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Microbiol Mol Biol Rev. 2009 March; 73(1): 134–154.
PMCID: PMC2650891

Eukaryotic Translesion Polymerases and Their Roles and Regulation in DNA Damage Tolerance


Summary: DNA repair and DNA damage tolerance machineries are crucial to overcome the vast array of DNA damage that a cell encounters during its lifetime. In this review, we summarize the current state of knowledge about the eukaryotic DNA damage tolerance pathway translesion synthesis (TLS), a process in which specialized DNA polymerases replicate across from DNA lesions. TLS aids in resistance to DNA damage, presumably by restarting stalled replication forks or filling in gaps that remain in the genome due to the presence of DNA lesions. One consequence of this process is the potential risk of introducing mutations. Given the role of these translesion polymerases in mutagenesis, we discuss the significant regulatory mechanisms that control the five known eukaryotic translesion polymerases: Rev1, Pol ζ, Pol κ, Pol η, and Pol ι.


The faithful replication of DNA and proper transmission of chromosomes are essential for inheritance of 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, that are 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 we will briefly introduce the many strategies a cell may employ to allow survival in the face of DNA damage before we turn to the contribution of damage tolerance mechanisms, in particular translesion synthesis (TLS). We describe the DNA polymerases that mediate TLS and highlight the unique properties of the Rev1 and Pol ζ families, which are together responsible for the majority of mutations in eukaryotes from yeast to humans. We 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. We conclude with a discussion of the two major models for the regulation of mutagenesis resulting from TLS.


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 (146). 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 (53, 146). Additionally, many environmental factors can cause DNA damage, such as ionizing or UV radiation and chemical agents, including methyl methanesulfonate, cisplatin, and benzo[a]pyrene (53). 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 (Fig. (Fig.1A).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 nonhomologous end joining, homologous recombination, or single-strand annealing. The reader is referred to reference 53 and the many excellent reviews that are available for further information on DNA repair.

FIG. 1.
DNA damage repair and bypass mechanisms. (A) DNA damage results in breakage of the sugar-phosphate backbone, DNA base loss (indicated by a gap in the DNA), or base alterations (as indicated by the gray star). This damage can be repaired/removed from the ...

Additionally, cells possess mechanisms to temporarily tolerate DNA damage until DNA repair processes can remove the damage (Fig. (Fig.1A).1A). In eukaryotes, tolerance includes an error-free pathway and a parallel, more mutagenic pathway, as reviewed by Andersen et al. (7). The type of posttranslational modification on the processivity clamp proliferating cell nuclear antigen (PCNA) 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 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 (Fig. (Fig.1B)1B) (53). 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 review, we 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 the complex regulatory pathways is that the mutations which do arise occur under conditions of stress when they might be evolutionarily advantageous through increasing genetic variability, as demonstrated in somatic hypermutation (SHM) of immunoglobulin genes.


TLS is the process by which a DNA lesion is bypassed by the incorporation of a nucleotide opposite to the lesion (53) (Fig. (Fig.1B).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 (15, 53). 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 that many modified bases may impose (53, 68, 217). TLS polymerases are found in organisms throughout all three domains of life. Most TLS polymerases are members of the Y family of DNA polymerases (199), 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, but as this activity is often weak or not the primary function of these polymerases (see below), we focus on the Y family of DNA polymerases, which are uniquely adapted for TLS. The Y family members include Rev1, Pol κ (DNA Pol IV in bacteria), Pol η, Pol ι, 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 Table1.1. Another eukaryotic DNA polymerase, Pol ζ, is a member of the B family of DNA polymerases, which includes replicative DNA polymerases, yet it 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.

Genes encoding catalytic subunits of eukaryotic DNA translesion polymerases

Discovery and 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 Saccharomyces cerevisiae that were unable to revert an auxotrophic marker to the wild-type allele after UV irradiation (137). Using this approach, REV1 (encoding the Rev1 DNA polymerase) and REV3 (encoding the catalytic subunit of Pol ζ) 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 Escherichia coli led to the identification of umuC (encoding the catalytic subunit of UmuD′2C, DNA Pol V) (109).

Although the REV1, REV3, and umuC genes were identified by their profound contributions to damage-induced mutagenesis, other translesion polymerase genes have more subtle effects on mutagenesis and were first identified primarily by homology searches with other TLS polymerases. For example, RAD30 and Pol ι (hRAD30B) were first identified solely by their homology to the TLS polymerase genes REV1, umuC, and dinB (initially identified in E. coli) (112, 165, 166, 225, 251). It was not until 1996 that the first biochemical description of a specialized translesion polymerase appeared (Pol ζ, a B family DNA polymerase [185]), followed rapidly by the demonstration that the Rev1 protein had a restricted DNA polymerase activity and was able to insert Cs opposite an abasic site (184). Even then, it was not until 1999, with the characterization of the translesion polymerase activity of DNA polymerase η (100, 156, 157), that it was recognized that all of the genes which shared homology with the eukaryotic REV1 and bacterial umuC were in fact genes for DNA polymerases with the unique ability to replicate over DNA lesions (extensively reviewed in references 68 and 217). 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 (199).

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 (147, 148, 180, 181, 232, 244, 245, 279), 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 (Fig. (Fig.2)2) (15, 217, 267). Like replicative polymerases, the catalytic aspartate and glutamate residues, which coordinate the divalent magnesium ions that stabilize the triphosphate group of the incoming deoxynucleoside triphosphate (dNTP), are located in the central palm region (15, 217, 266). The thumb and finger 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 (Fig. (Fig.2)2) (15, 217, 266).

FIG. 2.
Crystal structures of two Y family polymerases. (A) Cocrystal structure of the S. cerevisiae TLS Pol η with a DNA template containing a cisplatin cross-link. The structure is oriented to highlight the right-hand architecture as seen in both TLS ...

Although they share a common overall architecture, Y family polymerases differ from replicative polymerases in certain key ways that allow them to perform TLS. 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 (68, 267). A novel little-finger domain, also called the polymerase-associated domain (PAD) (148, 232) or wrist (244), is present only in Y family polymerases and extends from the classical finger domain to make extra contacts with the DNA (Fig. (Fig.2)2) (266, 267). Certain TLS polymerases also contain additional regions, such as the “N-clasp” of DNA Pol κ, that further contribute to DNA binding (150). These additional DNA-binding regions provide important stability for the ternary complex, since Y family members have short, stubby thumb and finger domains which make few contacts with the DNA backbone (217, 266). Y family polymerases generally have an open grip on the DNA (Fig. (Fig.2)2) and a greatly reduced processivity relative to replicative DNA polymerases (55, 217); truncations of the little-finger domain or N clasp reduces DNA binding and processivity even further (148, 150, 217). 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 (Fig. (Fig.2)2) (160, 162) and has been implicated in lesion specificity (23).

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 (217, 266, 267). A more spacious active site allows accommodation of large bulky adducts (149, 180) and even two covalently linked bases in a thymine-thymine dimer (147). Other Y family members appear to have more constrained active sites that nonetheless are specialized to accommodate particular classes of DNA lesions (150). Further promoting their ability to use modified DNA templates, Y family polymerases make fewer contacts with the forming base pair (266, 267) 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 (206). 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 (267); however, there is evidence to suggest a conformational change during catalysis by archaeal Dpo4, S. cerevisiae Pol η, and perhaps human Pol η as well (48, 102, 255).

As of the writing of this review, four eukaryotic Y family polymerases have been cocrystallized with DNA: human Pol ι (181), S. cerevisiae Pol η with a cisplatin adduct (Fig. (Fig.2A)2A) (6), human Pol κ (150), and S. cerevisiae Rev1 (Fig. (Fig.2B)2B) (179, 180). Each polymerase displays distinct and often unusual interactions with the DNA lesion and incoming nucleotide. For example, Rev1 uses an amino acid in the catalytic domain to base pair with an incoming dCTP in lieu of pairing with the DNA template (Fig. (Fig.2B)2B) (180), while Pol ι appears to use an unusual Hoogsteen base pairing mechanism (181). In contrast, Pol κ uses Watson-Crick base pairing to incorporate nucleotides and has a relatively constrained active site (150). 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 (68, 119, 217). Therefore, TLS polymerases have a potentially mutagenic activity inside the cell (52). The lack of a 3′-to-5′ proofreading domain reduces the fidelity of TLS polymerases operating on undamaged DNA by ~10−2 compared to replicative DNA polymerases (68, 119, 267), 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 (266, 267), and certain TLS polymerases, like DNA Pol ι and Rev1, do not use canonical Watson-Crick base pairing (180, 217). 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

However, despite their relatively low fidelity on undamaged DNA, a paradigm shift has reclassified TLS polymerases from generally being considered “error-prone” polymerases (68), as often initially described, to a more nuanced understanding of their role as lesion-specific bypass polymerases (52). 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 (97, 100, 253). 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 an efficiency equal to or higher than that on undamaged DNA (97, 100, 162, 253). This is strikingly seen in the case of DNA Pol η, which is specialized to bypass cis-syn TT dimers caused by UV irradiation (99, 100, 157). Although DNA Pol η exhibits among the lowest fidelities of any TLS polymerase on undamaged DNA (~10−1) (158), its highly accurate activity in bypassing this UV-induced lesion makes Pol η critical for the avoidance of sunlight-induced skin cancers in humans (discussed further below) (132). Additionally, Pol κ, and its archaeal and bacterial homologs, can bypass certain N2-dG adducts accurately and efficiently (97).


Phylogenetic analysis and extensive biochemical characterization have revealed that there are five subfamilies within the Y family of DNA polymerases: Rev1, UmuC, DinB/Pol κ, Pol ι, and Pol η (199), 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 ζ (Rev3/Rev7) (123). For this review, we 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, we need to introduce one recurring theme for the regulation of eukaryotic TLS polymerases involving the physical and genetic interactions with PCNA and the proteins involved in modifying PCNA. The homotrimeric clamp, PCNA, serves as the processivity factor for Pols δ and epsilon during replication (26). For its role in DNA damage tolerance, particular ubiquitin modifications of PCNA are involved. Specifically, the Rad6-Rad18 complex catalyzes the monoubiquitination of PCNA at K164, a modification that stimulates TLS, the more mutagenic branch of tolerance (238). Monoubiquitination of PCNA can be later extended to polyubiquitination by the Mms2-Ubc13-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 (reviewed by Ulrich [246]). Therefore, the interaction(s) between TLS polymerases and PCNA is key in comprehending TLS activity and regulation, as discussed frequently in the sections below.


Uniquely among eukaryotic Y family polymerases, Rev1 actively promotes the introduction of mutations in organisms ranging from unicellular yeast to multicellular organisms, including humans (65, 137). Cells bearing rev1 mutations display a drastic reduction in spontaneous and induced mutagenesis by a wide variety of DNA-damaging agents (53, 123). In multiple genetic backgrounds and in response to different types of DNA lesions, mutations of REV1 abolish most mutagenesis, indicating its fundamental importance to this biologically important process (123). For example, Rev1 is required for ~95% of all UV-induced base pair substitutions (124). Although only marginally correlated with the onset of cancer to date (85, 230), REV1 has been shown to modulate the frequencies with which cisplatin-resistant cells are generated from an ovarian carcinoma cell line (144, 201). Therefore, REV1 may contribute to cancer progression and could be an important target of cancer therapy.

Rev1's catalytic activity.

Rev1 was the first member of the Y family to be shown to have the capability of catalyzing the formation of a phosphodiester bond (184). However, because its activity was limited, Rev1 was described as a DNA polymerase only after the discovery of the DNA polymerase activity of other Y family members (199). Rev1 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 (123, 184, 254). 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 (Fig. (Fig.2B)2B) (180). 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 (Fig. (Fig.2B)2B) (180).

In contrast, a catalytically inactive mutant of Rev1 displays no reduction in levels of mutagenesis induced by a wide range of DNA-damaging agents, including UV light (80, 224), although a change in the mutation spectrum is observed (202, 224). Interestingly, DNA damage sensitivity and mutagenesis phenotypes are observable for the catalytic dead mutant after cells are exposed to 4-nitroquinoline 1-oxide, an alkylating agent that produces, among other lesions, N2-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 N2-dG adducts (M. E. Wiltrout and G. Walker, unpublished data).

Rev1's noncatalytic function(s) and protein interactions.

REV1 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 (183, 202, 276). Therefore, although Rev1's unique and highly specialized dCMP transferase catalytic activity is conserved from yeast to humans (142, 184), its DNA polymerase activity does not seem to be required for bypass of many lesions for which Rev1 function is required in vivo. Instead, the ability of Rev1 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 (BRCA1 C-terminal) domain, the C-terminal ~100 amino acids, the PAD, and the ubiquitin-binding motifs (UBMs) (Fig. (Fig.33).

FIG. 3.
Cartoon representation of the protein domains in the human B family TLS polymerase ζ and the Y family TLS polymerases Rev1, κ, ι, and η. aa, amino acids. (Based on data from references 58 and 268.)

The N-terminal BRCT domain of Rev1 was the first region to be characterized, since the original loss-of-function rev1-1 mutation in S. cerevisiae, whereby the REV1 gene was identified, is located in the BRCT domain (65, 122, 137). The BRCT 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 (29, 249). In striking contrast to mutations in the catalytic active site, mutations affecting the BRCT domain largely inactivate Rev1 in vivo. In yeast, BRCT mutants exhibit a severe defect in survival and mutagenesis after DNA damage (137). More recently, mutations affecting the BRCT domain have also been shown to reduce REV1 function in higher eukaryotes; however, the extent of the defect varies between studies (71, 96, 224). It was this finding that REV1 function could be inactivated by a mutation which left the polymerase activity of Rev1 intact that first led Nelson et al. to propose a “second function” for Rev1 (183). This idea was supported by the recognition that BRCT domains can mediate protein-protein or protein-DNA interactions (29, 66, 115, 259). 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 (70, 80, 243).

Recently, the BRCT domain of Rev1 has been shown to interact with two DNA-binding proteins: the replicative clamp PCNA (71) and the Rev7 subunit of DNA Pol ζ (44a). Since a BRCT domain is not found in other TLS polymerases, these interactions are likely to confer unique properties upon Rev1. It seems probable that the BRCT domain may promote a specialized interaction with damaged DNA to mark sites of incomplete replication that require TLS. Indeed, in subcellular localization studies, the BRCT domain is sufficient for nuclear localization (243) and is required for constitutive localization of Rev1 to replication foci (71). This may be because mouse Rev1 interacts with PCNA via the BRCT domain of Rev1 (71). 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, Rev1 likely interacts with a different surface of PCNA than the other TLS polymerases. As monoubiquitinated PCNA accumulates in response to agents that block replication forks (107), this unique interaction may have functional consequences for differential recruitment to lesions and/or, as one study indicates, stimulation of catalysis by Rev1 relative to Pol ζ (59). The biological relevance of stimulation of Rev1 catalysis by monoubiquitinated PCNA remains unresolved, though, since conflicting in vitro results have been reported (59, 82).

Moreover, as BRCT domains in other proteins have been found to interact with DNA at single-stranded regions or double-strand breaks (115, 259), the BRCT of Rev1 may also be directly involved in localizing Rev1 to aberrant DNA structures. Supporting this suggestion is a study demonstrating that both S. cerevisiae Pol ζ and Rev1 are recruited to the vicinity of an endonuclease-induced double-strand break (88). This property was found to require the Rev1 BRCT domain but not its catalytic activity.

In an activity unrelated to DNA binding, the BRCT domain of Rev1 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 (66, 154, 270). Although Rev1 has only a single BRCT domain, it has also been implicated in phosphopeptide binding in vitro (270). To date, an interaction between the Rev1 BRCT domain and phosphorylated target proteins has not been demonstrated in vivo. In principle, however, the ability to bind phosphorylated target proteins would allow unique opportunities for regulation (or activity) of or by Rev1 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. The last ~100 amino acids of mammalian Rev1 interact with the TLS Pols η, ι, κ, and ζ (70, 176, 197, 243). Initially, the polymerase interaction region at the C terminus of Rev1 did not seem to be conserved between higher eukaryotes and yeast (107, 176, 243). However, extensive sequence alignment and functional studies have revealed that S. cerevisiae Rev1 does interact with another TLS polymerase, DNA Pol ζ, through its C terminus (2, 4, 44a, 44b, 117, 224). 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 of its interaction with Rev3 that stimulates Pol ζ extension from a mismatch or opposite a DNA lesion (4). Importantly, mammalian and S. cerevisiae rev1 constructs lacking this C-terminal polymerase interaction region are unable to complement a rev1Δ strain for survival or mutagenesis after DNA damage (4, 44b, 117, 122, 224), showing that Rev1 functions in vivo through interactions with other TLS polymerases. Additionally, in vitro, the PAD region of S. cerevisiae Rev1 interacts with the Rev7 subunit of DNA Pol ζ (2), as well as DNA Pol η, an interaction that mildly stimulates the polymerase activity of Rev1 (3).

Finally, mouse Rev1 binds ubiquitin via a noncanonical UBM (73). Interaction with ubiquitin is necessary for localization of Rev1 into DNA damage-induced foci (73) and for the hyperstimulation of its catalytic activity by PCNA-ubiquitin (261). Mutants with mutations in the UBM display increased chromosomal aberrations, decreased viability, and decreased mutagenesis after exposure to DNA-damaging agents (44b, 73, 261), showing that they, like the other interaction motifs, are required for REV1 function in vivo. Murine Rev1 is monoubiquitinated itself, but the mechanism of ubiquitination, the position in the protein for this attachment, and the functional relevance of this modification remain unknown (73, 171).

Thus, multiple protein-protein interaction domains are critical for REV1 function in vivo. These findings have led to a model in which Rev1 functions primarily as a scaffold for various postreplication 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 (51, 132). Thus, Rev1 is thought to play a central role in TLS by regulating access of TLS polymerases to the primer terminus (51, 132).

Temporal and spatial regulation of Rev1.

Despite its importance in regulating TLS, precisely how, when, and where Rev1 functions in vivo is not yet well understood. Rev1 clearly functions in mitotically dividing cells. The fact that the REV1 transcript is upregulated during meiosis in S. cerevisiae (27, 37) and has the highest expression in human testis (176) suggests a meiotic function for Rev1 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 Rev1-dependent TLS acts after the replication machinery has reprimed downstream and generated a single-stranded DNA (ssDNA) gap opposite to a lesion. Strikingly, in S. cerevisiae, Rev1 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 (257). REV1 mRNA levels are cell cycle regulated to a lesser extent. Additionally, Rev1 is phosphorylated in a cell-cycle-dependent manner, as well as in response to DNA damage in S. cerevisiae, although how this affects Rev1 activity is not yet known (227). Together with a key study indicating that TLS can occur after replication in S. cerevisiae (151), this unexpected finding has led to a reevaluation 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 (257). Although it appears that Rev1 is associated with chromatin constitutively throughout the cell cycle (227), Rev1 binds to ssDNA with high affinity and can likely translocate on this substrate to find primer termini (155). This ssDNA targeting may allow Rev1 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 ζ, 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 Rev1 in S. cerevisiae, REV1 in DT40 cells is required for replication fork progression in the presence of DNA damage, indicating a role for Rev1 during S phase in this system (46). Taken together, these results suggest that the partitioning of Rev1 function between possible replicative and postreplicative roles may vary between biological systems.

Localization studies using ectopically overexpressed green fluorescent protein fusions have reported that Rev1 forms foci in vertebrate cells after DNA damage (71, 174, 175, 243). Damage-induced focus formation requires the UBMs (73), while the BRCT domain is sufficient for nuclear localization and basal focus formation but is not required for damage-induced focus formation (71, 243), highlighting the role of the protein-protein interactions in mediating Rev1 function. Under conditions of ectopic overexpression, colocalization of Rev1 foci with PCNA and Pol η has been interpreted to indicate that Rev1 associates with replication forks to enable continuous DNA synthesis on templates containing DNA lesions (71, 174, 243). This first led to a model in which Rev1 was thought to act mainly during replication (51, 132). However, Rev1 foci have also been observed in G1 (175). Additionally, one study observed no Rev1 foci, occurring either spontaneously or after DNA damage, when a more physiological expression level of Rev1 was used (224). These authors propose that lack of focus formation reflects the need of the cell for only one or a small number of molecules of Rev1 at sites of stalled replication (224).

Given that the cell cycle regulation of Rev1 is likely to be complex and that cells appear to keep the levels of Rev1 as well as Pol ζ low (124), it seems likely that high levels of Rev1, especially during G1 and S, would be detrimental. Additionally, Rev1 protein levels are under the control of proteasomal degradation throughout the cell cycle in S. cerevisiae (M. E. Wiltrout and G. Walker, unpublished data). Indeed, the fact that overexpressed Rev1 localizes to replication forks may help to explain why S. cerevisiae cells keep the level of Rev1 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 (124), future experiments taking advantage of new technology such as DNA combing may reveal phenotypes that are more challenging to observe. For example, DNA combing has been used to show that even mild overexpression of Pols β and κ in mammalian cells interferes with normal replication fork progression (212).

Potential relationship between diseases and proper Rev1 function.

Novel in vivo functions for Rev1 are only beginning to be uncovered. In higher eukaryotes, additional pathways regulating Rev1/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 cross-linking agents and are hypomutable (62, 87, 171, 187, 204, 215). More specifically, it has been reported that Rev1 and Rev3 are epistatic to FANCC with respect to survival after cisplatin exposure in DT40 cells and that Rev1 colocalizes with FANCD2 after the blockage of replication in HeLa cells (187). Another recent study shows that the FA core complex is required for mutagenesis and efficient Rev1 focus formation in mammalian cells in a manner that is independent of PCNA monoubiquitination (171). These results contribute to the expanding field of TLS regulatory mechanisms that are not necessarily related to PCNA modifications.

Rev1 has also been shown to have an unanticipated role in preventing trinucleotide repeat expansion, particularly for those with hairpin-forming capacity. This as-yet poorly understood role of Rev1 may be relevant to neurodegenerative diseases (38, 43). Furthermore, Rev1, as well as Rev3 and Rev7, participates in nuclear mutagenesis induced by mitochondrial dysfunction, localizes to the mitochondria, and contributes to mitochondrial mutagenesis in S. cerevisiae (220, 274). Given the connection of mitochondrial function to disease, these functions of TLS polymerases may be associated with human diseases.

Pol ζ (Rev3/Rev7)

DNA Pol ζ is a heterodimer composed of the Rev3 catalytic subunit and the Rev7 accessory subunit (185). REV3 was identified in the same screen for reversionless mutants in S. cerevisiae as REV1 (137). REV7 was isolated by a similar strategy a few years later (125). Like rev1 mutants, 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 (123). 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 (88, 123). Like REV1, DNA Pol ζ plays a key role in most mutagenesis from yeast to humans (34, 64, 137, 168) as well as in cisplatin resistance in human cancer cells (145). Together, Rev1 and DNA Pol ζ are thought to mediate the vast majority of the mutagenic class of DNA damage tolerance in vivo.

Pol ζ's catalytic activity.

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 δ, epsilon, and α (123, 173). In contrast to most other B family replicative polymerases, DNA Pol ζ lacks the motifs characteristic of a 3′-to-5′ exonuclease activity (123). Although it can bypass certain lesions such as a cis-syn TT dimer and perform both the insertion and extension steps opposite a thymine glycol lesion in an error-free manner (103, 185), Pol ζ 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 (123, 217). In combination with a relatively high error rate for base substitutions, this proficiency for extending mismatches is what allows Pol ζ to contribute significantly to mutagenesis (123, 278). The accessory subunit of Rev3, Rev7, significantly enhances the polymerase activity of Rev3 (185). Despite the lack of conserved PCNA interaction motifs, Pol ζ exhibits increased lesion bypass activity in the presence of PCNA (60). However, stimulation of Pol ζ activity is not observed with either monoubiquitinated PCNA or the alternative 9-1-1 processivity clamp (59, 190).

Other Pol ζ functions and protein interactions.

Although it is a very large protein, Rev3 does not contain any known protein-protein interaction modules or other regulatory motifs (Fig. (Fig.3).3). In S. cerevisiae, Rev3 interacts with the C-terminal 100 amino acids of Rev1 in vitro, and this interaction stimulates the ability of Pol ζ to extend mismatches and bypass specific lesions (4, 74). However, the majority of the regulation of Pol ζ activity appears to occur through the accessory factor of Rev7. Rev7 contains a HORMA (Hop1/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 ζ to sites of DNA damage (229). Additionally, Rev7 interacts with Rev1 (70, 176, 197, 240, 243), which seems likely to promote localization of DNA Pol ζ to DNA lesions.

The physical and genetic interactions of Rev1 with DNA Pol ζ are complex. Despite the fact that each of the three proteins interacts with the others (see above), a heterotrimer of Rev1, Rev3, and Rev7 does not appear to be formed between purified proteins, as binding of Rev1 to Rev7 inhibits interaction of purified Rev1 and Rev3 in vitro (4). These findings indicate that the architecture of the Rev1-Pol ζ complex is intricate and that several subcomplexes may exist, possibly in a regulated manner. It is also possible that the posttranslational modifications of Rev1 mentioned above may influence the nature of Rev1's interaction with DNA Pol ζ in vivo.

Although Rev1, Rev3, and Rev7 are generally believed to work together, the functions of Rev1 and Pol ζ are not entirely overlapping. For example, REV1 appears to act independently of REV3/7 in the generation of sister chromatid exchanges during the recombinational bypass mode of damage tolerance (200). Additionally, Rev1's role in preventing trinucleotide repeat expansion is independent of both its own catalytic activity and that of DNA Pol ζ. This suggests that for some cellular roles, Rev1 can also act alone (38). Moreover, REV7 appears to have a distinct and independent function in cell cycle control (see below).

Loss of Pol ζ causes embryonic lethality in mice (18, 47, 248, 260), indicating that during rapid proliferation, mammalian cells require a function of Pol ζ. The inability to study rev3 mutant cell lines in mammalian systems has hampered understanding of Pol ζ function. However, studies with the chicken DT40 line have provided insight into the role of Pol ζ in vivo, in particular, the contribution of REV1, REV3, and REV7 to chromosomal rearrangements during recombination and interstrand cross-link repair (200, 231). 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 subpathway of homologous recombination (90, 221).

Despite being a relatively small protein, Rev7 participates in many protein-protein interactions apart from its interactions with Rev1 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 Cdh1 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 (32, 177, 211). Interaction with Rev7 inhibits the ubiquitin ligase activity of the APC/C and prevents the onset of mitotic anaphase (32, 211). 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 (94). 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 (139), the Elk1 transcription factor affecting cell cycle progression and the DNA damage stress response (275), the PRCC cancer protein implicated in RNA splicing and mitotic progression (258), two metalloproteases involved in cell proliferation and signaling (186), and the adenovirus death protein (269).

Regulation of Pol ζ.

Multiple mechanisms collaborate to keep Pol ζ levels low (123), indicating that overexpression may be detrimental to cells. Indeed, overexpression of Pol ζ causes increased UV-induced mutagenesis and decreased UV resistance in S. cerevisiae (219). 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 (64, 65, 123, 142). 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 (123). REV3 transcript levels are upregulated above the normally low basal levels in late meiosis in yeast (234). Reminiscent of the cell cycle regulation of Rev1 in S. cerevisiae, Rev3 chromatin association in human cells has a cell cycle-regulated pattern, showing highest levels during the G1/S boundary, which decreased during S phase and increased again during late S and G2 (25).

Mammalian cells possess additional mechanisms to regulate TLS activity that are 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 REV1 mRNA levels (143). These backgrounds also exhibit increased rates of 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 (110, 129, 130, 235). Using a nondegradable p21, Soria et al. reported the inhibition of PCNA ubiquitination in the presence of stabilized p21 (236).

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 (139, 222). Rev7 overexpression has been found in colon cancer, and this correlates with chromosomal instability and patient mortality (222). Curiously, another study found that rev3 transcript levels were downregulated in colon carcinomas (25). The contradictory data reveal the complexity of cancer and suggest that TLS polymerases could have roles in cancer in specific contexts. Consequently, changes in the regulation of Rev7 and Rev3 levels in vivo may be connected to cellular events related to disease.

Pol κ (DinB)

In contrast to the REV genes, Pol κ was not identified by its involvement in mutagenesis or its resistance to DNA-damaging agents. Rather, Pol κ was identified by homology searches for eukaryotic orthologs of the E. coli dinB gene (101). Although dinB was first discovered in 1980 as a gene that was significantly upregulated during the bacterial SOS response (112), it was not until nearly 20 years later that its polymerase activity was demonstrated (251). Found in all domains of life (199), DinB/Pol IV (as it is known in E. coli) and Pol κ (as it is known in eukaryotes) is the most highly represented and most strongly conserved of all the TLS polymerases. The pervasiveness of Pol κ argues that this protein contributes to the normal functioning of all cells. It has been surprising, then, that loss of Pol κ generally reveals only mild phenotypes (see below). Although the bacterial ortholog, DinB, has been studied extensively (for a review, see reference 98), eukaryotic Pol κ has been less characterized, particularly in terms of its role(s) in TLS in vivo. This discrepancy derives in part from the fact that Pol κ is conspicuously absent from what is arguably the best-studied single-cell eukaryote, S. cerevisiae. Pol κ orthologs have been identified in other related fungal species, though, including the fission yeast Schizosaccharomyces pombe. Organisms that lack a DinB homolog may possess another protein that plays a functionally redundant role.

Pol κ's catalytic activity.

With regard to in vitro DNA replication on undamaged DNA, mammalian Pol κ is relatively accurate compared to other TLS polymerases; human Pol κ has a misinsertion frequency of 1 in every 102 to 103 nucleotides replicated (116). Although the bacterial and archaeal orthologs of DinB demonstrate a marked proclivity for −1 frameshifts in vitro and when overproduced in vivo (67, 113, 116, 216), mammalian Pol κ appears to be more restricted in this activity both in vitro and in vivo (196). With respect to DNA damage bypass, Pol κ has limited ability to synthesize across numerous DNA lesions (for reviews, see references 16 and 268); however, it can bypass many N2-adducted dG residues, including N2-dG-linked DNA-peptide cross-links, both efficiently and accurately (see, e.g., references 10, 97, 170, and 271). Indeed, it appears that Pol κ is specialized in its ability to bypass N2-adducted dG lesions; Pol κ operates with high accuracy and strikingly increased catalytic efficiency opposite N2-furfuryl-dG and N2-(1-carboxyethyl)-2-dG residues relative to an undamaged dG (97, 271). Significantly, like Pol ζ, Pol κ 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 (30, 150, 217). Furthermore, in vitro, the DNA synthesis activity of human Pol κ is stimulated in the presence of PCNA, replication factor C, and replication protein A but not by a single complex in the absence of the others (81).

Role of Pol κ in mutagenesis.

The role(s) of Pol κ in mutagenesis is enigmatic. In contrast to deletion of Rev1 and Pol ζ, deletion of Pol κ does not appear to have a profound effect on either spontaneous or damage-induced mutagenesis. In mammalian cells, loss of Pol κ sensitizes cells to the killing by benzo[a]pyrene and moderately increases mutagenesis induced by this agent, suggesting that Pol κ bypasses N2-benzo[a]pyrene dG adducts relatively accurately in vivo (10, 195). Loss of Pol κ is also associated with sensitivity to DNA-alkylating agents and to UV irradiation (193, 239), although sensitivity to UV in the absence of Pol κ seems likely to reflect its yet-to-be-defined role in nucleotide excision repair (193). Ectopic overexpression of human Pol κ in mammalian cell lines inhibits replication fork progression (212) and leads to general genomic instability, including increased DNA strand breaks, loss of heterozygosity, and aneuploidy (17).

Pol κ's protein interactions.

Multiple protein-protein interactions likely regulate Pol κ function. Eukaryotic Pol κ interacts with the PCNA processivity clamp (human [81]), ubiquitin (mouse [72]), the 9-1-1 checkpoint clamp (S. pombe [104]), and Rev1 (mouse [70]). Many, if not all, of these interactions are important for Pol κ's function in vivo. For example, as noted below, mutations of Pol κ that disrupt its interaction with ubiquitin or with PCNA result in aberrant nuclear localization after DNA damage (72, 191). Additionally, the 9-1-1 complex, which is involved in signaling the DNA damage checkpoint via the clamp loader, localizes Pol κ to chromatin in replication-compromised S. pombe strains (104). Also, a mutant of the 9-1-1 clamp that perturbs DNA binding by Pol κ displays a reduction in point mutations.

Regulation of Pol κ.

Pol κ relocalizes from a diffuse nuclear pattern into foci upon DNA damage (19, 20, 191). Focus formation by Pol κ requires both its PCNA interaction motif and its UBMs (72, 191) (Fig. (Fig.3).3). Interestingly, Pol κ relocalizes in response to DNA damage differently from the other Y family members, forming fewer spontaneous and damage-induced foci (20, 191). The reports disagree, however, on whether DNA Pol κ forms foci during S phase (20, 191).

A key source of regulation of Pol κ may be at the level of transcription. Murine Pol κ transcript levels increase after treatment with 3-methyl methylcholanthrene, a polycyclic aromatic hydrocarbon similar to benzo[a]pyrene (194). Notably, a change in Pol κ's transcript levels may be connected to cancer development in some contexts, as Pol κ transcripts are downregulated in some human colorectal tumors (136). Conversely, Pol κ transcripts are upregulated in non-small-cell lung cancers (202a). This increase in Pol κ transcripts correlates with the increased loss of heterozygosity in these tumors (17).

Pol η (Rad30A/XP-V)

Like Pol κ, the rad30 gene was identified not on the basis of its contribution to mutagenesis but rather by its homology to the genes encoding the Rev1, UmuC, and DinB proteins. Its name reflects the slight sensitization of a rad30 mutant in S. cerevisiae to UV irradiation (165). Indeed, under most circumstances, rad30 mutants display very limited reduction in mutagenesis in yeast.

The Rad30/Pol η subfamily is found only in eukaryotes, where it is broadly conserved. Pol η is perhaps the most thoroughly characterized TLS polymerase, since in humans, loss of Pol η 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 (111, 132, 135, 157). Clinically, XPV is very similar to other forms of xeroderma pigmentosum, which result from mutations in any of six key nucleotide excision repair genes, but XPV cells are not defective in nucleotide excision repair (140, 223). This phenotype highlights the predominantly nonmutagenic role of Pol η, setting it apart from the more mutagenic functions of Pol ζ and Rev1.

Pol η's catalytic activity and role in mutagenesis.

The phenotypes of mutants with DNA Pol η deficiencies and the in vitro activity of Pol η indicate that its major role is the nonmutagenic bypass of UV-induced DNA lesions. In particular, Pol η 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, 63, 263). In vitro, Pol η has been shown to bypass CPDs with high accuracy and efficiency (100), and in vivo, it is thought to be responsible for restarting stalled replication forks and allowing continuous DNA synthesis past sites of UV damage (132). In the absence of Pol η, 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 (61, 141). Furthermore, Pol η-independent CPD bypass, which is thought to involve other TLS polymerases such as Pol ζ and/or Pol ι, is significantly more mutagenic, presumably accounting for the increased frequency of cancer in XPV patients (126, 132, 157, 252). In addition to the severely distorting CPDs, in vitro, Pol η is also able to bypass a broad range of other DNA lesions, such as 7,8-dihydro-8-oxoguanine (84), (+)-trans-anti-benzo[a]pyrene-N2-dG (277), acetylaminofluorene-adducted guanine (272), O6-methylguanine (79), thymine glycol (120), and adducts derived from cisplatin and oxaliplatin (247). Aside from CPDs, purified Pol η is 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 η after cisplatin exposure in XPV cells (5, 6, 33).

Interestingly, though Pol η plays a major role in accurately bypassing particular types of DNA lesions, such as CPDs and 7,8-dihydro-8-oxoguanine, it exhibits among the lowest fidelity of any DNA polymerase on undamaged DNA in vitro (11, 42, 118, 127, 158, 217). In spite of its mutagenic potential, depletion of Pol η in human cells by small interfering RNA actually increases mutation frequency (36, 92), and S. cerevisiae rad30 mutants do not display a major reduction in spontaneous or induced mutagenesis (165, 225). Similarly, Pol η knockout mouse embryonic fibroblasts show an increased UV-induced mutation frequency (28). Taken together, these results indicate that Pol η's normal function in vivo primarily reduces mutagenesis. Therefore, regulation of Pol η activity is thought to play a crucial role in modulating the mutagenic potential of Pol η in living cells.

Intriguingly, overexpression of Pol η in human cells does not increase mutagenesis and causes only a weak mutator effect in S. cerevisiae (114, 208), suggesting that Pol η is largely restricted from accessing undamaged DNA by additional regulatory mechanisms even when overexpressed. Pol η's potential for introducing mutations, though normally inhibited in somatic cells, is harnessed in a specific context. Pol η is the major mutator of A·T base pairs during the SHM step in antibody diversification in B lymphocytes (159, 209).

Pol η's protein-protein interactions.

The regulation of Pol η's catalytic activity is directed in part through protein-protein interactions. Pol η interacts with the eukaryotic processivity clamp, PCNA, through its C-terminal PCNA-binding motif (PIP box) (Fig. (Fig.3)3) (108), and the interaction between PCNA and Pol η plays an important role in Pol η function. This is at least partially attributable to the stimulatory effect of PCNA on Pol η's TLS activity in vitro (76, 78, 105). Interestingly, the interaction between PCNA and Pol η is inhibited by p21, a protein discussed above with respect to regulation of TLS (236).

Although ubiquitinated PCNA is not required for Pol η to access stalled replication forks in vitro (189), Pol η's interaction with PCNA can be enhanced by the monoubiquitination of PCNA. Mammalian Pol η foci colocalize with foci of monoubiquitinated PCNA in the nucleus (108), and accumulation of Pol η foci in response to DNA damage is dependent upon monoubiquitinated PCNA (213), although a small proportion of cells (5 to 10%) do have Pol η foci in rad18−/− or pol30(K164R) mutants, in which PCNA is not monoubiquitinated (256). A similar proportion of cells contain Pol η foci in the absence of DNA damage, consistent with a model in which PCNA monoubiquitination induces Pol η's response to exogenous DNA damage, above a low level of uninduced DNA association by Pol η.

The dependence of Pol η's damage-induced foci on monoubiquitinated PCNA is attributed to Pol η's interactions with PCNA and ubiquitin (213), which appear to give Pol η a competitive advantage over the replicative Pol δ for PCNA association after DNA damage (256, 273). Pol η's interaction with monoubiquitinated PCNA is mediated by both the PCNA interaction motif (PIP box) and its ubiquitin-binding zinc finger (UBZ) domain (205) (Fig. (Fig.3).3). Mutants disrupting the UBZ, in either S. cerevisiae or mammalian Pol η, fail to complement the UV sensitivity of Pol η-deficient cells (21, 205), although at lower UV doses, Acharya et al. (3) have demonstrated partial complementation of the UV-sensitive phenotype of the rad30Δ strain. Monoubiquitinated PCNA may also promote TLS by enhancing Pol η's catalytic activity, but in vitro results have so far been inconsistent (59, 82).

Another protein which participates in the regulation of Pol η is Rad18, an E3 ubiquitin ligase that mediates PCNA monoubiquitination. Mouse Pol η has been found to have a direct physical interaction with Rad18, independent of the presence of DNA damage, via C-terminal regions of both proteins (256). Furthermore, in human cells, Pol η copurifies as a complex with Rad18, Rad6, and Rev1; the complex is enriched in the chromatin fraction in response to UV radiation or S-phase arrest (273), consistent with the model that Rad18 is involved in recruitment of Pol η to stalled replication forks. Pol η foci colocalize with Rad18 foci (256), and the formation and damage-dependent accumulation of Pol η foci are largely dependent on Rad18 (256).

Pol η may also be regulated by ubiquitination through a covalent attachment of a monoubiquitin moiety (21, 203, 205), although the functional significance of this modification is not yet understood. Ubiquitination of Pol η is dependent on the UBZ domain of Pol η. Intriguingly, the monoubiquitination of Pol η is not dependent on the postreplicative repair proteins Rad6 and Rad18, nor is it responsive to DNA damage (205).

There is also a robust physical interaction between DNA Pol η and Rev1 in vertebrates and flies but only a weak interaction, if any, between Pol η and Rev1 in budding yeast (3, 70, 117, 243, 262). Thus, the functional interactions between TLS polymerases are complex and, to some extent, species dependent.

Regulation of Pol η.

Transcriptional regulation of Pol η was demonstrated early. In S. cerevisiae, the RAD30 transcript is induced three- to fourfold in response to UV radiation (165, 225). In mouse, however, expression of the XPV gene (encoding Pol η) is not induced by UV radiation; instead, it has been found to increase about fourfold during cell proliferation (264). The RAD30 gene in S. cerevisiae has been placed in the RAD6 epistasis group (165) but appears to function independently of both the error-free pathway defined by RAD5 (165) and the error-prone TLS pathway which includes REV1, REV3, and REV7 (165, 262).

Pol η forms foci spontaneously in a small percentage of untreated cells, suggesting that Pol η 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 methyl methanesulfonate (105) and in cells subjected to hydroxyurea-induced replication stress (5, 20, 41, 105, 106). These foci are thought to form at sites of DNA damage since they colocalize with PCNA (5, 105, 106) and with Rad18 foci (256). Although it is assumed that the nuclear Pol η foci represent sites of TLS, focus formation does not necessarily imply activity. For example, a mutant form of Rad18 that is unable to form foci nonetheless activates DNA damage tolerance pathways (182). Recent data even reveal that Pol η is transiently immobilized in foci (228) supporting a model of TLS polymerases transiently probing the chromatin. Additionally, accumulation of Pol η foci is stimulated by the physical interaction of Pol η's UBZ domain with monoubiquitinated PCNA (21, 108, 213, 256). Together with the fact that Pol η mutants progress more slowly through S phase after DNA damage (5, 237), these findings have led to a model in which Pol η rescues replication forks that have stalled at sites of DNA damage by allowing continuous DNA synthesis past the lesion(s).

Pol ι (Rad30B)

Pol ι is most closely related to Pol η at the sequence level but is divergent enough to have distinct biochemical properties and function. In contrast to the wealth of information about Pol η, the role of Pol ι is less well understood. Pol ι is present not only in higher eukaryotes as initially thought (166, 199, 242) but in organisms scattered throughout the Eukaryota, including some yeasts (L. S. Waters, unpublished observation). Because Pol ι is 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.

The role of Pol ι is complex and not well understood; Pol ι is reported both to contribute to mutagenesis in some situations and not to be involved in mutagenesis in others (35, 45, 265). More recent data support that Pol ι is involved in UV-induced mutagenesis in Burkitt's lymphoma and XPV cell lines (69, 252). As for Pol η, however, mice and humans lacking Pol ι tend to develop cancer at an increased rate, suggesting that Pol ι in the presence of other DNA repair and tolerance pathways is not a major contributor to mutagenesis in vivo (35, 45, 128, 198, 265). Consistent with the lack of strong data indicating a mutagenic role in vivo, loss of Pol ι seems to have little consequence in SHM, as the 129/J strain of mice that possess a nonsense mutation in the Pol ι gene preventing its expression have normal immunoglobulin hypermutation (164). A recent report investigating the biological significance of Pol ι in human cells has demonstrated that cell lines depleted for Pol ι exhibit enhanced sensitivity to oxidative damage (210). Moreover, Pol ι interacts with the base excision repair factor XRCC1 and is recruited to sites of oxidative DNA damage in living cells, suggesting a role for Pol ι in the repair of DNA damaged by oxidative stress (210).

Pol ι's catalytic activity.

Biochemically, Pol ι has been thought to be one of the least accurate DNA polymerases, especially opposite pyrimidines (163). Interestingly, though, Pol ι was recently shown to exhibit significantly different catalytic activities on both damaged and undamaged DNA when manganese was used as a cofactor rather than magnesium. Importantly, these altered catalytic properties with manganese included a dramatic increase in the ability of Pol ι to bypass a variety of DNA lesions (50). The structure of Pol ι uniquely allows for Hoogsteen base pairing which helps explain the varying catalytic efficiencies opposite different template nucleotides (178, 181). As with the polymerases discussed above, the catalytic activity of Pol ι is stimulated in the presence of PCNA, replication factor C, and replication protein A in vitro (75, 77, 250).

Regulation of Pol ι.

Limited information exists for the regulation of Pol ι's protein expression or activity. Possible regulatory mechanisms can be inferred from gene expression and protein localization studies. For example, like for other TLS polymerases, RAD30B and mouse Rad30b mRNAs are highly expressed in the testis, with the mouse Rad30b expression occurring mostly in the postmeiotic round spermatids (166). Additionally, Pol ι interacts physically with Pol η, and this interaction is required for its localization into DNA damage-induced foci (106). Similar to the case for the other TLS polymerases, the PCNA interaction motif and UBMs of Pol ι (Fig. (Fig.3)3) are also required for localization into foci after DNA damage, indicating that recruitment to stalled replication forks by monoubiquitinated PCNA mediates the function of Pol ι (21, 250). However, it should be noted that the biological relevance of the interaction with monoubiquitinated PCNA has not yet been tested for Pol ι.

Other Non-Y Family DNA Polymerases Capable of TLS

It is worth noting that there are other nonreplicative DNA polymerases that have various abilities to bypass DNA lesions and that synthesize DNA with a range of fidelities (reviewed in reference 172). The members of the X family of DNA polymerases (DNA Pols β, λ, and μ in eukaryotes), in particular, can insert nucleotides opposite to certain lesions (22, 172). After the Y family, the X family polymerases display the next lowest replication fidelity of the six major DNA polymerase families (119). 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 θ, 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 base excision repair and nonhomologous end joining by the X family polymerases. Accordingly, the term TLS polymerases generally refers to the Y family and DNA Pol ζ, which clearly have specialized roles involved primarily in lesion bypass (53, 123).


The numerous genetic and biochemical data regarding the posttranslational regulatory strategies detailed above have been integrated into two models for DNA lesion bypass by TLS polymerases that are not mutually exclusive (Fig. (Fig.4):4): (i) the polymerase-switching model (51, 53, 133, 134, 161, 163, 214) and (ii) the gap-filling model (131, 133, 163, 257). 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.

FIG. 4.
Two nonexclusive models for TLS: the polymerase-switching model (A) and the gap-filling model (B). See text for details.

Polymerase-Switching Model

In the polymerase-switching model (Fig. (Fig.4A),4A), TLS polymerases act at the replication fork to enable replication to continue, bypassing DNA lesions that halt the forward progression of the replicative DNA polymerase. Access of TLS polymerases to the DNA is governed by protein-protein interactions that mediate a polymerase handoff at the primer-template terminus from the replicative polymerase to one or more TLS DNA polymerases. A further switch restores the replicative DNA polymerase to the primer terminus, and accurate DNA synthesis resumes.

The polymerase-switching mechanism likely comes into play during times of active DNA replication. Upon DNA damage in eukaryotes, Rad18 is thought to bind to the ssDNA generated when a replication fork is blocked at a lesion, bringing the E2 ubiquitin ligase Rad6 with it to sites of DNA damage (13, 14). Once localized at or near the lesion, the Rad6/Rad18 heterodimer monoubiquitinates PCNA (89) (Fig. (Fig.4A,4A, step 2). Many of the TLS polymerases have a basal affinity for unmodified PCNA in vitro through their PIP domains (76, 81, 83, 153, 250) or the BRCT domain, in the case of Rev1 (71). Thus, it is not entirely clear if the monoubiquitination of PCNA serves primarily to signal the need for TLS, simply serves to strengthen the interaction between PCNA and Y family TLS polymerases via their UBM or UBZ motifs, and/or facilitates a handoff between the replicative polymerase and the TLS polymerases (reviewed in reference 134) (Fig. (Fig.4A,4A, step 3). The specific requirement for the UBM and UBZ domains of TLS polymerases for survival after DNA damage (89, 233) and for damage-induced relocalization of TLS polymerases into foci in vivo (see the references cited above and references therein), however, makes a tempting argument that the interaction between TLS polymerases and monoubiquitinated PCNA functions as a mechanism to stimulate a switch between a replicative polymerase and a TLS polymerase.

Once the TLS polymerase is recruited to the site of the lesion, an exchange is thought to occur between the replicative polymerase and the incoming TLS polymerase (Fig. (Fig.4A,4A, step 3). The details of this process, however, are not well characterized. For example, it is unclear if the replicative polymerase must exit the primer terminus before the TLS polymerase can be recruited or if the replicative polymerase somehow aids in this transaction (as has been suggested for the Pol32 subunit of the replicative Pol δ [91, 169]). In addition, it has recently been demonstrated in vitro that S. cerevisiae Pol η cannot gain access to the primer terminus while it is occupied by a moving replicative polymerase, suggesting that the stalling of the replicative polymerase is required to allow entry of Pol η onto the DNA (280).

Following the handoff between the replicative and TLS polymerase on the primer terminus, the subsequent nucleotide insertion across and extension past the lesion may require the concerted action of two or more TLS polymerases (Fig. (Fig.4A,4A, step 3 to 4). In eukaryotes, extension from a lesion is thought to be primarily mediated by Pol ζ, and also to some extent by DNA Pol κ, due to the distortion or mismatched base pair often produced by TLS (217, 218). Finally, once the TLS polymerase(s) is able to bypass the lesion and extend past the distorting mismatch, a further polymerase switch, perhaps facilitated by the deubiquitination of PCNA (92), reinstates the replicative DNA polymerase at the primer terminus. Accurate DNA synthesis can therefore resume and the replication fork again moves forward (Fig. (Fig.4A,4A, step 5). Following bypass by TLS, the lesion would again be a substrate for removal by DNA repair mechanisms. The described mechanism likely has more layers of complexity in vivo given that monoubiquitinated PCNA persists for hours after the removal of UV damage in human cells (188).

While PCNA plays a significant role in TLS-assisted DNA lesion bypass, it is likely not the only factor involved in the recruitment and polymerase exchange reaction. For example, eukaryotes also contain an alternative processivity clamp, termed the 9-1-1 complex, which is loaded onto regions of ssDNA by an alternative clamp loader, and functions in the DNA damage checkpoint response (24). The 9-1-1 clamp physically interacts with the Rev7 subunit of DNA Pol ζ in S. cerevisiae (229) and with Pol κ in S. pombe (104). Loss of the alternative clamp and clamp loader lowers the mutation frequencies in both organisms (104, 207, 229), and loss of the alternative clamp loader prevents Pol κ from being recruited to chromatin in S. pombe (104). Additionally, in a striking report, Fu et al. have recently shown that the Rad6/Rad18 heterodimer monoubiquitinates the Rad17 subunit of the 9-1-1 complex to induce the DNA damage response in S. cerevisiae (54). Thus, the interplay of the two eukaryotic clamps in the recruitment of TLS polymerases and other factors is just beginning to be elucidated. Finally, a clamp may not be required in every circumstance of lesion bypass because, as noted above, Rev1 may interact with the DNA on its own by utilizing its BRCT domain (88) and, via its last 100 amino acids, may target other TLS polymerases to the site of the DNA lesion (44a, 70, 176, 197, 243).

Gap-Filling Model

The second model, gap filling (Fig. (Fig.4B),4B), has been proposed to account for TLS-assisted DNA lesion bypass outside the context of the replication fork, when DNA damage may reside in single-stranded gaps. In this situation, the purpose of TLS is not to restart a stalled replication fork but rather to seal gaps that have resulted from the replication machinery repriming downstream of the blocking lesion, thereby leaving behind a ssDNA gap opposite the lesion. Repriming events leading to single-stranded gaps were first suggested to arise in E. coli by Rupp and Howard-Flanders (226) and more recently suggested to arise in S. cerevisiae by Lopes et al. (151), either by initiating a new Okazaki fragment on the lagging strand or by repriming on the leading strand (86). Alternatively, gaps containing lesions can be formed as a result of processing of closely spaced lesions on opposite DNA strands or by the processing of interstrand cross-links. It would be expected that this gap-filling mechanism would be utilized primarily outside of S phase, during G1 or G2/M phases of the cell cycle. It could, however, also occur during the later stages of S phase.

In the gap-filling model, once the cell identifies a need for lesion bypass, the TLS polymerase(s) is presumably directed to the ssDNA gap. This localization may involve many of the same factors used in polymerase switching, including PCNA, the 9-1-1 alternative clamp, and Rev1, which in S. cerevisiae is upregulated after the bulk of replication is complete during late S/G2/M (227, 257). In this situation, handoffs with the replicative polymerase are not expected to contribute significantly in the recruitment of the TLS polymerase(s) but might come into play after the lesion is bypassed if the remaining gap after the lesion is large. Indeed, in S. cerevisiae, ssDNA gaps of hundreds of nucleotides have been observed after UV irradiation (151). Following DNA lesion bypass and subsequent resealing of the nick with DNA ligase, the lesion would now be a substrate for removal by DNA repair pathways and could be removed before the next round of replication.

Choosing the TLS Polymerase for Lesion Bypass

One of the most perplexing questions regarding TLS-assisted lesion bypass is how the “correct” TLS polymerase is selected to bypass a particular DNA lesion out of the pool of multiple possible TLS enzymes in vivo. It has been suggested that transient association of several polymerases with the DNA lesion at the primer terminus may occur sequentially until the best-suited polymerase is able to perform lesion bypass (163). In such models, specificity of TLS for bypass of a particular lesion is imparted by a passive trial-and-error approach based on the inherent efficiency of TLS polymerases opposite their cognate lesion. However, the many levels of regulatory controls and protein-protein interactions discussed in this review likely constrain which DNA polymerases are able to access a given primer terminus in a particular biological context. Examples include local increases in polymerase concentration by DNA damage induction (as in the case of Pol κ [194]) or by cell cycle regulation (as in the case of Rev1 [227, 257]), the ubiquitination state of PCNA, the interaction of TLS polymerases with monoubiquitin through UBMs, the ubiquitination/deubiquitination of TLS polymerases themselves, undiscovered modifications of these proteins, or the interaction of Rev1 with the other TLS polymerases. Ultimately, for both models of TLS lesion bypass, the differential primer/template affinities, processivities, and bypass activities of translesion polymerases may be the primary mechanism by which a particular DNA polymerase gains access to the DNA (56, 93, 121, 162).


Although they may have high accuracy opposite certain lesions, the fact that TLS polymerases have poor fidelity on undamaged DNA suggests that they are tightly regulated in vivo to avoid rampant mutagenesis. As detailed above, when overexpressed or inappropriately regulated, certain TLS polymerases confer a hypermutator phenotype (16, 19, 67, 113, 192) and/or change replication fork progression (15, 56, 93, 138, 212). Why, then, has there not been a stronger correlation between improper regulation of TLS and cancer? Perhaps the multiple overlapping systems for regulation of TLS provide the answer. For example, in the case of TLS polymerase overexpression, it may be the relative availability of accessory proteins used to direct TLS polymerases to the sites of DNA lesions that prevents rampant mutagenesis. In contrast, loss of TLS polymerases may have a greater effect upon multicellular organisms than overexpression. Indeed, loss of Pol η results in the variant form of XPV, a condition which renders one highly sensitive to the effects of UV light and susceptible to cancer (111, 132, 135, 157). Additionally, loss of Pol ζ leads to early embryonic lethality in mice (18, 47, 248, 260). While mouse models have not detected profound phenotypes upon single deletions of the TLS Pols κ and ι, there may be considerable redundancy between the roles of TLS polymerases in multicellular organisms that mask the effects of the loss of a single TLS polymerase. More profound phenotypes may be detected only upon the simultaneous loss of multiple 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, i.e., 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 (38), D-loop extension during homologous recombination by Pol η (111, 167), 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 (44), and the complex control of TLS DNA polymerases discussed in this review 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 emergence of a variant that is better able to withstand the stress (49, 57). 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 (49, 57). Additionally, in higher eukaryotes, the mutagenic capacity of TLS polymerases has been harnessed for SHM, the generation of mutations in the variable regions of antibodies produced by B-cell lymphocytes (31). 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.


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, 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.


We thank members of the Walker lab for helpful discussions.

This work was supported by National Institute of Environmental Health Sciences (NIEHS) grant 5-R01-ES015818 to G.C.W., NIEHS grant P30 ES002109 to the MIT Center of Environmental Health Sciences, National Institute of Health Fellowship grant 5-F32-GM078966 to B.K.M., and an American Cancer Society Research Professorship to G.C.W.


1. Abdulovic, A. L., and S. Jinks-Robertson. 2006. The in vivo characterization of translesion synthesis across UV-induced lesions in Saccharomyces cerevisiae: insights into Pol zeta- and Pol eta-dependent frameshift mutagenesis. Genetics 1721487-1498. [PubMed]
2. Acharya, N., L. Haracska, R. E. Johnson, I. Unk, S. Prakash, and L. Prakash. 2005. Complex formation of yeast Rev1 and Rev7 proteins: a novel role for the polymerase-associated domain. Mol. Cell. Biol. 259734-9740. [PMC free article] [PubMed]
3. Acharya, N., L. Haracska, S. Prakash, and L. Prakash. 2007. Complex formation of yeast Rev1 with DNA polymerase η. Mol. Cell. Biol. 278401-8408. [PMC free article] [PubMed]
4. Acharya, N., R. E. Johnson, S. Prakash, and L. Prakash. 2006. Complex formation with Rev1 enhances the proficiency of Saccharomyces cerevisiae DNA polymerase zeta for mismatch extension and for extension opposite from DNA lesions. Mol. Cell. Biol. 269555-9563. [PMC free article] [PubMed]
5. Albertella, M. R., C. M. Green, A. R. Lehmann, and M. J. O'Connor. 2005. A role for polymerase eta in the cellular tolerance to cisplatin-induced damage. Cancer Res. 659799-9806. [PubMed]
6. Alt, A., K. Lammens, C. Chiocchini, A. Lammens, J. C. Pieck, D. Kuch, K. P. Hopfner, and T. Carell. 2007. Bypass of DNA lesions generated during anticancer treatment with cisplatin by DNA polymerase eta. Science 318967-970. [PubMed]
7. Andersen, P. L., F. Xu, and W. Xiao. 2008. Eukaryotic DNA damage tolerance and translesion synthesis through covalent modifications of PCNA. Cell Res. 18162-173. [PubMed]
8. Arana, M. E., M. Seki, R. D. Wood, I. B. Rogozin, and T. A. Kunkel. 2008. Low-fidelity DNA synthesis by human DNA polymerase theta. Nucleic Acids Res. 363847-3856. [PMC free article] [PubMed]
9. Aravind, L., and E. V. Koonin. 1998. The HORMA domain: a common structural denominator in mitotic checkpoints, chromosome synapsis and DNA repair. Trends Biochem. Sci. 23284-286. [PubMed]
10. Avkin, S., M. Goldsmith, S. Velasco-Miguel, N. Geacintov, E. C. Friedberg, and Z. Livneh. 2004. Quantitative analysis of translesion DNA synthesis across a benzo[a]pyrene-guanine adduct in mammalian cells: the role of DNA polymerase kappa. J. Biol. Chem. 27953298-53305. [PubMed]
11. Avkin, S., and Z. Livneh. 2002. Efficiency, specificity and DNA polymerase-dependence of translesion replication across the oxidative DNA lesion 8-oxoguanine in human cells. Mutat. Res. 51081-90. [PubMed]
12. Avkin, S., Z. Sevilya, L. Toube, N. Geacintov, S. G. Chaney, M. Oren, and Z. Livneh. 2006. p53 and p21 regulate error-prone DNA repair to yield a lower mutation load. Mol. Cell 22407-413. [PubMed]
13. Bailly, V., J. Lamb, P. Sung, S. Prakash, and L. Prakash. 1994. Specific complex formation between yeast RAD6 and RAD18 proteins: a potential mechanism for targeting RAD6 ubiquitin-conjugating activity to DNA damage sites. Genes Dev. 8811-820. [PubMed]
14. Bailly, V., S. Lauder, S. Prakash, and L. Prakash. 1997. Yeast DNA repair proteins Rad6 and Rad18 form a heterodimer that has ubiquitin conjugating, DNA binding, and ATP hydrolytic activities. J. Biol. Chem. 27223360-23365. [PubMed]
15. Baker, T. A., and S. P. Bell. 1998. Polymerases and the replisome: machines within machines. Cell 92295-305. [PubMed]
16. Bavoux, C., J. S. Hoffmann, and C. Cazaux. 2005. Adaptation to DNA damage and stimulation of genetic instability: the double-edged sword mammalian DNA polymerase kappa. Biochimie 87637-646. [PubMed]
17. Bavoux, C., A. M. Leopoldino, V. Bergoglio, O. W. J., T. Ogi, A. Bieth, J. G. Judde, S. D. Pena, M. F. Poupon, T. Helleday, M. Tagawa, C. Machado, J. S. Hoffmann, and C. Cazaux. 2005. Up-regulation of the error-prone DNA polymerase κ promotes pleiotropic genetic alterations and tumorigenesis. Cancer Res. 65325-330. [PubMed]
18. Bemark, M., A. A. Khamlichi, S. L. Davies, and M. S. Neuberger. 2000. Disruption of mouse polymerase zeta (Rev3) leads to embryonic lethality and impairs blastocyst development in vitro. Curr. Biol. 101213-1216. [PubMed]
19. Bergoglio, V., C. Bavoux, V. Verbiest, J. S. Hoffmann, and C. Cazaux. 2002. Localisation of human DNA polymerase kappa to replication foci. J. Cell Sci. 1154413-4418. [PubMed]
20. Bi, X., D. M. Slater, H. Ohmori, and C. Vaziri. 2005. DNA polymerase kappa is specifically required for recovery from the benzo[a]pyrene-dihydrodiol epoxide (BPDE)-induced S-phase checkpoint. J. Biol. Chem. 28022343-22355. [PubMed]
21. Bienko, M., C. M. Green, N. Crosetto, F. Rudolf, G. Zapart, B. Coull, P. Kannouche, G. Wider, M. Peter, A. R. Lehmann, K. Hofmann, and I. Dikic. 2005. Ubiquitin-binding domains in Y-family polymerases regulate translesion synthesis. Science 3101821-1824. [PubMed]
22. Blanca, G., G. Villani, I. Shevelev, K. Ramadan, S. Spadari, U. Hubscher, and G. Maga. 2004. Human DNA polymerases lambda and beta show different efficiencies of translesion DNA synthesis past abasic sites and alternative mechanisms for frameshift generation. Biochemistry 4311605-11615. [PubMed]
23. Boudsocq, F., R. J. Kokoska, B. S. Plosky, A. Vaisman, H. Ling, T. A. Kunkel, W. Yang, and R. Woodgate. 2004. Investigating the role of the little finger domain of Y-family DNA polymerases in low fidelity synthesis and translesion replication. J. Biol. Chem. 27932932-32940. [PubMed]
24. Branzei, D., and M. Foiani. 2008. Regulation of DNA repair throughout the cell cycle. Nat. Rev. Mol. Cell Biol. 9297-308. [PubMed]
25. Brondello, J. M., M. J. Pillaire, C. Rodriguez, P. A. Gourraud, J. Selves, C. Cazaux, and J. Piette. 2008. Novel evidences for a tumor suppressor role of Rev3, the catalytic subunit of Pol zeta. Oncogene 276093-6101. [PubMed]
26. Burgers, P. M. 1991. Saccharomyces cerevisiae replication factor C. II. Formation and activity of complexes with the proliferating cell nuclear antigen and with DNA polymerases delta and epsilon. J. Biol. Chem. 26622698-22706. [PubMed]
27. Burns, N., B. Grimwade, P. B. Ross-Macdonald, E. Y. Choi, K. Finberg, G. S. Roeder, and M. Snyder. 1994. Large-scale analysis of gene expression, protein localization, and gene disruption in Saccharomyces cerevisiae. Genes Dev. 81087-1105. [PubMed]
28. Busuttil, R. A., Q. Lin, P. J. Stambrook, R. Kucherlapati, and J. Vijg. 2008. Mutation frequencies and spectra in DNA polymerase eta-deficient mice. Cancer Res. 682081-2084. [PubMed]
29. Callebaut, I., and J. P. Mornon. 1997. From BRCA1 to RAP1: a widespread BRCT module closely associated with DNA repair. FEBS Lett. 40025-30. [PubMed]
30. Carlson, K. D., R. E. Johnson, L. Prakash, S. Prakash, and M. T. Washington. 2006. Human DNA polymerase kappa forms nonproductive complexes with matched primer termini but not with mismatched primer termini. Proc. Natl. Acad. Sci. USA 10315776-15781. [PubMed]
31. Casali, P., Z. Pal, Z. Xu, and H. Zan. 2006. DNA repair in antibody somatic hypermutation. Trends Immunol. 27313-321. [PubMed]
32. Chen, J., and G. Fang. 2001. MAD2B is an inhibitor of the anaphase-promoting complex. Genes Dev. 151765-1770. [PubMed]
33. Chen, Y. W., J. E. Cleaver, F. Hanaoka, C. F. Chang, and K. M. Chou. 2006. A novel role of DNA polymerase eta in modulating cellular sensitivity to chemotherapeutic agents. Mol. Cancer Res. 4257-265. [PubMed]
34. Cheung, H. W., A. C. Chun, Q. Wang, W. Deng, L. Hu, X. Y. Guan, J. M. Nicholls, M. T. Ling, Y. Chuan Wong, S. W. Tsao, D. Y. Jin, and X. Wang. 2006. Inactivation of human MAD2B in nasopharyngeal carcinoma cells leads to chemosensitization to DNA-damaging agents. Cancer Res. 664357-4367. [PubMed]
35. Choi, J. H., A. Besaratinia, D. H. Lee, C. S. Lee, and G. P. Pfeifer. 2006. The role of DNA polymerase iota in UV mutational spectra. Mutat. Res. 59958-65. [PubMed]
36. Choi, J. H., and G. P. Pfeifer. 2005. The role of DNA polymerase eta in UV mutational spectra. DNA Repair (Amsterdam) 4211-220. [PubMed]
37. Chu, S., J. DeRisi, M. Eisen, J. Mulholland, D. Botstein, P. O. Brown, and I. Herskowitz. 1998. The transcriptional program of sporulation in budding yeast. Science 282699-705. [PubMed]
38. Collins, N. S., S. Bhattacharyya, and R. S. Lahue. 2007. Rev1 enhances CAG. CTG repeat stability in Saccharomyces cerevisiae. DNA Repair (Amsterdam) 638-44. [PubMed]
39. Reference deleted.
40. Reference deleted.
41. de Feraudy, S., C. L. Limoli, E. Giedzinski, D. Karentz, T. M. Marti, L. Feeney, and J. E. Cleaver. 2007. Pol eta is required for DNA replication during nucleotide deprivation by hydroxyurea. Oncogene 265713-5721. [PubMed]
42. de Padula, M., G. Slezak, P. Auffret van Der Kemp, and S. Boiteux. 2004. The post-replication repair RAD18 and RAD6 genes are involved in the prevention of spontaneous mutations caused by 7,8-dihydro-8-oxoguanine in Saccharomyces cerevisiae. Nucleic Acids Res. 325003-5010. [PMC free article] [PubMed]
43. Dixon, M. J., and R. S. Lahue. 2002. Examining the potential role of DNA polymerases eta and zeta in triplet repeat instability in yeast. DNA Repair (Amsterdam) 1763-770. [PubMed]
44. Drake, J. W. 1991. A constant rate of spontaneous mutation in DNA-based microbes. Proc. Natl. Acad. Sci. USA 887160-7164. [PubMed]
44a. D'Souza, S., and G. C. Walker. 2006. Novel role for the C terminus of Saccharomyces cerevisiae Rev1 in mediating protein-protein interactions. Mol. Cell. Biol. 268173-8182. [PMC free article] [PubMed]
44b. D'Souza, S., L. S. Waters, and G. C. Walker. 2008. Novel conserved motifs in Rev1 C-terminus are required for mutagenic DNA damage tolerance. DNA Repair (Amsterdam) 71455-1470. [PMC free article] [PubMed]
45. Dumstorf, C. A., A. B. Clark, Q. Lin, G. E. Kissling, T. Yuan, R. Kucherlapati, W. G. McGregor, and T. A. Kunkel. 2006. Participation of mouse DNA polymerase iota in strand-biased mutagenic bypass of UV photoproducts and suppression of skin cancer. Proc. Natl. Acad. Sci. USA 10318083-18088. [PubMed]
46. Edmunds, C. E., L. J. Simpson, and J. E. Sale. 2008. PCNA ubiquitination and REV1 define temporally distinct mechanisms for controlling translesion synthesis in the avian cell line DT40. Mol. Cell 30519-529. [PubMed]
47. Esposito, G., I. Godindagger, U. Klein, M. L. Yaspo, A. Cumano, and K. Rajewsky. 2000. Disruption of the Rev3l-encoded catalytic subunit of polymerase zeta in mice results in early embryonic lethality. Curr. Biol. 101221-1224. [PubMed]
48. Fiala, K. A., and Z. Suo. 2004. Mechanism of DNA polymerization catalyzed by Sulfolobus solfataricus P2 DNA polymerase IV. Biochemistry 432116-2125. [PubMed]
49. Foster, P. L. 2007. Stress-induced mutagenesis in bacteria. Crit. Rev. Biochem. Mol. Biol. 42373-397. [PMC free article] [PubMed]
50. Frank, E. G., and R. Woodgate. 2007. Increased catalytic activity and altered fidelity of human DNA polymerase iota in the presence of manganese. J. Biol. Chem. 28224689-24696. [PubMed]
51. Friedberg, E. C., A. R. Lehmann, and R. P. Fuchs. 2005. Trading places: how do DNA polymerases switch during translesion DNA synthesis? Mol. Cell 18499-505. [PubMed]
52. Friedberg, E. C., R. Wagner, and M. Radman. 2002. Specialized DNA polymerases, cellular survival, and the genesis of mutations. Science 2961627-1630. [PubMed]
53. Friedberg, E. C., G. C. Walker, W. Siede, R. D. Wood, R. A. Schultz, and T. Ellenberger. 2005. DNA repair and mutagenesis, 2nd ed. ASM Press, Washington, DC.
54. Fu, Y., Y. Zhu, K. Zhang, M. Yeung, D. Durocher, and W. Xiao. 2008. Rad6-Rad18 mediates a eukaryotic SOS response by ubiquitinating the 9-1-1 checkpoint clamp. Cell 133601-611. [PubMed]
55. Fuchs, R. P., S. Fujii, and J. Wagner. 2004. Properties and functions of Escherichia coli: Pol IV and Pol V. Adv. Protein Chem. 69229-264. [PubMed]
56. Fujii, S., and R. P. Fuchs. 2004. Defining the position of the switches between replicative and bypass DNA polymerases. EMBO J. 234342-4352. [PubMed]
57. Galhardo, R. S., P. J. Hastings, and S. M. Rosenberg. 2007. Mutation as a stress response and the regulation of evolvability. Crit. Rev. Biochem. Mol. Biol. 42399-435. [PMC free article] [PubMed]
58. Gan, G. N., J. P. Wittschieben, B. O. Wittschieben, and R. D. Wood. 2008. DNA polymerase zeta (pol zeta) in higher eukaryotes. Cell Res. 18174-183. [PubMed]
59. Garg, P., and P. M. Burgers. 2005. Ubiquitinated proliferating cell nuclear antigen activates translesion DNA polymerases eta and REV1. Proc. Natl. Acad. Sci. USA 10218361-18366. [PubMed]
60. Garg, P., C. M. Stith, J. Majka, and P. M. Burgers. 2005. Proliferating cell nuclear antigen promotes translesion synthesis by DNA polymerase zeta. J. Biol. Chem. 28023446-23450. [PubMed]
61. Garinis, G. A., J. R. Mitchell, M. J. Moorhouse, K. Hanada, H. de Waard, D. Vandeputte, J. Jans, K. Brand, M. Smid, P. J. van der Spek, J. H. Hoeijmakers, R. Kanaar, and G. T. van der Horst. 2005. Transcriptome analysis reveals cyclobutane pyrimidine dimers as a major source of UV-induced DNA breaks. EMBO J. 243952-3962. [PubMed]
62. German, J., S. Schonberg, S. Caskie, D. Warburton, C. Falk, and J. H. Ray. 1987. A test for Fanconi's anemia. Blood 691637-1641. [PubMed]
63. Gibbs, P. E., J. McDonald, R. Woodgate, and C. W. Lawrence. 2005. The relative roles in vivo of Saccharomyces cerevisiae Pol eta, Pol zeta, Rev1 protein and Pol32 in the bypass and mutation induction of an abasic site, T-T (6-4) photoadduct and T-T cis-syn cyclobutane dimer. Genetics 169575-582. [PubMed]
64. Gibbs, P. E., W. G. McGregor, V. M. Maher, P. Nisson, and C. W. Lawrence. 1998. A human homolog of the Saccharomyces cerevisiae REV3 gene, which encodes the catalytic subunit of DNA polymerase zeta. Proc. Natl. Acad. Sci. USA 956876-6880. [PubMed]
65. Gibbs, P. E., X. D. Wang, Z. Li, T. P. McManus, W. G. McGregor, C. W. Lawrence, and V. M. Maher. 2000. The function of the human homolog of Saccharomyces cerevisiae REV1 is required for mutagenesis induced by UV light. Proc. Natl. Acad. Sci. USA 974186-4191. [PubMed]
66. Glover, J. N., R. S. Williams, and M. S. Lee. 2004. Interactions between BRCT repeats and phosphoproteins: tangled up in two. Trends Biochem. Sci. 29579-585. [PubMed]
67. Godoy, V. G., D. F. Jarosz, S. M. Simon, A. Abyzov, V. Ilyin, and G. C. Walker. 2007. UmuD and RecA directly modulate the mutagenic potential of the Y family DNA polymerase DinB. Mol. Cell 281058-1070. [PMC free article] [PubMed]
68. Goodman, M. F. 2002. Error-prone repair DNA polymerases in prokaryotes and eukaryotes. Annu. Rev. Biochem. 7117-50. [PubMed]
69. Gueranger, Q., A. Stary, S. Aoufouchi, A. Faili, A. Sarasin, C. A. Reynaud, and J. C. Weill. 2008. Role of DNA polymerases eta, iota and zeta in UV resistance and UV-induced mutagenesis in a human cell line. DNA Repair (Amsterdam) 71551-1562. [PubMed]
70. Guo, C., P. L. Fischhaber, M. J. Luk-Paszyc, Y. Masuda, J. Zhou, K. Kamiya, C. Kisker, and E. C. Friedberg. 2003. Mouse Rev1 protein interacts with multiple DNA polymerases involved in translesion DNA synthesis. EMBO J. 226621-6630. [PubMed]
71. Guo, C., E. Sonoda, T. S. Tang, J. L. Parker, A. B. Bielen, S. Takeda, H. D. Ulrich, and E. C. Friedberg. 2006. REV1 protein interacts with PCNA: significance of the REV1 BRCT domain in vitro and in vivo. Mol. Cell 23265-271. [PubMed]
72. Guo, C., T. S. Tang, M. Bienko, I. Dikic, and E. C. Friedberg. 2008. Requirements for the interaction of mouse Polkappa with ubiquitin and its biological significance. J. Biol. Chem. 2834658-4664. [PubMed]
73. Guo, C., T. S. Tang, M. Bienko, J. L. Parker, A. B. Bielen, E. Sonoda, S. Takeda, H. D. Ulrich, I. Dikic, and E. C. Friedberg. 2006. Ubiquitin-binding motifs in REV1 protein are required for its role in the tolerance of DNA damage. Mol. Cell. Biol. 268892-8900. [PMC free article] [PubMed]
74. Guo, D., Z. Xie, H. Shen, B. Zhao, and Z. Wang. 2004. Translesion synthesis of acetylaminofluorene-dG adducts by DNA polymerase zeta is stimulated by yeast Rev1 protein. Nucleic Acids Res. 321122-1130. [PMC free article] [PubMed]
75. Haracska, L., N. Acharya, I. Unk, R. E. Johnson, J. Hurwitz, L. Prakash, and S. Prakash. 2005. A single domain in human DNA polymerase iota mediates interaction with PCNA: implications for translesion DNA synthesis. Mol. Cell. Biol. 251183-1190. [PMC free article] [PubMed]
76. Haracska, L., R. E. Johnson, I. Unk, B. Phillips, J. Hurwitz, L. Prakash, and S. Prakash. 2001. Physical and functional interactions of human DNA polymerase eta with PCNA. Mol. Cell. Biol. 217199-7206. [PMC free article] [PubMed]
77. Haracska, L., R. E. Johnson, I. Unk, B. B. Phillips, J. Hurwitz, L. Prakash, and S. Prakash. 2001. Targeting of human DNA polymerase iota to the replication machinery via interaction with PCNA. Proc. Natl. Acad. Sci. USA 9814256-14261. [PubMed]
78. Haracska, L., C. M. Kondratick, I. Unk, S. Prakash, and L. Prakash. 2001. Interaction with PCNA is essential for yeast DNA polymerase eta function. Mol. Cell 8407-415. [PubMed]
79. Haracska, L., S. Prakash, and L. Prakash. 2000. Replication past O(6)-methylguanine by yeast and human DNA polymerase eta. Mol. Cell. Biol. 208001-8007. [PMC free article] [PubMed]
80. Haracska, L., I. Unk, R. E. Johnson, E. Johansson, P. M. Burgers, S. Prakash, and L. Prakash. 2001. Roles of yeast DNA polymerases delta and zeta and of Rev1 in the bypass of abasic sites. Genes Dev. 15945-954. [PubMed]
81. Haracska, L., I. Unk, R. E. Johnson, B. B. Phillips, J. Hurwitz, L. Prakash, and S. Prakash. 2002. Stimulation of DNA synthesis activity of human DNA polymerase kappa by PCNA. Mol. Cell. Biol. 22784-791. [PMC free article] [PubMed]
82. Haracska, L., I. Unk, L. Prakash, and S. Prakash. 2006. Ubiquitylation of yeast proliferating cell nuclear antigen and its implications for translesion DNA synthesis. Proc. Natl. Acad. Sci. USA 1036477-6482. [PubMed]
83. Haracska, L., M. T. Washington, S. Prakash, and L. Prakash. 2001. Inefficient bypass of an abasic site by DNA polymerase eta. J. Biol. Chem. 2766861-6866. [PubMed]
84. Haracska, L., S. L. Yu, R. E. Johnson, L. Prakash, and S. Prakash. 2000. Efficient and accurate replication in the presence of 7,8-dihydro-8-oxoguanine by DNA polymerase eta. Nat. Genet. 25458-461. [PubMed]
85. He, X., F. Ye, J. Zhang, Q. Cheng, J. Shen, and H. Chen. 2008. REV1 genetic variants associated with the risk of cervical carcinoma. Eur. J. Epidemiol. 23403-409. [PubMed]
86. Heller, R. C., and K. J. Marians. 2006. Replication fork reactivation downstream of a blocked nascent leading strand. Nature 439557-562. [PubMed]
87. Hinz, J. M., P. B. Nham, E. P. Salazar, and L. H. Thompson. 2006. The Fanconi anemia pathway limits the severity of mutagenesis. DNA Repair (Amsterdam) 5875-884. [PubMed]
88. Hirano, Y., and K. Sugimoto. 2006. ATR homolog Mec1 controls association of DNA polymerase zeta-Rev1 complex with regions near a double-strand break. Curr. Biol. 16586-590. [PubMed]
89. Hoege, C., B. Pfander, G. L. Moldovan, G. Pyrowolakis, and S. Jentsch. 2002. RAD6-dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature 419135-141. [PubMed]
90. Holbeck, S. L., and J. N. Strathern. 1997. A role for REV3 in mutagenesis during double-strand break repair in Saccharomyces cerevisiae. Genetics 1471017-1024. [PubMed]
91. Huang, M. E., A. G. Rio, M. D. Galibert, and F. Galibert. 2002. Pol32, a subunit of Saccharomyces cerevisiae DNA polymerase delta, suppresses genomic deletions and is involved in the mutagenic bypass pathway. Genetics 1601409-1422. [PubMed]
92. Huang, T. T., S. M. Nijman, K. D. Mirchandani, P. J. Galardy, M. A. Cohn, W. Haas, S. P. Gygi, H. L. Ploegh, R. Bernards, and A. D. D'Andrea. 2006. Regulation of monoubiquitinated PCNA by DUB autocleavage. Nat. Cell Biol. 8339-347. [PubMed]
93. Indiani, C., P. McInerney, R. Georgescu, M. F. Goodman, and M. O'Donnell. 2005. A sliding-clamp toolbelt binds high- and low-fidelity DNA polymerases simultaneously. Mol. Cell 19805-815. [PubMed]
94. Iwai, H., M. Kim, Y. Yoshikawa, H. Ashida, M. Ogawa, Y. Fujita, D. Muller, T. Kirikae, P. K. Jackson, S. Kotani, and C. Sasakawa. 2007. A bacterial effector targets Mad2L2, an APC inhibitor, to modulate host cell cycling. Cell 130611-623. [PubMed]
95. Reference deleted.
96. Jansen, J. G., A. Tsaalbi-Shtylik, P. Langerak, F. Calleja, C. M. Meijers, H. Jacobs, and N. de Wind. 2005. The BRCT domain of mammalian Rev1 is involved in regulating DNA translesion synthesis. Nucleic Acids Res. 33356-365. [PMC free article] [PubMed]
97. Jarosz, D. F., V. G. Godoy, J. C. Delaney, J. M. Essigmann, and G. C. Walker. 2006. A single amino acid governs enhanced activity of DinB DNA polymerases on damaged templates. Nature 439225-228. [PubMed]
98. Jarosz, D. F., V. G. Godoy, and G. C. Walker. 2007. Proficient and accurate bypass of persistent DNA lesions by DinB DNA polymerases. Cell Cycle 6817-822. [PubMed]
99. Johnson, R. E., C. M. Kondratick, S. Prakash, and L. Prakash. 1999. hRAD30 mutations in the variant form of xeroderma pigmentosum. Science 285263-265. [PubMed]
100. Johnson, R. E., S. Prakash, and L. Prakash. 1999. Efficient bypass of a thymine-thymine dimer by yeast DNA polymerase, Poleta. Science 2831001-1004. [PubMed]
101. Johnson, R. E., S. Prakash, and L. Prakash. 2000. The human DINB1 gene encodes the DNA polymerase Poltheta. Proc. Natl. Acad. Sci. USA 973838-3843. [PubMed]
102. Johnson, R. E., J. Trincao, A. K. Aggarwal, S. Prakash, and L. Prakash. 2003. Deoxynucleotide triphosphate binding mode conserved in Y family DNA polymerases. Mol. Cell. Biol. 233008-3012. [PMC free article] [PubMed]
103. Johnson, R. E., S. L. Yu, S. Prakash, and L. Prakash. 2003. Yeast DNA polymerase zeta (ζ) is essential for error-free replication past thymine glycol. Genes Dev. 1777-87. [PubMed]
104. Kai, M., and T. S. Wang. 2003. Checkpoint activation regulates mutagenic translesion synthesis. Genes Dev. 1764-76. [PubMed]
105. Kannouche, P., B. C. Broughton, M. Volker, F. Hanaoka, L. H. Mullenders, and A. R. Lehmann. 2001. Domain structure, localization, and function of DNA polymerase eta, defective in xeroderma pigmentosum variant cells. Genes Dev. 15158-172. [PubMed]
106. Kannouche, P., A. R. Fernandez de Henestrosa, B. Coull, A. E. Vidal, C. Gray, D. Zicha, R. Woodgate, and A. R. Lehmann. 2003. Localization of DNA polymerases eta and iota to the replication machinery is tightly co-ordinated in human cells. EMBO J. 221223-1233. [PubMed]
107. Kannouche, P. L., and A. R. Lehmann. 2004. Ubiquitination of PCNA and the polymerase switch in human cells. Cell Cycle 31011-1013. [PubMed]
108. Kannouche, P. L., J. Wing, and A. R. Lehmann. 2004. Interaction of human DNA polymerase eta with monoubiquitinated PCNA: a possible mechanism for the polymerase switch in response to DNA damage. Mol. Cell 14491-500. [PubMed]
109. Kato, T., and Y. Shinoura. 1977. Isolation and characterization of mutants of Escherichia coli deficient in induction of mutations by ultraviolet light. Mol. Gen. Genet. 156121-131. [PubMed]
110. Kaur, M., M. Pop, D. Shi, C. Brignone, and S. R. Grossman. 2007. hHR23B is required for genotoxic-specific activation of p53 and apoptosis. Oncogene 261231-1237. [PMC free article] [PubMed]
111. Kawamoto, T., K. Araki, E. Sonoda, Y. M. Yamashita, K. Harada, K. Kikuchi, C. Masutani, F. Hanaoka, K. Nozaki, N. Hashimoto, and S. Takeda. 2005. Dual roles for DNA polymerase eta in homologous DNA recombination and translesion DNA synthesis. Mol. Cell 20793-799. [PubMed]
112. Kenyon, C. J., and G. C. Walker. 1980. DNA-damaging agents stimulate gene expression at specific loci in Escherichia coli. Proc. Natl. Acad. Sci. USA 772819-2823. [PubMed]
113. Kim, S. R., G. Maenhaut-Michel, M. Yamada, Y. Yamamoto, K. Matsui, T. Sofuni, T. Nohmi, and H. Ohmori. 1997. Multiple pathways for SOS-induced mutagenesis in Escherichia coli: an overexpression of dinB/dinP results in strongly enhancing mutagenesis in the absence of any exogenous treatment to damage DNA. Proc. Natl. Acad. Sci. USA 9413792-13797. [PubMed]
114. King, N. M., N. Nikolaishvili-Feinberg, M. F. Bryant, D. D. Luche, T. P. Heffernan, D. A. Simpson, F. Hanaoka, W. K. Kaufmann, and M. Cordeiro-Stone. 2005. Overproduction of DNA polymerase eta does not raise the spontaneous mutation rate in diploid human fibroblasts. DNA Repair (Amsterdam) 4714-724. [PubMed]
115. Kobayashi, M., F. Figaroa, N. Meeuwenoord, L. E. Jansen, and G. Siegal. 2006. Characterization of the DNA binding and structural properties of the BRCT region of human replication factor C p140 subunit. J. Biol. Chem. 2814308-4317. [PubMed]
116. Kobayashi, S., M. R. Valentine, P. Pham, M. O'Donnell, and M. F. Goodman. 2002. Fidelity of Escherichia coli DNA polymerase IV. Preferential generation of small deletion mutations by dNTP-stabilized misalignment. J. Biol. Chem. 27734198-34207. [PubMed]
117. Kosarek, J. N., R. V. Woodruff, A. Rivera-Begeman, C. Guo, S. D'Souza, E. V. Koonin, G. C. Walker, and E. C. Friedberg. 2008. Comparative analysis of in vivo interactions between Rev1 protein and other Y-family DNA polymerases in animals and yeasts. DNA Repair (Amsterdam) 7439-451. [PMC free article] [PubMed]
118. Kozmin, S. G., Y. I. Pavlov, T. A. Kunkel, and E. Sage. 2003. Roles of Saccharomyces cerevisiae DNA polymerases Poleta and Polzeta in response to irradiation by simulated sunlight. Nucleic Acids Res. 314541-4552. [PMC free article] [PubMed]
119. Kunkel, T. A. 2004. DNA replication fidelity. J. Biol. Chem. 27916895-16898. [PubMed]
120. Kusumoto, R., C. Masutani, S. Iwai, and F. Hanaoka. 2002. Translesion synthesis by human DNA polymerase eta across thymine glycol lesions. Biochemistry 416090-6099. [PubMed]
121. Kusumoto, R., C. Masutani, S. Shimmyo, S. Iwai, and F. Hanaoka. 2004. DNA binding properties of human DNA polymerase eta: implications for fidelity and polymerase switching of translesion synthesis. Genes Cells 91139-1150. [PubMed]
122. Larimer, F. W., J. R. Perry, and A. A. Hardigree. 1989. The REV1 gene of Saccharomyces cerevisiae: isolation, sequence, and functional analysis. J. Bacteriol. 171230-237. [PMC free article] [PubMed]
123. Lawrence, C. W. 2004. Cellular functions of DNA polymerase zeta and Rev1 protein. Adv. Protein Chem. 69167-203. [PubMed]
124. Lawrence, C. W. 2002. Cellular roles of DNA polymerase zeta and Rev1 protein. DNA Repair (Amsterdam) 1425-435. [PubMed]
125. Lawrence, C. W., G. Das, and R. B. Christensen. 1985. REV7, a new gene concerned with UV mutagenesis in yeast. Mol. Gen. Genet. 20080-85. [PubMed]
126. Lawrence, C. W., and D. C. Hinkle. 1996. DNA polymerase zeta and the control of DNA damage induced mutagenesis in eukaryotes. Cancer Surv. 2821-31. [PubMed]
127. Lee, D. H., and G. P. Pfeifer. 2008. Translesion synthesis of 7,8-dihydro-8-oxo-2′-deoxyguanosine by DNA polymerase eta in vivo. Mutat. Res. 64119-26. [PMC free article] [PubMed]
128. Lee, G. H., and H. Matsushita. 2005. Genetic linkage between Pol iota deficiency and increased susceptibility to lung tumors in mice. Cancer Sci. 96256-259. [PubMed]
129. Lee, H., S. X. Zeng, and H. Lu. 2006. UV induces p21 rapid turnover independently of ubiquitin and Skp2. J. Biol. Chem. 28126876-26883. [PubMed]
130. Lee, J. Y., S. J. Yu, Y. G. Park, J. Kim, and J. Sohn. 2007. Glycogen synthase kinase 3β phosphorylates p21WAF1/CIP1 for proteasomal degradation after UV irradiation. Mol. Cell. Biol. 273187-3198. [PMC free article] [PubMed]
131. Lehmann, A. R. 2006. New functions for Y family polymerases. Mol. Cell 24493-495. [PubMed]
132. Lehmann, A. R. 2005. Replication of damaged DNA by translesion synthesis in human cells. FEBS Lett. 579873-876. [PubMed]
133. Lehmann, A. R., and R. P. Fuchs. 2006. Gaps and forks in DNA replication: rediscovering old models. DNA Repair (Amsterdam) 51495-1498. [PubMed]
134. Lehmann, A. R., A. Niimi, T. Ogi, S. Brown, S. Sabbioneda, J. F. Wing, P. L. Kannouche, and C. M. Green. 2007. Translesion synthesis: Y-family polymerases and the polymerase switch. DNA Repair (Amsterdam) 6891-899. [PubMed]
135. Leibeling, D., P. Laspe, and S. Emmert. 2006. Nucleotide excision repair and cancer. J. Mol. Histol. 37225-238. [PubMed]
136. Lemee, F., C. Bavoux, M. J. Pillaire, A. Bieth, C. R. Machado, S. D. Pena, R. Guimbaud, J. Selves, J. S. Hoffmann, and C. Cazaux. 2007. Characterization of promoter regulatory elements involved in downexpression of the DNA polymerase kappa in colorectal cancer. Oncogene 263387-3394. [PubMed]
137. Lemontt, J. F. 1971. Mutants of yeast defective in mutation induced by ultraviolet light. Genetics 6821-33. [PubMed]
138. Lenne-Samuel, N., J. Wagner, H. Etienne, and R. P. Fuchs. 2002. The processivity factor beta controls DNA polymerase IV traffic during spontaneous mutagenesis and translesion synthesis in vivo. EMBO Rep. 345-49. [PubMed]
139. Li, L., Y. Shi, H. Wu, B. Wan, P. Li, L. Zhou, H. Shi, and K. Huo. 2007. Hepatocellular carcinoma-associated gene 2 interacts with MAD2L2. Mol. Cell. Biochem. 304297-304. [PubMed]
140. Lichon, V., and A. Khachemoune. 2007. Xeroderma pigmentosum: beyond skin cancer. J. Drugs Dermatol. 6281-288. [PubMed]
141. Limoli, C. L., E. Giedzinski, W. F. Morgan, and J. E. Cleaver. 2000. Inaugural article: polymerase eta deficiency in the xeroderma pigmentosum variant uncovers an overlap between the S phase checkpoint and double-strand break repair. Proc. Natl. Acad. Sci. USA 977939-7946. [PubMed]
142. Lin, W., H. Xin, Y. Zhang, X. Wu, F. Yuan, and Z. Wang. 1999. The human REV1 gene codes for a DNA template-dependent dCMP transferase. Nucleic Acids Res. 274468-4475. [PMC free article] [PubMed]
143. Lin, X., and S. B. Howell. 2006. DNA mismatch repair and p53 function are major determinants of the rate of development of cisplatin resistance. Mol. Cancer Ther. 51239-1247. [PubMed]
144. Lin, X., T. Okuda, J. Trang, and S. B. Howell. 2006. Human REV1 modulates the cytotoxicity and mutagenicity of cisplatin in human ovarian carcinoma cells. Mol. Pharmacol. 691748-1754. [PubMed]
145. Lin, X., J. Trang, T. Okuda, and S. B. Howell. 2006. DNA polymerase zeta accounts for the reduced cytotoxicity and enhanced mutagenicity of cisplatin in human colon carcinoma cells that have lost DNA mismatch repair. Clin. Cancer Res. 12563-568. [PubMed]
146. Lindahl, T., and D. E. Barnes. 2000. Repair of endogenous DNA damage. Cold Spring Harbor Symp. Quant. Biol. 65127-133. [PubMed]
147. Ling, H., F. Boudsocq, B. S. Plosky, R. Woodgate, and W. Yang. 2003. Replication of a cis-syn thymine dimer at atomic resolution. Nature 4241083-1087. [PubMed]
148. Ling, H., F. Boudsocq, R. Woodgate, and W. Yang. 2001. Crystal structure of a Y-family DNA polymerase in action: a mechanism for error-prone and lesion-bypass replication. Cell 10791-102. [PubMed]
149. Ling, H., J. M. Sayer, B. S. Plosky, H. Yagi, F. Boudsocq, R. Woodgate, D. M. Jerina, and W. Yang. 2004. Crystal structure of a benzo[a]pyrene diol epoxide adduct in a ternary complex with a DNA polymerase. Proc. Natl. Acad. Sci. USA 1012265-2269. [PubMed]
150. Lone, S., S. A. Townson, S. N. Uljon, R. E. Johnson, A. Brahma, D. T. Nair, S. Prakash, L. Prakash, and A. K. Aggarwal. 2007. Human DNA polymerase kappa encircles DNA: implications for mismatch extension and lesion bypass. Mol. Cell 25601-614. [PubMed]
151. Lopes, M., M. Foiani, and J. M. Sogo. 2006. Multiple mechanisms control chromosome integrity after replication fork uncoupling and restart at irreparable UV lesions. Mol. Cell 2115-27. [PubMed]
152. Madril, A. C., R. E. Johnson, M. T. Washington, L. Prakash, and S. Prakash. 2001. Fidelity and damage bypass ability of Schizosaccharomyces pombe Eso1 protein, comprised of DNA polymerase eta and sister chromatid cohesion protein Ctf7. J. Biol. Chem. 27642857-42862. [PubMed]
153. Maga, G., and U. Hubscher. 2003. Proliferating cell nuclear antigen (PCNA): a dancer with many partners. J. Cell Sci. 1163051-3060. [PubMed]
154. Manke, I. A., D. M. Lowery, A. Nguyen, and M. B. Yaffe. 2003. BRCT repeats as phosphopeptide-binding modules involved in protein targeting. Science 302636-639. [PubMed]
155. Masuda, Y., and K. Kamiya. 2006. Role of single-stranded DNA in targeting REV1 to primer termini. J. Biol. Chem. 28124314-24321. [PubMed]
156. Masutani, C., M. Araki, A. Yamada, R. Kusumoto, T. Nogimori, T. Maekawa, S. Iwai, and F. Hanaoka. 1999. Xeroderma pigmentosum variant (XP-V) correcting protein from HeLa cells has a thymine dimer bypass DNA polymerase activity. EMBO J. 183491-3501. [PubMed]
157. Masutani, C., R. Kusumoto, A. Yamada, N. Dohmae, M. Yokoi, M. Yuasa, M. Araki, S. Iwai, K. Takio, and F. Hanaoka. 1999. The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase eta. Nature 399700-704. [PubMed]
158. Matsuda, T., K. Bebenek, C. Masutani, F. Hanaoka, and T. A. Kunkel. 2000. Low fidelity DNA synthesis by human DNA polymerase-eta. Nature 4041011-1013. [PubMed]
159. Mayorov, V. I., I. B. Rogozin, L. R. Adkison, and P. J. Gearhart. 2005. DNA polymerase eta contributes to strand bias of mutations of A versus T in immunoglobulin genes. J. Immunol. 1747781-7786. [PubMed]
160. McCulloch, S. D., R. J. Kokoska, O. Chilkova, C. M. Welch, E. Johansson, P. M. Burgers, and T. A. Kunkel. 2004. Enzymatic switching for efficient and accurate translesion DNA replication. Nucleic Acids Res. 324665-4675. [PMC free article] [PubMed]
161. McCulloch, S. D., R. J. Kokoska, and T. A. Kunkel. 2004. Efficiency, fidelity and enzymatic switching during translesion DNA synthesis. Cell Cycle 3580-583. [PubMed]
162. McCulloch, S. D., R. J. Kokoska, C. Masutani, S. Iwai, F. Hanaoka, and T. A. Kunkel. 2004. Preferential cis-syn thymine dimer bypass by DNA polymerase eta occurs with biased fidelity. Nature 42897-100. [PubMed]
163. McCulloch, S. D., and T. A. Kunkel. 2008. The fidelity of DNA synthesis by eukaryotic replicative and translesion synthesis polymerases. Cell Res. 18148-161. [PubMed]
164. McDonald, J. P., E. G. Frank, B. S. Plosky, I. B. Rogozin, C. Masutani, F. Hanaoka, R. Woodgate, and P. J. Gearhart. 2003. 129-derived strains of mice are deficient in DNA polymerase iota and have normal immunoglobulin hypermutation. J. Exp. Med. 198635-643. [PMC free article] [PubMed]
165. McDonald, J. P., A. S. Levine, and R. Woodgate. 1997. The Saccharomyces cerevisiae RAD30 gene, a homologue of Escherichia coli dinB and umuC, is DNA damage inducible and functions in a novel error-free postreplication repair mechanism. Genetics 1471557-1568. [PubMed]
166. McDonald, J. P., V. Rapic-Otrin, J. A. Epstein, B. C. Broughton, X. Wang, A. R. Lehmann, D. J. Wolgemuth, and R. Woodgate. 1999. Novel human and mouse homologs of Saccharomyces cerevisiae DNA polymerase eta. Genomics 6020-30. [PubMed]
167. McIlwraith, M. J., A. Vaisman, Y. Liu, E. Fanning, R. Woodgate, and S. C. West. 2005. Human DNA polymerase eta promotes DNA synthesis from strand invasion intermediates of homologous recombination. Mol. Cell 20783-792. [PubMed]
168. McNally, K., J. A. Neal, T. P. McManus, J. J. McCormick, and V. M. Maher. 2008. hRev7, putative subunit of hPolzeta, plays a critical role in survival, induction of mutations, and progression through S-phase, of UV((254nm))-irradiated human fibroblasts. DNA Repair (Amsterdam) 7597-604. [PubMed]
169. Minesinger, B. K., and S. Jinks-Robertson. 2005. Roles of RAD6 epistasis group members in spontaneous polzeta-dependent translesion synthesis in Saccharomyces cerevisiae. Genetics 1691939-1955. [PubMed]
170. Minko, I. G., K. Yamanaka, I. D. Kozekov, A. Kozekova, C. Indiani, M. E. O'Donnell, Q. Jiang, M. F. Goodman, C. J. Rizzo, and R. S. Lloyd. 2008. Replication bypass of the acrolein-mediated deoxyguanine DNA-peptide cross-links by DNA polymerases of the DinB family. Chem. Res. Toxicol. 211983-1990. [PMC free article] [PubMed]
171. Mirchandani, K. D., R. M. McCaffrey, and A. D. D'Andrea. 2008. The Fanconi anemia core complex is required for efficient point mutagenesis and Rev1 foci assembly. DNA Repair (Amsterdam) 7902-911. [PMC free article] [PubMed]
172. Moon, A. F., M. Garcia-Diaz, V. K. Batra, W. A. Beard, K. Bebenek, T. A. Kunkel, S. H. Wilson, and L. C. Pedersen. 2007. The X family portrait: structural insights into biological functions of X family polymerases. DNA Repair (Amsterdam) 61709-1725. [PMC free article] [PubMed]
173. Morrison, A., R. B. Christensen, J. Alley, A. K. Beck, E. G. Bernstine, J. F. Lemontt, and C. W. Lawrence. 1989. REV3, a Saccharomyces cerevisiae gene whose function is required for induced mutagenesis, is predicted to encode a nonessential DNA polymerase. J. Bacteriol. 1715659-5667. [PMC free article] [PubMed]
174. Mukhopadhyay, S., D. R. Clark, N. B. Watson, W. Zacharias, and W. G. McGregor. 2004. REV1 accumulates in DNA damage-induced nuclear foci in human cells and is implicated in mutagenesis by benzo[a]pyrenediolepoxide. Nucleic Acids Res. 325820-5826. [PMC free article] [PubMed]
175. Murakumo, Y., S. Mizutani, M. Yamaguchi, M. Ichihara, and M. Takahashi. 2006. Analyses of ultraviolet-induced focus formation of hREV1 protein. Genes Cells 11193-205. [PubMed]
176. Murakumo, Y., Y. Ogura, H. Ishii, S. Numata, M. Ichihara, C. M. Croce, R. Fishel, and M. Takahashi. 2001. Interactions in the error-prone postreplication repair proteins hREV1, hREV3, and hREV7. J. Biol. Chem. 27635644-35651. [PubMed]
177. Murakumo, Y., T. Roth, H. Ishii, D. Rasio, S. Numata, C. M. Croce, and R. Fishel. 2000. A human REV7 homolog that interacts with the polymerase zeta catalytic subunit hREV3 and the spindle assembly checkpoint protein hMAD2. J. Biol. Chem. 2754391-4397. [PubMed]
178. Nair, D. T., R. E. Johnson, L. Prakash, S. Prakash, and A. K. Aggarwal. 2005. Human DNA polymerase iota incorporates dCTP opposite template G via a G.C + Hoogsteen base pair. Structure 131569-1577. [PubMed]
179. Nair, D. T., R. E. Johnson, L. Prakash, S. Prakash, and A. K. Aggarwal. 2008. Protein-template-directed synthesis across an acrolein-derived DNA adduct by yeast Rev1 DNA polymerase. Structure 16239-245. [PubMed]
180. Nair, D. T., R. E. Johnson, L. Prakash, S. Prakash, and A. K. Aggarwal. 2005. Rev1 employs a novel mechanism of DNA synthesis using a protein template. Science 3092219-2222. [PubMed]
181. Nair, D. T., R. E. Johnson, S. Prakash, L. Prakash, and A. K. Aggarwal. 2004. Replication by human DNA polymerase-iota occurs by Hoogsteen base-pairing. Nature 430377-380. [PubMed]
182. Nakajima, S., L. Lan, S. Kanno, N. Usami, K. Kobayashi, M. Mori, T. Shiomi, and A. Yasui. 2006. Replication-dependent and -independent responses of RAD18 to DNA damage in human cells. J. Biol. Chem. 28134687-34695. [PubMed]
183. Nelson, J. R., P. E. Gibbs, A. M. Nowicka, D. C. Hinkle, and C. W. Lawrence. 2000. Evidence for a second function for Saccharomyces cerevisiae Rev1p. Mol. Microbiol. 37549-554. [PubMed]
184. Nelson, J. R., C. W. Lawrence, and D. C. Hinkle. 1996. Deoxycytidyl transferase activity of yeast REV1 protein. Nature 382729-731. [PubMed]
185. Nelson, J. R., C. W. Lawrence, and D. C. Hinkle. 1996. Thymine-thymine dimer bypass by yeast DNA polymerase zeta. Science 2721646-1649. [PubMed]
186. Nelson, K. K., J. Schlondorff, and C. P. Blobel. 1999. Evidence for an interaction of the metalloprotease-disintegrin tumour necrosis factor alpha convertase (TACE) with mitotic arrest deficient 2 (MAD2), and of the metalloprotease-disintegrin MDC9 with a novel MAD2-related protein, MAD2beta. Biochem. J. 343673-680. [PubMed]
187. Niedzwiedz, W., G. Mosedale, M. Johnson, C. Y. Ong, P. Pace, and K. J. Patel. 2004. The Fanconi anaemia gene FANCC promotes homologous recombination and error-prone DNA repair. Mol. Cell 15607-620. [PubMed]
188. Niimi, A., S. Brown, S. Sabbioneda, P. L. Kannouche, A. Scott, A. Yasui, C. M. Green, and A. R. Lehmann. 2008. Regulation of proliferating cell nuclear antigen ubiquitination in mammalian cells. Proc. Natl. Acad. Sci. USA 10516125-16130. [PubMed]
189. Nikolaishvili-Feinberg, N., G. S. Jenkins, K. R. Nevis, D. P. Staus, C. O. Scarlett, K. Unsal-Kacmaz, W. K. Kaufmann, and M. Cordeiro-Stone. 2008. Ubiquitylation of proliferating cell nuclear antigen and recruitment of human DNA polymerase eta. Biochemistry 474141-4150. [PubMed]
190. Northam, M. R., P. Garg, D. M. Baitin, P. M. Burgers, and P. V. Shcherbakova. 2006. A novel function of DNA polymerase zeta regulated by PCNA. EMBO J. 254316-4325. [PubMed]
191. Ogi, T., P. Kannouche, and A. R. Lehmann. 2005. Localisation of human Y-family DNA polymerase kappa: relationship to PCNA foci. J. Cell Sci. 118129-136. [PubMed]
192. Ogi, T., T. Kato, Jr., T. Kato, and H. Ohmori. 1999. Mutation enhancement by DINB1, a mammalian homologue of the Escherichia coli mutagenesis protein dinB. Genes Cells. 4607-618. [PubMed]
193. Ogi, T., and A. R. Lehmann. 2006. The Y-family DNA polymerase kappa (pol kappa) functions in mammalian nucleotide-excision repair. Nat. Cell Biol. 8640-642. [PubMed]
194. Ogi, T., J. Mimura, M. Hikida, H. Fujimoto, Y. Fujii-Kuriyama, and H. Ohmori. 2001. Expression of human and mouse genes encoding polkappa: testis-specific developmental regulation and AhR-dependent inducible transcription. Genes Cells 6943-953. [PubMed]
195. Ogi, T., Y. Shinkai, K. Tanaka, and H. Ohmori. 2002. Polkappa protects mammalian cells against the lethal and mutagenic effects of benzo[a]pyrene. Proc. Natl. Acad. Sci. USA 9915548-15553. [PubMed]
196. Ohashi, E., K. Bebenek, T. Matsuda, W. J. Feaver, V. L. Gerlach, E. C. Friedberg, H. Ohmori, and T. A. Kunkel. 2000. Fidelity and processivity of DNA synthesis by DNA polymerase kappa, the product of the human DINB1 gene. J. Biol. Chem. 27539678-39684. [PubMed]
197. Ohashi, E., Y. Murakumo, N. Kanjo, J. Akagi, C. Masutani, F. Hanaoka, and H. Ohmori. 2004. Interaction of hREV1 with three human Y-family DNA polymerases. Genes Cells 9523-531. [PubMed]
198. Ohkumo, T., Y. Kondo, M. Yokoi, T. Tsukamoto, A. Yamada, T. Sugimoto, R. Kanao, Y. Higashi, H. Kondoh, M. Tatematsu, C. Masutani, and F. Hanaoka. 2006. UV-B radiation induces epithelial tumors in mice lacking DNA polymerase eta and mesenchymal tumors in mice deficient for DNA polymerase iota. Mol. Cell. Biol. 267696-7706. [PMC free article] [PubMed]
199. Ohmori, H., E. C. Friedberg, R. P. Fuchs, M. F. Goodman, F. Hanaoka, D. Hinkle, T. A. Kunkel, C. W. Lawrence, Z. Livneh, T. Nohmi, L. Prakash, S. Prakash, T. Todo, G. C. Walker, Z. Wang, and R. Woodgate. 2001. The Y-family of DNA polymerases. Mol. Cell 87-8. [PubMed]
200. Okada, T., E. Sonoda, M. Yoshimura, Y. Kawano, H. Saya, M. Kohzaki, and S. Takeda. 2005. Multiple roles of vertebrate REV genes in DNA repair and recombination. Mol. Cell. Biol. 256103-6111. [PMC free article] [PubMed]
201. Okuda, T., X. Lin, J. Trang, and S. B. Howell. 2005. Suppression of hREV1 expression reduces the rate at which human ovarian carcinoma cells acquire resistance to cisplatin. Mol. Pharmacol. 671852-1860. [PubMed]
202. Otsuka, C., N. Kunitomi, S. Iwai, D. Loakes, and K. Negishi. 2005. Roles of the polymerase and BRCT domains of Rev1 protein in translesion DNA synthesis in yeast in vivo. Mutat. Res. 57879-87. [PubMed]
202a. O-Wang, J., K. Kawamura, Y. Tada, H. Ohmori, H. Kimura, S. Sakiyama, and M. Tagawa. 2001. DNA polymerase kappa, implicated in spontaneous and DNA damage-induced mutagenesis, is overexpressed in lung cancer. Cancer Res. 615366-5369. [PubMed]
203. Pabla, R., D. Rozario, and W. Siede. 2008. Regulation of Saccharomyces cerevisiae DNA polymerase eta transcript and protein. Radiat. Environ. Biophys. 47157-168. [PubMed]
204. Papadopoulo, D., C. Guillouf, B. Porfirio, and E. Moustacchi. 1990. Decreased mutagenicity in Fanconi's anemia lymphoblasts following treatment with photoactivated psoralens. Prog. Clin. Biol. Res. 340A241-248. [PubMed]
205. Parker, J. L., A. B. Bielen, I. Dikic, and H. D. Ulrich. 2007. Contributions of ubiquitin- and PCNA-binding domains to the activity of polymerase eta in Saccharomyces cerevisiae. Nucleic Acids Res. 35881-889. [PMC free article] [PubMed]
206. Patel, P. H., M. Suzuki, E. Adman, A. Shinkai, and L. A. Loeb. 2001. Prokaryotic DNA polymerase I: evolution, structure, and “base flipping” mechanism for nucleotide selection. J. Mol. Biol. 308823-837. [PubMed]
207. Paulovich, A. G., C. D. Armour, and L. H. Hartwell. 1998. The Saccharomyces cerevisiae RAD9, RAD17, RAD24 and MEC3 genes are required for tolerating irreparable, ultraviolet-induced DNA damage. Genetics 15075-93. [PubMed]
208. Pavlov, Y. I., D. Nguyen, and T. A. Kunkel. 2001. Mutator effects of overproducing DNA polymerase eta (Rad30) and its catalytically inactive variant in yeast. Mutat. Res. 478129-139. [PubMed]
209. Pavlov, Y. I., I. B. Rogozin, A. P. Galkin, A. Y. Aksenova, F. Hanaoka, C. Rada, and T. A. Kunkel. 2002. Correlation of somatic hypermutation specificity and A-T base pair substitution errors by DNA polymerase eta during copying of a mouse immunoglobulin kappa light chain transgene. Proc. Natl. Acad. Sci. USA 999954-9959. [PubMed]
210. Petta, T. B., S. Nakajima, A. Zlatanou, E. Despras, S. Couve-Privat, A. Ishchenko, A. Sarasin, A. Yasui, and P. Kannouche. 2008. Human DNA polymerase iota protects cells against oxidative stress. EMBO J. 272883-2895. [PubMed]
211. Pfleger, C. M., A. Salic, E. Lee, and M. W. Kirschner. 2001. Inhibition of Cdh1-APC by the MAD2-related protein MAD2L2: a novel mechanism for regulating Cdh1. Genes Dev. 151759-1764. [PubMed]
212. Pillaire, M. J., R. Betous, C. Conti, J. Czaplicki, P. Pasero, A. Bensimon, C. Cazaux, and J. S. Hoffmann. 2007. Upregulation of error-prone DNA polymerases beta and kappa slows down fork progression without activating the replication checkpoint. Cell Cycle. 6471-477. [PubMed]
213. Plosky, B. S., A. E. Vidal, A. R. de Henestrosa, M. P. McLenigan, J. P. McDonald, S. Mead, and R. Woodgate. 2006. Controlling the subcellular localization of DNA polymerases iota and eta via interactions with ubiquitin. EMBO J. 252847-2855. [PubMed]
214. Plosky, B. S., and R. Woodgate. 2004. Switching from high-fidelity replicases to low-fidelity lesion-bypass polymerases. Curr. Opin. Genet. Dev. 14113-119. [PubMed]
215. Poll, E. H., F. Arwert, H. Joenje, and A. H. Wanamarta. 1985. Differential sensitivity of Fanconi anaemia lymphocytes to the clastogenic action of cis-diamminedichloroplatinum (II) and trans-diamminedichloroplatinum (II). Hum. Genet. 71206-210. [PubMed]
216. Potapova, O., N. D. Grindley, and C. M. Joyce. 2002. The mutational specificity of the Dbh lesion bypass polymerase and its implications. J. Biol. Chem. 27728157-28166. [PubMed]
217. Prakash, S., R. E. Johnson, and L. Prakash. 2005. Eukaryotic translesion synthesis DNA polymerases: specificity of structure and function. Annu. Rev. Biochem. 74317-353. [PubMed]
218. Prakash, S., and L. Prakash. 2002. Translesion DNA synthesis in eukaryotes: a one- or two-polymerase affair. Genes Dev. 161872-1883. [PubMed]
219. Rajpal, D. K., X. Wu, and Z. Wang. 2000. Alteration of ultraviolet-induced mutagenesis in yeast through molecular modulation of the REV3 and REV7 gene expression. Mutat. Res. 461133-143. [PubMed]
220. Rasmussen, A. K., A. Chatterjee, L. J. Rasmussen, and K. K. Singh. 2003. Mitochondria-mediated nuclear mutator phenotype in Saccharomyces cerevisiae. Nucleic Acids Res. 313909-3917. [PMC free article] [PubMed]
221. Rattray, A. J., B. K. Shafer, C. B. McGill, and J. N. Strathern. 2002. The roles of REV3 and RAD57 in double-strand-break-repair-induced mutagenesis of Saccharomyces cerevisiae. Genetics 1621063-1077. [PubMed]
222. Rimkus, C., J. Friederichs, R. Rosenberg, B. Holzmann, J. R. Siewert, and K. P. Janssen. 2007. Expression of the mitotic checkpoint gene MAD2L2 has prognostic significance in colon cancer. Int. J. Cancer 120207-211. [PubMed]
223. Robbins, J. H., K. H. Kraemer, and B. A. Flaxman. 1975. DNA repair in tumor cells from the variant form of xeroderma pigmentosum. J. Investig. Dermatol. 64150-155. [PubMed]
224. Ross, A. L., L. J. Simpson, and J. E. Sale. 2005. Vertebrate DNA damage tolerance requires the C-terminus but not BRCT or transferase domains of REV1. Nucleic Acids Res. 331280-1289. [PMC free article] [PubMed]
225. Roush, A. A., M. Suarez, E. C. Friedberg, M. Radman, and W. Siede. 1998. Deletion of the Saccharomyces cerevisiae gene RAD30 encoding an Escherichia coli DinB homolog confers UV radiation sensitivity and altered mutability. Mol. Gen. Genet. 257686-692. [PubMed]
226. Rupp, W. D., and P. Howard-Flanders. 1968. Discontinuities in the DNA synthesized in an excision-defective strain of Escherichia coli following ultraviolet irradiation. J. Mol. Biol. 31291-304. [PubMed]
227. Sabbioneda, S., I. Bortolomai, M. Giannattasio, P. Plevani, and M. Muzi-Falconi. 2007. Yeast Rev1 is cell cycle regulated, phosphorylated in response to DNA damage and its binding to chromosomes is dependent upon MEC1. DNA Repair (Amsterdam) 6121-127. [PubMed]
228. Sabbioneda, S., A. M. Gourdin, C. M. Green, A. Zotter, G. Giglia-Mari, A. Houtsmuller, W. Vermeulen, and A. R. Lehmann. 2008. Effect of proliferating cell nuclear antigen ubiquitination and chromatin structure on the dynamic properties of the Y-family DNA polymerases. Mol. Biol. Cell 195193-5202. [PMC free article] [PubMed]
229. Sabbioneda, S., B. K. Minesinger, M. Giannattasio, P. Plevani, M. Muzi-Falconi, and S. Jinks-Robertson. 2005. The 9-1-1 checkpoint clamp physically interacts with polzeta and is partially required for spontaneous polzeta-dependent mutagenesis in Saccharomyces cerevisiae. J. Biol. Chem. 28038657-38665. [PubMed]
230. Sakiyama, T., T. Kohno, S. Mimaki, T. Ohta, N. Yanagitani, T. Sobue, H. Kunitoh, R. Saito, K. Shimizu, C. Hirama, J. Kimura, G. Maeno, H. Hirose, T. Eguchi, D. Saito, M. Ohki, and J. Yokota. 2005. Association of amino acid substitution polymorphisms in DNA repair genes TP53, POLI, REV1 and LIG4 with lung cancer risk. Int. J. Cancer 114730-737. [PubMed]
231. Shen, X., S. Jun, L. E. O'Neal, E. Sonoda, M. Bemark, J. E. Sale, and L. Li. 2006. REV3 and REV1 play major roles in recombination-independent repair of DNA interstrand cross-links mediated by monoubiquitinated proliferating cell nuclear antigen (PCNA). J. Biol. Chem. 28113869-13872. [PubMed]
232. Silvian, L. F., E. A. Toth, P. Pham, M. F. Goodman, and T. Ellenberger. 2001. Crystal structure of a DinB family error-prone DNA polymerase from Sulfolobus solfataricus. Nat. Struct. Biol. 8984-989. [PubMed]
233. Simpson, L. J., A. L. Ross, D. Szuts, C. A. Alviani, V. H. Oestergaard, K. J. Patel, and J. E. Sale. 2006. RAD18-independent ubiquitination of proliferating-cell nuclear antigen in the avian cell line DT40. EMBO Rep. 7927-932. [PubMed]
234. Singhal, R. K., D. C. Hinkle, and C. W. Lawrence. 1992. The REV3 gene of Saccharomyces cerevisiae is transcriptionally regulated more like a repair gene than one encoding a DNA polymerase. Mol. Gen. Genet. 23617-24. [PubMed]
235. Soria, G., O. Podhajcer, C. Prives, and V. Gottifredi. 2006. P21Cip1/WAF1 downregulation is required for efficient PCNA ubiquitination after UV irradiation. Oncogene 252829-2838. [PubMed]
236. Soria, G., J. Speroni, O. L. Podhajcer, C. Prives, and V. Gottifredi. 2008. p21 differentially regulates DNA replication and DNA-repair-associated processes after UV irradiation. J. Cell Sci. 1213271-3282. [PubMed]
237. Stary, A., P. Kannouche, A. R. Lehmann, and A. Sarasin. 2003. Role of DNA polymerase eta in the UV mutation spectrum in human cells. J. Biol. Chem. 27818767-18775. [PubMed]
238. Stelter, P., and H. D. Ulrich. 2003. Control of spontaneous and damage-induced mutagenesis by SUMO and ubiquitin conjugation. Nature 425188-191. [PubMed]
239. Takenaka, K., T. Ogi, T. Okada, E. Sonoda, C. Guo, E. C. Friedberg, and S. Takeda. 2006. Involvement of vertebrate Polkappa in translesion DNA synthesis across DNA monoalkylation damage. J. Biol. Chem. 2812000-2004. [PubMed]
240. Takeuchi, R., M. Oshige, M. Uchida, G. Ishikawa, K. Takata, K. Shimanouchi, Y. Kanai, T. Ruike, H. Morioka, and K. Sakaguchi. 2004. Purification of Drosophila DNA polymerase zeta by REV1 protein-affinity chromatography. Biochem. J. 382535-543. [PubMed]
241. Tanaka, K., T. Yonekawa, Y. Kawasaki, M. Kai, K. Furuya, M. Iwasaki, H. Murakami, M. Yanagida, and H. Okayama. 2000. Fission yeast Eso1p is required for establishing sister chromatid cohesion during S phase. Mol. Cell. Biol. 203459-3469. [PMC free article] [PubMed]
242. Tissier, A., E. G. Frank, J. P. McDonald, S. Iwai, F. Hanaoka, and R. Woodgate. 2000. Misinsertion and bypass of thymine-thymine dimers by human DNA polymerase iota. EMBO J. 195259-5266. [PubMed]
243. Tissier, A., P. Kannouche, M. P. Reck, A. R. Lehmann, R. P. Fuchs, and A. Cordonnier. 2004. Co-localization in replication foci and interaction of human Y-family members, DNA polymerase pol eta and REVl protein. DNA Repair (Amsterdam) 31503-1514. [PubMed]
244. Trincao, J., R. E. Johnson, C. R. Escalante, S. Prakash, L. Prakash, and A. K. Aggarwal. 2001. Structure of the catalytic core of S. cerevisiae DNA polymerase eta: implications for translesion DNA synthesis. Mol. Cell 8417-426. [PubMed]
245. Uljon, S. N., R. E. Johnson, T. A. Edwards, S. Prakash, L. Prakash, and A. K. Aggarwal. 2004. Crystal structure of the catalytic core of human DNA polymerase kappa. Structure 121395-1404. [PubMed]
246. Ulrich, H. D. 2005. The RAD6 pathway: control of DNA damage bypass and mutagenesis by ubiquitin and SUMO. Chembiochem 61735-1743. [PubMed]
247. Vaisman, A., C. Masutani, F. Hanaoka, and S. G. Chaney. 2000. Efficient translesion replication past oxaliplatin and cisplatin GpG adducts by human DNA polymerase eta. Biochemistry 394575-4580. [PubMed]
248. Van Sloun, P. P., I. Varlet, E. Sonneveld, J. J. Boei, R. J. Romeijn, J. C. Eeken, and N. De Wind. 2002. Involvement of mouse Rev3 in tolerance of endogenous and exogenous DNA damage. Mol. Cell. Biol. 222159-2169. [PMC free article] [PubMed]
249. Venkitaraman, A. R. 2002. Cancer susceptibility and the functions of BRCA1 and BRCA2. Cell 108171-182. [PubMed]
250. Vidal, A. E., P. Kannouche, V. N. Podust, W. Yang, A. R. Lehmann, and R. Woodgate. 2004. Proliferating cell nuclear antigen-dependent coordination of the biological functions of human DNA polymerase iota. J. Biol. Chem. 27948360-48368. [PubMed]
251. Wagner, J., P. Gruz, S. R. Kim, M. Yamada, K. Matsui, R. P. Fuchs, and T. Nohmi. 1999. The dinB gene encodes a novel E. coli DNA polymerase, DNA pol IV, involved in mutagenesis. Mol. Cell 4281-286. [PubMed]
252. Wang, Y., R. Woodgate, T. P. McManus, S. Mead, J. J. McCormick, and V. M. Maher. 2007. Evidence that in xeroderma pigmentosum variant cells, which lack DNA polymerase eta, DNA polymerase iota causes the very high frequency and unique spectrum of UV-induced mutations. Cancer Res. 673018-3026. [PubMed]
253. Washington, M. T., R. E. Johnson, S. Prakash, and L. Prakash. 1999. Fidelity and processivity of Saccharomyces cerevisiae DNA polymerase eta. J. Biol. Chem. 27436835-36838. [PubMed]
254. Washington, M. T., I. G. Minko, R. E. Johnson, L. Haracska, T. M. Harris, R. S. Lloyd, S. Prakash, and L. Prakash. 2004. Efficient and error-free replication past a minor-groove N2-guanine adduct by the sequential action of yeast Rev1 and DNA polymerase zeta. Mol. Cell. Biol. 246900-6906. [PMC free article] [PubMed]
255. Washington, M. T., L. Prakash, and S. Prakash. 2001. Yeast DNA polymerase eta utilizes an induced-fit mechanism of nucleotide incorporation. Cell 107917-927. [PubMed]
256. Watanabe, K., S. Tateishi, M. Kawasuji, T. Tsurimoto, H. Inoue, and M. Yamaizumi. 2004. Rad18 guides poleta to replication stalling sites through physical interaction and PCNA monoubiquitination. EMBO J. 233886-3896. [PubMed]
257. Waters, L. S., and G. C. Walker. 2006. The critical mutagenic translesion DNA polymerase Rev1 is highly expressed during G(2)/M phase rather than S phase. Proc. Natl. Acad. Sci. USA 1038971-8976. [PubMed]
258. Weterman, M. A., J. J. van Groningen, L. Tertoolen, and A. G. van Kessel. 2001. Impairment of MAD2B-PRCC interaction in mitotic checkpoint defective t(X;1)-positive renal cell carcinomas. Proc. Natl. Acad. Sci. USA 9813808-13813. [PubMed]
259. Wilkinson, A., A. Smith, D. Bullard, M. Lavesa-Curto, H. Sayer, A. Bonner, A. Hemmings, and R. Bowater. 2005. Analysis of ligation and DNA binding by Escherichia coli DNA ligase (LigA). Biochim. Biophys. Acta 1749113-122. [PubMed]
260. Wittschieben, J., M. K. Shivji, E. Lalani, M. A. Jacobs, F. Marini, P. J. Gearhart, I. Rosewell, G. Stamp, and R. D. Wood. 2000. Disruption of the developmentally regulated Rev3l gene causes embryonic lethality. Curr. Biol. 101217-1220. [PubMed]
261. Wood, A., P. Garg, and P. M. Burgers. 2007. A ubiquitin-binding motif in the translesion DNA polymerase Rev1 mediates its essential functional interaction with ubiquitinated proliferating cell nuclear antigen in response to DNA damage. J. Biol. Chem. 28220256-20263. [PubMed]
262. Xiao, W., B. L. Chow, S. Broomfield, and M. Hanna. 2000. The Saccharomyces cerevisiae RAD6 group is composed of an error-prone and two error-free postreplication repair pathways. Genetics 1551633-1641. [PubMed]
263. Yagi, Y., D. Ogawara, S. Iwai, F. Hanaoka, M. Akiyama, and H. Maki. 2005. DNA polymerases eta and kappa are responsible for error-free translesion DNA synthesis activity over a cis-syn thymine dimer in Xenopus laevis oocyte extracts. DNA Repair (Amsterdam) 41252-1269. [PubMed]
264. Yamada, A., C. Masutani, S. Iwai, and F. Hanaoka. 2000. Complementation of defective translesion synthesis and UV light sensitivity in xeroderma pigmentosum variant cells by human and mouse DNA polymerase eta. Nucleic Acids Res. 282473-2480. [PMC free article] [PubMed]
265. Yang, J., Z. Chen, Y. Liu, R. J. Hickey, and L. H. Malkas. 2004. Altered DNA polymerase iota expression in breast cancer cells leads to a reduction in DNA replication fidelity and a higher rate of mutagenesis. Cancer Res. 645597-5607. [PubMed]
266. Yang, W. 2003. Damage repair DNA polymerases Y. Curr. Opin. Struct. Biol. 1323-30. [PubMed]
267. Yang, W. 2005. Portraits of a Y-family DNA polymerase. FEBS Lett. 579868-872. [PubMed]
268. Yang, W., and R. Woodgate. 2007. What a difference a decade makes: insights into translesion DNA synthesis. Proc. Natl. Acad. Sci. USA 10415591-15598. [PubMed]
269. Ying, B., and W. S. Wold. 2003. Adenovirus ADP protein (E3-11.6K), which is required for efficient cell lysis and virus release, interacts with human MAD2B. Virology 313224-234. [PubMed]
270. Yu, X., C. C. Chini, M. He, G. Mer, and J. Chen. 2003. The BRCT domain is a phospho-protein binding domain. Science 302639-642. [PubMed]
271. Yuan, B., H. Cao, Y. Jiang, H. Hong, and Y. Wang. 2008. Efficient and accurate bypass of N2-(1-carboxyethyl)-2′-deoxyguanosine by DinB DNA polymerase in vitro and in vivo. Proc. Natl. Acad. Sci. USA 1058679-8684. [PubMed]
272. Yuan, F., Y. Zhang, D. K. Rajpal, X. Wu, D. Guo, M. Wang, J. S. Taylor, and Z. Wang. 2000. Specificity of DNA lesion bypass by the yeast DNA polymerase eta. J. Biol. Chem. 2758233-8239. [PubMed]
273. Yuasa, M. S., C. Masutani, A. Hirano, M. A. Cohn, M. Yamaizumi, Y. Nakatani, and F. Hanaoka. 2006. A human DNA polymerase eta complex containing Rad18, Rad6 and Rev1; proteomic analysis and targeting of the complex to the chromatin-bound fraction of cells undergoing replication fork arrest. Genes Cells 11731-744. [PubMed]
274. Zhang, H., A. Chatterjee, and K. K. Singh. 2006. Saccharomyces cerevisiae polymerase zeta functions in mitochondria. Genetics 1722683-2688. [PubMed]
275. Zhang, L., S. H. Yang, and A. D. Sharrocks. 2007. Rev7/MAD2B links c-Jun N-terminal protein kinase pathway signaling to activation of the transcription factor Elk-1. Mol. Cell. Biol. 272861-2869. [PMC free article] [PubMed]
276. Zhang, Y., X. Wu, O. Rechkoblit, N. E. Geacintov, J. S. Taylor, and Z. Wang. 2002. Response of human REV1 to different DNA damage: preferential dCMP insertion opposite the lesion. Nucleic Acids Res. 301630-1638. [PMC free article] [PubMed]
277. Zhang, Y., F. Yuan, X. Wu, O. Rechkoblit, J. S. Taylor, N. E. Geacintov, and Z. Wang. 2000. Error-prone lesion bypass by human DNA polymerase eta. Nucleic Acids Res. 284717-4724. [PMC free article] [PubMed]
278. Zhong, X., P. Garg, C. M. Stith, S. A. Nick McElhinny, G. E. Kissling, P. M. Burgers, and T. A. Kunkel. 2006. The fidelity of DNA synthesis by yeast DNA polymerase zeta alone and with accessory proteins. Nucleic Acids Res. 344731-4742. [PubMed]
279. Zhou, B. L., J. D. Pata, and T. A. Steitz. 2001. Crystal structure of a DinB lesion bypass DNA polymerase catalytic fragment reveals a classic polymerase catalytic domain. Mol. Cell 8427-437. [PubMed]
280. Zhuang, Z., R. E. Johnson, L. Haracska, L. Prakash, S. Prakash, and S. J. Benkovic. 2008. Regulation of polymerase exchange between Poleta and Poldelta by monoubiquitination of PCNA and the movement of DNA polymerase holoenzyme. Proc. Natl. Acad. Sci. USA 1055361-5366. [PubMed]

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