interaction with ubiquitin is necessary for localization of Rev1 into
DNA damage induced foci (64) and for the hyperstimulation of its catalytic activity by
PCNA-Ub (230). Mutants in the UBM display increased chromosomal aberrations,
decreased viability, and decreased mutagenesis after exposure to DNA damaging agents
(36, 64, 230), showing that they, like the other interaction motifs, are required for REV]
function in vivo. Murine Rev1 is monoubiquitinated itself but the mechanism of
ubiquitination, the position of Ub attachment, and the functional relevance of this
modification remain unknown (64, 149).
Thus, multiple protein-protein interaction domains are critical for RE V function
in vivo. These findings have led to a model in which Revl functions primarily as a
scaffold for various post-replication repair proteins to localize mutagenic translesion
complexes to sites of DNA damage and/or to modulate polymerase switching at the site
of a DNA lesion (45, 114). Thus, Revl is thought to play a central role in translesion
synthesis by regulating access of TLS polymerases to the primer terminus (45, 114).
Temporal and spatial regulation of Revl.
Despite its importance in regulating TLS, precisely how, when, and where Revl
functions in vivo is not yet well understood. Revi clearly functions in mitotically
dividing cells. The fact that the REV] transcript is upregulated during meiosis in S.
cerevisiae (24, 33) and has the highest expression in human testis (154) suggests a
meiotic function for Revl as well.
Although TLS is commonly considered to occur during replication/S-phase at a
stalled replication fork, recent evidence has led to the proposal that Revi-dependent TLS
acts after the replication machinery has reprimed downstream and generated a ssDNA
gap opposite to a lesion. Strikingly, in S. cerevisiae, Rev levels fluctuate throughout the
cell cycle and are maximal, not during S-phase as might have been anticipated for a DNA
polymerase, but rather during G2 and throughout mitosis (226). REV] mRNA levels are
cell cycle regulated to a lesser extent. Additionally, Rev1 is phosphorylated in a cellcycle dependent manner, as well as in response to DNA damage in S. cerevisiae,
although how this affects Revl activity is not yet known (197). Together with a key
study indicating that TLS can occur after replication in S. cerevisiae (132), this
unexpected finding has led to a re-evaluation of the implicit assumption that TLS
polymerases act during replication to restart DNA synthesis by replacing the replicative
DNA polymerase at stalled replication forks. Instead, one model is that Rev1 functions
(primarily or in addition to during S phase) after the bulk of replication has been
completed, binding to the aberrant primer termini located in gaps opposite DNA lesions
(226). Although it appears that Rev1 is associated with chromatin constitutively
throughout the cell cycle (197), Rev binds to single-stranded DNA with high affinity
and can likely translocate on this substrate to find primer termini (135). This ssDNA
targeting may allow Revl to identify and localize to sites of incomplete replication
opposite DNA lesions that persist into late S/G2/M. It could then be able to recruit Pol 5,
other TLS polymerases, or additional factors to bypass the lesion and fill in the ssDNA
gap. In contrast to this evidence for a late S/G2/M role for Rev in S. cerevisiae, REV1 in
chicken DT40 cells is required for replication fork progression in the presence of DNA
damage indicating a role for Revl during S-phase in this system (41). Taken together,
these results suggest that the partitioning of Rev1 function between possible replicative
and post-replicative roles may vary between biological systems.
Localization studies using ectopically overexpressed GFP fusions have reported
that Rev1 forms foci in vertebrate cells after DNA damage (62, 152, 153, 212). Damageinduced foci formation requires the UBMs (64), whereas the BRCT domain is sufficient
for nuclear localization and basal foci formation but not required for damage-induced foci
formation (62, 212), highlighting the role of the protein-protein/protein-DNA interactions
in mediating Revl function. Under conditions of ectopic overexpression, colocalization
of Revl foci with PCNA and Pol q has been interpreted to indicate that Rev 1 associates
with replication forks to enable continuous DNA synthesis on templates containing DNA
lesions (62, 152, 212). This first led to a model in which Rev1 was thought to act mainly
during replication (45, 114). However, Revl foci have also been observed in G1 (153).
Additionally, one study observed no Revl foci, either spontaneously or after DNA
damage, when a more physiological expression level of Revl was used (195). These
authors propose that lack of foci formation reflects the need of the cell for only one or a
small number of molecules of Rev1 at sites of stalled replication (195).
Given that the cell cycle regulation of Rev 1 is likely to be complex and that cells
appear to keep the levels of Revl, as well as Pol
low (107), it seems likely that high
levels of Rev , especially during Gi and S, would be detrimental. Additionally, Revl
protein levels are under the control of proteasomal degradation throughout the cell cycle
in S. cerevisiae (Chapter 2). Indeed, the fact that overexpressed Revl localizes to
replication forks may help to explain why S. cerevisiae cells keep the amount of Rev 1
low during S-phase. Even though overexpression of Rev1 in S. cerevisiae did not lead to
any change in cell cycle length or the spontaneous mutation rate, most likely due to the
multiple mechanisms regulating activity (107), future experiments taking advantage of
new technology like DNA combing may reveal additional phenotypes. For example,
DNA combing has been used to show that even mild overexpression of Pol 3 and I in
mammalian cells interferes with normal replication fork progression (184).
The potential relationship between diseases and proper Revl function.
Novel in vivo functions for Rev1 are only beginning to be uncovered. In higher
eukaryotes, additional pathways regulating Revl/Rev3-dependent TLS activity are
emerging that involve the genes implicated in the chromosome instability syndrome,
Fanconi anemia (FA). Interestingly, like TLS-deficient cells, FA-deficient cells exhibit
hypersensitivity to DNA crosslinking agents and are hypomutable (54, 74, 149, 164, 178,
187). More specifically, it has been reported that Rev] and Rev3 are epistatic to FANCC
with respect to survival after cisplatin exposure in DT40 cells and that Revi colocalizes
with FANCD2 after the blockage of replication in HeLa cells (164). Another recent study
shows that the FA core complex is required for mutagenesis and efficient Revl foci
formation in mammalian cells in a manner that is independent of PCNA
monoubiquitination (149). These results contribute to the expanding field of TLS
regulatory mechanisms that are not necessarily related to PCNA modifications.
Revi has also been shown to have unanticipated roles in other events. For
example, Rev1 mutants suppress trinucleotide repeat expansion, particularly those repeats
with hairpin forming capacity. This as-yet poorly understood role of Rev may be
relevant to neurodegenerative diseases (34, 39). Furthermore, Rev1, as well as Rev3 and
Rev7, participate in nuclear mutagenesis induced by mitochondrial dysfunction, localize
to the mitochondria, and contribute to mitochondrial mutagenesis in S. cerevisiae (191,
242). Given the connection of mitochondrial function to disease, these functions of TLS
polymerases may be associated with human diseases.
Pol t (Rev3/Rev7):
DNA Pol
is a heterodimer composed of the Rev3 catalytic subunit and the Rev7
accessory subunit (162). REV3 was identified in the same screen for reversionless
mutants in S. cerevisiae as REV] (119). REV7 was isolated by a similar strategy a few
years later (108). Like rev], rev3 and rev7 mutants are severely defective for spontaneous
mutagenesis, as well as for mutagenesis induced by a wide variety of DNA damaging
agents, and for mutations induced in various DNA repair and tolerance pathway mutant
backgrounds (106). REV1, REV3, and REV7 are considered to be in the same branch of
the RAD6 epistasis group based on phenotypic similarity and limited epistasis analysis
(75, 106). Like REV1, DNA Pol
plays a key role in most mutagenesis from yeast to
humans (31, 56, 119, 147) as well as in cisplatin resistance in human cancer cells (126).
Together, Rev and DNA Pol
are thought to mediate the vast majority of the mutagenic
class of DNA damage tolerance in vivo.
The catalytic activity of Pol t.
Unlike most of the TLS polymerases, which are Y family DNA polymerases,
Rev3 is a member of the B family, which includes the highly accurate replicative DNA
polymerases, DNA Pols 8, F, and t (106, 151). In contrast to most other B family
replicative polymerases, DNA Pol
lacks the motifs characteristic of a 3' to 5'
exonuclease activity (106). Although it can bypass certain lesions like a cis-syn TT
dimer and perform both the insertion and extension steps opposite a thymine glycol
lesion in an error-free manner (87, 162), Pol
appears to be particularly specialized to
extend distorted base pairs, such as mismatches that might result from inaccurate base
insertion by a TLS polymerase or a base pair involving a bulky DNA lesion (106, 189).
In combination with a relatively high error rate for base substitutions, this proficiency for
extending mismatches is what allows Pol
to contribute significantly to mutagenesis
(106, 246). The accessory subunit of Rev3, Rev7, significantly enhances the polymerase
activity of Rev3 (162). Despite the lack of conserved PCNA interaction motifs, Pol (
exhibits increased lesion bypass activity in the presence of PCNA (52). However,
31
stimulation of Pol
activity is not observed with either monoubiquitinated PCNA or the
alternative 9-1-1 processivity clamp (51, 166).
Other Pol t functions and protein interactions.
Although a very large protein, Rev3 does not contain any known protein-protein
interaction modules or other regulatory motifs (Figure 3). In S. cerevisiae, Rev3 interacts
with the C-terminal 100 amino acids of Rev 1 in vitro and this interaction stimulates the
ability of Pol
to extend mismatches and bypass specific lesions (4, 65). However, the
majority of the regulation of Pol ; activity appears to occur through the accessory factor
of Rev7. Rev7 contains a HORMA (Hopl/Rev7/Mad2) domain known to interact with
chromatin (9). Due to its homology with Mad2, Rev7 is also known as Mad2L2 and
Mad2B in higher eukaryotes. In yeast, Rev7 binds to the 9-1-1 alternative DNA
processivity clamp, which participates in DNA damage signaling and checkpoint, and
this interaction may recruit DNA Pol
to sites of DNA damage (199). Additionally,
Rev7 interacts with Revl (61, 154, 172, 209, 212), which seems likely to promote
localization of DNA Pol
to DNA lesions.
The physical and genetic interactions of Revl with DNA Pol
are complex.
Despite the fact that each of the three proteins interacts with the other two (see above), a
heterotrimer of Rev 1, Rev3 and Rev7 does not appear to be formed between purified
proteins, as binding of Revl to Rev7 inhibits interaction of purified Revl and Rev3 in
vitro (4). These findings indicate that the architecture of the Revl-Pol
complex is
intricate, and that several subcomplexes may exist, possibly in a regulated manner. It is
also possible that the post-translational modifications of Rev1 mentioned above may
influence the nature of Revi 's interaction with DNA Pol
in vivo.
Although Rev 1, Rev3, and Rev7 are generally believed to work together, the
functions of Revi and Pol
are not entirely overlapping. For example, REV] appears to
act independently of REV3/7 in the generation of sister chromatid exchanges during the
recombinational bypass mode of damage tolerance (174). Additionally, Revl's role in
preventing trinucleotide repeat expansion is independent of both its own catalytic activity
and that of DNA Pol
,.
This suggests that for some cellular roles, Rev1 can also act alone
(34). Moreover, REV7 appears to have a distinct and independent function in cell cycle
control (see below).
Loss of Pol
causes embryonic lethality in mice (16, 42, 217, 229), indicating
that during proliferation, mammalian cells require a function of Pol . The inability to
study rev3 mutant cell lines in mammalian systems has hampered understanding of Pol C
function. However, studies in the chicken DT40 line have provided insight into the role
of Pol
in vivo, in particular, the contribution of REV], REV3, and REV7 to
chromosomal rearrangements during recombination and interstrand crosslink repair (174,
201). In S. cerevisiae, an organism in which rev3 mutants are viable, REV3 has also been
shown to participate in homologous recombination by mediating the mutagenesis
observed in the break-induced replication (BIR) subpathway of HR (76, 192).
Despite being a relatively small protein, Rev7 participates in many protein-protein
interactions apart from its interactions with Rev and Rev3. Many of these additional
Rev7 interactions are with cell-cycle proteins, indicating a potential link between TLS
and regulation of cell growth. In higher eukaryotes, Rev7 has been shown to interact
with the specificity factors Cdhl and/or Cdc20 of the anaphase promoting
complex/cyclosome (APC/C), as well as the spindle checkpoint protein Mad2, both key
regulators of mitotic progression (29, 155, 183). Interaction with Rev7 inhibits the
ubiquitin ligase activity of the APC/C and prevents the onset of mitotic anaphase (29,
183). Interestingly, Rev7 was recently shown to be the target of a bacterial effector
protein during Shigella infection. Upon delivery of the bacterial IpaB protein into the
cytoplasm, human epithelial cells arrest in G2/M due to aberrant activation of the APC/C
by the removal of the Rev7 inhibition (78). Therefore, Rev7 plays a key in vivo role in
cell cycle regulation. Rev7 also interacts with a variety of other proteins involved in the
cell cycle and regulation of cell growth: the HCCA2 transcriptional activator involved in
cell cycle control (120), the Elkl transcription factor affecting cell cycle progression and
the DNA damage stress response (243), the PRCC cancer protein implicated in RNA
splicing and mitotic progression (227), two metalloproteases involved in cell proliferation
and signaling (163), and the adenovirus death protein (ADP) (237).
Regulation of Pol t.
Multiple mechanisms collaborate to keep Pol
levels low (106), indicating that
overexpression may be detrimental to cells. Indeed, overexpression of Pol
causes
increased UV-induced mutagenesis and decreased UV resistance in S. cerevisiae (190).
Both yeast and human REV3 transcripts contain small upstream open reading frames,
which presumably reduce the translational efficiency of the major open reading frame
encoding the Rev3 protein (56, 57, 106, 123). Additionally, alternative splicing of the
human REV3 gene produces an in-frame stop codon in -40% of REV3 transcripts, further
reducing the levels of Rev3 protein (106). REV3 transcript levels are upregulated above
the normally low basal levels in late meiosis in yeast (203). Reminiscent of the cell cycle
regulation of Rev in S. cerevisiae, Rev3 chromatin association in human cells has a cell
cycle regulated pattern showing highest levels during the G1/S boundary that decreased
during S phase and increased again during late S and G2 (22).
Mammalian cells possess additional mechanisms to regulate TLS activity not
found in S. cerevisiae such as those involving the p53 and p21 proteins, which are
emerging as regulators of TLS. In human colon carcinoma cells, loss of p53 or DNA
mismatch repair causes an increase in REV3 and REV mRNA levels (124). These
backgrounds also exhibit increased rates for the development of cisplatin resistance likely
caused by enhanced TLS-induced mutagenesis. In mammalian cells, p53 and p21
suppress TLS activity and, counterintuitively, stimulate UV-induced monoubiquitination
of PCNA (12).
Subsequent studies shed light on this contradiction exposing the problem
of p21 degrading after exposure to UV damage (94, 111, 112, 204). Using a nondegradable p21, Soria et al. report the inhibition of PCNA ubiquitination in the presence
of stabilized p 2 1 (205).
Given its roles in mutagenic TLS and cell cycle control, it is not surprising that
Rev3 and Rev7 have been studied with respect to cancer (120, 193). Rev7 overexpression
has been found in colon cancer, and this correlates with chromosomal instability and
patient mortality (193). Curiously, another study found that rev3 transcript levels were
downregulated in colon carcinomas (22). The contradicting data reveal the complexity of
cancer and suggest that TLS polymerases could have roles in cancer under specific
contexts. Consequently, changes in the regulation of Rev7 and Rev3 levels in vivo may
be connected to cellular events related to disease.
Pol K (DinB):
In contrast to the REV genes, Pol K was not identified by its involvement in
mutagenesis nor its resistance to DNA damaging agents. Rather, Pol K was identified by
homology searches for eukaryotic orthologs of the E. coli dinB gene (85). Although dinB
was first discovered in 1980 as a gene that was significantly upregulated during the
bacterial SOS response (96), it was not until nearly 20 years later that its polymerase
activity was demonstrated (219). Found in all domains of life (173), DinB/Pol IV (as it is
known in E. coli) and Pol K (as it is known in eukaryotes) is the most highly represented
and most strongly conserved of all the TLS polymerases. The pervasiveness of Pol K
argues that this protein contributes to the normal functioning of all cells. It has been
surprising then that loss of Pol K generally reveals only mild phenotypes (see below).
Although the bacterial ortholog, DinB, has been studied extensively [for review see
reference (82)], eukaryotic Pol Khas been less characterized, particularly in terms of its
role(s) in TLS in vivo. This discrepancy derives in part from the fact that Pol K is
conspicuously absent from what is arguably the best-studied single-celled eukaryote, S.
cerevisiae. Pol K orthologs have been identified in other related fungal species though,
including the fission yeast Schizosaccharomycespombe. Organisms that lack a DinB
homolog may possess another protein that plays a functionally redundant role.
The catalytic activity Pol 1K.
With regard to in vitro DNA replication on undamaged DNA, mammalian Pol Kis
relatively accurate compared to other TLS polymerases; human Pol K has a misinsertion
frequency of I in every 102 to 103 nucleotides replicated (100). Although the bacterial
and archaeal orthologs of DinB demonstrate a marked proclivity for -1 frameshifts in
vitro and when overproduced in vivo (59, 97, 100, 188), mammalian Pol K appears more
restricted in this activity both in vitro and in vivo (171). With respect to DNA damage
bypass, Pol Khas limited ability to synthesize across numerous DNA lesions [for review
see (14, 236)], however it can bypass many N2 -adducted dG residues, including N2-dGlinked DNA-peptide crosslinks, both efficiently and accurately [e.g. (10, 81, 148, 239)].
Indeed, it appears that Pol K is specialized in its ability to bypass N2-adducted dG lesions;
Pol Koperates with high accuracy and strikingly increasedcatalytic efficiency opposite
N2-furfuryl-dG and N2-(1-carboxyethyl)-2-dG [N2-CEdG] residues relative to an
undamaged dG (81, 239). Significantly, like Pol , Pol K appears to be specialized to
extend mismatched primer termini and thus seems likely to function as a second
"extender" polymerase when two TLS polymerases are required in concert to bypass a
lesion (27, 131, 189). Furthermore, in vitro, the DNA synthesis activity of human Pol K
is stimulated in the presence of PCNA, replication factor C (RFC), and replication protein
A (RPA) but not by a single complex in the absence of the others (70).
Role of Pol K in mutagenesis.
The role(s) of Pol K in mutagenesis is enigmatic. In contrast to Revl and Pol ,
deletion of Pol K does not appear to have a profound effect on either spontaneous or
damage-induced mutagenesis. In mammalian cells, loss of Pol K sensitizes cells to the
killing by benzo[a]pyrene and moderately increases mutagenesis induced by this agent,
suggesting that Pol K bypasses N 2- benzo[a]pyrene dG adducts relatively accurately in
vivo (10, 170). Loss of Pol K is also associated with sensitivity to DNA alkylating agents
and to UV irradiation (168, 208), although sensitivity to UV in the absence of Pol K
seems likely to reflect its yet-to-be-defined role in nucleotide excision repair (NER)
(168). Ectopic overexpression of human Pol K in mammalian cell lines inhibits
replication fork progression (184) and leads to general genomic instability, including
increased DNA strand breaks, loss of heterozygosity and aneuploidy (15).
The protein interactions of Pol K.
Multiple protein-protein interactions likely regulate Pol K function. Eukaryotic
Pol K interacts with the PCNA processivity clamp [human (70)], ubiquitin [mouse (63)],
the 9-1-1 checkpoint clamp [S. pombe (88)], and Revl [mouse (61)]. Many, if not all, of
these interactions are important for Pol K's function in vivo. For example, as noted
below, mutations of Pol Kthat disrupt its interaction with ubiquitin or with PCNA result
in aberrant nuclear localization after DNA damage (63, 167). Additionally, the 9-1-1
complex, involved in the DNA damage checkpoint, localizes Pol Kto chromatin in
replication-compromised S. pombe strains (88). Also, a mutant of the 9-1-1 clamp that
perturbs DNA binding by Pol K displays a reduction in point mutations.
Regulation of Pol K.
Pol K relocalizes from a diffuse nuclear pattern into foci upon DNA damage (17,
18, 167). Focus formation of Pol Krequires both its PCNA-interaction motif and its
ubiquitin-binding motifs (63, 167) (Figure 3). Interestingly, Pol K relocalizes in response
to DNA damage differently from the other Y family members, forming fewer
spontaneous and damage-induced foci (18, 167). These reports disagree, however, on
whether DNA Pol K forms foci during S-phase (18, 167).
A key source of regulation of Pol Kmay be at the level of transcription. Murine
Pol K transcript levels increase after treatment with 3-methyl methylcholanthrene, a
polycyclic aromatic hydrocarbon similar to benzo[a]pyrene (169). Notably, a change in
Pol K's transcript levels may be connected to cancer development in some contexts, as
Pol K transcripts are downregulated in some human colorectal tumors (118). Conversely,
Pol K transcripts are upregulated in non-small cell lung cancers (79). This increase in Pol
Ktranscripts correlates with the increased loss of heterozygosity in these tumors (15).
Pol Yj (Rad30A/XP-V):
Like Pol K, the rad30 gene was not identified on the basis of its contribution to
mutagenesis, but rather by its homology to the genes encoding the Rev 1, UmuC, and
DinB proteins. Its name reflects the slight sensitization of a rad30 mutant in S. cerevisiae
to UV irradiation (144). Indeed, under most circumstances, rad30 mutants display very
limited reduction in mutagenesis in yeast.
The Rad30/Pol r subfamily is found only in eukaryotes, where it is broadly
conserved. Pol il is perhaps the most thoroughly characterized TLS polymerase since, in
39
humans, loss of Pol q activity results in a cancer-prone syndrome known as xeroderma
pigmentosum variant (XPV), which is characterized by an increased incidence of skin
cancers and sensitivity to sunlight (95, 114, 117, 137). Clinically, XPV is very similar to
other forms of xeroderma pigmentosum, which result from mutations in any of six key
NER genes, but XPV cells are not defective in NER (121, 194). This phenotype
highlights the predominantly non-mutagenic role of Poli
mutagenic functions of Pol
,
setting it apart from the more
and Revl.
The catalytic activity and role in mutagenesis for Pol q.
The phenotypes of mutants with DNA Pol rl deficiencies and the in vitro activity
of Pol q indicate that its major role is the non-mutagenic bypass of UV-induced DNA
lesions. In particular, Pol f is the primary TLS polymerase responsible in many
organisms for error-free bypass of cis-syn cyclobutane pyrimidine dimers (CPDs), one of
the major lesions resulting from UV radiation (1, 55, 232). In vitro, Pol rJ has been
shown to bypass CPDs with high accuracy and efficiency (84), and in vivo, it is thought
to be responsible for restarting stalled replication forks and allowing continuous DNA
synthesis past sites of UV damage (114). In the absence of Pol r, double strand DNA
breaks develop after UV radiation when unrepaired lesions are encountered during DNA
replication, which can ultimately cause cell death or genomic rearrangements (53, 122).
Furthermore, Pol r-independent CPD bypass, which is thought to involve other TLS
polymerases such as Pol ' and/or Pol u,is significantly more mutagenic, presumably
accounting for the increased frequency of cancer in XPV patients (109, 114, 137, 220).
In addition to the severely distorting CPDs, in vitro, Pol r is also able to bypass a broad
range of other DNA lesions such as: 7, 8-dihydro-8-oxoguanine (8-oxoG) (72), (+)-transanti-benzo[a]pyrene-N2 -dG (245), acetylaminofluorene-adducted guanine (240), 06methylguanine (68), thymine glycol (104), and adducts derived from cisplatin and
oxaliplatin (216). Aside from CPDs, purified Pol 1is able to bypass other large and
distorting lesions such as the cisplatin-induced 1,2-d(GpG) adduct (Pt-GG) and evidence
exists for the importance of Pol 1 after cisplatin exposure in XPV cells (5, 6, 30).
Interestingly, though Pol I plays a major role in accurately bypassing particular
types of DNA lesions, such as CPDs and 8-oxoG, it exhibits among the lowest fidelity of
any DNA polymerase on undamaged DNA in vitro (11, 38, 102, 110, 138, 189). In spite
of its mutagenic potential, depletion of Pol ri in human cells by siRNA actually increases
mutation frequency (32, 77), and S. cerevisiae rad30mutants do not display a major
reduction in spontaneous or induced mutagenesis (144, 196). Similarly, Pol 1 knockout
mouse embryonic fibroblasts show an increase UV-induced mutation frequency (25).
Taken together, these results indicate that Pol l's normal function in vivo primarily
reduces mutagenesis. Therefore, regulation of Pol l activity is thought to play a crucial
role in modulating the mutagenic potential of Pol fl in living cells.
Intriguingly, overexpression of Pol r in human cells does not increase
mutagenesis and only causes a weak mutator effect in S. cerevisiae (98, 181), suggesting
that Pol
Y1 is
largely restricted from accessing undamaged DNA by additional regulatory
mechanisms even when overexpressed. Pol i's potential for introducing mutations,
though normally inhibited in somatic cells, is harnessed in a specific context. Pol 1 is the
major mutator of A:T base pairs during the SHM step in antibody diversification in B
lymphocytes (139, 182).
The protein-protein interactions of Pol vl.
The regulation of Pol r's catalytic activity is directed in part through proteinprotein interactions. Pol ri interacts with the eukaryotic processivity clamp, PCNA,
through its C-terminal PCNA-binding motif (PIP-box) (Figure 3) (92), and the interaction
between PCNA and Poli plays an important role in Pol r function. This is at least
partially attributable to the stimulatory effect of PCNA on Pol rl's TLS activity in vitro
(66, 67, 89). Interestingly, the interaction between PCNA and Pol Tr is inhibited by p21,
a protein discussed above with respect to regulation of TLS (205).
Although ubiquitinated PCNA is not required for Poli to access stalled
replication forks in vitro (165), Pol r's interaction with PCNA can be enhanced by the
monoubiquitination of PCNA. Mammalian Pol r foci co-localize with foci of
monoubiquitinated PCNA in the nucleus (92), and accumulation of Pol r foci in response
to DNA damage is dependent upon monoubiquitinated PCNA (185), although a small
proportion of cells (5-10%) do have Pol Tr foci in radl8-/- orpol30(K]64R) mutants, in
which PCNA is not monoubiquitinated (224). A similar proportion of cells contain Pol ri
foci in the absence of DNA damage, consistent with a model in which PCNA
monoubiquitination induces Pol T's response to exogenous DNA damage, above a low
level of uninduced DNA-association by Pol rl.
The dependence of Pol r's damage-induced foci on monoubiquitinated PCNA is
attributed to Pol r's interactions with PCNA and ubiquitin (185), which appear to give
Pol rl a competitive advantage over the replicative Pol 8 for PCNA association after DNA
damage (224, 241). Pol r's interaction with monoubiquitinated PCNA is mediated by
42
both the PCNA interaction motif (PIP-box) and its ubiquitin binding zinc finger (UBZ)
domain (179) (Figure 3). Mutants disrupting the UBZ, in either S. cerevisiae or
mammalian Pol rI,fail to complement the UV sensitivity of Pol r~ deficient cells (19,
179), although at lower UV doses, Acharya et al. (3) have demonstrated partial
complementation of the UV-sensitive phenotype of the rad30A strain. Monoubiquitinated
PCNA may also promote TLS by enhancing Pol I's catalytic activity, but in vitro results
have so far been inconsistent (51, 71).
Another protein which participates in the regulation of Pol r is Radl8, an E3
ubiquitin ligase that mediates PCNA monoubiquitination. Mouse Pol l has been found to
have a direct physical interaction with Rad 18, independent of the presence of DNA
damage, via C-terminal regions of both proteins (224). Furthermore, in human cells, Pol
i
co-purifies as a complex with Rad 18, Rad6, and Rev 1: the complex is enriched in the
chromatin fraction in response to UV radiation or S phase arrest (241), consistent with
the model that Radl8 is involved in recruitment of Pol r to stalled replication forks. Pol
rl foci co-localize with Radl8 foci (224), and the formation and damage-dependent
accumulation of Pol Ilfoci is largely dependent on Radl8 (224).
Pol Tj may also be regulated by ubiquitination through a covalent attachment of a
monoubiquitin moiety (19, 177, 179), although the functional significance of this
modification is not yet understood. Ubiquitination of Pol Iris dependent on the UBZ
domain of Pol q. Intriguingly, the monoubiquitination of Pol I1is not dependent on the
post-replicative repair (PRR) proteins Rad6 and Radl8, nor is it responsive to DNA
damage (179).
There is also a robust physical interaction between DNA Pol r and Revi in
vertebrates and flies, but only a weak interaction, if any, between Pol ri and Rev 1 in
budding yeast (3, 61, 101, 212, 231). Thus, the functional interactions between TLS
polymerases are complex and, to some extent, species-dependent.
Regulation of Pol 1.
Transcriptional regulation of Pol rl was demonstrated early on. In S. cerevisiae,
the RAD30 transcript is induced 3-4 fold in response to UV radiation (144, 196). In
mouse, however, expression of the XPV gene (encoding Poli ) is not induced by UV
radiation; instead, it has been found to increase about 4-fold during cell proliferation
(233). The RAD30 gene in S. cerevisiae has been placed in the RAD6 epistasis group
(144) but appears to function independently of both the error-free pathway defined by
RAD5 (144) and the error-prone TLS pathway that includes REV], REV3, and REV7
(144, 231).
Pol r1 forms foci spontaneously in a small percentage of untreated cells suggesting
that Pol r is localized to these sites to perform its lesion bypass activity. These foci
accumulate in the majority of cells that have been treated with DNA damaging agents
such as UV or MMS (89), and in cells subjected to hydroxyurea-induced replication
stress (5, 18, 37, 89, 90). These foci are thought to form at sites of DNA damage since
they colocalize with PCNA (5, 89, 90) and with Rad18 foci (224). Although it is assumed
that the nuclear Pol rl foci represent sites of TLS, focus formation does not necessarily
imply activity. For example, a mutant form of Radl 8 that is unable to form foci
nonetheless activates DNA damage tolerance pathways (159). Recent data even reveal
that Pol r is transiently immobilized in foci (198) supporting a model of TLS
polymerases transiently probing the chromatin. Additionally, accumulation of Pol r foci
is stimulated by the physical interaction of Pol 1's UBZ domain with monoubiquitinated
PCNA (19, 92, 185, 224). Together with the fact that Pol 1mutants progress more
slowly through S-phase after DNA damage (5, 206), these findings have led to a model in
which Pol r rescues replication forks that have stalled at sites of DNA damage by
allowing continuous DNA synthesis past the lesion(s).
Pol L (Rad30B):
Pol t is most closely related to Pol
Yr
at the sequence level, but is divergent
enough to have distinct biochemical properties and function. In contrast to the wealth of
information about Pol r1 , the role of Pol Lis less well understood. Pol Lis present not only
in higher eukaryotes as initially thought (145, 173, 211), but in organisms scattered
throughout the Eukaryota including some yeasts (L.S. Waters, unpublished observation).
Because Pol Lis lacking in S. cerevisiae, in which most genetic studies of TLS DNA
polymerases have been performed, little is known about its genetic relationships to other
DNA damage tolerance pathways. A complete description of Pol L's functions are beyond
the scope of this thesis and can be found in (225).
Other non-Y family DNA polymerases capable of translesion synthesis:
It is worth noting that there are other non-replicative DNA polymerases that have
varying abilities to bypass DNA lesions and that synthesize DNA with a range of
fidelities [as reviewed in (150)]. The members of the X family of DNA polymerases
(DNA Pols P, k, t in eukaryotes), in particular, can insert nucleotides opposite to certain
lesions (20, 150). After the Y family, the X family polymerases display the next lowest
replication fidelity of the six major DNA polymerase families (103). The X family
polymerases are occasionally referred to as translesion polymerases and, indeed, can lay a
claim to the name. One A family member, Pol 0, exhibits a reduced fidelity relative to the
other A family members, and has been suggested to participate in TLS and SHM in vivo
(8). Even the highly stringent replicative DNA polymerases have very weak abilities to
replicate over certain lesions. In general though, these non-Y family polymerases have
other primary physiological functions, such as participation in BER and NHEJ by the X
family polymerases. Accordingly, the term TLS polymerases generally refers to the Y
family and DNA Pol , which clearly have specialized roles primarily involved in lesion
bypass (47, 106).
GLOBAL MODELS FOR THE MECHANISM OF LESION BYPASS BY TLS
POLYMERASES
The numerous genetic and biochemical data regarding the post-translational
regulatory strategies detailed above have been integrated into two models for DNA lesion
bypass by TLS polymerases that are not mutually exclusive (Figure 4): i) the polymeraseswitching model (45, 47, 115, 116, 141, 143, 186) and ii) the gap-filling model (113, 115,
143, 226). There is compelling evidence for both models. It is likely that TLS
polymerases act in a manner consistent with both models when spatially and temporally
appropriate, dependent, for example, on the context of the DNA lesion or phase of the
cell cycle. For a more in depth discussion of the two models, see (225).
THE EVOLUTIONARY SIGNIFICANCE OF TLS POLYMERASES
Why are TLS polymerases that can actively cause mutagenesis so conserved
throughout all domains of life? The risk to the cell of potential mutations and replication
perturbation is presumably outweighed by the fact that TLS polymerases confer a
measure of resistance to DNA damaging agents. In general, the type of mutations created
by TLS, base pair substitutions, are less detrimental to the integrity of the genome than
translocations and other gross chromosomal rearrangements that can occur in the absence
of TLS.
Evidence exists to show that the use of TLS polymerases is not trivial. In
mammals, TLS polymerases contribute significantly to lesion bypass, as it has been
estimated that -50% of DNA damage tolerance events occur through TLS rather than the
more error-free recombinational bypass pathways (10). Furthermore, the striking
phenotypes associated with XPV dramatically underscore the significance of TLS to
human health. Some modest phenotypes observed for TLS-deficient cells may be a result
of overlapping functionality. For example, under certain conditions error-free tolerance
may compensate for the loss of TLS, masking the true involvement of TLS polymerases
in DNA damage resistance in cells. In addition, TLS polymerases may provide important
functions to cells by aiding in replication of undamaged but difficult DNA substrates,
such as the recently observed contribution of Rev1 to trinucleotide repeat stability (34),
D-loop extension during homologous recombination by Pol r~ (95, 146), or other as-yet
unknown structures.
Since the majority of mutations are deleterious, most organisms have evolved
mechanisms that keep their mutation rates extremely low (40), and the complex control
of TLS DNA polymerases discussed in this introduction help achieve that end.
Nevertheless, an increase in the genetic variation within a population can be beneficial
under adverse conditions as it increases the chance of a variant emerging that is better
able to withstand the stress (44, 49). Thus, the mutations introduced by TLS polymerases
can be an important factor in evolution by increasing the genetic variability in response to
stresses that damage DNA. In bacteria, TLS polymerases have been implicated in
adaptive mutagenesis--the ability to induce mutations upon cellular stress (44, 49).
Additionally, in higher eukaryotes, the mutagenic capacity of TLS polymerases has been
harnessed for somatic hypermutation, the generation of mutations in the variable regions
of antibodies produced by B cell lymphocytes (28). Thus, despite potentially deleterious
mutagenic effects, TLS polymerases presumably provide more benefits than
disadvantages to cells, consistent with the observation that TLS polymerases have been
found in all organisms whose genomes have been sequenced to date.
THESIS SUMMARY
Cells have developed specialized translesion polymerases to complete replication
in the face of DNA damage, either at stalled replication forks or at sites of gaps
containing lesions. The use of TLS polymerases to bypass DNA lesions provides
resistance to DNA damaging agents through the ability to restart stalled replication forks
or fill in ssDNA gaps found in the genome after DNA damage. However, this comes at
the potential cost of increased mutation frequencies. To counteract the mutagenic risk of
using TLS polymerases, cells have developed elaborate regulation strategies. The
regulation mechanisms detailed here are likely to increase in complexity as our
knowledge in this field grows. The past decade has seen a profound increase in our
knowledge of TLS polymerases and the future promises to reveal further insights into the
mechanism of action of these intriguing enzymes.
In this thesis, I provide data to advance our understanding regarding the
regulation of TLS polymerases. Specifically, in Chapter 2, my results indicate that Rev1
and Pol
are targeted for proteasomal degradation as a means to keep protein levels low.
In Chapter 3, I describe how Revl has a lesion-specific catalytic activity in vivo that is
masked by error-prone tolerance. Finally, I present some preliminary data in the appendix
that help show the instrumental roles of Revl and Pol
protein levels affect this resistance.
in cisplatin resistance and how
ACKNOWLEDGEMENTS
I thank members of the Walker lab for helpful discussions. This work was
supported by a National Institute of Environmental Health Sciences (NIEHS) grant 5RO1-ES015818 to G.C.W., a NIEHS grant P30 ES002109 to the MIT Center of
Environmental Health Sciences, and an American Cancer Society Research Professorship
to G.C.W.
TABLE 1: Genes Encoding Catalytic Subunits of Eukaryotic DNA Translesion
Polymerases
cerevisiae
S. pombe
D. melanogaster
Mouse
Human
Pol
REV3
rev3
mus205/dmREV3
Rev3
REV3L
RevI
REV1
rev]+
revl
Revl
REV1
Pol K
---
dinB /mug4O
---
PolKIDinBi
DINBJ
Pol 11
RAD30
esol +*
DNApol-Ir
Polq
RAD3OA/XPV
---
---
DNApol-
Pol
RAD30B
Pol
*S.mbe esol+contains two separable protein domains. The amino-terminal end is
homologous to Pol rl and exhibits similar in vivo phenotypes and in vitro activities to Pol
1)homologs in other organisms (133, 210). The carboxy-terminal end is comprised of an
essential sister chromatid cohesion protein (133).
FIGURE 1.- DNA damage repair and bypass mechanisms. (A) DNA damage results in
breakage of the sugar-phosphate backbone (not shown) or DNA base loss (indicated by a
gap in the DNA) or base alterations (as indicated by the grey star). This damage can be
repaired/removed from the DNA strand or tolerated, in which the DNA lesion remains,
but cellular processes continue. (B) An example of the DNA damage tolerance
mechanism translesion synthesis (TLS), whereby a damaged DNA template is replicated
using a TLS polymerase and the damage remains in the genome. A more detailed
mechanism of TLS can be found in the text and represented in Figure 4.
sugaf-phoI !4haL1L
Baeos.-
iis
hikoeji
Deueunti-uo.n or
crosIanks. ch,.ericid
ilepy-ninom
\44
I
11'
DNA
DNA
repair
tolerance
~F11
gon~nien
Trawk-uia. Svotbaesis ITISj
NML~ntch ibkghsd
Dwk
Rcv%4
TIT
I
TLS
S~~Jl
TEJ*11111~m
FIGURE 2.- Crystal structures of two Y family polymerases. (A) Co-crystal structure of
the S. cerevisiae TLS Pol 'i with a DNA template containing a cisplatin crosslink. The
structure is oriented to highlight the right-hand architecture as seen in both TLS and
replicative polymerases. Adapted from (6); PDB ID number 2r8j. (B) Close-up view of
the unique lesion-bypass mechanism of Rev1 from S. cerevisiae. Highlighted are the
novel leucine (L325) that helps to flip out the template guanine and the catalytic arginine
(R324) that hydrogen bonds to stabilize the incoming dCTP. The domains of Revl are
colored as in (A) with the exception of the DNA, which is shown in black. Adapted from
(157); PDB ID number 2aq4.
A
Palm
DNA
Thumb
B
Incoming dCTP
L325
Displaced template G
R324
FIGURE 3.- Cartoon representation of the protein domains in the human B-family TLS
polymerase
and the Y-family TLS polymerases Rev1, K, L, and '1. Adapted from Yang
and Woodgate (236) and Gan et al. (50).
3130aa Pol
M
125 laa Rev I
-
.......
A
870aa Pol K
ddbb.-
713aa Pol q
I
--V-
BRCT domain
Polymerase domain
Pot K, Potl , Potl
ldgft
D
300
715aa Pol t
M
Amalk
'MW
-
---
interaction domain
Rev7 interaction domain
1
Ask
nook,
-
UBM
1 PCNA interaction domain (PIP)
Revl interaction domain
0
UBZ
FIGURE 4.- Two non-exclusive models for TLS: the polymerase-switching model (A)
and the gap-filling model (B). See (225) for details.
B.
A.
I
1.
#
1
1.
I
111111 111
111
1111
.............
"]lllllllll
5'
3
0
5'
53.
3. 3E
. 11111 5'
2.
5 rmmnl
%FLt
llTiI I
S1111111
3511111
I
.ll 111111111
5'
5.
0111111
6. 3
AM
III
U11111111117
AW
Ubiquitin
DNA lesion
111111111ILL
K')
9Il
complex
Replicalive
t'nslesion
Polymrnerase I
m
Translesion
Polymncrase 2
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76
Chapter 2
Proteasomal Regulation of the Mutagenic Translesion DNA
Polymerases, Saccharomyces cerevisiae Revi and Pol
This chapter will be submitted for publication. The authors are Mary Ellen Wiltrout and
Graham C. Walker.
ABSTRACT
Translesion DNA synthesis (TLS) functions as a tolerance mechanism for DNA
damage at a potentially mutagenic cost. Three TLS polymerases (Pols) are known to exist
in S. cerevisiae: Rev , Pol , a heterodimer of the Rev3 and Rev7 proteins, and Pol
i
(Rad30). Our lab has shown that Rev1 protein levels are under dramatic cell cycle
regulation, being -50-fold higher during G2/M than during Gi and much of S phase.
REV1 transcript levels just vary -3-fold in a similar cell cycle pattern, suggesting a
posttranscriptional mechanism controls protein levels. Here, we find that the S. cerevisiae
Revl protein is unstable during both the Gi and G2/M phases of the cell cycle. The
protein's half-life is shorter in GI arrested cells than in G2/M arrested cells, indicating
that the rate of proteolysis contributes to Revl's cell cycle regulation. In the presence of
the proteasome inhibitor, MG132, the steady-state levels and half-life of Rev1 increase
during G 1 and G2/M. Through the use of a viable proteasome mutant, we confirm that
the levels of Rev 1 protein are dependent on proteasome-mediated degradation. The
accumulation of higher migrating forms of Revl under certain conditions suggests that
the degradation of Rev 1 is likely directed through the addition of a polyubiquitination
signal. In addition to Rev1, we find that the non-cell cycle regulated TLS polymerase, Pol
, is subject to proteasomal degradation of Rev3 but not Rev7. These results support a
model that proteasomal degradation acts as a regulatory system of mutagenic translesion
synthesis mediated by Rev1 and Pol .
INTRODUCTION
Cells constantly face the challenge of maintaining genomic integrity as a result of
DNA damage arising from endogenous and exogenous sources. To prevent the negative
consequences of DNA damage such as replication fork stalling or collapse, mutations, or
cell death, the cell is equipped with repair and tolerance mechanisms. DNA repair
restores the original state of the DNA. DNA damage tolerance, however, allows DNA
lesions to remain in the genome even during replication.
When the cell employs translesion DNA synthesis (TLS) to tolerate DNA
damage, specialized DNA polymerases catalyze replication opposite lesions that
normally prevent the replicative DNA polymerases' activity. Most TLS polymerases
belong to the Y-family of DNA polymerases that has members from all domains of life
(27). The active site of these Y-family DNA polymerases are better able to accommodate
bulky DNA lesions because they are structurally more open and less sterically
constrained than those of the high-fidelity, replicative polymerases (34). Given these
structural properties of TLS polymerases and their lack of any proofreading activity, TLS
polymerases can exhibit high error rates. TLS across from lesions can be relatively errorfree or quite error-prone depending on the lesion and polymerase involved (12, 13, 17).
Following bypass, the DNA repair pathways can later remove the DNA lesion, which
remains in the DNA.
There are three known TLS polymerases in S. cerevisiae: Rev and Pol rl (Rad30)
of the Y-family and the B-family member, Pol t, a heterodimer of Rev3 and Rev7. All
three are highly conserved among eukaryotes. The REV1, REV3, and REV7 genes were
discovered in screens for reversionlessmutants in yeast (a phenotype indicating loss of a
mutagenic activity) (19, 21). The rev1A mutant phenotypes include an increased
sensitivity to certain DNA damaging agents and a decreased damage-induced mutation
frequency, indicating Rev I's instrumental role in DNA damage resistance and
mutagenesis (18).
Rev l's DNA polymerase activity exhibits unique properties that include a
preference for inserting dCMPs and a specificity for a template G (10, 26). Despite this
clear and evolutionarily conserved activity, the non-catalytic functions of Rev I appear to
be more critical for DNA damage tolerance and mutagenesis in vivo based on known
mutant phenotypes. In S. cerevisiae, the revl-1 (G193R) mutant of the BRCA1 C
terminus (BRCT) domain leads to almost null phenotypes in vivo, but the mutant protein
retains about 60% of the catalytic activity in vitro (25). Additionally, the newly
discovered ubiquitin binding motif (UBM2) and the conserved region of Rev l's Cterminus that interacts with other TLS polymerases are critical for cellular survival and
mutagenesis after DNA damage [as reviewed in (44)]. Therefore, beyond its DNA
polymerase function, Rev1 serves to regulate the other TLS polymerases through proteinprotein interactions or direct interaction with the DNA.
The mutagenic nature of Revl indicates that the activity must be tightly regulated.
The conservation of Rev1 in higher eukaryotes suggests that the evolutionary benefits
outweigh the risks of its potentially mutagenic activity, although it is possible that all of
Rev I's in vivo functions are not known. Aside from the requirement of Rev for cellular
survival after DNA damage, TLS polymerases clearly have a beneficial role in vertebrate
immunity (6). Specifically, these polymerases participate in somatic hypermutation of the
immunoglobin variable regions to produce high-affinity antibodies for specific antigens.
Not surprisingly, disrupting the normal protein levels of TLS polymerases has
negative consequences. In S. cerevisiae, ectopic overexpression of Pol t's Rev3 and
Rev7 proteins leads to a greater sensitivity to UV radiation and an increase in UVinduced mutation frequency (35). In one study, a 2 to 4.5-fold overexpression of the base
excision repair DNA Pol 3 or the TLS DNA Pol Kinterferes with replication fork
progression in CHO cell lines (31). In another report, overexpression of human REV I in
ovarian carcinoma cells demonstrates the potential danger of misregulated Rev 1 levels
(22). When these cells are treated with cisplatin, a drug widely used in the treatment of
tumors, they develop more resistant clones after sequential cycles of drug exposure, an
effect that potentially hinders clinical efficacy in patients. Therefore, understanding how
the regulation of REV] properly balances survival and mutagenesis in the cell is crucial.
Currently, limited data exists regarding the regulation of REV] gene expression.
Unlike some other genes encoding DNA repair proteins, REV] is not inducible by DNA
damage or heat shock (16). REV1 transcript levels are, however, upregulated during
sporulation in S. cerevisiae (2, 4, 40). At the protein level, our lab has shown that Revl is
under dramatic cell cycle control with protein levels peaking during G2/M rather than S
phase when the bulk of replication occurs (45). Despite the approximately 50-fold change
at the protein level, REV] transcript levels only increase 3-fold during G2/M relative to
GI. Interestingly, RevI is phosphorylated in a similar cell cycle-dependent manner,
demonstrating another potential method of regulation (38). The molecular means
controlling the unexpected cell cycle regulation of Rev 1, however, are not yet fully
understood.
Several genetic studies indicate that TLS may be subject to regulation by the
proteasome. These studies took advantage of the umplA strain, which is a viable mutant
of a gene encoding a maturation factor for the 20S catalytic core of the 26S proteasome
(36). The spontaneous and UV-induced mutator phenotype of the umplA strain is
dependent on the TLS polymerase gene, REV3, which is generally placed in the same
pathway as REVI (23, 32). The umplA strain is hypermutable, whereas rev3A and umplA
rev3A strains are hypomutable, suggesting that Umpl may act as a negative regulator of
Rev3 activity, possibly through Revl's interaction with Pol . The authors of this study,
however, did not examine REVI's genetic interactions with UMP]. In an umplA strain,
short-lived proteins are stabilized and ubiquitin-protein conjugates accumulate (36).
Therefore, we hypothesized the involvement of proteasomal degradation in TLS
regulation as a means for control of this potentially mutagenic process.
Selective protein turnover through ubiquitination and subsequent proteasomal
degradation represents an essential regulatory mechanism in eukaryotic cells. The
irreversibility of protein degradation ensures both spatial and temporal control and
eliminates improper reactivation of the protein. Ubiquitin, a 76-residue protein, is
covalently attached to other proteins through the action of a ubiquitin-activating enzyme
(El), a ubiquitin-conjugating enzyme (E2), and a ubiquitin-ligating enzyme (E3). The
attachment of monoubiquitin or polyubiquitin chains to specific proteins is critical for a
variety of cellular processes from DNA repair to gene silencing, in addition to protein
degradation (11). In general, the attachment of a polyubiquitin chain of at least four
Lys48-linked ubiquitins will target proteins for degradation by the 26S proteasome (14).
The 26S proteasome contains a 20S catalytic core that possesses chymotrypsin-like,
trypsin-like, and peptidylglutamyl peptide hydrolytic activities (42).
Here, we studied the role that proteasomal degradation has in regulating
mutagenic TLS polymerases, especially Rev , the levels of which are cell cycle
regulated. We show that RevI is a moderately short-lived protein throughout the cell
cycle. Our data indicate that Revi undergoes proteasome-mediated degradation during
G1 and G2/M arrest that is potentially targeted through a polyubiquitin modification.
Despite Revi proteolysis occurring during GI and G2/M, there is an expected shorter
half-life for Revi during GI when RevI protein levels are the lowest. Additionally, Rev3
but not Rev7 of the non-cell cycle regulated Pol
is prone to proteasomal degradation.
Overall, these results indicate that proteasomal degradation serves as an efficient and
irreversible mechanism of regulating the potentially mutagenic effects of Rev 1 and Pol
action.
,
MATERIALS AND METHODS
Yeast strains:
A strain list for this study is described in Table 1. All strains are derivatives of the
W1588-4C (MATaleu2-3,112 ade2-1 canl-100 his3-11,15 ura3-1 trpl-1 RAD5) (47)
parent strain. The Revl, Rev3, Rev7, and Rad30 proteins were tagged at their respective
locus with a C-terminal -TEV-ProA-His 7 epitope tag (marked with HIS3) using pYM10
(15), similar to that previously described (5, 45). UMP1 and ERG6 (also called ISE1)
were each separately deleted via a one-step replacement, amplifying the umpl..kanMX4
or erg6::kanMX4 cassette from the deletion library and transforming the product into the
appropriate strain background (43). The BAR] gene was disrupted by a one-step gene
replacement using digested pZV77 to aid in arresting cells with a factor (gift from S.
Bell). The multicopy vector, pMRT7 (pCK322), contains the Pcupl-myc-UBI expression
cassette and the URA3 marker (37) (gift from C. Kaiser). All cassettes and plasmids were
introduced through a standard lithium acetate protocol (9). Oligonucleotide sequences
that were used in strain construction are available up request.
Cell cycle arrest:
Cells were grown in YEPD at 30 C with the exception of umplA strains that
were grown at 25 'C. When the culture reached an OD of 0.5, the cells were split into
two cultures for arrest, one Gl arrested with a factor (50 ng/ml) and the other G2/M
arrested with nocodazole (15 tg/ml). Cells were treated for 3 to 4 hours prior to starting
the assays.
Immunoblots:
Protein extracts were made using a trichloroacetic acid (TCA) procedure similar
to that published (15) with the exception of the Pol rl (Rad30) cycloheximide experiment
that required a SDS boiling method as described by Skoneczna et al. (41). TCA
precipitations were run on 7.5% SDS-PAGE gels (Lonza), and the immunoprecipitation
samples were run on NuPAGE 3-8% tris-acetate gels (Invitrogen) before being
transferred to polyvinylidene difluoride membranes (PVDF, Immobilon-P; Millipore).
PVDF membranes were probed with peroxidase-anti-peroxidase soluble complex (PAP,
Sigma) for ProA-tagged proteins and anti-3-phosphoglycerate kinase (yeast), mouse IgG,
monoclonal antibody (anti-PGK, Molecular Probes) with mouse secondary for the Pgkl
control.
Flow cytometry:
Cells were prepped as in (1) and analyzed on a Becton Dickinson FACSCalibur
flow cytometer.
Cycloheximide chase assay:
GI or G2/M arrested cells were treated with cycloheximide (Sigma) (50 tg/ml)
after the full 3 to 4 hours for arrest. For logarithmic growing cells, cultures were grown to
an equivalent O.D. of 0.7, and then cycloheximide was added at 50 tg/ml to start the
time course. At specific time points, cells were collected for flow cytometry (0.5 ml) or
TCA precipitations (1.5 ml). Cells for TCA precipitations were immediately spun down,
frozen in liquid nitrogen, and stored at -20 C.
Proteasome inhibitor assays:
Cultures were treated with MG132 (Z-Leu-Leu-Leu-al, 50 tM, Sigma) for GI
and G2/M arrested cells. All experiments involving MG132 were completed in an erg6A
(isel A) strain background to allow for MG132 permeability (20). The cells were
collected as in the cycloheximide experiments.
Immunoprecipitations:
Lysis and immunoprecipitations were carried out as described (5) with the
following modifications. The immunoprecipitated strains were subcultured into 500 ml of
SC media lacking uracil (for selection of pMRT7) or histidine (for the strain lacking the
plasmid) and grown to an OD of 0.7 at 25 'C. Copper sulfate (0.5 M) was added to
induce the expression of Myc-tagged ubiquitin. Cells were harvested in 50 ml tubes,
washed in cold water, transferred to and pelleted in 2 ml screw cap tubes, and stored in
lysis buffer at -80 oC until the remaining steps of the lysis and immunoprecipitation
protocol were completed.
RESULTS
S. cerevisiae Revi is unstable during the G1 and G2/M phases of the cell cycle:
Given Rev l's profound cell cycle regulation, we wanted to know to what extent
protein degradation contributes to the significant drop in Rev levels during the G stage
of the cell cycle. Proteolysis influences the cell cycle regulation of many proteins in S.
cerevisiae (29). If protein degradation dictates Revi protein fluctuations throughout the
cell cycle, the protein would have a shorter half-life during the GI stage of the cell cycle
when Revl levels are typically low and a longer half-life during G2/M when protein
levels are the highest. To monitor Revl protein stability, we inhibited translation by
adding cycloheximide to arrested cells, collected samples at subsequent time points, and
visualized ProA-tagged Rev by western blot. REV] was expressed under its native
promoter at the endogenous locus, and the protein produced had a tag at its C-terminus
that does not affect Rev l's contribution to survival and mutagenesis after UV damage
(45). Cells were arrested during the Gi stage of the cell cycle with ca factor and during
the G2/M stages with nocodazole. Contrary to our expectations, the half-life of Revi is
relatively short both during G2/M when protein levels are highest and during G 1 when
protein levels are the lowest (Figure 1, A and B). The flow cytometry data depicts that the
cells remain in the arrested state even after the addition of cycloheximide (IN DNA
content for G1 arrested cells, 2N DNA content for G2/M arrested cells) (Figure 1, C and
D). From our estimations, the half-life of Rev during a GL arrest is shorter than during
the G2/M arrest (approximately 18 minutes versus 32 minutes, respectively) (Figure 1, E
and F). These results suggest that protein degradation acts as a means to keep Revl levels
low throughout the cell cycle in addition to serving as a necessary component of Rev I's
cell cycle regulation.
Inhibition of the proteasome causes an increase in Rev1 protein levels:
Knowing that Revl protein is unstable throughout the cell cycle, we asked
whether this stability was dependent on proteasomal degradation. We used the
proteasome inhibitor, MG132, and assessed Rev protein levels following treatment. All
experiments involving MG 132 were performed in an erg6A background to allow the drug
to enter the cells. The steady-state level of Rev protein increased in the presence of the
proteasome inhibitor for Gi and G2/M arrested cells (Figure 2, A and B). Flow cytometry
analysis confirmed that the erg6A cells arrest normally in the absence or presence of
proteasome inhibitor (data not shown, also see Figure 3, C and D).
The proteasome is responsible for Rev1's relatively short half-life:
To monitor the effect that the disruption of proteasome function has on the halflife of Revl protein, G1 and G2/M arrested cells were pre-incubated with MG132 for 30
minutes. The time course began with the addition of cycloheximide. The half-life of Rev1
during G 1 or G2/M arrest is longer in the presence of proteasome inhibitor than when
translation is inhibited in its absence (Figure 3, A and B). The flow cytometry data does
not show any abnormalities for the arrests (Figure 3, C and D). As seen in another report
(8), MG 132 does not completely prevent degradation of Revl in this cycloheximidechase assay.
Rev1 steady-state levels increase when proteasome function is defective:
To further support our proteasome inhibitor results with a genetic approach, we
utilized one of the viable mutants associated with the assembly of the proteasome
(umpl A), which lacks the gene encoding a maturation factor for the 20 S proteasome. In
cells that have been arrested by a factor or nocodazole, the steady-state levels of Revl
are significantly greater in the umplA cells than in wild type cells during GI and are
moderately increased during G2/M (Figure 4, A and B at time points 0) similar to the
effect observed after the addition of the proteasome inhibitor drug. These experiments
were carried out at 250 C instead of 300 C to avoid problems with the temperature
sensitivity of an umplA strain. After the addition of cycloheximide, the half-life of Revl
in the umplA strain during G2/M arrest does not seem to significantly differ from the
estimated half-life in wild type from Figure IF (Figure 4B). Interestingly, this result
differs from the noticeable change in the half-life of Rev1 after treating cells with
MG132. Since Rev1 levels are much greater in the umplA background during the GI
arrest, Rev1 is not shown in the wild type blot at the selected exposure (Figure 4A). The
flow cytometry reveals that the umplA cells fail to properly arrest in nocodazole (Figure
4, C and D). Since more cells accumulate with IN DNA content (when Revl levels are
the lowest) in an umplA background than in a wild type background, Revl protein levels
for these time points are, if anything, an underestimate. These results support that the
proteasome is involved in the degradation of RevI and are consistent with the data using
MG 132. The differences seen between the proteasome inhibitor and proteasome
assembly mutant experiments can be attributed to the pleotropic effects of deleting
UMP1.
Higher migrating forms of Revi indicate targeting of the protein to the proteasome:
After learning that disruption of proteasome function affected Rev1 protein levels,
we decided to assess if Revi is targeted to the proteasome. In general, the attachment of a
polyubiquitin chain of at least four Lys48-linked ubiquitins will target proteins for
degradation by the 26S proteasome (14). To detect higher migrating forms of Revl and
subsequently test if these forms represent polyubiquitinated Rev 1, we
immunoprecipitated ProA-tagged Revl in an umplA strain background (Figure 5, lane 2).
This strain also included myc-tagged ubiquitin under a copper-inducible promoter. We
compared this immunoprecipitation to one of ProA-tagged Revl in an UMPI strain
lacking myc-tagged ubiquitin (Figure 5, lane 1) and to another immunoprecipitation of
non-tagged Revl in an umplA strain with myc-tagged ubiquitin (Figure 5, lane 3). When
probing for ProA-tagged Rev 1, a significant smear appears above the Revl band for the
immunoprecipitation done in the presence of myc-tagged ubiquitin in an umplA
background (Figure 5, lane 2). As expected, no RevI is detected when the protein lacks
the ProA tag (Figure 5, lane 3).
The blot for myc-tagged ubiquitin with anti-myc after immunoprecipitation
showed no bands and for anti-ubiquitin did not show any distinct bands or smears
corresponding to Revl 's migration or higher that were specific to the ProA-tagged Rev I
immunoprecipitations (data not shown). In both cases, this could be due to Revl protein
levels still being very low even after the immunoprecipitation, given that they are only
detectable with by immunoblot and not silver stain. The anti-myc or anti-ubiquitin may
not be sensitive enough to detect the fraction Rev that is modified. Also, the anti-
ubiquitin blot is not ideal, since a lot of background exists on the immunoblot despite
immunoprecipitating the protein of interest. Alternatively, carrying out the experiment in
the presence of MG 132 instead of using the umplA strain may yield better detection of
polyubiquitination.
Although these results do not conclusively reveal polyubiquitination of S.
cerevisiae Rev 1, the higher migrating smear in lane 2 (Figure 5) implies that a modified
form of RevI exists and is more easily detected in the umplA strain background. Since
the deletion of umpl is known to cause an accumulation of ubiquitin-conjugated proteins
(36), the smear in lane 2 (Figure 5) may represent polyubiquitinated Rev . Aside from
technical difficulty, another explanation for the lack of a band or smear on the ubiquitin
immunoblot is that RevI's degradation is ubiquitin-independent and possibly dependent
on another protein modification.
Pol
is also subject to proteasomal degradation:
Since Rev I protein levels are under the control of degradation by the proteasome,
we next examined whether the other TLS polymerases, Pol
and Pol fl (Rad30), were
also subjected to proteasome-dependent degradation. To address this question, we
compared the levels of ProA-tagged Rev3 (of Pol ) expressed from its endogenous locus
in wild type and umpl A cells in the presence of cycloheximide. We used asynchronous
cultures, since Rev3 protein levels are not cell cycle regulated (45). At time 0, Rev3
steady-state levels are greater in an umpl A background (Figure 6A). Moreover, the halflife of Rev3 appears to increase in an umpl A background. These results indicate that
proteasomal degradation functions to control the levels of an additional protein involved
in TLS that is not cell cycle regulated.
The protein levels of ProA-tagged Rev7, a subunit of Pol , were compared
between asynchronous wild type and umpl A cells as well. As with Rev3 and Rev7, the
protein is expressed from the endogenous locus and tagged at the C-terminus without
interfering with function. Unlike the catalytic subunit of Pol , Rev3, Rev7 protein levels
do not change in an umplA strain background (Figure 6B). Proteasomal degradation,
therefore, does not control the protein levels for both components of Pol 's heterodimer.
Skoneczna et al. previously reported that Pol ql (Rad30) is short-lived due to
proteasomal degradation (41). We see the decrease in Pol i1 (Rad30) protein levels after
the addition of cycloheximide only when using a boiling method for protein extraction
(Figure 6C). The degradation of Rev 1, however, is observable with TCA precipitations
(Figures 1, 3, and 4) and the boiling method (data not shown). The extent to which
method better represents the actual in vivo fate of Pol 'r (Rad30) protein is uncertain.
DISCUSSION
Since Revl protein levels fluctuate -50-fold as cells go from the GI to G2/M
stages of the cell cycle and transcript levels only undergo a -3-fold change (45), we
hypothesized that proteolysis contributes to the cell cycle regulation of Rev 1. Indeed, we
find that Revl has a relatively short half-life during G and G2/M that is dependent on
the proteasome function with the half-life during GI being less than during G2/M.
Degradation by the proteasome is not the sole mechanism for cell cycle regulation. Since
the transcriptional and proteasomal factors of regulation are not dramatic, translational
control or another posttranscriptional mechanism also must contribute to the significant
increase of Rev1 protein levels during G2/M.
Degradation by the proteasome serves as an excellent mechanism to ensure the
proper timing and positioning of a protein for action. Proteolysis eliminates the protein in
an effective and irreversible way to prevent action and can destroy aberrant proteins. For
potentially mutagenic TLS polymerases, ensuring that these polymerases do not interfere
with the replicative DNA polymerases and only function when needed is critical to avoid
widespread mutagenesis. These results demonstrate that S. cerevisiae uses proteasomal
degradation to keep Rev1 and Pol t protein levels low in general or low at specific times
in the cell cycle. Similarly, TLS proteins are regulated by degradation in E. coli. For
example, UmuC, the catalytic subunit of Pol V, undergoes proteolysis by the Lon
protease (7).
In higher eukaryotes, more recent data is emerging that other DNA polymerases
involved in base excision repair and capable of TLS, Pols k and [3, are targeted for
proteasome-mediated degradation (28, 46). Interestingly, phosphorylation of Pol k
stabilizes the protein during late S and the G2/M stages of the cell cycle. These are the
same cell cycle stages that Rev1 protein levels are the highest and phosphorylated in S.
cerevisiae (38). Future work will be required to know if Revl degradation is modulated
by phosphorylation or if Revl is targeted for degradation through another protein
modification. In fact, another explanation for the higher migrating band in Figure 5 and
for the inability to detect ubiquitin is that this smear is actually hyperphosphorylated
Rev from cells in their G2/M stages of the cell cycle that are stabilized in the umplA
strain.
A few examples of ubiquitin-independent proteasomal degradation exist such as
Spel degradation mediated by the interaction with Oazl in S. cerevisiae (33). If not
polyubiquitination, the higher migrating form of Revl could represent another protein
interacting with Rev1 or phosphorylation of specific residue(s) or the addition of
alternative protein modifications. If the higher migrating form of Rev1 is due to
polyubiquitination, then the attachment of polyubiquitin on Revl will involve an E2 and
E3 ubiquitin ligase. The timing of the anaphase-promoting complex/cyclosome (APC/C)
activity coincides with the lowest levels of Revl occurring during G1. The APC/C, a
multisubunit ubiquitin-protein ligase, controls cell cycle progression by targeting key
proteins for 26S proteasome degradation during late mitosis and Gl (29). As shown here
though, Revl is degraded throughout the cell cycle and therefore may not be a substrate
for classical ubiquitin ligases associated with cell cycle regulation.
Curiously, proteolysis regulates the protein levels of Rev3 but not Rev7. Since
Rev3 is the catalytic subunit of Pol t, degradation of Rev3 is more significant in
preventing error-prone DNA synthesis. Rev3 is much larger than Rev7, and REV3 has
upstream, out-of-frame ATG codons, so the translation of REV3 is likely less efficient
than REV7 as well (18). Potentially, the additional protein-protein interactions or
functions of Rev7 may prevent its degradation or make Rev7 undesirable to degrade at
the same time as Rev3. Aside from the interaction with Rev3 in S. cerevisiae, Rev7 also
binds Ddc 1 and Mec3 of the 9-1-1 clamp, which is involved in the DNA damage
checkpoint (39). In higher eukaryotes, Rev7 interacts with several cell cycle proteins (3,
24, 30).
ACKNOWLEDGEMENTS
I thank members of the Walker lab for helpful discussions, Kevin Wang for
critically reading the manuscript, and members of Drs. S. P. Bell and C. Kaiser's groups
for strains and materials. This work was supported by a National Institute of
Environmental Health Sciences (NIEHS) grant 5-R01-ES015818 to G.C.W., a NIEHS
grant P30 ES002109 to the MIT Center of Environmental Health Sciences, and an
American Cancer Society Research Professorship to G.C.W.
Table 1. Yeast strains used in this study
Strain
YLW70
Revl-ProA
Revl-ProA umplA
Revl-ProA erg6A
Rev3-ProA
Rev3-ProA umplA
Rev7-ProA
Rev7-ProA umplA
Rad30-ProA
W1588-4C umplA +
pMRT7
Revl-ProA umplA +
pMRT7
Relevant Genotype
W1588-4C barlA::LEU2
W1588-4C barlA::LEU2
REVI-TEV-ProA- 7HIS
same as Rev -ProA but
umpl A::kanMX4
same as Revl-ProA but
erg6A::kanMX4
W1588-4C barlA::LEU2
REV3-TEV-ProA- 7HIS
same as Rev3-ProA but
umplA::kanMX4
W1588-4C barlA::LEU2
REV7-TEV-ProA-7HIS
same as Rev7-ProA but
umplA::kanMX4
W1588-4C barlA::LEU2
RAD30-TEV-ProA- 7HIS
W1588-4C barlA::LEU2
umplA::kanMX4 pPcup,myc-UBI
Same as Revl-ProA umplA
with pPcup;-myc-UBI
Source
(5)
(45)
This study
This study
(gift from S. D'Souza)
This study
(gift from S. D'Souza)
This study
(45)
This study
This study
All strains are derivatives of W1588-4C (MATa leu2-3,112 ade2-1 canl-100 his3-11,15
ura3-1 trpl-1 RAD5) (47).
FIGURE 1. - Rev 1 protein experiences turnover during the G and G2/M phases of the
cell cycle. (A) Revl levels decrease during GI arrest after cycloheximide treatment. The
cells from the Revl-ProA strain were arrested with ca factor at 300 C, and then split into a
cycloheximide treated culture and a non-treated culture before time points were collected
each hour. Immunoblots are probed with PAP for the ProA-tagged Rev and with antiPgkl for the Pgkl loading control. (B) Revl levels decrease during G2/M arrest after
cycloheximide treatment. The assay was performed as in (A), except that the arrest was
done with nocodazole. (C) Revl-ProA cells stay arrested after cycloheximide treatment
with IN DNA content. Flow cytometry data is shown for c factor arrested cells with and
without cycloheximide. (D) Revl-ProA cells remain in G2/M arrest after the addition of
cycloheximide. Flow cytometry data is given for nocodazole arrested cells in the
presence and absence of cycloheximide. (E) The half-life of Rev in GI arrested cells is
less than in G2/M arrested cells. The half-life was estimated to be 18 minutes. The assay
for the imminoblot was carried out as in (A), except that time points were taken at
smaller intervals. (F) The half-life of Rev1 in G2/M arrested cells is greater than in GI
arrested cells. The half-life is estimated to be 32 minutes. The assay was completed as in
(E), except that cells were arrested with nocodazole.
Figure 1
nocodazole arrest (G2/M)
a factor arrest (Gi)
no drug + cycloheximide
0 1 2 3 0 1 2 3hours
no drug + cycloheximide
0 1 2 3 0 1 2 3hours
Revl
Pgkl
I ---
-.
P
I
Revl
Pgkl
c
3
+ cycloheximide
no drug
2
+ cycloheximide 2
0
0
3
3
2
2
no drug
1
0
Ime
0
Time
1N2N
a factor arrest (G1)
no drug
1N 2N
(hours) DNA Content
)urs) DNA Content
nocodazole arrest (G2/M)
no drug + cycloheximide
+ cycloheximide
0 15 30 60 120 0 15 30 60 120 minutes
0 15 30 60 120 0 15 30 60 120 minutes
Revl
Revl
Pgkl
Pgkl
FIGURE 2. - Proper proteasome function regulates Revl protein levels. (A) Revl
protein levels significantly increase in the presence of the proteasome inhibitor, MG132,
in GI arrested cells. Cells were arrested with ca factor at 300 C, and then divided into a
MG 132 treated and non-treated cultures before time points were taken. The strain
background is erg6A. The immunoblot was probed with PAP for ProA-tagged Rev and
anti-Pgkl for the Pgkl loading control. (B) Revl protein levels are also greater after
MG 132 treatment in G2/M arrested cells. The assay was carried out as in (A), apart from
the arrest being done with nocodazole.
100
Figure 2
nocodazole arrest (G2/M)
a factor arrest (Gi)
no drug
no drug
+ MG132
Revl
Pgkl
+ MG132
0 15 30 60 120 0 15 30 60 120 minutes
0 15 30 60 120 0 15 30 60 120 minutes
Revl
Pgkl
101
FIGURE 3. - Inhibiting proteasome action greatly prolongs the half-life of Revl. (A)
Revl protein levels are stabilized when the proteasome is inhibited during Gl. After a
pre-incubation of ca factor-arrested cells with MG 132 at 300 C, cycloheximide was added
to start the time course. The strain background is erg6A. The immunoblot shows Protagged Revl and the loading control, Pgkl. (B) The half-life of Revl during G2/M is also
lengthened in the presence of MG132. The assay was performed as in (A), but cells were
arrested with nocodazole. (C) Cells maintain IN DNA content after MG132 and
cycloheximide treatment. Flow cytometry data is shown for select time points in the
presence of cycloheximide alone or cycloheximide and MG132. (D) MG132 does not
affect the nocodazole arrest. Flow cytometry data is depicted as in (C), apart from the
cells being in a G2/M arrest.
102
Figure 3
nocodazole arrest (G2/M)
a factor arrest (Gi)
+ MG132/
+ cycloheximide cycloheimide
cyclohexmide
+ MG132/1
+ cycloheximide cychexmde
cycloheximide
0 15 30 60 120 0 15 30 60 120 minutes
0 15 30 60 120 0 15 30 60 120 minutes
Revl
Revl
Pgkl
Pgkl k~L""slsls9ii$sl"~'~
120
120
+ MG132/
cycloheximide 60
+ MG132/
60
cycloheximide
o
0
0i
120
120
+ cyclohexlmide0
+ cycloheximide 6
0
Time
(hours)
Time
D1 n
1N 2N
(hours) DNA Content
DNA Content
103
FIGURE 4. - The steady-state levels of Rev are greater in a proteasome-defective strain
background. (A) Revl protein levels increase in the umplA background during GI arrest.
Cells were arrested with c factor at 250 C, and then treated with cycloheximide to start
the time course using the Revl-ProA or Revl-ProA umplA strains. The immunoblot
shows ProA-tagged Rev1 and Pgkl as a loading control. The difference in half-life
cannot be judged from this immunoblot. (B) Revl protein levels are greater in the umplA
background during G2/M arrest. The assay was completed as in (A), except that the cells
were arrested with nocodazole. The half-life does not seem to be affected much. (C) The
umplA strain background does not change the ability of cells to arrest with 1N DNA
content. Flow cytometry is shown for the Revl-ProA or Revl-ProA umplA strains
during a factor arrest. (D) The umplA cells accumulate more with IN DNA content
during nocodazole arrest than the wild type background. Flow cytometry shows the DNA
content for G2/M arrested cells of the Revl-ProA or Revl-ProA umplA strains.
104
Figure 4
nocodazole arrest (G2/M)
a factor arrest (G1)
umplA
wild type
+ cycloheximide + cycloheximide
umplA
wild type
+ cycloheximide + cycloheximide
0 15 30 60 120 0 15 30 60 120 minutes
0 15 30 60 120 0 15 30 60 120 minutes
Rev1
Revi
Pgkl
Pgkl
C
120
ump lA
umpl A
o60
30
15
0
120
60
wild type 30
wild tvp
15
0
1N 2N
Time
(hours) DNA Content
105
FIGURE 5. - A higher migrating form of Revl exists when UMP1 is deleted and myctagged ubiquitin is overexpressed. Strains are Revl-ProA (lane 1), Revl-ProA umplA +
pMRT7 (lane 2), and W1588-4C umplA + pMRT7 (lane 3). The immunoblot for the
Rev 1-ProA immunoprecipitation samples were probed with PAP for ProA-tagged Rev1.
106
Figure 5
123
- 250 kDa
- 150 kDa
Revl
- 100 kDa
107
FIGURE 6. - Proteasome function also affects the protein levels of Rev3 (of Pol t). (A)
Rev3 protein levels increase in the umplA strain background. At an equivalent O.D.,
cycloheximide was added to asynchronous cells at 25 C to start the time course. Rev3ProA is probed with PAP in the immunoblot, and Pgkl is shown as the loading control.
Strains are Rev3-ProA and Rev3-ProA umplA. (B) Rev7 protein levels do not change in
the umplA strain background. The assay was carried out as in (A), except Rev7-ProA is
shown in the immunoblot. Strains are Rev7-ProA and Rev7-ProA umplA. (C) The
protein levels of Pol r (Rad30) are unstable under certain conditions. Asynchronous cells
were treated with cycloheximide at 30 oC and collected at specific time points. In the
immunoblot, Rad30-ProA is probed with PAP for detection and Pgkl with anti-Pgkl as
the loading control. The Rad30-ProA strain was used.
108
Figure 6
ump1A
wild type
+ cycloheximide + cycloheximide
0 15 30 60 120 0 15 30 60 120 minutes
Rev3
Pgkl
umplA
wild type
+ cycloheximide + cycloheximide
0 15 30 60 120 0 15 30 60 120 minutes
140,
Rev7
wild type
+ cycloheximide
0 15 30 60 120 minutes
Rad30
Pgkl
109
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114
Chapter 3
The DNA Polymerase Activity of Saccharomyces cerevisiae
Revi is Biologically Significant in a Lesion-Specific Manner
This chapter will be submitted for publication. The authors are Mary Ellen Wiltrout and
Graham C. Walker.
115
ABSTRACT
The cell's ability to tolerate DNA damage relies on error-prone translesion
synthesis (TLS) and an error-free, template-switching pathway. The primary proteins
mediating TLS in Saccharomyces cerevisiae are three DNA polymerases (Pols), Rev1,
Pol
(Rev3/7), and Pol r (Rad30). Revl's non-catalytic role in recruiting other DNA
polymerases is known to be important for TLS. The biological significance of Rev l's
conserved DNA polymerase activity is much less well understood. Here, we demonstrate
that inactivating Rev1's DNA polymerase function sensitizes cells to both chronic and
acute exposure to 4-nitroquinoline- 1-oxide (4-NQO) but not to other DNA damaging
agents (UV or cisplatin). Full Revl-dependent resistance to 4-NQO, however, also
requires the additional Rev1 functions. When error-free tolerance is disrupted through the
deletion of MMS2, Rev 1's catalytic activity is more vital for survival after 4-NQO
damage. In the presence or absence of error-free tolerance, the catalytic dead strain of
Revl exhibits a lower 4-NQO-induced mutation frequency than wild-type. Furthermore,
Pol , but not Pol r, also contributes to 4-NQO resistance. These results show that
Revl's catalytic activity is important in vivo when the cell has to cope with specific DNA
lesions, likely N2 -dG adducts commonly produced by 4-NQO and methylglyoxal. Our
data reveal that the cell relies on Mms2-dependent error-free tolerance for 4-NQO
resistance when Rev 1-polymerase-dependent TLS is unavailable, possibly explaining
why the biological significance of Rev1's catalytic activity has been largely overlooked.
116
INTRODUCTION
The integrity of a cell's genome is constantly threatened by endogenous and
exogenous sources of DNA damage that range from byproducts of metabolism to UV
radiation from sunlight (11). Cells possess multiple DNA repair and tolerance
mechanisms to avoid the negative consequences of DNA damage such as stalls in
replication, strand breaks, mutations, or even cell death. Whereas DNA repair restores the
DNA to the original state, DNA damage tolerance allows DNA lesions to remain in the
genome without disturbing cellular processes. As a means to ensure the complete
transmission of genetic material to the daughter cells, two major classes of DNA damage
tolerance are available: translesion synthesis (TLS) and an error-free, template-switching
pathway.
During TLS, specialized DNA polymerases (Pols) catalyze DNA synthesis across
from DNA lesions. TLS may take place during S phase or later in the cell cycle to fill in
persistent gaps in the DNA [as reviewed in (56)]. TLS polymerases are found in all
domains of life (45). Of the three TLS polymerases in S. cerevisiae, only Pol
(Rev3/Rev7 heterodimer) belongs to the B family of DNA polymerases, which also
includes the highly processive, replicative DNA polymerases (Pols 6 and e in S.
cerevisiae). In contrast, Rev1 and Pol T1 (Rad30) are members of the Y family of DNA
polymerases that includes only TLS polymerases (45). TLS DNA polymerases are
structurally different from their replicative counterparts in that they lack a 3' to 5'
exonuclease domain and possess a more open active site (18, 49). These differences
contribute to TLS polymerases having a reduced fidelity and lower processivity
117
compared to the replicative DNA polymerases and allow for action on suboptimal DNA
templates (38, 49).
Rev1, Pol
,,
and Pol rq are specialized to perform different molecular functions.
For example, although TLS polymerases increase a cell's resistance to DNA damage,
Rev and Pol
are notable for their prominent roles in mutagenesis as characterized by
the reversionless (unmutable) phenotypes caused by REV1/3/7 mutants in spontaneous
and damage-induced mutagenesis assays (15, 30-32). Furthermore, unlike Revi and Pol
,1, Pol
is primarily thought to extend after catalytic insertion by another TLS
polymerase or even itself, especially after the incorporation of mismatches (2, 30, 31, 64).
Pol ri is specialized to perform error-free bypass of UV cis-syn cyclobutane pyrimidine
dimers (25). In humans, loss of Pol r function leads to a variant form of the cancer-prone
disease xeroderma pigmentosum (XPV) (34, 37, 60).
The in vivo role for this catalytic function remains ill-defined. Rev was
originally described as a deoxycytidyl transferase (43) but was later reclassified as a
DNA polymerase of limited function (59) and a dG template-specific DNA polymerase
(22, 62) in in vitro studies. The in vivo phenotypes of cells lacking Revl's DNA
polymerase activity are not as abundant as cells completely missing Rev1. For example,
Revl's DNA polymerase activity is not required for resistance to cisplatin and UV
damage in DT40 cells (51) and is dispensable for methyl methanesulfonate (MMS)induced mutagenesis in S. cerevisiae (23). In contrast, Rev1's catalytic activity
contributes to immunoglobin diversification (50). Other studies in S. cerevisiae provide
evidence that the catalytic activity of Rev1 inserts dCMPs across from abasic sites (4, 13,
118
14, 42, 46-48). This in vivo activity of RevI is consistent with in vitro studies showing
insertion across from abasic sites (43).
The mechanism and capabilities of Rev1 as a DNA polymerase have been
structurally and biochemically characterized. The crystal structure of Revl's polymerase
domain bound to an undamaged DNA template revealed a unique mechanism of proteinmediated selection of the template base and incoming nucleotide, rather the DNA
template dictating the incoming nucleotide by base-pairing (41). The incoming dCTP
hydrogen bonds with Arg324 in S. cerevisiae Revl, while the template dG is flipped out
to interact with another region of Rev 1. A more recent structure confirms that Rev l's
active site can accommodate a bulky adduct at the N 2 position of guanine (40). This is
consistent with multiple in vitro studies displaying efficient catalysis of RevI across from
a variety of N 2-dG lesions (6, 40, 55, 62). After insertion by Revl, these reports also
show extension of DNA synthesis past the N 2-dG lesion by Pol t (55) or Pol K for human
Revl (62). The insertion of dCMP across from N2-dG lesions by RevI is an error-free
process, unlike insertion across from abasic sites.
Although curious, the lack of strong evidence for Rev 1's action as a DNA
polymerase in contributing to cellular viability after DNA damage could be explained by
Revl's "second function" as originally termed by Nelson et al. (42). Since this
observation that Revl is required for bypass of T-T (6-4) photoadducts, a situation in
which dCMP is rarely inserted, many studies have elucidated Revl's roles in TLS by
addressing the importance of the other non-catalytic protein domains in Revl,
particularly the BRCT (BRCA1 C-Terminus), UBM2 (ubiquitin-binding motif), and Cterminal domains. The analysis of the original loss-of-function mutation, rev]-1
119
(G 193R), gave the first clue that the functionality of the N-terminal BRCT domain of
Rev was significant for DNA damage resistance and mutagenesis in S. cerevisiae
despite the protein maintaining 60% of the catalytic activity in vitro (42). The BRCT
domain of RevI interacts with PCNA (20) and promotes the affinity of Revl for ssDNA
(36). Mutants in the UBM2 of Revl also display decreased survival and mutagenesis in
response to DNA damage that is likely caused by a less efficient interaction with
monoubiquitinated PCNA (9, 21, 58). Additional studies found that the last -100 amino
acids of Revl could interact with the other TLS polymerases in mammalian cells (19, 39,
44, 53) and in S. cerevisiae (1, 2, 8, 9, 27). Not surprisingly, truncations of, or point
mutations in, the C-terminus cause severe survival and mutagenesis defects after DNA
damage (2, 9, 27, 29, 51). Given that these domains bind other proteins and contribute to
survival and mutagenesis for a spectrum of DNA damage, it has been proposed that Revl
acts a scaffolding protein that promotes the recruitment of TLS polymerases to sites of
DNA adducts and/or aids in the recognition of DNA lesions (56).
In addition to TLS, another tolerance mechanism is available when DNA
polymerases encounter a DNA lesion, referred to as error-free tolerance. The description
as error-free originated from the fact that mutants of the genes involved in this pathway
lead to an increase in mutagenesis (rather than a decrease, as for REV1/3/or 7 mutants)
(28). Unlike the TLS branch of tolerance, error-free tolerance involves template
switching for bypassing lesions. The mechanism of action and identities of the
participating proteins are not fully understood, but a particular modification on
proliferating cell nuclear antigen (PCNA) is instrumental in the process. Whereas TLS is
associated with the monoubiquitination of PCNA by Rad6/Radl 8, the error-free tolerance
120
branch relies on the Lys-63 linked polyubiquitination of PCNA (3). This
polyubiquitination of PCNA is completed through the action of Rad5 and the
Mms2/Ubc 13 heterodimer. Despite the distinctions in how the tolerance pathways bypass
a lesion, there is clear functional redundancy between TLS and error-free tolerance. For
example, Rev3 and Mms2 have a synergistic relationship in response to DNA damage
(5).
The circumstances that require Rev1 catalytic activity remain unclear. Here, we
report that the DNA polymerase activity of Rev1 provides resistance to 4-nitroquinoline1-oxide (4-NQO) but not UV or cisplatin exposure. This resistance also requires the
functionality of the BRCT, UBM2, and C-terminal domains of Revl. The necessity of the
DNA polymerase domain of Revl for tolerance of 4-NQO lesions and the endogenously
occurring aldehyde, methylglyoxal, is more striking in the absence of the error-free
tolerance pathway (mms2A background). The catalytic dead mutant of Rev1 also displays
a lower 4-NQO-induced mutation frequency in cells proficient or deficient for error-free
tolerance. Additionally, Rev1 depends on Pol
to contribute to cellular viability after 4-
NQO damage. Overall, these results provide evidence that Rev1 acts catalytically to
bypass DNA lesions produced by 4-NQO or methylglyoxal exposure, agents that cause
N 2-dG adducts. In wild-type S. cerevisiae, however, this catalytic function is masked by
the redundancy between TLS and the error-free DNA damage tolerance pathway.
121
MATERIALS AND METHODS
Yeast strains:
Table 1 displays a complete list of strains used in this study. A W1588-4C (MA4 Ta
leu2-3,112 ade2-1 canl-100 his3-11,15 ura3-1 trpl-1 RAD5) (63) derivative (also
barlA::LEU2 and rev1A::kanMX4) (9, 57) is the isogenic parent of all strains involving
the integrated REV1 and mutated revl alleles. These integrations at the REV1 locus were
tagged with a C-terminal -TEV-ProA-His 7 epitope tag (marked with HIS3) using pYM10
(26), similar to that previously described (9, 57). Vector DNA was removed by selecting
for 5-fluoroorotic acid (5-FOA) resistant colonies (ura auxotrophs). Final strains were
G418s, 5-FOAR, and HIS3 and were sequenced to ensure that the intended REV]
genotype was integrated. S288C (MA Ta SUC2 gal2 mal melflol flo8-1 hapl ho biol
bio6) (35) served as the parent for all plasmid (pRS416)-bearing strains. REV] was
deleted via a one-step replacement, amplifying the revl..'kanMX4 cassette from the
deletion library and transforming the product into the S288C strain (54). Additional
deletions were made with a one-step replacement using PCR-generated cassettes
containing hygMX4 (MMS2) or natMX4 (RAD30, REV3, REV7) with 5' and 3' homology
to the gene of interest (17). The BAR1 gene was disrupted by a one-step gene replacement
using digested pZV77 (gift from S. Bell). All cassettes and plasmids were introduced
through a standard lithium acetate protocol (16).
Plasmids:
The construction of the pRS306-REV1 (for integration) and pRS416-REV1
plasmid were previously described (9). The pRS306-REV1 plasmid and site-directed
122
derivatives were digested at the SexAI site in the REV] promoter and transformed into
the W1588-4C barlA rev1A strain. Integrated constructs were selected on synthetic
complete medium lacking uracil plus 2% glucose (SC-Ura) and verified by PCR. The
integrated REV1 and mutant revl alleles were then tagged as described above.
Site-directed mutagenesis:
The site-directed mutagenesis of pRS306-REV1 and pRS416-REV1 was
performed following the QuikChange Site-Directed Mutagenesis Kit (Stratagene)
protocol with the exceptions of a 2 min/kb extension time and 50 'C annealing
temperature. The mutations were all verified by sequencing. The resulting amino acid
changes are summarized in Table 2.
Oligonucleotides:
The sequences of oligonucleotides used for strain construction are available upon
request.
Survival assays:
All cultures were inoculated with individual colonies. The calculated values for
percent survival represent the results from at least three independent clones of each strain.
The cultures were grown to saturation (-48 hours) while rotating at 30 oC in 5 ml SC-Ura
media for chronic exposure or in 25 ml SC-Ura media for acute exposure in liquid. For
chronic DNA damage, the cells were diluted as 10-fold serial dilutions to 10-4 in sterile
water. 100 cl of the appropriate dilution to achieve single colonies was plated in
duplicate on SC-Ura plates alone or SC-Ura plates containing a concentration of drug as
indicated. For acute exposure in liquid, the specified drug dose was added to 5 ml
123
aliquots of saturated cells and allowed to rotate at 30 C for 1 hour. Cells were washed in
sterile water, diluted, and then plated on SC-Ura plates.
Cells for the methylglyoxal experiments were plated on YEPD plates with and
without methylgloxal. UV-treated cells were exposed to UV at 1 J/m 2s with a G 15T8 UV
lamp (General Electric) at 254nm. Stock solutions of 4-NQO at 2 mg/ml (Sigma) in N, N
- dimethylformamide or freshly prepared cis-diammineplatinum (II) dichloride (Sigma)
at 0.8 mg/ml in sterile water were added directly to the desired liquid media or agar
media for plates at the appropriate concentration. Methylgloxal (Sigma) was added to
agar media directly from the 40% solution as sold. When necessary, drug-containing
plates were stored at 4 oC overnight (at most) before the day of the experiment. After
plating, individual colonies were counted after 3 days of growth at 30 oC, with the
exception of methylgloxal plates that were counted after 5 days due to the slow growth of
some strains. Percent survival was calculated as the number of survivors after exposure to
drug divided by the number of colony forming units without any drug exposure.
Mutagenesis assays:
Generally, the procedure for mutagenesis assays was similar to that of survival
assays and included a survival assay for use in calculating the frequency of canavanine
resistant mutants. In addition to the survival plates with and without DNA damage
exposure, 100 tl of saturated culture was plated in duplicate on SC-Ura-Arg
supplemented with canavanine (30 [tg/ml) (Sigma) plates for each culture. These cells
were in the presence and absence of DNA damage in the plates or in liquid as noted in the
figures. Colonies were counted after five days of growth at 30 oC. The mutation
frequency was calculated as the average number of colonies on the canavanine SC-Ura-
124
Arg plates divided by the calculated total number of survivors for the same dose of drug
exposure. The mutation frequency was expressed as the number of canavanine resistant
colonies per one million survivors.
Preparation of cells for immunoblots:
For cell cycle arrests, a similar procedure was followed as previously published
(9, 57). Briefly, cells were collected after 4 hours of arrest in a factor or nocodazole at 30
C. Logarithmically growing cells were harvested at an equivalent O.D. for each strain.
Immunoblots:
To examine the level of protein expression from different revl alleles relative to
the REV1 gene integrated into the chromosome, protein extracts were made using a
trichloroacetic acid (TCA) procedure similar to that published (26). Samples were run on
a 7.5% SDS-PAGE gel (Lonza) and transferred to polyvinylidene difluoride membranes
(PVDF, Immobilon-P; Millipore) using a Mini-PROTEAN II transfer apparatus (BioRad). PVDF membranes were probed with rabbit peroxidase-anti-peroxidase soluble
complex (PAP, Sigma) for protein A-tagged proteins and anti-3-phosphoglycerate kinase
(yeast), mouse IgG, monoclonal antibody (anti-PGK, Molecular Probes) with mouse
secondary for the Pgkl control.
125
RESULTS
The DNA polymerase function of S. cerevisiae Rev1 has lesion-specific requirements
in vivo:
To gain insight into the circumstances under which Rev acts as a DNA
polymerase in the cell, we took advantage of an allele of RE V that encodes a
catalytically inactive protein (23). This mutant substitutes alanine for Asp467 and
Glu468, which are the acidic residues that coordinate the two metal ions required for
Rev1 catalysis. We refer to this allele and protein product as "catalytic dead". The wild
type, catalytic dead (integrated and tagged allele at the native locus), and rev1A strains
were exposed to DNA damaging agents in the agar growth medium, and the percent
survival was calculated for each strain at several doses. Previous studies have shown that
the epitope-tagged version of Revl used here retains wild type function for DNA damage
resistance and mutagenesis (57). If the catalytic dead strain is sensitive to a particular
DNA damaging agent relative to the wild type strain, then the DNA polymerase activity
of Revl plays a physiologically important role in tolerating the lesions created by that
agent in vivo. We were particularly interested in whether Revl's DNA polymerase
activity might be critical for the bypass of N2-dG adducts that commonly result in vivo
from the reaction of lipid peroxidation products with DNA (7). We chose 4-NQO as a
representative chemical that primarily causes N2 -dG lesions unlike UV or MMS (11).
We found that the integrated catalytic dead mutant had a dose-dependent
increased sensitivity to 4-NQO. Although this sensitivity is modest, the catalytic dead
strain shows no increased sensitivity to UV damage (Figure 1, A and B). UV light
introduces lesions such as cyclobutane pyrimidine dimers and T-T (6-4) photoadducts
126
(11) that Revl cannot bypass in vitro (62). The catalytic dead strain, however, is not as
sensitive to killing by 4-NQO as the rev1A strain, indicating that the other non-catalytic
functions of Revi contribute to 4-NQO resistance independent of the DNA polymerase
activity. These data demonstrate that DNA damage tolerance after exposure to 4-NQO is
enhanced by the DNA polymerase function of Revi and suggests that the DNA
2
polymerase function could be important for other agents that make N -dG lesions.
The assay for UV sensitivity exposes cells to the DNA damaging agent for a set
amount of time. In contrast, the 4-NQO survival assays involved chronic exposure to
DNA damage during growth. To ensure that the decreased resistance that we observed for
the catalytic dead mutant was not due to the chronic exposure conditions, we acutely
damaged the chromosomally integrated strains with a range of 4-NQO doses for one hour
in liquid culture before plating. The results confirmed that the catalytic dead strain loses
viability as a consequence of the DNA lesion type and not the conditions under which the
damage was received (Figure IC).
We next tested if the sensitivity caused by the catalytic dead version of Rev1 to 4NQO also occurs when a rev1A strain contains the wild-type REV] or catalytic dead
alleles on plasmids, since the levels of expression and regulation of plasmid copies of
REV] may differ from chromosomal copies. However, as in the case of the integrated
allele, the rev1A strain carrying the catalytic dead revl mutant is more sensitive to 4NQO treatment compared to the derivative carrying the wild type REV] gene but not as
sensitive as the derivative carrying the empty vector (Figure 1D). In contrast, the killing
curve for the catalytic dead strain was virtually the same as the REV] strain after UV
radiation (Figure lE) and cisplatin exposure (Figure IF). Cisplatin introduces lesions
127
such as 1,2-intrastrand linkages between the N7 positions of adjacent guanines and
monoadducts into the DNA (11). This decreased resistance of catalytic dead rev] strains
to 4-NQO and not to these other DNA damaging agents suggests that the DNA
polymerase activity of Revl is employed in a lesion-specific manner in vivo. Moreover,
Revl is a DNA polymerase with a propensity to use a template dG (22). Consequently,
based on these in vitro studies, an agent that creates adducts on dGs would be expected to
cause a loss in viability for cells that lack the DNA polymerase function of Rev1.
Since we were able to detect the physiological importance of Revl's DNA
polymerase activity regardless of whether REV1 was in the chromosome or carried on a
plasmid, we continued to use the plasmid containing strains for two reasons. First, we
were able to test the effects of various revl mutations much more rapidly by introducing
them into the plasmid-borne REVI than by constructing integrated mutant allele strains.
Secondly, to examine the effects of rev] alleles on mutagenesis by assaying for
canavanine resistance, we needed to use a second strain background (S288C) with the
CAN] gene intact. Our original strain background (W1588), in which REV] and the
mutant copies were integrated, possesses the canl-100 allele.
To further explore the role that specific amino acids located in the conserved Yfamily polymerase domain of Rev have in tolerance of 4-NQO lesions, two additional
strains bearing the revl-R324K and revl-R324T alleles on plasmids were evaluated in the
4-NQO survival assay. Arg324 of RevI forms hydrogen bonds with the incoming dCTP,
thereby directing the preference of Revl for inserting dCs (41). Similar to the catalytic
dead mutant, both the revl-R324K and revl-R324T strains have an increased sensitivity
to 4-NQO than the REV] strain (Figure 2A). These mutations do not cause the same
128
degree of killing after 4-NQO exposure as the D467A, E468A alteration of the catalytic
dead allele, which completely inactivates all DNA polymerase activity (23).
In addition to the residues mentioned above, two amino acids located in the
catalytic domain of Revl, Phe367 and Phe441, are conserved as an aromatic
phenylalanine or tyrosine in most organisms that possess Revl. A similarly positioned
aromatic amino acid in the TLS Pol Dpo4 of S. solfataricus(comparable by alignment to
the Phe367 residue in Revl) acts as a steric gate that excludes rNTPs from the active site
(10). More notably, analogous residues in the TLS polymerase DinB (F13 and Y79 in E.
coli) are critical for DinB's TLS function and cellular resistance to 4-NQO damage in E.
coli (24). Given their importance for DinB catalysis, we measured cellular viability after
4-NQO exposure for strains carrying the revl-F367L or rev]-F441L alleles. Unlike
DinB, these residues do not significantly affect Revl function in vivo after 4-NQO
damage (Figure 2B). This difference is understandable considering the uniqueness of
Rev 1's polymerase mechanism (41).
The non-catalytic functions of Rev1 also contribute to 4-NQO tolerance:
The sensitivity of the catalytic dead strain is not as severe as when the REV] gene
is entirely missing (Figure 1, A, C, and D), indicating that Revl promotes 4-NQO
resistance in the absence of its DNA polymerase activity. To investigate the role of the
other Revi domains, we compared the survival of revl mutant strains with mutations in
the BRCT domain, UBM2 domain, and C-terminus after exposure to 4-NQO and UV
irradiation. Consistent with published data (21), when rev] alleles with mutations in the
BRCT or UBM2 domains are integrated at the endogenous locus, the cellular resistance
129
to chronic and acute exposure to 4-NQO decreases relative to a REV] strain (Figures 3, A
and B). Unlike the catalytic mutants, these mutations also reduced resistance to UV
(Figure 3C). The same trends for survival after 4-NQO exposure are observed for the
BRCT and UBM2 mutants that we tested in plasmid strains, except that the relative
magnitude of the sensitization to killing varies (Figure 3D). The normal function of the
TLS polymerase interaction region located in the C-terminus appears to be the most
critical for 4-NQO resistance. The phenotypes of the integrated catalytic domain, BRCT,
and UBM2 mutant strains are not due to problems with protein expression; the protein
levels during log phase or G and G2/M arrests did not significantly differ for the mutant
forms of Revi compared to the wild-type protein (Figure 4, A and B). Therefore, even
though the DNA polymerase function of Revi contributes to 4-NQO resistance, the noncatalytic functions provided by the protein interaction domains of Rev1 are also required
for full 4-NQO resistance and are especially important for bypass of non-N 2-dG lesions.
The BRCT and UBM2 domains of Revl can serve in the tolerance of 4-NQO damage
through interacting with PCNA or DNA to assist in Revl gaining access to the DNA
lesion, whereas the C-terminus's likely contributes by recruiting Pol
for extension past
a lesion.
The DNA polymerase action of Revi is even more critical in the absence of errorfree DNA damage tolerance:
Since the sensitivity of the catalytic dead strain is modest at some doses of 4NQO, we hypothesized that the data might reflect the action of another repair or tolerance
pathway masking the loss of catalytic insertion by Revl. To investigate what pathway
130
might be involved, we measured the survival after DNA damage in cells with deletions in
genes involved in nucleotide excision repair [NER] (RAD14) or error-free tolerance
(MMS2) in combination with the rev] deletion. NER is the cell's best option to repair the
bulky N 2-dG adducts created by 4-NQO, and error free tolerance serves as another means
to tolerate such lesions.
Deleting both RAD14 and REV has an additive effect on survival after 4-NQO
damage with respect to the deletion of each gene individually (Figure 5A). Therefore,
NER and Rev -mediated TLS act independently for 4-NQO resistance. Conversely, in
cells lacking both Rev 1 and Mms2, there is a dramatic increase in the sensitivity to
killing by 4-NQO relative to cells only missing the REV] or MMS2 gene alone (Figure
5B), indicating that significant functional redundancy exists between the two tolerance
pathways.
In the rev]Aradl4A background, the killing curve for the strain carrying the
catalytic dead plasmid is very close to the killing curve of the strain bearing the REV]
gene (Figure 5B). When the cells were deficient in error-free tolerance, however, the
strain carrying the catalytic dead plasmid shows a substantial sensitivity to 4-NQO
relative to the strain with the REV1 plasmid in that same background (Figure 5B).
Furthermore, the susceptibility of the catalytic dead strain to killing by 4-NQO damage is
much more striking in the mms2A background than in the MMS2 background (Figures 5B
versus 1D). Therefore, the Mms2-dependent error-free tolerance pathway seems to
significantly compensate for the loss of Rev1's DNA polymerase activity in vivo to
tolerate lesions produced after 4-NQO damage.
131
The DNA polymerase activity of Revl aids in bypass of DNA lesions produced by a
naturally occurring aldehyde, methylgloxal:
The data presented so far implicates a role for the DNA polymerase function of
Rev in surviving exposure to a representative exogenous DNA damaging agent, 4-NQO,
which causes a substantial fraction of N2-dG adducts. Considering the apparently absolute
conservation of the specific residues in the polymerase domain of Revi (such as Arg324
and the metal-binding Asp467 and Glu468), it seems likely that there may be a naturally
occurring DNA lesion caused by endogenous agents that similarly require Revl's
polymerase activity for bypass. An example of such an endogenous agent produced by
normal metabolism is methlyglyoxal, an aldehyde that can react with DNA to produce
N2-(1-carboxyethyl)-2'-deoxyguanosine (N2-CEdG) as the major lesion (12, 61). This
lesion is also detectable in human samples (33, 52). Thus, methylglyoxal serves as a good
candidate aldehyde for testing the importance of the DNA polymerase function of Rev 1
in surviving endogenous DNA damage. We assessed the percent survival of our plasmid
strains in the presence of chronic exposure to methylglyoxal. We chose the revlAmms2A
strain background for our experiments to maximize any observable phenotypes for the
catalytic dead Revi strain. The catalytic dead strain is sensitive to methylglyoxal
exposure, indicating that Revi inserts across from the lesions that develop when
methylgloxal naturally reacts with DNA (Figure 6). As seen with the 4-NQO survival
assays, the greatest dose-dependent decrease in survival is observed when the cell is
devoid of any Revi protein.
132
The DNA polymerase activity of Rev1 is partially required for 4-NQO-induced
mutagenesis:
Given the propensity of Revi to insert dCs across from dGs in vitro (22, 43), the
step of Rev catalytically inserting dCs across from N2-dG lesions produced by 4-NQO
would not be expected to be a major source of the mutations induced by 4-NQO.
Disruption of the DNA polymerase domain of Revl, thus, might be expected to prevent
this relatively error-free bypass with the consequence that other TLS polymerases might
insert across from the N2 -dG adducts in a more mutagenic manner. To determine the level
of 4-NQO-induced mutagenesis caused by to Revi function, we used the CAN] forward
mutation assay in which cells become resistant to canavanine when a loss-of-function
mutation occurs in the CAN] gene. In a strain carrying the catalytic dead allele of REV1,
loss of RevI catalytic function decreases, rather than increases, the mutation frequency
compared to the REV1 strain (Figure 7A). Nevertheless, the strain carrying the catalytic
dead revl allele exhibits greater mutagenesis than the strain without any Rev1. This
observation contrasts with the absence of any effect of Rev1 polymerase activity on UVinduced (Figure 7B) or MMS-induced mutagenesis as reported previously in S. cerevisiae
(23). Thus a subset of mutations induced by 4-NQO require Revi 's polymerase activity,
2
despite the fact that Revi potentially inserting dCs across from 4-NQO-induced N -dG
lesions would be expected to be non-mutagenic. Possible explanations (discussed in more
detail below) for this data include loss of Pol t extension, loss of Revi insertion across
from non-N2-dG lesions produced by 4-NQO, loss of a likely infrequent misincorporation
by Revl across from N 2-dG adducts, and/or an increased bias for use of error-free
tolerance.
133
We also assayed the 4-NQO-induced mutagenesis of the other catalytic domain
mutants in the revlA strain background. Both Arg324 mutants could potentially disrupt
the hydrogen bonding capability of Arg324 with the incoming dCTP and potentially
cause Rev to have an altered (incorporating another nucleotide) or no DNA polymerase
activity. Both the rev1-R324K and revl-R324T mutants modestly decrease 4-NQOinduced mutagenesis but not to the same extent as the catalytic dead mutant (Figure 8A).
The rev]-F367L and revl-F441L mutants also had minor effects on 4-NQO-induced
mutagenesis (Figure 8B). These results indicate that the Arg324, Phe367, and Phe441
residues of Rev 1 do not greatly influence mutagenesis in vivo for this particular type of
DNA damage, and, as with survival, these mutant proteins function somewhat differently
from the catalytic dead protein.
We also measured the frequency of 4-NQO-induced canavanine resistant cells in
the BRCT, UBM2, and the C-terminus mutant strains. Relative to the catalytic dead
mutation, mutations in the BRCT, UBM2, or C-terminus diminish the mutation frequency
to levels very close to the empty vector strain (Figure 9). Despite these mutations
conferring 4-NQO survival phenotypes that are better than a strain lacking any Rev1, the
mutagenesis phenotypes are equivalent to a revlA strain (Figure 3D versus 9). These data
suggest that Revl can promote survival in a manner that is error-free when the BRCT,
UBM2, or C-terminus is not functional either through additional protein interactions and
their pathways such as error-free tolerance or through the accurate DNA polymerase
activity that remains.
134
The 4-NQO-induced mutation frequency is also reduced in the absence of error-free
tolerance when Rev1 is catalytically dead:
The decreased mutation frequency after 4-NQO damage of the catalytic dead
strain in the rev1A background might result from the cell compensating for loss of Rev1
polymerase activity by making a greater use of the error-free tolerance pathway. Given
that the survival results of the catalytic dead strain were significantly affected when cells
were deficient in Mms2-dependent error-free tolerance, we measured the CAN] mutation
frequency in the rev]Amms2A strain background. Due to the dramatic drop in percent
survival for the rev]Amms2A strain, we chose to acutely expose the cells to 4-NQO
damage in liquid media for this assay. The catalytic dead Rev1 strain displays a decrease
in CAN] mutagenesis at higher doses of 4-NQO compared to the REV1 strain in the
rev]Amms2A background that is similar to that observed when the catalytic dead Revl is
in the rev]A background (Figure 10A). Since error-free tolerance does not seem to be the
cause of this reduction in mutation frequency, this result likely reflects one of our other
proposed mechanisms for loss of mutagenesis mentioned above and described in the
Discussion. The killing curve of the catalytic dead strain for acute 4-NQO exposure in
this background follows a similar pattern to the killing curve after chronic damage, but
the catalytic activity seems to play a greater role under conditions of chronic exposure
(Figures 10B versus 5B).
The Involvement of Pol vTand Pol t in 4-NQO Resistance:
Because Pol ql (Rad30) and Pol
(Rev3/7), the other TLS polymerases in S.
cerevisiae, are plausible candidates for acting with Rev1 to bypass bulky N2-dG adducts,
135
we examined the percent survival after 4-NQO damage in cells lacking RAD30, REV3, or
REV7 in the revlAmms2A background. The additional deletion of RAD30 did not affect
survival for the REV], catalytic dead, or revlA strains in response to 4-NQO damage
(Figure 11, bars 4, 5, and 6 versus 1, 2, and 3). The deletion of REV3 or REV7, however,
eliminated the protection conferred by Revi or the catalytic dead Revl (Figure 11, bars 8,
9, 11, and 12 versus 2 and 3). These data imply that Pol r has little to no role in the
bypass of lesions produced by 4-NQO, whereas, Pol ( is indispensable for Revl's
contribution to 4-NQO resistance when the error-free pathway cannot compensate for
defects in the TLS pathway. Similarly, Pol
requires the presence of Rev1 to aid in DNA
damage tolerance after 4-NQO exposure, since bars 1, 7, and 10 are equivalent but differ
from lane 2 when Revi is available (Figure 11).
136
DISCUSSION
I show in this Chapter that Revl's catalytic activity does influence the ability of
wild type cells to survive exposure to the carcinogen 4-NQO. In contrast, loss of Rev l's
catalytic activity has no effect on the ability of cells to survive exposure to UV radiation
or cisplatin. Furthermore, loss of the catalytic activity causes relatively greater
sensitization to killing by 4-NQO in an mms2A strain, suggesting that the Mms2dependent error-free DNA damage tolerance can normally partially compensate for the
2
loss of Revl's catalytic activity. Since 4-NQO causes a significant fraction of N -dG
adducts, these data are consistent with Rev1 helping cells to survive 4-NQO-induced
DNA damage by inserting dC opposite these N 2-dG lesions. These results suggest that the
reason for the current confusion in the literature concerning the in vivo importance of
Revl's catalytic activity has arisen because Revl's catalytic activity is important for
DNA damaging agents that introduce a substantial fraction ofN2 -dG adducts but is not
important for agents that do not. The extensive conservation of Revl's catalytic activity
throughout eukaryotes suggests that the catalytic activity is needed to bypass DNA
2
damage acquired from ubiquitous endogenous or exogenous agents that produce N -dG
lesions.
The likelihood of the cell tolerating the 4-NQO lesion through the error-free
tolerance branch is greater in the presence of the catalytic dead Revl than in the complete
absence of Rev 1. This point is deduced from the trends of the killing curves in Figures
lD and 5B. In Figure iD, the killing curve of the catalytic dead strain is much closer to
that of the REVI strain than the rev1A strain. On the other hand, the killing curve of the
catalytic dead strain is closer to the rev1A strain rather than the REV1 in Figure 5B when
137
Mms2 is not present. Therefore, the survival after chronic 4-NQO exposure of the
catalytic dead strain in the rev1A background is contingent on a functional Mms2
pathway. The possible crosstalk between Revl-dependent TLS and Mms2-mediated
error-free tolerance was unexpected since the synergistic increase in sensitivity to killing
by 4-NQO would be consistent with the two pathways being completely independent
modes of tolerating DNA damage. In contrast, the catalytic dead Revl does not seem to
have a role in facilitating NER, because the catalytic dead strain is not much more
sensitive in the rev]Aradl4A background relative to the rev1A background (Figure 5A).
These results suggest that the non-catalytic functions of Revl must assist in Mms2mediated error-free tolerance for some fraction of the 4-NQO lesion bypass events. In
principle, this could happen either through direct protein interactions or indirectly by
stabilizing the replication fork. Since little is known about the mechanistic details of
error-free tolerance, it is conceivable that Rev1 is capable of recruiting proteins involved
in error-free tolerance in addition to the influence that polyubiquitinated PCNA has in
signaling that pathway.
The discovery of RE V and initial characterization was based on the fundamental
role that Rev1 has in mutagenesis. Unexpectedly, the catalytic dead strain displayed a
reduced mutation frequency with 4-NQO (Figures 7 and 10). Consistent with the
reversionless quality of the rev A strain though, the 4-NQO-induced mutation
frequencies are also lower for all rev] mutants tested (Figures 7-10). If Pol performed
the insertion step instead of the catalytic dead Revl, there might be a greater chance of
not introducing a dC across from the N2 -dG lesion, and the mutation frequency might
138
have been suspected to increase relative to the REV] strain. That is not what was
observed in Figure 7A.
Several hypotheses could explain the loss of the mutagenic activity generated by
the normal catalytic function of Rev 1 in Figures 7A and 10A that may account for the
lower 4-NQO-induced mutation frequency alone or in combination. First, Revi needs Pol
2
for extension once a nucleotide had been inserted opposite the N -dG lesion. This
extension will be more error-prone than synthesis by the replicative DNA polymerase.
Loss of the insertion step by Revi (assuming error-free) would also mean loss of
extension by Pol
and the mutagenesis created by extension. Secondly, one must also
consider the other potential DNA lesions produced after a cell is exposed to 4-NQO such
as the N6-adducted adenine (11), in which Revl insertion of dCMP would be error-prone.
So a catalytic dead strain will lose mutagenesis from Revi insertion over these lesions,
depending on what other DNA polymerases may insert over this adduct when Rev1
cannot act. The third possible contributing factor to 4-NQO-induced mutagenesis is that
Rev may insert a nucleotide other than dCTP across from N2-dG lesions at some low
frequency. Finally, when the DNA polymerase activity of Revl is disturbed, some more
error-free process may compensate, thus leading to overall less mutagenesis. The
decreased mutation frequency of the catalytic dead strain in the rev] A background
(Figure 7A) may in part result from the cell using the error-free tolerance pathway more
than when Revi is available as a DNA polymerase. The catalytic dead mutant does not
exhibit an increased, 4-NQO-induced mutation frequency in the revlAmms2A
background though (Figure 10A). So the action of the Mms2-dependent error-free
tolerance branch does not fully account for the decrease in 4-NQO-induced mutagenesis.
139
The 4-NQO-induced mutation frequencies for all of the non-catalytic (the BRCT,
UBM2, and C-terminus) mutants tested are almost at levels equivalent to the rev1A strain
carrying the empty vector (Figure 9). In Figure 3D though, the non-catalytic mutant
strains survive significantly better than the rev1A strain bearing the empty vector.
Interestingly, a defect in any one of the non-catalytic domains alone leads to a drastic
drop in the 4-NQO-induced mutation frequency. This result indicates that the remaining
functional domains of Revl in these mutants are able to promote survival after 4-NQO
exposure without greatly increasing mutagenesis. This is not due to the catalytic activity
of Rev 1, since the data reported here showed that Rev1 catalytic activity actually adds to
4-NQO-induced mutagenesis. Alternatively, the cell may discourage TLS and favor
error-free tolerance in the presence of a partially functional Rev I to improve survival
without increasing mutagenesis. Taken together with our conclusions from Figure 5,
these observations further support that Rev1 of the TLS branch enhances the ability of the
error-free tolerance pathway to tolerate DNA lesions produced by 4-NQO.
As with the catalytic dead mutant strain but not to the same degree, the Arg324,
Phe367, and Phe441 mutants of Revl exhibit a decrease in 4-NQO-induced mutagenesis
(Figure 8, A and B). These mutant forms of Rev1 probably act as less efficient
polymerases rather than losing all their ability for catalysis. The decrease in mutagenesis,
as in the catalytic dead mutant strain, may reflect less Rev1 bypass of non-N2 -dG lesions
or less Pol t extension past N2 -dG adducts. For mutants of Arg324 that hydrogen bonds
with the incoming dCTP, especially the replacement of arginine with threonine, the
residue change may alter Revl's selectivity for dC. Even though we did not observe an
increased mutation frequency in the presence of 4-NQO for either Arg324 mutant, the
140
data could indicate that the less efficient polymerase function outweighs any possible
increase in the misincorporation frequency across from N2 -dG lesions. Considering that
Rev l's mechanism of insertion is very unique among DNA polymerases, the results for
survival and mutagenesis after 4-NQO exposure of the Phe367 and Phe441 mutants are
not surprising and differ from the phenotypes observed for similar mutants of E. coli
DinB.
These results provide evidence of a lesion-specific catalytic activity for RevI to
bypass N2-dG adducts in the cell. The propensity of Revi to insert a dC opposite a
template dG makes Revi apt to use template N2 -dG lesions similar to Pol vI's
specialization for bypassing cis-syn cyclobutane pyrimidine dimers. Like Pol rl,the
conserved catalytic activity of Rev 1 provides an error-free mechanism of insertion across
from the N2-dG adduct, and the lesion specificity serves as an example of another means
for regulation of TLS activity. The non-catalytic functions of Revl are important for
gaining access to the lesion and recruiting Pol t for extension past N 2-dG adducts. Aside
from Rev l's non-catalytic domains functioning to tolerate DNA damage, the modest
sensitivity of the catalytic dead strains to 4-NQO is largely due to error-free tolerance
masking the phenotype. Lastly, the results reveal a partial dependence of error-free
tolerance on Rev 1 that requires more studies in the future.
141
ACKNOWLEDGEMENTS
The authors thank members of the Walker lab for helpful discussions particularly
Brenda Minesinger for her critical reading of the manuscript. We also thank Michael
Onwugbufor for his contributions to this project during the summer as an MSRP
undergraduate. This work was supported by a National Institute of Environmental Health
Sciences (NIEHS) grant 5-R01-ES015818 to G.C.W., a NIEHS grant P30 ES002109 to
the MIT Center of Environmental Health Sciences, and an American Cancer Society
Research Professorship to G.C.W.
142
TABLE 1
Strains used in this study
source
Relevant genotype
Strain
revlAbarlA
W1588-4C barlA::LEU2 revlA::kanMX
(9, 57)
YMEW1
W1588-4C barlA::LEU2 rev A::kanMX4::pRS306-
Modified from (9)
REV1-TEV-ProA-His 7
YMEW2
W1588-4C barlA::LEU2 rev1A::kanMX4::pRS306-
Modified from (9)
revi-D467A E468A-TEV-ProA-His 7
YMEW3
W1588-4C barlA::LEU2 rev A::kanMX4::pRS306-
Modified from (9)
rev 1-G 193R-TEV-ProA-His 7
YMEW4
W1588-4C barlA::LEU2 rev1A::kanMX4::pRS306-
This study
rev1-L821A L822A-TEV-ProA-His 7
YMEW5
W1588-4C barlA::LEU2 revliA::kanMX4::pRS306-
This study
rev 1-F367L- TEV-ProA-His 7
YMEW6
W1588-4C barlA::LEU2 re vl A::kanMX4::pRS306rev1-F441L-TEV-ProA-His
This study
7
YMEW7
S288C barlA::LEU2 rev1A::kanMX4 pRS416
This study
YMEW8
S288C barlA::LEU2 revlA::kanMX4 pRS416-REV1
This study
YMEW9
S288C barlA::LEU2 revlA::kanMX4 pRS416-rev 1-
This study
D467A E468A
YMEW10
S288C barlA::LEU2 revlA::kanMX4 pRS416-rev1G 193R
143
This study
YMEW11
S288C barlA::LEU2 revlA::kanMX4 pRS416-revi-
This study
E820A L821A P822A T823A Q824A
YMEW12
S288C barlA::LEU2 revlA::kanMX4 pRS416-revi-
This study
L889A V890A K891A W893A V894A
YMEW13
S288C barlA::LEU2 revlA::kanMX4 pRS416-revi-
This study
R324T
YMEW14
S288C barlA::LEU2 revlA::kanMX4 pRS416-revl-
This study
R324K
YMEW15
S288C barlA::LEU2 revlA::kanMX4 pRS416-revl-
This study
F367L
YMEW16
S288C barlA::LEU2 revlA::kanMX4 pRS416-revl-
This study
F441L
YMEW17
S288C barlA::LEU2revlA::kanMX4
This study
mms2A::hygM4 pRS416
YMEW18
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 pRS416-REV1
YMEW19
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 pRS416-revl-D467A E468A
YMEW20
S288C barlA:: LEU2 revlA::kanMX4
This study
radl4A::hygM4 pRS416
YMEW21
S288C barlA:: LEU2 revlA::kanMX4
This study
radl4A::hygM4 pRS416-REV1
YMEW22
S288C barlA:: LEU2 revlA::kanMX4
radl4A::hygM4 pRS416-revl-D467A E468A
144
This study
YMEW23
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 rad30A::natMX4 pRS416
YMEW24
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 rad30A::natMX4 pRS416-REV1
YMEW25
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 rad30A::natMX4 pRS416-re v D467A E468A
YMEW26
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 rev3A::natMX4 pRS416
YMEW27
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 rev3A::natMX4 pRS416-REV1
YMEW28
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 rev3A::natMX4 pRS416-rev 1D467A E468A
YMEW29
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 rev7A::natMX4 pRS416
YMEW30
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 rev7A::natMX4 pRS416-REV1
YMEW31
S288C barlA::LEU2 revlA::kanMX4
This study
mms2A::hygMX4 rev7A::natMX4 pRS416-rev1D467A, E468A
All strains are derivatives of W1588-4C (MATa leu2-3,112 ade2-1 can 1-100 his3-11,15 ura3-1
trpl-1 RAD5) (63) or S288C (MATa SUC2 gaI2 mal mel flol flo8-1 hapi ho biol bio6) (35) as
indicated.
145
TABLE 2
Site-directed revi mutants used in this study
Name in this
Published Allele
Report
Name
Catalytic dead
revi Ala4 6 7 -
Amino Acid Change
Protein
Reference
Location
D467A E468A
Ala468
DNA
(23)
Polymerase
Domain
BRCT
revl-1
G193R
BRCT
(29, 32)
domain
UBM2a
UBM2*
L821A P822A
UBM2
(21)
UBM2 b
revl-106
E820A L821A P822A
UBM2
(9)
C-terminus
(9)
DNA
This study
T823A Q824A
C-terminus
revl-108
L889A V890A K891A
W893A V894A
revl-R324K
R324K
Polymerase
Domain
146
rev]-R324T
R324T
DNA
This study
Polymerase
Domain
revl-F367L
F367L
DNA
This study
Polymerase
Domain
rev]-F441L
F441L
DNA
Polymerase
Domain
a
This allele was used for the integration into the chromosome.
b This allele was used for pRS416.
147
This study
FIGURE 1. - The catalytic activity of Rev1 is biologically relevant in a lesion-specific
manner. (A) The integrated catalytic dead strain is susceptible to killing by chronic 4NQO exposure. Strains are * revlAbarlA, E YMEW1 (REV]), and A YMEW2 (revlcatalytic dead). (B) The integrated catalytic dead strain is not sensitive to UV radiation.
(C) Acute 4-NQO damage also causes increased cell death when Revl is catalytic dead.
(D) Plasmid expression of catalytic dead Revl leads to a similar level of 4-NQO
resistance as found with the integrated strain. Strains are 0 YMEW7 (revlA pRS416), N
YMEW8 (revlA pRS416-REV1), and A YMEW9 (revlA pRS416-revl-catalytic dead).
(E) The plasmid bearing catalytic dead strain is not sensitive to UV radiation or (F)
cisplatin exposure.
148
FIGURE 1
C
B
A
g10
iO
js
601
o
75
c4o Doe (nm"r
0
i
0
S
0
25
woo0
4
0
1 mwouEmpor O)
F
E
D
2
mnoo ow
so
4 M
I
i
6.1T(L
0
to
10
4. co De.* (n
N
N
S
to
25
mN
Doew (am)
149
40
N
0
2
1
T.me I mm
3
.U (HourW)
FIGURE 2. - Other mutations in the catalytic domain perturb Revl function. (A)
Mutating Arg324 mildly sensitizes cells to chronic 4-NQO exposure. Strains are O
YMEW7 (revlA pRS416), U YMEW8 (revlA pRS416-REV1), A YMEW9 (revlA
pRS416-revl-catalyticdead), 0 YMEW13 (revlA pRS416-revl-R324T), and 4
YMEW14 (revlA pRS416-revl-R324K). (B) Mutating Phe367 and Phe441 mildly affects
survival after chronic 4-NQO exposure. Strains are @ rev]AbarlA, N YMEW1 (REV]),
A YMEW2 (revl-catalyticdead), + YMEW5 (rev]-F367L), and 0 YMEW6 (revlF441L).
150
FIGURE 2
10
0
30
4-NOO Dose (ngI'm)
60
151
0
50
75
4-NQO Dose (ng/mi)
100
FIGURE 3. - Revl's non-catalytic domains also function to tolerate 4-NQO lesions. (A)
Cellular survival for the integrated BRCT and UBM2 mutants decreases after chronic 4NQO exposure. Strains are S revlAbarlA, U YMEW1 (REV]), A YMEW2 (revlcatalytic dead), * YMEW3 (rev]-BRCT), and 0 YMEW4 (revl-UBM2). (B) The
BRCT and UBM2 mutants are also sensitive to acute 4-NQO damage. (C) The BRCT
and UBM2 mutants are sensitive to UV radiation. (D) The C-terminus mutant is more
sensitive to chronic 4-NQO exposure than the BRCT and UBM2 mutants on plasmids.
Strains for B, C, and D are
YMEW7 (revlA pRS416), N YMEW8 (rev]A pRS416-
REV1), A YMEW9 (revlA pRS416-revl-catalytic dead), * YMEW10 (rev] A pRS416rev]-BRCT), O YMEW11 (rev1A pRS416-rev1-UBM2), and O YMEW12 (revlA
pRS416-rev l-C-terminus).
152
FIGURE 3
A
10I
0.1
50
0
4-NQO 0Do.
75
100
4-No0 Dos for I Hour ExposUre (pgml)
(ngld)
Dy
10
lO
0
10
40
25
Uv Do (Jn)
0
50
153
30
4-NO Do" (ngAn)
00
FIGURE 4. - Integrated mutant proteins of Revl have a similar expression pattern to
wild-type Rev1. (A) Protein levels of logarithmically growing cells are normal for
mutant proteins of Revl as shown by western blot. Strains are YMEW1 (REV]) - lane 3,
YMEW2 (rev]-catalytic dead) - lane 4, YMEW3 (rev]-BRCT)- lane 2, YMEW4 (rev]UBM2) - lane 1, YMEW5 (revl-F367L)- lane 5, and YMEW6 (revl-F441L) - lane 6.
(B) Mutant proteins of Revl are at low levels during Gi arrest and high levels during
G2/M arrest like wild-type Revl.
154
FIGURE 4
A
Log Phase Cells
123456
123456 123456
Rev1
Rev1
Pgkl
G1 Arrested G2M Arrested
155
FIGURE 5. - The enhanced requirement for Revl's catalytic activity occurs in the
absence of error-free tolerance after 4-NQO damage. (A) The catalytic dead strain is
significantly more sensitive to 4-NQO in the mms2A background. Strains are O YMEW7
(revlA pRS416), N YMEW8 (rev1A pRS416-REV1), 0 YMEW17 (rev]Amms2A
pRS416), 0 YMEW18 (revlAmms2A pRS416-REV1), and V YMEW19 (revlAmms2A
pRS4]6-revl-catalyticdead). (B) The deletion of radl4 does not increase the divergence
of the catalytic dead strain's 4-NQO killing curve from the REV1 strain's curve. Strains
are S YMEW7 (revlA pRS416), U YMEW8 (revlA pRS416-REV1), 0 YMEW20
(rev1Aradl4A pRS416), 0 YMEW21 (rev]Aradl4ApRS416-REV1), and V YMEW22
(rev1Aradl 4A pRS416-revl-catalyticdead).
156
FIGURE 5
I
10
1
0
0
2.5
4-N00 Doe (ngE
157
2.5
4-NO Doo (nE
FIGURE 6. - Revl's catalytic activity helps tolerate exposure to an endogenously found
aldehyde. Cells were exposed to a chronic dose of methylglyoxal. Strains are @
YMEW17 (revlAmms2A pRS416), E YMEW18 (rev]Amms2A pRS416-REV1), and A
YMEW 19 (revl Amms2A pRS416-rev] -catalytic dead).
158
FIGURE
110
I
0
12.5
10
Muthygyoxal Doe (mrM)
159
15
FIGURE 7. - Loss of Rev1's catalytic activity decreases 4-NQO-induced CAN]
mutagenesis. (A) The mutation frequency of the catalytic dead mutant is lower than the
REV] strain in response to chronic 4-NQO exposure. Strains are O YMEW7 (rev1A
pRS416), N YMEW8 (revlA pRS416-REV1), and A YMEW9 (rev1A pRS416-revlcatalytic dead). (B) The mutation frequency of the catalytic dead strain does not differ
from wild-type for UV damage.
160
SJGAAMS 90&'d
NUIO
*1
I
woSjOAng
~ot Jd VtW
FIGURE 8. - Additional mutants in the catalytic domain of Revl modestly decrease the
4-NQO-induced mutation frequency. (A) The Arg324 mutants display slightly altered
mutations frequencies after 4-NQO exposure. Strains are @ YMEW7 (rev1A pRS416), U
YMEW8 (revlA pRS416-REV1), A YMEW9 (revlA pRS416-rev-catalytic dead), 0
YMEW13 (rev1A pRS416-rev-R324T), and 4 YMEW14 (revlA pRS416-revl-R324K).
(B) The 4-NQO-induced mutation frequencies of the Phe367 and Phe441 mutants are
somewhat lower than the REV] strain. Strains are O YMEW7 (revlA pRS416), U
YMEW8 (rev1A pRS416-REV1), A YMEW9 (revlA pRS416-revl-catalytic dead), *
YMEW15 (rev]A pRS416-rev1-F367L), and P YMEW16 (revlA pRS416-revl-F441L).
162
FIGURE 8
250
S200
150
P10
4-NO
30
Do" (nghnQ)
50
163
0
10
20
30
4NQO Dose (ngml)
FIGURE 9. - The functions of Rev l's non-catalytic domains play a significant role in
mutagenesis after chronic 4-NQO exposure. Strains are @ YMEW7 (revlA pRS416), U
YMEW8 (revlA pRS416-REV1), A YMEW9 (revlA pRS4]6-revl-catalyticdead), *
YMEW10 (rev]A pRS416-revl-BRCT), O YMEW11 (revlA pRS416-revl-UBM2), and
O YMEW12 (revl A pRS416-revl-C-terminus).
164
FIGURE 9
200
250
30
4NOO Dose (nghr
165
60
FIGURE 10. - The 4-NQO-induced mutagenesis for the catalytic dead mutant is also
lower in Mms2-deficient cells. (A) The 4-NQO induced mutation frequency of the
catalytic dead strain deviates from the REV] strain at higher doses of 4-NQO for acute
exposure. Strains are @ YMEW17 (revlAmms2A pRS416), U YMEW18 (revAmms2A
pRS416-REV1), and A YMEW19 (rev1Amms2A pRS416-revl-catalytic dead). (B) The
catalytic dead mutant is sensitive to acute killing by 4-NQO in the absence of Mms2.
166
FIGURE10
A
B
1001
160
140
120
10
100
so
~60
a40
20
0
0.1
0
00
200
foar 1 Hour Exposure (nf)q
so60
4-NOO Do
167
0
soo
200
so50
440NODose for 1 Hour Exposure (ngl)
FIGURE 11. - Revl requires Pol
to contribute to 4-NQO resistance. Percent survival is
shown for 4-NQO exposure at 1 ng/ml. Strains harboring the
pRS416,
pRS416-
rev]-catalyticdead, and EpRS416-REV1 are indicated. Strains are YMEW17
(revlAmms2A pRS416), YMEW1 8 (revlAmms2A pRS416-REV1), YMEW1 9
(revlAmms2A pRS416-revl-catalytic dead), YMEW23 (rev]Amms2Arad30A pRS416),
YMEW24 (rev1Amms2Arad30A pRS416-REV1), YMEW25 (revl Amms2Arad30A
pRS416-rev]-catalyticdead), YMEW26 (rev]Amms2Arev3A pRS416), YMEW27
(revl Amms2Arev3A pRS416-REV), YMEW28 (revlAmms2Arev3A pRS416-revlcatalytic dead), YMEW29 (rev1Amms2Arev7A pRS416), YMEW30 (revlAmms2Arev7A
pRS416-REV]), and YMEW31 (rev]Amms2Arev7A pRS4]6-rev]-catalytic dead).
168
FIGURE 11
1
2
4
3
5 6
7
8
9 10 11 12
100
10
0.1
0.01 •
.
revlA
mmns2A
revlA
mms2A
rad30A
169
revlA
mms2A
rev3A
revlA
mns2A
rev7A
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Woodgate, R. 1999. A plethora of lesion-replicating DNA polymerases. Genes
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173
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polymerase in vitro and in vivo. Proc Natl Acad Sci U S A 105:8679-84.
Zhang, Y., X. Wu, O. Rechkoblit, N. E. Geacintov, J. S. Taylor, and Z. Wang.
2002. Response of human REV to different DNA damage: preferential dCMP
insertion opposite the lesion. Nucleic Acids Res 30:1630-8.
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174
Chapter 4
Conclusion and Future Directions
175
Summary of Results
DNA damage can alter a cell's fate by causing stalls in replication, a loss of
genetic material, or alterations to the DNA that lead to a change in a protein's function. In
addition to DNA repair, DNA damage tolerance works to prevent or minimize these
events through the error-prone pathway of translesion DNA synthesis (TLS) and the
error-free template-switching branch. The experiments described in this thesis address
how some of these TLS polymerases are regulated to limit their mutagenic activity and
provide examples of the redundancy between the two tolerance pathways.
As described in Chapter 2, the protein levels of the S. cerevisiae TLS
polymerases, Revl and Rev3 (of Pol ), are kept low through proteasomal degradation.
Since a higher migrating form of Revi exists in the proteasome-compromised umplA
background, the mechanism for Rev degradation is likely due to a direct protein
modification. These results further our understanding of the molecular mechanisms
behind Rev l's cell cycle regulation and open new avenues of research. For example,
greater insight into Revl's cell cycle regulation could be determined, including the
identity of the modification, additional proteins involved in the regulation, investigating
the potential for a translational control, and understanding exactly why the protein levels
follow a cell cycle-dependent pattern.
Chapter 3 details the exciting findings for the biological significance of Revl's
catalytic activity. The DNA polymerase activity of Revl is mostly ignored in vivo due to
the lack of phenotypes for the catalytic dead mutant, but the results described in Chapter
3 show that the catalytic activity is required for the full resistance to the lesions produced
by 4-NQO and methylglyoxal, agents known to cause N2 -dG adducts. The specificity of
176
Revl's catalytic activity illustrates another layer of regulation for this TLS polymerase.
Interestingly, the catalytic activity also contributes to 4-NQO-induced mutagenesis, even
though the insertion of a C across from N2 -dG lesions by Revi should be error-free based
on what we know from biochemical work. This loss of mutagenesis at least in part likely
represents the lack of the extension function of Pol , since Revl cannot tolerate 4-NQO
lesions in the absence of Pol . The data in Chapter 3 also detail how the functional
redundancy of TLS with error-free tolerance makes Rev1's catalytic activity less obvious
in vivo and alludes to a new direction of study focusing on the interplay between TLS and
the error-free tolerance branch.
In the following paragraphs, I will discuss the data presented in Chapters 2 and 3
in more detail and provide specific examples of the potential experiments based on these
data.
The molecular mechanisms governing Rev1's cell cycle regulation:
In Chapter 2, I discuss the findings that the protein levels of Revl and Rev3 (of
Pol ) are regulated by proteasomal degradation. Since our lab discovered that Rev1
protein levels fluctuate in a cell cycle-dependent pattern (11), we wanted to learn more
about the molecular means controlling the cell cycle regulation. Understanding how Revl
levels are controlled throughout the cell cycle can provide insight as to why the protein
undergoes cell cycle regulation. To initially explore this question, we took a candidate
gene approach to look for novel regulators of Rev1. Candidate genes were selected based
on a genetic or physical link to Rev1 and their potential involvement in a pathway that
177
influences gene expression. Using this method, I was able to determine that Rev1 levels
are elevated in the proteasome defective strain, umplA (Chapter 2).
Through my work and that of a previous graduate student, multiple candidate
genes have been tested for affecting Rev1 protein levels or the half-life of Rev1 in a cell
cycle-dependent manner. Table 1 summarizes the results of the work from Laurie Waters
and myself (10). In a cycloheximide-chase assay, I compared the half-life of Revl in G1
or G2/M arrested cells for deletion backgrounds of the genes listed in Table 1 and
compared those to that in the wild type strain background. The genes were selected for
various reasons. For example, we were informed that Jab 1, a catalytic subunit of the
COP9 signalosome that cleaves ubiquitin-like Nedd8 from SCF ubiquitin ligases,
interacted with Revi (E. Friedberg, personal communication). For some, given what is
now known in the literature, the selection of that gene is less valid. After examining
Rev1's protein levels in these various mutant backgrounds, however, we determined that
only the loss of one of these proteins (Tom 1) revealed a mild effect on Revl.
Specifically, loss of Toml resulted in a slightly longer half-life for Revi than that seen in
a TOM] strain. Originally, Toml, an E3 ubiqutin ligase, came to our attention for having
regions of homology to Revl. Later, these regions in Revl became known as UBMs for
their ability to bind ubiquitin (1). Since the change in Rev1's half-life is not dramatic in
the tomlA strain and no clear connection between Toml and Revl is evident at this point,
these results may indicate an indirect effect. Also, we may not see a noticeable difference
in Rev1's half-life in a strain missing the E2 conjugating enzyme or E3 ubiquitin ligase if
polyubiquitin is not the modification on Rev1 or if other E2s or E3s can compensate for
the loss of one.
178
Another promising future direction that has not yet been tested is the involvement
of CDC7 in the regulation of Revl protein levels or activity. CDC7 encodes an essential
kinase that has a cell cycle regulated binding partner, Dbf4 (5). Curiously, strains
carrying specific alleles of CDC7 exhibit increased or decreased mutation frequencies
after DNA damage (2). Furthermore, CDC7 and REV3 are epistatic for resistance to UV
damage (8). Taken together with the evidence that Revi is phosphorylated (9),
investigating the possible relationship between Cdc7 and the regulation of RevI or other
members of the TLS pathway appears to be a worthwhile endeavor.
Possible experiments to examine Revl regulation includes determining what if
any contribution does translational regulation have on the cell cycle fluctuation of Rev 1
protein levels. Given that Revi levels become unstable after the addition of
cycloheximide (as described in Chapter 2) indicates that translation of Revl takes place
during both G 1 and G2/M. However, it is unclear if translation occurs at different rates
throughout the cell cycle or if an unknown factor limits translation during G1. More
involved experiments looking at translation rates of RevI at particular points in the cell
cycle will help answer these queries. Additionally, genetic screens may facilitate finding
the particular transcription factors or other proteins involved in the regulation of Revl's
transcription, translation, or degradation. For example, the reporter for such a screen
could be a lacZ translational fusion to the 3' end of the chromosomal copy of REV1.
Following optimization of screening colonies by color and testing controls, one could
identify novel regulators of REV1 expression through transposon mutagenesis.
The lesion specificity of Rev1's catalytic activity:
179
Although I show that Revl's catalytic activity is required for the full resistance to
4-NQO and methylgloxal in Chapter 3, the exact lesion(s) that Revi bypasses in a cell
during normal cellular growth still remain unknown. Since methylglyoxal is a naturally
occurring aldehyde, the N 2-dG adduct produced by methylglyoxal's reaction with DNA,
N2-(1-carboxyethyl)-2'-deoxyguanosine, serves as a good starting point in designing
assays. A combination of in vivo and in vitro assays will help identify what endogenously
created DNA lesions have led to the conservation of Revl's catalytic domain.
To assess Revl's role in the tolerance of a candidate lesion, lesion-bypass assays
involving DNA constructs containing one known lesion as described in Yuan et al. (12)
could be designed. In these experiments, one can quantitate the lesion bypass capabilities
of cells with a revl, rev3, or rev7 mutant background. Lesions that require Revl's
catalytic activity will be recognizable when a Revi catalytic dead strain has the reduced
ability to bypass one lesion. Any particular lesion can then be analyzed using purified
Rev and Pol
to look at the accuracy and efficiency of bypass over an adduct. The best
DNA lesions to use for the in vivo and in vitro assays would be those that are normally
found in the cell, especially N2 -dG adducts. Furthermore, finding another compound that
sensitizes the Revi catalytic dead strain to exogenous exposure but is also an agent that is
normally found in vivo, could give clues about other DNA lesions that Revl's catalytic
function is significant in tolerating and possibly even more so than 4-NQO or
methylglyoxal lesions. Knowing other compounds that sensitize cells in the absence of
Revl's catalytic activity may also help in designing lesions for the experiments
mentioned earlier. Potentially, these experiments could reveal a cognate lesion, an adduct
that RevI is optimized to bypass. Such experiments have determined that cyclobutane
180
thymine dimers and N2-dG adducts are cognate lesions for Pol rl and E. coli DinB/
mammalian Pol K,respectively (3, 4).
Another direction of study that resulted from my data in Chapter 3 involves the
relationship between S. cerevisiae Rev and E. coli DinB. DinB is absent from S.
cerevisiae, even though DinB is found in all domains of life (7). On the other hand, Revl
2
is not present in E. coli. Since DinB was found to efficiently and accurately bypass N -dG
adducts in vitro (3) and Revl's catalytic activity is critical for DNA damage tolerance
after exposure to agents that produce N2-dG lesions (discussed in Chapter 3), we
proposed that Rev1 and DinB have redundant functions in vivo. If Rev1 and DinB can
bypass the same or similar endogenous lesions, then an organism could evolve to have
only DinB or Revl. To test this hypothesis, we took advantage of the model organism, S.
pombe, in which both Revi and DinB are present. Indeed, we find that the Arev]Adinb
strain is synergistically more sensitive to the N2 -dG producing agent, 4-NQO, compared
to either single mutant (B. K. Minesinger, unpublished data), indicating an overlapping
function for Rev1 and DinB. Also, the expression of S. pombe DinB in a rev1A in S.
cerevisiae partially rescues the sensitivity of the rev1A strain to 4-NQO damage (M. E.
Wiltrout, unpublished data). Future work will involve expanding this work to mammalian
cells to understand the relationship of Revl and DinB in higher eukaryotes.
The genetic and physical interactions with the error-free tolerance pathway:
Given our results in Chapter 3, an attractive direction of future study is to learn
more about how the TLS and error-free tolerance branches interact genetically and
possibly physically. The cell likely has mechanisms to determine which pathway is
181
chosen for a specific lesion and under certain conditions. New experiments may explain
how the actions of one pathway may regulate the activity of the other pathway or how
they are both regulated by another common method such as the ubiquitination status of
PCNA.
Connections between various DNA repair and tolerance pathways are already
emerging. For example, in higher eukaryotes, the proteins associated with Fanconi
Anemia (FA), the chromosome instability syndrome, regulate TLS. The FA core complex
is required for spontaneous and UV-induced point mutagenesis, and cells deficient in the
FA core complex fail to form nuclear Rev1 foci (6).
Based on the results discussed in Chapter 3, catalytic dead Revl aids in the
resistance to 4-NQO significantly more when the error-free tolerance pathway is intact.
Therefore, Rev confers 4-NQO resistance in the MMS2 background through the noncatalytic domains. These results lead to many questions regarding how a cell tolerates
DNA damage. Is there a direct physical interaction between Rev1 and the proteins
involved in a template-switching mode of tolerance? Does Revl help recruit error-free
tolerance proteins to the site of the lesion? Or does the catalytic dead Rev 1 stabilize the
replication fork while attempting to insert, which gives the proteins involved in error-free
tolerance more time to get to the site of DNA damage? How does the cell decide when to
use error-free tolerance or TLS? Is this decision lesion-specific? We already know that
the ubiquitination state of PCNA operates as a signal for TLS or error-free tolerance.
Does monoubiqitination occur before polyubiquitination, meaning TLS attempts bypass
before template-switching? Why would the cell opt for the more error-prone process first
though? Are the ubiquitin modifications dynamic then? These questions are just the
182
starting point for designing the many possible experiments that will provide insight into
how the cell tolerates DNA damage.
183
TABLE 1
Genes tested for affecting Revi protein in a cycloheximide-chase assay.
Gene
RAD6
RAD18
UBC13
MMS2
TOM
UFO1
JAB]
UBM2 mutant of REV]
Protein Function
E2 ubiquitin-conjugating
enzyme
E3 ubiquitin ligase
E2 ubiquitin-conjugating
enzyme with Mms2
E2 ubiquitin-conjugating
enzyme with Ubcl3
E3 ubiqutin ligase
E3 ubiqutin ligase
Catalytic subunit of the
COP9 signalosome
Ubiquitin-binding motif in
Revl
184
Effect on Rev 1 Protein
Levels or Half-life in
Deletion Strain
None
None
None
None
Slight increase in half-life
None
None
None
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polymerase Rev1 is highly expressed during G(2)/M phase rather than S phase.
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185
186
Appendix A
The Role of Saccharomyces cerevisiae Revi in the Cellular
Resistance to Cisplatin
187
INTRODUCTION
As discussed throughout this thesis, Rev1 and Pol
(the heterodimer of Rev3 and
Rev7) have central roles in DNA damage tolerance, primarily through the translesion
synthesis (TLS) branch. In this section, I will focus on further understanding why Revi
protein levels undergo a 50-fold increase from G1 to the G2/M stages of the cell cycle.
Rev1 is cell cycle regulated; yet the rev1A strain has no known defects in cell cycle
progression during normal growth. The revlA cells do arrest, however, in the budded
state more than rad30A cells when measured 24 hours following UV irradiation (12).
The second question addressed in this appendix is why Revi is so important for
the cellular resistance to cross-linking agents. For most DNA damaging agents, TLS
mutants exhibit modest sensitivities relative to mutants in repair pathways. Curiously,
some TLS mutants are particularly sensitive to the cross-linking agent, cisplatin (7). In a
ranking of the 50 most sensitive deletion strains to cisplatin, the revlA (ranked 11th) and
rev3A (ranked
1 0 th)
were as sensitive or more sensitive than many repair mutants (14).
For comparison, the same strains were respectively the
5 5th
and
8 2 nd
least resistant strains
to UV irradiation. This bias toward requiring TLS for cisplatin resistance seems to be a
general need for DNA damage tolerance, since the error-free tolerance deletion strains,
rad5A (ranked 6th) and mms2A (ranked
13th),
were also quite sensitive. On the other hand,
Pol ~ (Rad30), the third TLS polymerase in S. cerevisiae, does not significantly
contribute to the tolerance of cisplatin lesions (6).
The preliminary results in this appendix are a starting point for studying the
functional purpose of Rev l's cell cycle regulation and learning more about the important
role that Rev1 has in cisplatin resistance. Future experiments will help determine whether
188
or not there is a connection between these two points of interest as described in the
discussion of this appendix.
189
MATERIALS AND METHODS
Yeast Strains:
Table 1 displays a complete list of strains used in this study. A W1588-4C (MA Ta
leu2-3,112 ade2-1 canl-100 his3-11,15 ura3-1 trpl-1 RAD5) (15) derivative (also
barlA.::LEU2 and rev1A:.:kanMX4) (2, 13) is the isogenic parent of all strains bearing the
pAS311 plasmids. S288C (MATa SUC2 gal2 mal mel flol flo8-1 hap] ho biol bio6) (8)
served as the parent for all pRS416-bearing strains. REVI was deleted via a one-step
replacement, amplifying the rev1::kanMX4 cassette from the deletion library and
transforming the product into the S288C strain (11). The BAR] gene was disrupted by a
one-step gene replacement using digested pZV77 (gift from S. Bell). All cassettes and
plasmids were introduced through a standard lithium acetate protocol (4).
Spotting Assay for Survival:
Cultures were inoculated with a single colony and grown while rotating at 30 C
in 5 ml SC-Ura plus 2% glucose or SC-Trp plus 2% raffinose for two overnights. In a 96well plate, the saturated cultures were diluted in sterile water as 10-fold serial dilutions to
10-4. For the overexpression strains containing the pAS311 vectors, 10 [tl of each dilution
was spotted onto SC-Trp plus 2% galactose plates with and without the indicated DNA
damaging agent. For the rev]A strains containing the pRS416 vectors, 10 dtl
of each
dilution was spotted onto SC-Ura plus glucose plates with and without the indicated
DNA damaging agent. UV-treated cells were exposed to UV at 1 J/m 2 s with a G15T8 UV
lamp (General Electric) at 254nm. Stock solutions of 4-NQO at 2 mg/ml (Sigma) in N, N
- dimethylformamide, carmustine (Sigma) in DMSO at 2.5 mg/ml, 10 mM nitrogen
190
mustard in sterile water, or freshly prepared cis-diammineplatinum (II) dichloride
[cisplatin] (Sigma) at 0.8 mg/ml in sterile water were added directly to the desired agar
media for plates at the appropriate concentration. Methyl methane-sulfonate (Sigma) was
added to agar media directly from the 99% solution as sold.
191
RESULTS
Overexpression of Rev1 particularly increases cellular resistance to cisplatin:
Given that Revl's protein levels are cell cycle regulated and peak during G2/M
(13), I wanted to know more about why Rev levels dramatically increase during that
stage of the cell cycle. In Chapter 3, I explain that Revl's DNA polymerase activity is
important in a lesion-specific manner. Therefore, I hypothesized that the increase in Rev 1
protein levels during the cell cycle may be for the tolerance of a particular lesion or may
be required to enhance a certain function of Rev1. If increasing Revl protein levels alone
could confer a phenotypic advantage or disadvantage in response to a DNA damaging
agent, we would learn more about why Rev1 protein levels are cell cycle regulated.
To explore this question, I exposed wild type strains that carried galactoseinducible REV] alleles to a variety of DNA damaging agents that cause different lesions.
Surprisingly, the strains carrying REVI or the catalytic dead rev] allele for
overexpression show approximately a 1000-fold protection to killing in response to
cisplatin damage relative to the strains bearing the empty vector or BRCT mutant allele
(Figure 1, columns 1 and 2 versus 3 and 4). There also is a slight increase in resistance to
methylmethane sulfonate (MMS) when REV1 is overexpressed (Figure 1). Even though
this experiment creates the non-natural situation of overexpressing REV] and does not
measure survival in a cell cycle-dependent manner, these results provide a clue as to
under what conditions Rev1 protein levels matter. This is especially intriguing because
this is the first phenotype for overexpression of full length REV] that we are aware of in
our lab.
192
The data in Figure 1 suggest that increasing Revi protein levels helps in DNA
damage tolerance for certain lesions in a way that is independent of Rev l's catalytic
activity. Cisplatin introduces lesions such as 1,2-intrastrand linkages between the N7
positions of adjacent guanines and monoadducts into the DNA (3). Since 4nitroquinoline-1-oxide (4-NQO), MMS, and UV also cause monoadducts, I chose to test
other agents that create cross-links when reacting with DNA. Interestingly,
overexpression of REVI does not enhance survival for cells exposed to carmustine or
nitrogen mustard (Figure 2), possibly reflecting a difference in the type of cross-links
produced.
Cisplatin resistance is dependent on Revl's non-catalytic functions:
Although the rev1A strain is known to be quite sensitive to cisplatin treatment,
how RevI functions in the resistance to cisplatin is still unknown. From Figure 1 in this
appendix and Chapter 3 of this thesis, we know that the Rev 1's catalytic activity is not
important for the tolerance of cisplatin lesions in S. cerevisiae as found in DT40 cells (9).
I further support those findings here and expand on the experiment to include additional
catalytic domain mutant strains as well as the BRCT domain mutant and the UBM2
mutant strains. In the rev1A background, strains carrying the catalytic dead or the other
catalytic domain mutant alleles (rev]-F367L or revl-F441L) are as resistant to cisplatin
as the REV]-bearing strain (Figure 3, columns 3,4, and 5 versus 2). The UBM2 or BRCT
mutant strains show sensitivity to cisplatin (Figure 3, columns 6 and 7 versus 2),
indicating that the non-catalytic functions of RevI are participating in cisplatin tolerance.
Furthermore, Revl's C-terminus contributes the most to cisplatin and nitrogen mustard
193
resistance (Figure 4, column 6 versus 2). The C-terminus mutant is far more sensitive to
these cross-linking agents than the BRCT and UBM2 mutants and has a similar level of
survival as the empty vector strain (Figure 4, column 6 versus 4, 5, and 2). As with
cisplatin, the catalytic activity also does not contribute to nitrogen mustard resistance
(column 3 versus 2). Therefore, the non-catalytic functions mediated through the BRCT
and UBM2 domains of Revi aid in the resistance to these cross-linking agents, but the Cterminus, likely through its interaction with Pol , is more critical for survival.
194
DISCUSSION
The results presented here reveal the novel phenotype of enhanced survival in the
presence of cisplatin when Revl protein levels are greater than under normal
circumstances. Since REV] was overexpressed using a galactose-inducible promoter in
these strains, the protein levels of Revl are high throughout the cell cycle in the presence
of galactose, meaning that the phenotype could reflect the effect of overall greater Revl
levels or having more RevI around during the GI and S stages of the cell cycle. The
mechanism of how this protection to killing is being conferred through Revl remains
unknown, but the data in Figures 1 and 4 suggest that the non-catalytic function provided
by the C-terminal region of Revi that interacts with Pol
is critical. Data from a first-
pass experiment supports that the cisplatin protection phenotype is dependent on Pol (
(data not shown).
Does this phenotype give us a clue as to when and how increasing RevI protein
levels affects Revl activity? At this stage, the almost 1000-fold difference in survival
after cisplatin exposure for the strain overexpressing REV1 compared to the strain
carrying the empty vector could be used as an experimental tool. For example, one could
measure survival for cells arrested during the GI or G2/M stages of the cell cycle and
exposed to cisplatin during the arrest. By comparing a wild type strain to a strain
overexpressing REVI in this experiment, we could learn if increasing Rev1 during a
certain stage of the cell cycle caused the enhancement in cisplatin resistance observed in
Figure 1.
Interestingly, this phenotype seems specific to cisplatin and not other crosslinking agents. One possible explanation is that cisplatin causes the cytotoxic lesion of
195
intrastrand cross-links, while the nitrogen mustard's cytotoxic lesion appears to be
interstrand cross-links, even though nitrogen mustard primarily produces monoadducts
and can also cause intrastrand cross-links (1, 3). Carmustine exposure leads to interstrand
cross-links in the DNA as well (1).
Another potential reason for the difference in survival phenotypes between
cisplatin and nitrogen mustard when REV] is overexpressed is related to the cell cycle.
Interestingly, nitrogen mustard and cisplatin cause cells to arrest or delay cell cycle
progression during different stages of the cell cycle. While exposing cells to nitrogen
mustard causes a G1/S delay in wild type cells, cisplatin treatment leads to a G2/M arrest
(5, 10), corresponding to when Rev protein levels are the highest in S. cerevisiae.
Moreover, the rev3A strain is more sensitive to nitrogen mustard when treated during a
G 1 arrest than during an S-phase or G2-phase arrest (10), and arrest permanently in G2
after cisplatin treatment (unlike rad30A cells) (6, 7). Possibly, Revl levels peaking
during G2/M help recruit its partner, Pol , to promote tolerance of intrastrand crosslinks.
Finally, the data presented here emphasize the significance of the C-terminus in
tolerating cisplatin or nitrogen mustard lesions (Figure 4) in agreement with the DT40
cell studies (9). This information is not unexpected given our results in Chapter 3 that
show the C-terminus is also critical for 4-NQO resistance. The large difference in
sensitivity between the C-terminus mutant and the BRCT and UBM2 mutants in Figure 4,
though, is noteworthy. Maybe the BRCT and UBM2-dependent functions of Rev1 are
needed less in the tolerance of cross-links compared to other types of damage.
196
ACKNOWLEDGEMENTS
I am grateful to Sanjay D'Souza for supplying the pAS311 plasmids and
Elizabeth Wiltrout for editing this section.
197
TABLE 1
Strains used in this study
Source
Relevant genotype
Strain
YMEW7
S288C barlA::LEU2 revlA::kanMX4 pRS416
Chapter 3
YMEW8
S288C barlA::LEU2 revlA::kanMX4 pRS416-REV1
Chapter 3
YMEW9
S288C barlA::LEU2 revlA::kanMX4 pRS416-revl-
Chapter 3
D467A E468A
YMEW10
S288C barlA::LEU2 revlA::kanMX4 pRS416-revl-
Chapter 3
G193R
YMEW11
S288C barlA::LEU2 revlA::kanMX4 pRS416-revl-
Chapter 3
E820A L821A P822A T823A Q824A
YMEW12
S288C barlA::LEU2 revlA::kanMX4 pRS416-revl-
Chapter 3
L889A V890A K891A W893A V894A
YMEW15
S288C barlA::LEU2 revlA::kanMX4 pRS416-revl-
Chapter 3
F367L
YMEW1 6
S288C barlA::LEU2 revlA::kanMX4 pRS416-revl-
Chapter 3
F44 1L
YMEW32
W1588-4C barlA::LEU2 pAS311
This study
YMEW33
W1588-4C barlA::LEU2pAS311-REVI
This study
YMEW34
W1588-4C barlA::LEU2 pAS311-rev1-D467A
This study
E468A
YMEW35
W1588-4C bar1A::LEU2 pAS311-rev1-G193R
This study
The parental strains are W1588-4C (MA Ta leu2-3,112 ade2-1 can 1-100 his3-11,15 ura3-1 trpl-1
RAD5) (15) or S288C (MATa SUC2 gal2 mal mel flol flo8-1 hapi ho biol bio6) (8) as indicated.
198
FIGURE 1. - Ectopic overexpression of RE V causes an agent-specific protection to
cisplatin damage. Cells grown to saturation were diluted in 10-fold serial dilutions, and
then spotted onto SC-Trp plus galactose plates containing the indicated DNA damaging
agent or exposed to UV. DNA damage agents shown are 4-NQO (0.2 tg/ml), cisplatin
(64 [tg/ml), MMS (0.025%), and UV (75 J/m 2). Plates were incubated at 30 oC for 4 days.
Strains are YMEW33 (W1588-4C pAS311-REV1) - column 1, YMEW34 (W1588-4C
pAS311-rev-catalytic dead) - column 2, YMEW32 (W1588-4C pAS311) - column 3,
and YMEW35 (W1588-4C pAS31 I -revi-BRCT) - column 4.
199
FIGURE 1
No damage
4-NQO
Cisplatin
MMS
UV
1
101
102
10-3
10,4
1234 123412344 1234
200
123
1234
FIGURE 2. - Ectopic overexpression of RE V does not enhance resistance to other
crosslinking agents. Cells were treated and plated as in Figure 1 except that the DNA
damaging agents are nitrogen mustard at 5 [tM (dose 1) and 50 [tM (dose 2) and
carmustine at 50 [tM (dose 1) and 150 RM (dose 2). Strains are YMEW33 (W1588-4C
pAS311-REV1) - column 1 and YMEW32 (W1588-4C pAS311) - column 2.
201
FIGURE 2
No Damage
Nitrogen Mustard
Dosel
Dose2
Carmustine
Dosel
Dose2
0-0
12
12
12
202
12
12
FIGURE 3. - Strains are extremely sensitive to cisplatin in the complete absence of
Rev due to polymerase-independent functioning. Cultures were grown to saturation,
serial diluted, spotted onto SC-Ura plus glucose plates containing the indicated drug dose,
and grown at 30 C for 4 days. Plates shown in order from left to right contain no
damage, cisplatin (16 tg/ml), cisplatin (40 tg/ml), and cisplatin (64 tg/ml). Strains are
YMEW7 (revlA pRS416) - column 1, YMEW8 (revlA pRS416-REV1) - column 2,
YMEW9 (revlA pRS416-rev1-catalytic dead) - column 3, YMEW15 (rev A pRS416rev]-F367L) - column 4, YMEW16 (revl A pRS416-rev]-F441L) - column 5, YMEW 11
(revlA pRS416-rev1-UBM2) - column 6, and YMEW10 (revl A pRS416-revl-BRCT) column 7.
203
FIGURE 3
No damage
Cisplatin Dose 1
1234567
1234567
Cisplatin Dose 2 Cisplatin Dose 3
1234567 1234567
1
10.1
10-2
10-3
104
204
FIGURE 4. - The function conferred by the C-terminus of Rev1 is the most critical for
resistance to crosslinking agents. Cells were treated as in Figure 1 except the type and
dose of DNA damage in the plate differed. Plates shown in order from left to right
contain no damage, cisplatin (40 [tg/ml), nitrogen mustard (1 [M), and nitrogen mustard
(3 [M). Strains are YMEW8 (rev1A pRS416-REV1) - column 1, YMEW7 (revlA
pRS416) - column 2, YMEW9 (revlA pRS416-revl-catalytic dead) - column 3,
YMEW10 (revlA pRS416-rev1-BRCT) - column 4, YMEW11 (revlA pRS416-revlUBM2) - column 5, and YMEW12 (rev] A pRS416-revl-C-terminus)- column 6.
205
FIGURE 4
No damage
Cisplatin
Nitrogen Mustard
Nitrogen Mustard
12 3456
123456
12 3456
12 3456
1
10.1
10-2
10-3
10-4
206
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