Roles of trans-lesion synthesis (TLS) DNA polymerases in tumorigenesis and cancer therapy

Jay Anand; Lilly Chiou; Carly Sciandra; Xingyuan Zhang; Jiyong Hong; Di Wu; Pei Zhou; Cyrus Vaziri

doi:10.1093/narcan/zcad005

. 2023 Feb 6;5(1):zcad005. doi: 10.1093/narcan/zcad005

Roles of trans-lesion synthesis (TLS) DNA polymerases in tumorigenesis and cancer therapy

Jay Anand ^1,², Lilly Chiou ^2,^3,², Carly Sciandra ^4,², Xingyuan Zhang ^5,², Jiyong Hong ⁶, Di Wu ⁷, Pei Zhou ⁸, Cyrus Vaziri ^9,^✉

PMCID: PMC9900426 PMID: 36755961

Abstract

DNA damage tolerance and mutagenesis are hallmarks and enabling characteristics of neoplastic cells that drive tumorigenesis and allow cancer cells to resist therapy. The ‘Y-family’ trans-lesion synthesis (TLS) DNA polymerases enable cells to replicate damaged genomes, thereby conferring DNA damage tolerance. Moreover, Y-family DNA polymerases are inherently error-prone and cause mutations. Therefore, TLS DNA polymerases are potential mediators of important tumorigenic phenotypes. The skin cancer-propensity syndrome xeroderma pigmentosum-variant (XPV) results from defects in the Y-family DNA Polymerase Pol eta (Polη) and compensatory deployment of alternative inappropriate DNA polymerases. However, the extent to which dysregulated TLS contributes to the underlying etiology of other human cancers is unclear. Here we consider the broad impact of TLS polymerases on tumorigenesis and cancer therapy. We survey the ways in which TLS DNA polymerases are pathologically altered in cancer. We summarize evidence that TLS polymerases shape cancer genomes, and review studies implicating dysregulated TLS as a driver of carcinogenesis. Because many cancer treatment regimens comprise DNA-damaging agents, pharmacological inhibition of TLS is an attractive strategy for sensitizing tumors to genotoxic therapies. Therefore, we discuss the pharmacological tractability of the TLS pathway and summarize recent progress on development of TLS inhibitors for therapeutic purposes.

INTRODUCTION

Mutability has long been recognized as a key hallmark and enabling characteristic of cancer (1,2). Identification of trans-lesion synthesis (TLS) DNA polymerases that perform error-prone replication of damaged DNA templates was once heralded as a major breakthrough in the field of chemical carcinogenesis: TLS polymerases provided a new molecular mechanism for mutagenesis and thus were hypothesized to be potential drivers of carcinogenesis (3). The extent to which this hypothesis has been validated is discussed here.

Historically, TLS was first viewed largely as a mechanism for replicative bypass of bulky DNA adducts and was presumed to be restricted to the S-phase of the cell cycle. We now know that TLS polymerases perform diverse functions in genome maintenance far beyond mediating replicative bypass of damaged DNA templates. TLS is an important gap-filling process that eliminates single-stranded DNA (ssDNA) discontinuities in the genome. TLS DNA polymerases are not always restricted to S-phase and may also be deployed in G0, G1 and G2 (4,5,6,7,8,9,10). Moreover, TLS is closely integrated with the replicative cell cycle and other genome maintenance mechanisms. It is now appreciated that neoplastic cells experience considerable DNA damage from intrinsic and therapeutic sources. Thus, in addition to promoting mutagenesis during carcinogenesis, the DNA damage-tolerance conferred by TLS DNA polymerases has the potential to sustain proliferation and viability of cancer cells. Accordingly, TLS is a dependency of some cancer cells and represents an appealing target pathway for therapeutic development. Small molecule inhibitors of TLS polymerases sensitize cancer cells to genotoxic therapeutic agents and preferentially inhibit the viability of neoplastic cell lines when compared with untransformed cells.

Here, we review the molecular anatomy of the TLS pathway and describe its proposed roles in genome maintenance. We emphasize potential mechanisms by which the TLS pathway activity may be dysregulated in cancer. We survey the evidence that TLS shapes the genomic landscape of cancer cells and contributes to carcinogenesis. We discuss emerging evidence that TLS is a dependency and vulnerability of cancer cells. Finally, we describe ongoing strategies and efforts to target the TLS pathway for therapeutic purposes.

TLS PATHWAY IMBALANCE DUE TO POLη-DEFICIENCY CAUSES SKIN CANCER

Our understanding of TLS and its relationship with cancer is founded largely on the Hanaoka group's seminal discovery of the TLS DNA polymerase eta (Polη) as the mutated gene product of xeroderma pigmentosum-Variant (XPV) patients (11,12). XPV is a sunlight-sensitivity and skin cancer-propensity syndrome. Solar UVB (290–320 nm) exposure is causally linked to skin cancer (13). Two frequent DNA lesions induced by UVB are cis–syn cyclobutane pyrimidine dimers (CPD) and pyrimidine (4,5,6) pyrimidone photoproducts (14). In XPV patients nucleotide excision repair (NER) of CPD is intact yet post-replicative DNA repair (to be defined later) is compromised. Using biochemical complementation, Masutani et al. identified Polη as the factor that corrects defective replication of CPD-damaged DNA templates by XPV cell extracts (11). Those workers subsequently showed that the POLH gene is mutated in XPV patients and that Polη is the human homologue of yeast Rad30. How then are the clinical features of XPV explained by Polη-deficiency?

Polη has low processivity and low fidelity when replicating undamaged DNA templates. However, this enzyme is specialized to perform efficient and error-free replicative bypass of templates containing helix-distorting CPD (12). Thus, Polη is important for ongoing DNA replication of genomes harboring CPD and confers ‘DNA damage tolerance’. XPV patients lacking Polη cannot replicate their UV-damaged genomes and lose viability, thereby explaining the sunlight-sensitivity phenotype. XPV patients develop carcinoma (both squamous- and basal cell carcinoma) with frequencies that are 10,000- and 2,000-fold higher, respectively, when compared with the general population (15). XPV patients are also prone to melanoma (16,17). The high skin cancer incidence of XPV patients is attributed to increased rates of UV-induced mutagenesis in those individuals. Mechanistically, when Polη is absent, compensatory, yet error-prone TLS of CPD lesions by other ‘inserter’ Y-family DNA polymerases, Pol kappa (Polκ) and Pol iota (Polι) working cooperatively with the B-family ‘extender’ DNA polymerase Polζ leads to hypermutability (18,19) (Figure 1). There is also evidence for compensatory usage of other genome-altering DNA repair pathways in UV-irradiated XPV cells, including Homologous Recombination (HR) which leads to increased sister chromatid exchanges, or SCE (20). Therefore, XPV is a malignancy resulting from deployment of alternative, more error-prone genome maintenance mediators when an individual Y-family TLS polymerase (Polη) is absent (Figure 1). Such ‘imbalance’ and altered DNA repair pathway choice is a general conceptual framework for explaining mechanisms of genome instability in cancer (21). A good example of how imbalanced DNA repair pathway choice may lead to genetic change is provided by BRCA1-deficient ovarian cancers in which HR is defective. D’Andrea and colleagues showed that those HRD tumors rely on compensatory overexpression and deployment of the error-prone DNA polymerase POLQ for DSB repair (22). POLQ mediates error-prone microhomology-mediated end-joining (MMEJ) of DSB which helps explain the genomic landscape of BRCA-mutated cancers (23,24).

Figure 1. — Aberrant activation of TLS affects genome stability. (A) In normal cells, Polη performs error-free bypass of CPD. TLS is restrained and used sparingly to minimize error-prone DNA synthesis and mutagenesis. (B) In XPV cells, Polη is absent. Compensatory error-prone TLS of Polη-cognate lesions by alternative Y-family inserter DNA polymerases and the B-family extender DNA polymerase Polζ leads to mutagenesis. (C) HR-deficient cancer (such as BRCA1/2 mutant cells) have increased dependency on TLS for ssDNA gap-filling, which may lead to increased error-prone DNA synthesis and mutagenesis. The mutagenic outcome will depend on the type of DNA damage and the choice of TLS polymerase deployed. (D) Cancer-associated increase (green arrow) or decrease (red arrow) in expression of TLS mediators might affect TLS polymerase selection. The upregulation of RAD18 and the TLS polymerase imbalances illustrated here represent the mRNA expression patterns observed in endometrial cancers when compared with normal adjacent tissues (Figure 3).

Each TLS polymerase is specialized to perform relatively accurate and efficient bypass of a specific type of DNA damage, referred to as their ‘cognate lesion’ (25,26). For example CPD are cognate lesions for Polη. Given the error-propensity of TLS polymerases on undamaged DNA templates or non-cognate lesions, their imbalanced activities in relation to each other might promote mutagenesis. XPV is a situation in which Polη activity is reduced relative to other TLS polymerases. By analogy, there are other possible ways in which imbalance and altered pathway choice may develop between the different TLS DNA polymerases (or between TLS polymerases and other DNA repair mechanisms). Figure 1 illustrates possible ways in which TLS might be altered in cancer. Later, we consider some of these possibilities and review evidence that altered TLS impacts DNA damage-sensitivity and the genomic features of cancer cells. First, we describe mechanisms for activation of the Y-family DNA polymerases and integration of TLS with the cell cycle and other elements of the DNA damage response.

THE TLS POLYMERASE SWITCH: ROLES OF RAD18-MEDIATED UBIQUITIN SIGNALING

To minimize the risk of mutagenesis, TLS DNA polymerases must be tightly regulated and employed only when necessary to replicate damaged templates. During S-phase, encounters between leading strand DNA polymerases and bulky DNA lesions (or other hard-to-replicate helix-distorting structures) lead to DNA replication fork stalling, triggering replacement of replicative DNA polymerases with TLS polymerase(s) at the replisome. This ‘polymerase switch’ is stimulated by ubiquitin modification of Proliferating Cell Nuclear Antigen (PCNA, a DNA polymerase processivity factor). In response to DNA replication fork stalling, PCNA is mono-ubiquitinated at lysine residue 164 by the RAD18-RAD6 complex (comprising two molecules of the E3 ubiquitin ligase RAD18 and one molecule of the E2 ubiquitin-conjugating enzyme RAD6) (27,28).

RAD18 is recruited to sites of DNA damage through direct interactions with RPA-coated single-stranded DNA (29). During S-phase, ssDNA can arise via helicase-polymerase uncoupling at stalled replication forks (30). The kinetics of RAD18-mediated PCNA mono-ubiquitination typically coincide with ATR/CHK1 signaling which is also initiated by accumulation of RPA-ssDNA. ssDNA can also be generated outside of S-phase, (for example as an intermediate during NER or BER), explaining how both PCNA-mono-ubiquitination and CHK1 activation may occur independently of DNA replication (6,7,31,32).

The four Y-family TLS DNA polymerases (Polη, Polκ, Polι, REV1) associate preferentially with the mono-ubiquitinated form of PCNA through their Ubiquitin-Binding Zinc Finger (UBZ) and Ubiquitin-Binding Motif (UBM) domains (33). Polη, Polκ, and Polι also contain PCNA-interacting Peptide (PIP) domains that interact with PCNA directly to facilitate replisome-binding (33,34). REV1 lacks a PIP-box and instead binds PCNA via its N-terminal BRCA1 C-terminus (BRCT) domain or polymerase-associated domain (PAD) (35,36,37). PCNA-association of Polη is also facilitated by a direct interaction with RAD18 which helps to chaperone the polymerase to the vicinity of stalled DNA replication forks (38,39).

Different Y-family TLS polymerases have different preferred ‘cognate lesions’ that are bypassed with relative accuracy and efficiency. For example, Polη performs relatively error-free bypass of UV-damaged DNA templates but is error-prone when replicating Benzo[a]pyrene (B[a]P)-adducted DNA (40,41). Conversely, Polκ can bypass B[a]P adducts in a relatively error-free manner (42,43) yet is mutagenic when replicating DNA harboring CPD lesions. Unlike other Y-family enzymes, REV1 cannot synthesize DNA polymers, but adds single C residues to primer termini (44). The main role of REV1 in TLS is non-catalytic, as discussed below. TLS DNA polymerases all lack proofreading exonuclease activity and are viewed as DNA damage tolerance factors. Collectively, recruitment of Y-family TLS DNA Polymerases to mono-ubiquitinated PCNA helps cells to replicate damaged genomes and tolerate diverse forms of genotoxicity. Depending on the nature of DNA damage and the choice of Y-family DNA polymerase(s) deployed, TLS may be error-free or mutagenic (Figure 1).

PCNA-ubiquitination is a trigger for recruitment of all Y-family DNA polymerases to stalled DNA replication forks. It is unclear how the DNA polymerases are selectively recruited in response to their cognate DNA lesions. One possibility is a ‘trial-and-error’ mechanism whereby all TLS DNA polymerases are recruited to mono-ubiquitinated PCNA but only the relevant polymerase will stably engage the stalled replisome. This model implies that TLS DNA polymerases may compete for mono-ubiquitinated PCNA at stalled replication forks. If Y-family DNA polymerases compete for binding mono-ubiquitinated PCNA, altered expression of any individual DNA polymerase in cancer cells may influence DNA polymerase selection, DNA damage tolerance and mutagenesis.

TLS DNA polymerases also have different affinities for PCNA, a feature that may impact selection of TLS polymerases at stalled DNA replication forks (38,45). For example, the Polη PIP box binds PCNA with much higher affinity than the Polκ PIP box (38). Polη is a versatile enzyme that can bypass diverse species of DNA lesions. The high-affinity PIP box may explain why Polη is generally the default TLS polymerase recruited in response to wide variety of DNA lesions.

For some adducts, a single TLS DNA polymerase can perform both nucleotide incorporation opposite the DNA lesion and the subsequent extension from the inserted nucleotide. For example, Polη alone performs both insertion and extension phases of TLS when replicating UV-induced CPD (46,47). However, for many DNA lesions, particularly those that significantly distort the DNA double helix, TLS is mediated by the sequential actions of an ‘inserter’ and then a separate ‘extender’ DNA polymerase. First, a Y-family TLS enzyme (usually Polη, Polι, or Polκ) inserts a nucleotide across a DNA lesion. In the second step, the 4-subunit B-family DNA polymerase Polζ (comprising Rev3/Rev7/PolD2/PolD3) performs extension from the initial nucleotide when inserted across from a diverse array of DNA lesions (48,49). Sometimes, Polζ can act as both the inserter and extender, for example when replicating over thymine glycol lesions generated by free-radicals (50).

The requirement for dual DNA polymerases to perform TLS necessitates a second switch from inserter to extender. The inserter/extender TLS polymerase switch is mediated by REV1. REV1 has very limited DNA polymerase enzymatic activity. Instead, the main role of REV1 in TLS is non-catalytic. Consistent with a non-enzymatic role for REV1 in TLS, a catalytically-inactive human REV1 mutant rescues DNA damage-sensitivity of Δrev1 DT40 cells (51). Moreover, human full length REV1, catalytically-inactive REV1, and REV1 lacking the BRCT correct replication fork progression defects of NQO-treated Δrev1 cells to wild-type levels (52). However, a REV1 mutant lacking the C-terminal domain is inactive for DNA damage tolerance (52). The major role of REV1 in TLS is to facilitate dynamic exchange of DNA polymerase partners with PCNA. The C-terminal of REV1 (REV1-CT) can simultaneously bind the REV7 subunit of Polζ (the extender) and REV1-interacting regions (RIRs) from Polη, Polκ and Polι (inserters) (53,54,55). The POLD3 subunit of Polζ also contains an RIR that interacts with the REV1-CT (56).

Taken together, analysis of TLS polymerase-PCNA association dynamics and structure/function studies suggest that the default inserter Polη interacts with monoubiquitinated PCNA via its UBZ domain and high-affinity PIP motif. Simultaneously REV1 associates with a different monoubiquitinated subunit of the same PCNA trimer via BRCT (57,58), PAD and UBM domains (33). If Polη fails to perform bypass (e.g. due to a non-cognate DNA lesion), a more appropriate Y-family enzyme may substitute, also associating with mono-ubiquitinated PCNA via UBZ/PIP-box interactions in a manner that is facilitated by RIR-REV1-CT interactions. Following insertion, Polζ displaces the inserter RIR/Rev1-CT interaction with the RIR motif of its POLD3 subunit. The REV1-CT interactions with REV7 and PolD3 subunits of Polζ promote an ‘extender’ Rev1/Polζ assembly that is tethered to mono-ubiquitinated PCNA via the BRCT, PAD and UBM domains of REV1 and the PIP motif of PolD3 (56). According to this model, non-enzymatic scaffold functions of REV1 are critically important for survival after DNA damage. Targeting the non-catalytic domains of REV1 may be an effective approach for sensitizing cancer cells to therapy-induced genotoxicity. In particular, the TLS polymerase-interacting C-terminal region represents an appealing target for small molecules that disrupt protein-protein interactions.

RAD18-INDEPENDENT MECHANISMS OF TLS PATHWAY ACTIVATION

RAD6/RAD18 activity accounts for most of the PCNA K164 mono-ubiquitination in mammalian cells and represents the canonical mechanism of TLS activation. However, PCNA can also be monoubiquitinated at K164 by other E3 ligases that are redundant with RAD18 (59). For example, the E3 ubiquitin ligase cullin-4 in complex with the substrate recognition factor CDT2 (CRL4^CDT2, which targets multiple cell cycle-regulated proteins for degradation) mono-ubiquitinates PCNA at K164 to activate mutagenic TLS. Moreover, CDT2-depleted cells are cisplatin-sensitive, consistent with a role for CRL4^CDT2-mediated PCNA ubiquitination in DNA damage tolerance (59). In some cell lines, CDT2 is dispensable for PCNA mono-ubiquitination following UV treatment, suggesting a minor role of CDT2 in TLS when compared with RAD18 (59). The RAD5 homologue HLTF also plays a back-up role in mediating PCNA mono-ubiquitination when RAD18 is absent (60).

TLS pathway activation can also occur in the complete absence of PCNA K164 mono-ubiquitination. For example, PCNA ubiquitination does not disrupt Polδ-PCNA interactions or enhance interaction of TLS DNA polymerases with PCNA (61). It has been suggested that K164 mono-ubiquitination displaces putative inhibitors of PCNA–TLS Polymerase interactions (61). A UBZ-deficient Polη mutant retaining the PCNA-interacting Peptide (‘PIP’ domain) corrects UV-sensitivity defects of XPV cells, again suggesting that PCNA mono-ubiquitination is dispensable for TLS polymerase activation (62).

On the other hand, mouse embryonic fibroblasts (MEFs) harboring a PCNA^K164R/K164R mutant show reduced colony-forming ability after UV irradiation when compared to wild-type cells, indicating that TLS is defective when PCNA cannot be mono-ubiquitinated (63). However, knockdown of Rev3l, Rev1 and Polh in Pcna^K164R/K164R cells results in additive UV-sensitivity, suggesting existence of a secondary PCNA-ubiquitination independent TLS pathway (63). Polκ is the main TLS polymerase responsible for bypassing DNA lesions induced by the alkylating agent methyl methanesulfonate (MMS). Double mutant Pcna^K164R Polk⁻^/⁻ MEFs show greater sensitivity to MMS treatment when compared with MEFs harboring individual Pcna^K164R or Polk⁻^/⁻ mutations. Therefore, Polκ functions in a PCNA-mono-ubiquitination independent pathway for tolerance of MMS-induced DNA damage (64). Double mutant Pcna^K164R Polη^+/⁻ mouse pre-B cells show greater sensitivity to UV and cisplatin treatment than either single mutant alone, again suggesting a PCNA-ubiquitination independent role of Polη in TLS (65). Taken together, PCNA mono-ubiquitination-independent mechanisms contribute to TLS under some circumstances.

REGULATED EXPRESSION AND ACTIVITY OF TLS FACTORS

Genome maintenance factors are often regulated at the level of expression, and via post-translational modifications that control subcellular distribution, stability, activity, and interactions with binding partners. Such regulatory mechanisms are often important for integrating genome maintenance with the cell cycle. For example, HR is a relatively accurate DSB repair mechanism when compared with NHEJ. However, HR must be restricted to S-phase and G2 when DNA has been replicated and an intact sister chromatid is available as template for repair. Cyclin-dependent kinases (CDKs) play an important role in the HR versus NHEJ pathway choice, particularly by controlling DSB resection, which is required for HR but not NHEJ. Many of the DNA DSB resection and repair factors involved in HR and NHEJ are subject to phosphorylation by CDKs to restrict error-prone NHEJ to G1 and promote relatively error-free HR in S/G2 (66,67,68,69,70,71,72,73).

Transcriptional regulation of resection factors also coordinates DSB repair pathway choice with the cell cycle (74). Cyclins and CDKs are highly dysregulated in neoplastic cells and likely affect DNA repair pathways that impact tumorigenesis and cancer therapy. By comparison with DSB repair factors, little is known regarding transcriptional and post-translational control of TLS proteins, although existence of such regulatory mechanisms seems likely.

Cell cycle-dependent regulation at the level of mRNA and protein expression is reported for REV1 and RAD18. In fission yeast, Rev1 protein levels peak during G1 and decrease during the G1/S transition in a proteasome- dependent manner (75). In contrast, in budding yeast Saccharomyces cerevisiae, Rev1 protein levels are low in G1 and early S phase, increase in late S-phase and peak during G2/M (76). REV1 mRNA levels only change 3-fold throughout the cell cycle whereas Rev1 protein levels are 50-fold higher in G2/M than in G1, suggesting that Rev1 protein is regulated post-transcriptionally (76). DNA damage by UV exposure does not significantly alter Rev1 protein expression (76). Rev1 protein expression is relatively low in S-phase, perhaps to limit mutagenic TLS during DNA replication. Rev1 protein expression peaking during G2/M suggests that TLS plays a role outside of active DNA replication in S-phase. Expression of RAD18 is also cell cycle regulated. In synchronized and untransformed human fibroblasts, RAD18 protein levels are high in S-phase when compared with G0 (quiescent) and G1 cells (6). RAD18 expression is under transcriptional control of E2F3 (77), possibly explaining the elevated expression of RAD18 in S-phase when RB is phosphorylated and E2F proteins are de-repressed. Loss of RB and excess E2F activity are common in cancer and may lead to elevated RAD18 expression. As discussed later, RAD18 mRNA levels are typically higher in cancer when compared with normal tissue. The RAD18 protein can also be stabilized and overexpressed due to its cancer-specific binding partner, the Cancer/Testes Antigen MAGEA4 (78). Over-production of RAD18 in cultured cells can lead to increased PCNA mono-ubiquitination and DNA damage-independent recruitment of Y-family TLS polymerases to chromatin (79,80). Therefore, high-level expression of RAD18 in cancer (via E2F activity, MAGEA4-binding or other processes) represents a potential mechanism for aberrant TLS activation and mutagenic DNA synthesis. The POLH promoter contains a p53-response element (81) and DNA damage-inducible expression of Polη is p53-mediated in several human cell lines (82). Importantly, p53-inducible Polη expression contributes to accelerated bypass of UV-lesions (83).

Integration of TLS factors with the cell cycle and other branches of the DDR is also achieved via phosphorylation-based mechanisms. For example, basal and DNA damage-inducible interaction between RAD18 and Polη is dependent upon RAD18 phosphorylation in the Polη-interacting domain by S-phase kinase CDC7 (84). CDC7 is both a biomarker and drug target in cancer (85,86). Therefore, it is likely that CDC7-mediated RAD18 phosphorylation contributes to TLS pathway activation in tumor cells.

The Polη-interacting domain of RAD18 is also phosphorylated in a DNA damage-inducible manner by c-Jun N-terminal kinase (JNK) to facilitate TLS (87). JNK controls many adaptive responses to both intracellular stresses and environmental exposures including UV light (88). Therefore, RAD18 phosphorylation by JNK might integrate TLS with other stress response pathways. During an unperturbed cell cycle, the RAD18-associated E2 ubiquitin-conjugating enzyme RAD6 is also phosphorylated by CDKs, leading to an increase in its catalytic activity (89). In synchronized cells, RAD6 phosphorylation peaks in G2/M. The extent to which TLS is affected by CDK-induced RAD6 phosphorylation has not been determined. However, RAD6 phosphorylation is a possible mechanism for promoting TLS after completion of S-phase.

Polη is phosphorylated by ATR and Protein Kinase C in response to UV and therapeutic agents (90). The Polη site targeted by ATR (Ser 601) is necessary for normal recovery of DNA replication and DNA damage tolerance after UV-irradiation (91). Polη is also phosphorylated at Ser 687 by CDK2 in a cell cycle-dependent manner. Ser 687 phosphorylation regulates Polη stability during the cell cycle, allowing accumulation in late S and G2 (92).

Whether Polι and Polκ are regulated via direct phosphorylation has not been determined. Nevertheless, multiple core components of the TLS pathway (RAD18, REV1, RAD6, Polη) are directly phosphorylated by kinases whose activities are altered in cancer (CDKs, CDC7, ATR, JNK). Protein phosphorylation represents a potential mechanism for creating TLS pathway imbalances that contribute to mutability and altered DNA damage tolerance of cancer cells.

TLS OCCURS BOTH AT STALLED LEADING DNA REPLICATION FORK AND POST-REPLICATIVELY

TLS may directly alleviate replication blockades on stalled leading strands (termed TLS ‘on-the-fly’), and may operate post-replicatively (i.e. behind an active leading strand) to eliminate daughter strand discontinuities.

TLS on-the-fly

Seminal work by Lehmann and colleagues revealed that the conversion of the low molecular weight nascent DNAs synthesized in UV-irradiated cells into high-molecular-weight DNA is compromised in XPV cells (93,94). The term post-replication repair (PRR) was used to describe growth of low molecular weight DNA to high molecular weight species. Hanaoka's work eventually revealed that Polη-mediated TLS accounted for PRR (11,12). Subsequent work from many groups has shown that Polη contributes to replication fork progression in cells harboring CPD and other lesions (95,96,97). Polη travels with the replication fork in unperturbed cells (98), is recruited to the vicinity of replication forks in UV-irradiated cells, and resolves helicase/polymerase uncoupling (99). In summary, for UV-induced CPD, on-the-fly direct lesion bypass by Polη is the default response. In the absence of Polη-mediated TLS, other compensatory mechanisms are deployed to sustain DNA synthesis.

TLS at post-replicative ssDNA gaps

It has long been recognized that UV-irradiation leads to ssDNA-gapped daughter strand DNA on both leading and lagging strands (100,101). Potentially such gaps may result from persistence of Okazaki fragments on the discontinuous lagging strand, and/or due to repriming downstream of a blocked leading strand. Two excellent recent reviews provide a historical perspective on mechanisms of ssDNA gap formation and repair (102,103).

TLS is an important mechanism for filling post-replicative ssDNA gaps. Lopes et al. analyzed DNA replication dynamics of TLS-compromised S. cerevisiae mutant strains following UV-irradiation (104). Those studies showed that strains lacking rev1, rev3 and rad30 (yeast genes encoding homologues of human REV1, REV3 and Polη respectively) maintain normal rates of fork progression following UV-treatment. However, TLS mutant strains accumulate more ssDNA gaps along replicated duplexes. It was concluded that post-replicative gaps accumulate along replicated daughter strand DNA due to repriming events downstream of the lesions on both leading and lagging strands. Furthermore, TLS (together with HR) counteract gap accumulation without affecting replication fork progression (104). Karras and Jentsch showed that restricting expression of rev3, rad30 or rad18 to G2/ M-phase is sufficient to confer lesion tolerance, further supporting the notion that TLS acts on ssDNA gaps behind newly re-primed replication forks (9). In a related study, Daigaku et al. directly visualized and quantified PRR tracts after UV-irradiation in TLS-mutant yeast strains. Those workers showed that TLS is temporally and spatially separable from global genomic DNA replication (10). Similar PRR roles have been proposed for TLS in human cells (95,105,106). Notably, UVC-irradiated XPV fibroblasts accumulate longer ssDNA stretches both at the replication fork, and behind the leading strand when compared with isogenic Polη-complemented cells (99). Therefore, Polη resolves helicase/polymerase uncoupling and also prevents post-replicative gap accumulation in cells harboring CPD.

The type of DNA damage might determine whether TLS occurs directly at or behind the stalled replication fork (95). For example, repriming may be favored when TLS is absent (99), when template switching is suppressed (107) or when replication fork obstacles are too bulky to be accommodated by DNA polymerases (103).

Insight into a re-priming/restart process that allows continued elongation of nascent daughter strands copying UV-damaged templates was revealed by the discovery of Primase-Polymerase (PRIMPOL; CCDC111). PRIMPOL is an archaeal-eukaryotic primase (AEP) in eukaryotic cells that performs lesion bypass and additionally re-primes DNA replication downstream of bulky lesions and other barriers to DNA synthesis (108,109,110). PRIMPOL is recruited to ssDNA via its interaction with RPA. PRIMPOL does not interact with PCNA, yet contains two RPA-binding motifs, one of which mediates recruitment to stalled replication forks. RPA also stimulates PRIMPOL primase activity (110,111). Therefore, direct associations of PRIMPOL with RPA-coated ssDNA at stalled DNA replication forks may stimulate restart of DNA synthesis and generate gaps behind the newly-reprimed daughter strand. PRIMPOL-mediated repriming activity provides a possible explanation for the presence of post-replicative ssDNA gaps that are filled by the Y-family TLS polymerases (112). Consistent with this idea, PRIMPOL depletion impedes fork progression to a higher extent in XPV fibroblasts when compared with isogenic POLH-complemented cells, and suppresses formation of post-replicative ssDNA gaps (99). Interestingly, PRIMPOL primase catalytic activity is dispensable for generation of ssDNA gaps in UV-irradiated XPV cells (99). Moreover, PRIMPOL is not the sole mediator of repriming when Polη is absent (99). RAD51 may also contribute to repriming downstream of stalled replication forks, independently of its roles in recombination-based mechanisms (99).

Post-replicative ssDNA gaps can also be filled by an error-free DNA damage-avoidance mechanism termed ‘template switching’ (TS). TS relies on DNA replication fork reversal to generate ‘chicken-foot structures’ and uses the newly synthesized strand as a template to replicate across damaged DNA and suppress discontinuities (113,114,115). TS is initiated by HLTF and SHPRH, mammalian homologues of the yeast E3 ubiquitin ligase and SWI/SNF ATPase family member RAD5 (116). HLTF and SHPRH recruit the ubiquitin-conjugating enzyme UBC13 and a non-canonical UBC variant MMS2 to damaged chromatin, to form a complex that cooperates with RAD6–RAD18 to poly-ubiquitinate PCNA at K164 (27). The two RAD5 homologues poly-ubiquitinate PCNA via distinct mechanisms. SHPRH mediates extension of mono-ubiquitinated K164 (117), while HLTF forms a thiol-linked Ub chain on UBC13 that is transferred to RAD6. RAD18 then transfers the pre-conjugated Ub chain to K164 of unmodified PCNA (118). HLTF and SHPRH promote TS in a DNA damage-specific manner (60). SHPRH mediates responses to alkylating agents (e.g. MMS) whereas HLTF mediates PCNA poly-ubiquitination and confers DNA damage tolerance in response to bulky DNA lesions (including UV, 4-NQO and MMC) (119,120). Poly-ubiquitinated PCNA generated via the concerted actions of RAD18 and RAD5 provides a platform for recruiting ZRANB3 (Zn finger, RAN-binding domain containing 3, also known as Annealing Helicase 2 or AH2) (121). Once recruited to the stalled fork, ZRANB3 facilitates fork regression, replication fork restart and confers DNA damage tolerance (92–94). TS is intimately coordinated with the TLS pathway. For example, in response to UV-irradiation, HLTF promotes PCNA mono-ubiquitination and Polη recruitment, while inhibiting SHPRH function. Conversely, MMS promotes formation of a RAD18-SHPRH complex and induces HLTF degradation (60). It is unclear how cells choose between error-prone TLS and error-free TS. It is possible that TS is deployed for DNA damage that is too severe to be bypassed via TLS and results in persistence of 3′ ends at stalled DNA replication forks (122). Like TLS, the TS pathway in S cerevisiae is also functional when RAD5 is restricted to G2. Therefore, template switching may also play a role in ssDNA gap maintenance in G2 phase (9). Figure 2 illustrates the canonical mechanisms of PRR gap-filling via TS and TLS.

Figure 2. — Post-Replication Repair (PRR) is mediated by TS and TLS. (A) Stalling of replicative DNA polymerases leads to recruitment of the primase/polymerase PRIMPOL and re-initiation of DNA synthesis downstream (3′) of the stalled DNA polymerase. PRIMPOL-mediated restart of DNA synthesis generates ssDNA gaps or discontinuities in the daughter strand. (B) TS is activated by PCNA poly-ubiquitination, and is dependent upon RAD5 homologues HLTF and SHPRH. (C) TLS is mediated by PCNA K164 mono-ubiquitination, which recruits Y-family TLS polymerases in a process termed ‘polymerase switching’. The RAD6/RAD18 (E2/E3 ubiquitin ligase) complex accounts for most (∼95%) of the mono-ubiquitinated PCNA in mammalian cells.

Tirman et al. showed that in cisplatin-treated U2OS cells, TLS factors are involved in temporally distinct mechanisms of gap filling: during G2 phase, ssDNA gap filling is dependent upon RAD18-mediated PCNA mono-ubiquitination and subsequent recruitment of REV1-Polζ (8). However, gap-filling during S-phase is RAD18-independent and requires RAD51, UBC13 and REV1-Polζ, (8). In avian DT40 cells, RAD18 and PCNA ubiquitination, but not REV1, are essential for post-replicative gap filling after UV exposure (52). Like cisplatin-treated U2OS cells, REV1 (but not RAD18 or mono-ubiquitinated PCNA) is required to maintain replication fork progression following DNA damage. In DT40 cells, mutation of REV1, but not RAD18 or PCNA, results in more frequent fork stalling after UV or NQO exposure (52), perhaps reflecting a role of REV1 in template-switching during S-phase. Taken together, these studies underscore the complexity of TLS and demonstrate temporally distinct mechanisms of gap-filling involving different mediators.

The specific factors involved in gap filling, and whether gap filling involves canonical TLS or TS (as may be the case for REV1/Polζ) may depend on multiple factors such as cell type, the species of DNA damage, and availability of other DDR pathways (103). In this regard, oncogenes and the intrinsic replication stresses of neoplastic cells are emerging as key causes of ssDNA gaps that require remediation via the TLS pathway. Yang et al. showed that RAD18 and Polκ are important for preventing both accumulation of ssDNA gaps and mitotic catastrophe in primary cells expressing CCNE1 or KRAS (123). In a related study, Nayak et al. showed that pharmacological inhibition of REV1 interferes with DNA replication and induces ssDNA gap formation in cancer cells (124). HR-deficient cancer cells may be particularly dependent on TLS for gap-filling (125,126). For example in BRCA1 mutant breast cancer cells, spontaneous ssDNA gaps arising from PRIMPOL-mediated repriming are filled by REV1 and Polζ-dependent TLS (127). It is important to determine precisely how global DNA repair status as well as other factors impact TLS-dependency of cancer cells. This knowledge will enable precision medicine strategies for targeting cancer cells using TLS inhibitors based on their unique genome maintenance characteristics. A more detailed discussion of mechanisms of gap formation may be found in two excellent reviews by Menck and Vindigni (102,103).

EVIDENCE THAT TLS POLYMERASES CONTRIBUTE TO CANCER

Since mutagenesis is a hallmark of cancer cells and drives carcinogenesis, it is intuitive to hypothesize that error-prone TLS polymerases promote cancer. Since cancer cells rely on TLS for ssDNA gap suppression and tolerance of intrinsic replication stresses (123,124) it is also tempting to hypothesize that TLS sustains carcinogenesis. Here we review evidence that TLS polymerases shape cancer genomes. We describe how TLS polymerases and their regulators are altered in cancer, and we summarize experimental evidence that TLS drives carcinogenesis.

Contributions of TLS to cancer based on analysis of mutation signatures

Each cancer cell harbors a composite ‘mutational portrait’ comprising many individual mutations generated by various error-prone genome maintenance processes (128). The ‘mutation signature’ concept provides a convenient way to analyze large cancer genome sequence data sets and annotate different classes of superimposed mutations in tumors. Studies of multiple cancer types have revealed over 30 single-base substitution (SBS) mutation signatures. Classification of SBS signatures is based upon relative frequencies of the six base substitutions (C>A, C>G, C>T, T>A, T>C and T>G) in tri-nucleotide sequence settings that consider every possible combination of flanking 5′ and 3′ nucleotides. Additionally, methods have been developed for classification of doublet-base substitutions (DBSs) and insertion/deletion mutations (indels): Indels are classified as deletions or insertions. Single-based indels are classified based upon the length of the mononucleotide repeat tract in which they occur. Longer indels are classified based on whether they reside at repeats or with overlapping microhomology at deletion boundaries, and based upon indel-size, repeat and microhomology (128). The etiologies of many mutational signatures have been attributed to specific genotoxic exposures and mutagenic processes. It is proposed that TLS polymerases mediate four of the SBS signatures, namely signatures 2, 5, 13 and 9.

Mutational signature 13, found in breast and bladder cancers, is attributed to the concerted activities of APOBEC (apolipoprotein B mRNA editing enzyme, catalytic) and REV1. APOBEC proteins deaminate Cytosine to Uracil. Subsequent excision of Uracils formed via cytosine deamination generates non-coding AP sites. REV1 is postulated to insert cytosines across the AP sites that lead to C>T or C>G mutations (129,130,131). A recent study experimentally validated the role of REV1 in APOBEC3-mediated mutagenesis. Petljak et al. annotated mutation signatures in human cancer cells that accumulate APOBEC3-associated mutations over time (131). Using REV1-knockout derivative lines, these workers demonstrated that REV1 has a critical role in the generation of signatures SBS2 and SBS13, and might also contribute to a low-fidelity TLS process underlying SBS5 (131). Therefore, there is strong evidence that REV1 contributes to mutagenesis in human tumors.

The other Y-family TLS polymerase strongly implicated in generating mutation signatures in human cancers is Polη. Mutation signature 9, found in Chronic Lymphocytic Leukemias (CLL) and malignant B-cell lymphomas, is attributed to Polη-dependent somatic hypermutation (SHM) activity. Consistent with a role for Polη in mediating signature 9, Polh⁻^/− mice have a GC-biased mutation profile demonstrating that Polη is a major contributor to A/T mutations during SHM (132). Supek and Lehner identified a prevalent mutational signature in solid tumors that is directed toward the 3′ end of active chromatin, associated with carcinogen exposure, and attributed to Polη (133). However, there has been no direct experimental validation that signature 9 is dependent upon Polη. There remains a need for experiments that test how specific TLS polymerase deficiencies impact mutational signatures of cancer genomes.

Altered expression of TLS polymerases and their regulators in cancer

There are several reports that TLS polymerases and their regulators are altered in cancer cells or patient-derived tumors. Polymorphisms in TLS polymerase genes are documented in a small set of studies and these alterations may be associated with cancer predisposition. For example, mutations in REV1 are associated with lung cancer (134), POLK mutations are linked with lung and breast cancer (134,135), and POLI mutations correlate with risk of adenocarcinoma and squamous cell carcinoma (136). Whether such TLS polymerase mutations necessarily affect their biochemical activities and participation in TLS, or threaten genome stability is not clear. However, some-cancer associated REV1 mutants reportedly have altered DNA polymerase and DNA-binding activities that lead to increased mutagenesis (137).

Altered expression of TLS polymerases represents another possible mechanism for TLS imbalances and mutagenesis. Potentially, over-expressed or aberrantly activated TLS polymerases might compete with replicative DNA polymerases for access to the replisome leading to mutagenesis (138). Improperly-expressed (both over-expressed and under-expressed) TLS polymerases may also compete with each other for replisome association via their PIP boxes and ubiquitin-binding motifs. Elevated expression of individual TLS polymerases may increase error-prone DNA synthesis on an undamaged template or might lead to aberrant deployment for bypass of non-cognate DNA lesions, leading to increased mutagenesis. For example, overexpression of POLK is reported to increase N-methyl-N-nitrosurea (MNU)-induced mutagenesis and occurrence of intestinal adenomas (139).

Pathological changes in TLS modulators can also lead to XPV-like imbalances that impact genome stability. The E3 ubiquitin ligase MDM2 which is overexpressed in many cancers degrades Polη and recapitulates hallmark TLS defects of XPV cells (140). The NPM1 gene, which encodes the Polη-binding protein Nucleophosmin, is frequently mutated in Acute myeloid leukemia (AML). A prevalent NPM1 gene mutation in AML patients leads to excessive Polη degradation and reduces error-free TLS activity (141).

The proximal TLS activator RAD18 is often overexpressed in cancer both at the mRNA level (Figure 3) and at the protein level, owing to its stabilization by a cancer selective binding partner Melanoma Antigen A4 (MAGEA4, a Cancer/Testes Antigen or CTA) (78). TLS polymerase activation and mutagenesis are highly sensitive to changes in RAD18 expression levels. Ectopically-expressed RAD18 induces PCNA mono-ubiquitination and recruitment of TLS polymerases to replicating DNA, even in the absence of DNA damage (78,79,80). DNA damage-independent activation of TLS polymerases due to excessive RAD18 could be a source of mutagenesis. Different TLS polymerases have different affinity for PCNA (45) and may be differentially reliant on RAD18-mediated PCNA mono-ubiquitination or chaperone activity for engaging the replisome (38). Excessive RAD18-mediated chaperone activity or PCNA-mono-ubiquitination favoring any individual TLS polymerase could create a selection bias and TLS imbalance that contributes to mutagenesis in cancer cells. The E3 ubiquitin ligase RNF168 is another potential TLS modulator which is often overexpressed in cancer (142). RNF168 increases levels of ubiquitinated H2A in the vicinity of replicating DNA which recruits Polη via direct interactions (143).

However, there is no single paradigm for how TLS polymerases (or other TLS pathway components) are altered in cancer and whether such changes correlate with mutation burden or other tumorigenic properties. Therefore, we interrogated Cancer Genome Atlas (TCGA) gene expression datasets and determined the extent to which TLS pathway gene expression levels are altered in representative cancers. Additionally, we asked whether expression levels of TLS pathway genes correlate with mutation load. We analyzed expression levels of RAD18 (upstream activator of Y-family TLS polymerases), POLH, POLK, POLI and REV1 (the four Y-family TLS polymerases), and REV3L and MAD2L2 (encoding the two subunits of Polζ, which mediates the extension phase of TLS).

We examined data from three different cancer types with a smoking-related etiology: Bladder cancer (BLCA), Lung Adenocarcinoma (LUAD), and Lung squamous cell carcinoma (LUSC). We reasoned that tobacco smoke genotoxicity may stimulate mutagenesis and reveal associations with TLS status for these cancers. For comparison, we also examined TCGA data from tumors whose etiology is unrelated to tobacco smoke exposure, namely prostate cancer (PRAD), glioblastoma (GBM), pancreatic cancer (PAAD) and endometrial cancer (UCEC).

Figure 3 and Supplementary Figure S1 summarize expression patterns of TLS genes in tumor samples relative to adjacent normal tissue. Also indicated are correlations between TLS gene expression and total Single Nucleotide Variation (SNV) burden (which is a direct measure of genome-wide point mutations). Several key patterns and trends are revealed by our analyses: Of the TLS genes studied here, only RAD18 is overexpressed in most tumor types. The only tumor not significantly overexpressing RAD18 is pancreatic cancer (for which datasets were available from only 4 normal samples, limiting our ability to accurately determine fold changes in tumor samples). Interestingly, RAD18 expression shows a strong positive correlation with the total SNV numbers in most cancer types, both in smokers and non-smokers. The overexpression of RAD18 in most tumors is consistent with the hypothesis that excessive RAD18 stimulates error-prone TLS and increases mutation burden. MAD2L2 (encoding REV7) is another TLS gene that is also overexpressed in most tumors. However, unlike RAD18, MAD2L2 expression does not show a strong correlation with SNV burden. REV3L is typically expressed at low levels in tumor cells when compared with adjacent normal tissues. Moreover, in some tumors (e.g. PAAD, PRAD), REV3L expression is negatively correlated with SNVs. Rev3l loss in mice leads to spontaneous tumorigenesis (144). Therefore, it is possible that the reduced REV3L expression in human tumors contributes to tumorigenicity.

There are reports that the Y-family polymerases are expressed at higher levels in tumors than normal cells and tissues (145,146,147,148). Surprisingly however, our expression analyses reveal that POLH, POLK, POLI and REV1 expression levels in cancers are typically equivalent to or reduced when compared with adjacent normal tissues. In LUAD, the expression levels of Y-family DNA polymerases are negatively correlated with the total SNV counts in tumors from smokers. In contrast, in PAAD, the general trend is that tumors with reduced expression of POLH, POLK, POLI and REV1 have a high number of SNV counts.

Taken together, these analyses reveal remarkable complexity in expression patterns of the core TLS pathway genes and how they are altered in different cancers. With the important caveat that mRNA expression is an imperfect surrogate for protein level and activity, the expression patterns described in Figure 3 suggest many ways in which imbalance between TLS polymerases and their activators might affect DNA damage tolerance and mutability in different cancers. The impact of such TLS pathway alterations is also likely to depend on the availability of other genome maintenance mechanisms. For example, HR-compromised cells have increased reliance on TLS (127). Accordingly, altered expression of TLS genes might be far more consequential in a BRCA-mutant cancer cell than in a BRCA-sufficient background. There is a need to determine the ways in which TLS and its intersecting DNA repair pathways are rewired in all individual tumor settings, and how those changes impact mutagenesis and DNA damage tolerance. For example, it is reported that HLTF promoter methylation and loss of HLTF expression are observed in some cancers (149). Changes in relative expression or activities of RAD5 homologues could affect the selection of (error-free) TS vs (error-prone) TLS and influence mutagenic outcomes. To investigate potential TS pathway alterations in cancer, we also analyzed expression patterns of HLTF, ZRANB3, SHPRH, MMS2 and UBC13 for all the malignancies described in Figure 3. In contrast with reported findings (149), HLTF expression was not reduced in tumors. Interestingly, we observed several trends: HLTF was overexpressed in tumors and positively correlated with SNVs in most cancer types. However, SHPRH was downregulated in tumors of most cancer types.

It is important to recognize that cancer constitutes many different diseases in which tumor types and subtypes are highly divergent based on proteogenomic characteristics. It is overly-simplistic and imprecise to conclude that any specific TLS factor is universally over- or under-expressed in cancer when compared to normal tissues. The analyses and a corresponding heatmap in Figure 3 illustrate that even despite general trends, every individual tumor is unique and expresses a distinct repertoire of TLS factors (as well as other genome maintenance genes). The goal of targeting TLS for therapy may require a precision medicine approach which is contingent on understanding the unique DNA repair characteristics of every individual patient tumor. The correlations we have identified between TLS factors and SNV burden in cancer provide an essential source of hypothesis. The results of Figure 3 could help prompt future studies to model pathologically-relevant TLS polymerase imbalances and test the impact of those imbalances on mutagenesis.

TLS is a dependency of some cancer cells

During multistep tumorigenesis, neoplastic cells acquire extensive DNA damage from intrinsic sources including oncogenes. Oncogene signaling leads to DNA damage via multiple mechanisms including: (a) dysregulated replication origin licensing and firing leading to under- or over-replicated DNA (b) increased transcription factor activation leading to replication-transcription conflicts (RTCs) (c) altered nucleotide metabolism leading to reduced dNTP pools and (d) increased ROS production causing oxidative DNA damage (150,151,152). Oncogenes may also deregulate R-loop homeostasis which further leads to RTCs (153). Taken together, oncogene signaling impedes replication fork advancement via multiple mechanisms and causes DNA replication stress.

Accumulating evidence suggests that TLS averts formation of lethal DNA damage in cells experiencing various forms of oncogene-induced DNA replication stress. TLS can be error-prone on undamaged DNA or templates containing non-cognate lesions. Therefore, it is possible that error-prone TLS sustains tumorigenesis by promoting both DNA damage tolerance and mutagenesis. Here we discuss key evidence that supports a role of TLS in cancer cell survival.

A cancer-essential gene (also termed a cancer-dependent essential gene) is required for proliferation or survival of cancer cells (but not normal cells) (154). Several reports suggest that TLS factors fulfill criteria for being genetic dependencies of cancer cells: TLS facilitates continued replication in cells challenged by oncogene-induced replication stress (123), prevents accumulation of ssDNA gaps and promotes cell survival (124). Inhibition of TLS by a small molecule, JH-RE-06 (which targets REV1) reduces survival of several cancer cell lines even in absence of an external stress (124). JH-RE-06 also suppresses growth of A549 and H1299 lung cancer cells grown as xenografts in vivo (155). Further consistent with a role for TLS in sustaining tumors, RAD18 overexpression elevates colonization of esophageal cancer cells in the mice lung (156), while RAD18-deficient TNBC xenografts show reduced tumor volume (157). Taken together, these studies suggest that TLS is a cancer-dependency at least under select conditions. TLS limits fork slowing by restricting fork reversal without altering replication restart or dormant origin firing through its gap filling function (124). Recent studies suggest that the cancer cell reliance on TLS for survival is contingent on ssDNA gap suppression (124,158).

Not all cancer cells rely on TLS for viability (124). According to the cancer dependency map (DepMap, a database of loss-of-function CRISPR screens in large number of cancer cell lines), REV1 is a dependency in only few cell lines (CRISPR 16/1086; RNAi 0/597). Similarly, other TLS polymerases are rarely identified as dependencies in DepMap. It remains to be determined what factor(s) determine whether TLS is a cancer dependency. Different oncogenes might induce distinct species and levels of DNA damage that are differentially reliant on TLS for remediation. For example, ectopic expression of CCNE1 in primary untransformed human fibroblasts induces a more robust expression of DDR markers (including mono-ubiquitinated PCNA) when compared with ectopically-expressed oncogenic RAS (123). Moreover, multiple DNA repair pathways (including HR, TMEJ, NHEJ) contribute to tolerance of oncogene-induced replication stress and DNA damage (123,152). Every cancer cell has a distinct DNA repair landscape. Therefore, the repertoire of redundant pathways available to repair intrinsic DNA damage might determine the extent to which any cancer cell depends on TLS.

Defining the factors and biomarkers that accurately predict TLS-dependency of cancer cells is critical if we are to develop precision medicine approaches based upon TLS inhibition. One factor that has been shown to create TLS-dependency in cancer cells is loss of BRCA1. Homologous recombination-deficient BRCA1 mutant breast cancer cells develop spontaneous ssDNA gaps which are repaired via PRIMPOL and REV1-Polζ-dependent gap-filling mechanism required for viability (127). Tutt and colleagues showed that cancer cells mis-expressing the germ cell protein HORMAD1 (a Cancer Testes Antigen or CTA) have increased dependency on several TLS mediators including POLH, POLK, REV1, REV3L and REV7 (159). Functions of HORMAD1 in cancer cells are not well understood. It is reported that HORMAD1 can both inhibit HR (160) and activate HR (161,162) when mis-expressed in cancer cells. Nevertheless, aberrant expression of HORMAD1 could generate species of DNA damage that are repaired via TLS. More work is needed to reveal the mechanistic underpinnings of TLS-dependencies in neoplastic cells. The availability of the REV1 inhibitor JH-RE-06 has been a valuable tool for demonstrating that some cancer cell lines rely on TLS for survival. The use of orthogonal methods such as CRISPR and RNAi to ablate REV1 (and other TLS genes) will help validate the TLS pathway as a cancer dependency. It is also crucial to test the TLS-dependency of cancer cells in pathologically-relevant models (including GEMM, PDX, organoids) that better recapitulate the characteristics and dependencies of tumors in patients.

Contributions of TLS to cancer based on mouse models

Mouse models are important for validating hypothetical roles of oncogenic drivers or tumor suppressors in carcinogenesis in vivo. Here, we consider the evidence that TLS polymerases and RAD18 are physiologically-relevant modifiers of spontaneous or induced carcinogenesis in vivo.

Spontaneous tumorigenesis

Deletion of the TLS activator Rad18 in mice does not affect rates of spontaneous mutagenesis or incidence of spontaneous tumors (163,164). Polk-deficiency elevates mutagenesis at G:C base pairs in the mouse lung, liver and kidney (165), while Rev1-deficient mice have reduced weight and lifespan (166,167). However, loss of Rad18 (164), TLS polymerases Polh (168), Poli (168,169), Polk (165), Rev1 (166)) or Rev7/Mad2l2 (170) does not affect spontaneous tumorigenesis in mice. Intriguingly, Rev3l-deficient mice suffer from embryonic lethality (171,172,173) and conditional deletion of Rev3l in adult epidermal or lymphoid cells leads to spontaneous squamous cell carcinomas in specialized sebaceous glands and skins, and lymphomas, respectively (144,174). Although the cancer-propensity of Rev3L-deficient mice may point to a tumor-suppressive role for Polζ, this phenotype is not seen in Rev7-deficient mice. Therefore, Rev3L may have additional Rev7-independent functions in tumor-suppression, possibly unrelated to TLS. Unfortunately, in vivo experiments assessing the tumorigenic consequences of TLS imbalance due to overexpressed TLS polymerases are scarce. In a one-of-a-kind experiment, Sasatani et al. did not observe spontaneous tumorigenesis in the intestine of Rev1-overexpressing mice (139). Collectively, the studies described above suggest that altered expression of TLS polymerases and Rad18, under unperturbed conditions is not oncogenic in mice.

Oncogene-induced tumorigenesis

Although TLS is implicated in mediating tolerance of oncogene-induced DNA replication stress, there is no published information regarding the requirement for TLS in oncogene-induced tumorigenesis in vivo. In our unpublished study, Rad18-defciency did not affect Kras^G12D-driven lung carcinogenesis in either p53^+/+ or p53⁻^/⁻ C57BL6 mice. This result could indicate that TLS is dispensable for oncogenic Kras-driven lung carcinogenesis, or that Rad18-independent TLS mechanisms are sufficient to sustain Kras^G12D-driven lung tumors. Rad18 loss also did not affect the mutation burden of Kras-driven lung tumors in p53-null mice. RAD18 is typically overexpressed (not ablated) in human lung tumors. It would also be interesting to determine whether Rad18 overexpression in the mouse lung affects rates of oncogene-induced lung carcinogenesis.

Orthotopic tumor models

Implanting cancer cells orthotopically in mice provides a convenient approach to study tumor growth in a physiologically relevant environment. Orthotopic injection of esophageal cancer cells overexpressing RAD18 increases colonization (156), while RAD18-deficient TNBC cell xenografts show reduced tumor growth (157). TLS inhibition by the REV1 inhibitor JH-RE-06 decreases colonization of lung cancer cells in mice (155). Therefore, studies with orthotopically-implanted cells support a role for TLS in tumor growth and as a cancer target.

Carcinogen-induced tumorigenesis

Polh-/- mice fully phenocopy the UV-induced skin cancer-propensity of XP-V patients. Heterozygous Polh^+/− mice are also sensitive to UV-induced skin carcinogenesis, but succumb at a lower rate when compared with Polh^-/- (175). Although Poli^-/- mice are not susceptible to UV-induced skin carcinogenesis; Poli^−/− Polh^−/− double knockout mice have increased rates of UV radiation-induced skin carcinogenesis, compared with Polh^-/- animals (168,169). Therefore, both Polη and Polι guard against UV-induced carcinogenesis. Interestingly though, UVB-treatment of Poli^−/− deficient animals induces mesenchymal tumors that are not observed in Polh^−/− mice (169). Polk^−/− mice are viable, but have a shorter lifespan than Polk^+/− and Polk^+/+ mice (165). Moreover, Polk^−/− mice have a spontaneous mutator phenotype in the kidney, liver and lung, which is attributed to a role of Polκ for accurate lesion bypass of bulky DNA adducts generated endogenously by cholesterol and its metabolites. Therefore, Polκ protects against spontaneous mutagenesis. Rev1^−/− mutant mice are viable on a 129/OLA background, but not on a C57BL/6 background. Rev1⁻^/⁻ mice show reduced weight, but no other gross abnormalities (166). Whether Rev1 mutation impacts carcinogenesis has not been reported.

Although Rad18 is proximal to Y-family TLS polymerases, Rad18-deficient mice do not phenocopy the UV-induced skin tumor-susceptibility of XPV (176). One possible explanation is that Polη bypasses UV-induced CPD in a Rad18-independent manner in the mouse skin. Interestingly, Chk2⁻^/⁻Rad18⁻^/⁻ double knockout mice develop spontaneous lymphomagenesis whereas Chk2⁻^/⁻ or Rad18⁻^/⁻ mice do not. Chk2-deficiency alleviates UV-induced apoptosis of Rad18⁻^/⁻ cells but leads to increased genomic instability (176). Therefore, the genetic interaction of Rad18 and checkpoint genes is important for maintaining genome stability and preventing carcinogenesis.

In a DMBA-ingestion tumorigenesis model, the incidence of leukemia and liver tumors is reduced in Rad18^−/− mice when compared with Rad18^+/+ littermates (164). DMBA-induced skin tumor incidence is unaffected by Rad18, yet the mutation signatures of Rad18^+/+ and Rad18^−/− skin tumor genomes are profoundly different: Rad18-deficiency is associated with reduced overall SNV burdens, increased levels of insertion/deletion (indels) and different contributions of COSMIC mutation signatures to the overall tumor mutational portrait (164). Therefore, Rad18 promotes error-prone TLS of DMBA-induced lesions, resulting in mutagenesis (SNV). However, in the absence of Rad18, TLS polymerases may not be recruited to sites of PAH-induced DNA damage efficiently, leading to replication stalling and fork collapse. Error-prone repair of DBSs resulting from collapsed replication forks could explain the insertions and deletions present in Rad18^-/- tumors.

In summary, most TLS polymerase- or Rad18-deficiencies do not lead to spontaneous tumorigenesis in mice. However, Rad18 and TLS pol-defects lead to altered mutagenesis and carcinogenesis following genotoxic exposures (UV, DMBA, cholesterol metabolites). It is unknown whether over-production of Rad18 or TLS polymerases impacts mutagenesis and carcinogenesis in vivo. Whether Rad18 and TLS polymerase-deficiencies compromise growth of oncogene-driven tumors in vivo has not been addressed comprehensively. However, accumulating evidence supports roles for TLS in conferring resistance to cancer therapy.

HOW DOES TLS AFFECT RESPONSE TO CANCER THERAPY?

Genotoxic cancer therapies are intended to interfere with DNA-templated processes and ultimately cause permanent replicative arrest and cell death. Many therapeutic genotoxins induce primary lesions that cause DNA replication fork stalling. Processing of stalled DNA replication forks may lead to secondary forms of DNA damage such as ssDNA and DSB (the latter often being viewed as the species of DNA damage responsible for lethality). The integrity of DNA repair and DDR signaling in both cancer cells and normal healthy cells critically impacts the overall response to therapeutic agents and patient outcomes. DDR is a protective mechanism and its deployment in cancer cells will antagonize the desired effects of therapy. Accordingly, inhibiting DNA repair processes and compromising DNA damage tolerance in cancer cells is predicted to sensitize tumors to therapy. We exemplify three classes of therapeutic agents whose mechanisms of action are highly dependent on TLS including platinum drugs, checkpoint protein kinase inhibitors and poly(ADP-ribose) polymerase (PARP) inhibitors, as described below.

Platinum drugs

Platinum compounds such as cisplatin and carboplatin are important genotoxic drugs used for the treatment of many cancers (177). Platinum drugs induce DNA damage (primarily intra-strand bulky DNA adducts and ∼1% inter-strand cross-links or ICL) that interfere with DNA synthesis in proliferating cancer cells. Platinating agents primarily induce formation of DNA mono-adducts that induce replication stress, mitotic catastrophe and cell death via apoptosis.

However, many cancer cells have innate resistance to platinum drugs, or acquire cisplatin-resistance during treatment and become refractory to therapy. Mechanisms of cisplatin-resistance include decreased uptake and increased efflux of the platinating agents, and increased DNA repair and increased DNA damage tolerance (178,179). Cisplatin therapy also leads to serious side effects including ototoxicity, nephrotoxicity and neurotoxicity due to cisplatin-induced ROS that cause apoptosis of neuronal and renal cells (180,181,182). Therefore, it is important to devise means to overcome the current limitations of cisplatin therapy.

TLS can help cancer cells acquire resistance to platinating agents and other genotoxic chemotherapies (183). Polη allows replication of cisplatin-damaged DNA templates and the structural basis for Polη-mediated chemoresistance to cisplatin has been elucidated (184,185,186). TLS-deficient cells lacking Polη (187,188,189) or RAD18 (190,191) fail to replicate cisplatin-damaged genomes and instead accumulate unfilled post-replicative gaps, collapsed replication forks and lethal DNA double stranded breaks (DSB). Moreover, high-level Polη expression is correlated with poor survival of platinum-treated non-small cell lung cancer and gastric adenocarcinoma patients (192,193). Polη expression and activity is elevated in cancer stem cells (CSC), the rare progenitors that both self-renew, repopulate tumors and account for chemoresistance (194). Other TLS polymerases are also implicated in mediating cisplatin-resistance. Overexpression of REV1 is strongly correlated with reduced survival probability of prostate cancer patients (195,196,197). Similarly, expression of REV1 is significantly upregulated in lung tumors compared with matched adjacent tissues, and such upregulation is associated with poor prognosis (155). Suppression of REV1 or REV3L not only sensitizes drug-resistant cancers to cisplatin, but also prevents acquisition of drug resistance in murine tumor models (198,199). Inhibition of TLS through RNAi-mediated depletion of REV1 and REV3L profoundly sensitizes human prostate cancer cells to a cisplatin prodrug and dramatically extends animal survival in vivo (200). Similarly, knockout of Rev7 in KRAS^G12D, TP53⁻^/⁻ non-small cell lung cancer (NSCLC) renders cancer cells hypersensitive to cisplatin treatment with significant improvement of animal survival over the REV7 proficient tumor (201).

Inhibiting TLS also represents an opportunity for overcoming toxic side effects of cisplatin therapy: high doses of cisplatin induce ROS and cause apoptosis of neuronal and renal cells leading to neurotoxicity and ototoxicity (180,181). Thus, many toxic side effects of cisplatin therapy are mediated via a DNA damage-independent mechanism that is not affected by TLS. Inhibiting TLS could lower the doses required for cisplatin therapy and reduce the toxic side effects associated with high doses of platinating agents. Rad18⁻^/⁻ and Polh⁻^/⁻ mice are viable and display no overt developmental defects. Thus, specific inhibition of TLS is unlikely to cause toxicity or be detrimental to cancer patients. In summary, there is very strong rationale to explore the TLS pathway as a therapeutic target whose inhibition will confer killing of cancer cells by cisplatin and reduce toxic side effects of cisplatin. By analogy, combining other genotoxic therapeutic agents with TLS inhibition will likely be an effective strategy to inhibit DNA damage tolerance in neoplastic cells.

Checkpoint kinase inhibitors

Cell cycle checkpoints integrate DNA repair with cell cycle progression to promote cell viability. The ATM/CHK2 and ATR/CHK1 pathways represent two major branches of the DDR and mediate DNA damage-inducible checkpoints that arrest cells in G1, S and G2 phases (202). Protein kinase inhibitors targeting ATM, ATR, CHK1 and CHK2 provide a strategy for uncoupling DNA repair from cell cycle progression and sensitizing cancer cells to therapy-induced genotoxicity (182,202,203,204,205,206,207).

ATR/CHK1 signaling and RAD18-mediated TLS occur simultaneously in response to replication stalling. ATR and RAD18 pathways are temporally-coincident because both are activated by RPA-coated ssDNA generated via helicase-polymerase uncoupling (205,208). There is also extensive cross-talk between the ATR and TLS pathways. RAD18 and Polη activities are regulated by ATR/CHK1-dependent phosphorylation (84,87,91). Conversely, when TLS is compromised (e.g. in XPV cells), there is a compensatory increase in ATR/CHK1 signaling which helps preserve cell viability (209). The compensatory ATR response of TLS-deficient cells following genotoxin exposure explains early observations that XPV cells are not UV-sensitive unless treated with caffeine (210). It is now appreciated that caffeine inhibits ATR, and that the ATR/CHK1 and TLS pathways are partially-redundant (209). Therefore, synthetic lethality resulting from inhibiting ATR/CHK1 together with TLS might represent a therapeutic strategy for sensitizing cancer cells to intrinsic or therapy-induced DNA damage.

Pharmacological inhibition of the WEE1 protein kinase also leads to bypass of cell cycle checkpoints and lethality. WEE1 performs inhibitory phosphorylation of Y15 on Cyclin-Dependent Kinases 1 and 2, thereby restricting S-phase and G2/M progression (211). WEE1 inhibitors de-repress CDK2 and CDK1 activities and promote bypass of S-phase and G2/M cell cycle checkpoints (212,213). Adavosertib is an inhibitor of WEE1 that is currently in clinical trials to treat multiple cancers and has been shown to have promising results in overcoming the resistance to platinum-based drugs, such as cisplatin (214).

In addition to promoting bypass of S/G2 checkpoints, the aberrant activation of CDK2 due to WEE1 inhibition in cancer cells causes excessive origin firing that triggers a DDR (211). WEE1 inhibitors are pharmacological equivalents of Cyclin E overexpression which is often used to experimentally model oncogene-induced DNA replication stress. In WEE1 inhibitor-treated cells, the combination of S-phase damage (due to excess CDK2 activity) and a compromised G2 checkpoint (due to excess CDK1 activity) allows S-phase cells to enter mitosis inappropriately and succumb to lethal mitotic catastrophe (211). Interestingly, TLS prevents WEE1 inhibitor-induced lethality (123,215). The role of TLS in protecting against the lethal effect of WEE1 inhibition is likely due to post-replicative gap filling since Rad18 and Polk-ablated cells aberrantly accumulate ssDNA in G2 after WEE1 inhibition (123). Therefore, concurrent inhibition of WEE1 and TLS represents a potential strategy for inducing mitotic catastrophe in cancer cells.

Poly(ADP-ribose) polymerase (PARP) inhibitors

PARP inhibition has received extensive attention as a synthetic lethal approach for killing BRCA-mutant HR-compromised tumors. PARP enzymes are DNA damage sensors and transducers that bind DNA at ssDNA breaks and synthesize negatively-charged, branched poly(ADP-ribose) (PAR) chains on target proteins in the vicinity of the damage (216). This PTM (termed PARylation) leads to the recruitment of DNA repair enzymes such as ERCC1 and remodeling of damaged DNA (217). In 2005, two groups reported the seminal observation that BRCA2-deficient cells, because of their deficiency in Homologous Recombination, are acutely sensitive to PARP inhibitors (218,219). Therefore, targeting PARPs is an attractive strategy for killing BRCA2-deficient tumors (220).

The synthetic lethal interaction between PARP inhibition and BRCA-deficiency was originally proposed to be due to persistent SSBs which are repaired by HR during S-phase. Thus, HR was originally proposed to avert fork collapse and lethal DSB. Subsequently it has become evident that PARPi not only block PARP-mediated SSB repair but also trap the PARP enzyme on damaged DNA. Thus, the trapped PARP protein itself is also an obstacle to replisome movement which cannot be resolved in the absence of HR (220,221,222).

Several recent studies suggest that PARPi toxicity in HR-deficient (HRD) cells is caused by DNA replication-associated single-stranded DNA (ssDNA) gaps (125,126,158,223) - a vulnerability which also suggests interesting and potential tractable roles of the TLS pathway as a therapeutic target in HRD tumors. Cantor's excellent recent review describes the various ways in which ssDNA gaps arise in BRCA1 and BRCA2-mutant cells (126). Interestingly, repriming mediated by PRIMPOL is a major source of spontaneous ssDNA gaps when HR is compromised (127). Consistent with TLS as a post-replicative gap-filling process that operates behind newly-reprimed forks, BRCA1/2-deficient cells are dependent upon RAD18 and Polζ-for gap suppression and viability (127). The demonstration that HR-deficiency creates a reliance on TLS suggests that patient stratification based on ‘BRCA-ness’ might reveal clinical contexts in which TLS inhibition has most therapeutic benefit.

ROLE OF TLS IN THERAPY-INDUCED MUTAGENESIS

The action of TLS polymerases on non-cognate lesions could plausibly play a role in acquisition of therapy-induced mutations. Therapy-induced mutations are of great potential clinical significance because they might help drive acquired chemoresistance. A good example of a clinical setting in which therapy-induced genotoxicity is linked to acquired chemoresistance is during Temozolomide (TMZ) treatment of brain cancers.

Temozolomide (TMZ) is a DNA methylating chemotherapeutic agent used in the treatment of Glioblastomas (GBM) and is the only FDA-approved first-line chemotherapeutic drug for this disease (224). All GBM eventually become TMZ-refractory and recur (225). The extent to which TMZ- induced hypermutation causes GBM recurrence is debated, but it has been proposed that therapy-induced mutations account for the adaptations that allow GBM to resist TMZ (226,227,228). Tumors from TMZ-treated GBM patients harbor a specific mutational signature (designated Single Base Substitution 11 or SBS 11) which is characterized by a preponderance of G:C>A:T transitions at non-CpG sites (229). Moreover, the therapy-associated mutational signature of hypermutated gliomas has been recapitulated by TMZ-treatment (230). However, the underlying error-prone DNA repair mechanisms that mediate de novo and therapy-induced mutation patterns and hypermutability in GBM are not fully understood. GBM cells do rely on RAD18 and Polκ to tolerate TMZ treatment (231,232,233). Overexpression of POLK in GBM patients is associated with TMZ-resistance and is a prognostic indicator for shorter survival (234,235). However, the contribution of TLS polymerases to TMZ-induced mutagenesis is unknown. To address the contribution of TLS to TMZ-induced hypermutation, Yang et al. annotated TMZ-induced mutation signatures in genomes from TMZ-treated isogenic RAD18^+/+ and RAD18^−/− GBM. In those experiments, analysis of mutation signatures from TMZ-treated GBM reveals a role for RAD18 in error-free bypass of O⁶mG (the most toxic TMZ-induced lesion), and error-prone bypass of other TMZ-induced lesions. However, the RAD18-dependent TLS polymerases responsible for mutagenic bypass of TMZ-induced DNA lesions have not yet been identified.

Cisplatin-induced mutation patterns resemble the cancer mutation signatures attributed to smoking and other sources of guanine-directed base damage. TLS has been implicated in generating cisplatin-induced mutational spectra at sites of guanine-directed base damage (236). Mutagenesis of cisplatin-induced DNA lesions is most likely due to the inserter/extender properties of Polη and Polζ which cooperate to perform bypass of GG cisplatin adducts (237). Further work is needed to elucidate how TLS polymerases impact therapy-induced mutations associated with acquired chemoresistance (238). Platinum-based and other chemotherapies for solid tumors are associated with increased risk of secondary hematological neoplasms (notably myelodysplastic syndrome, MDS and acute nonlymphocytic leukemia, ANLL) (239,240). Presumably these therapy-induced hematological malignancies are caused by genotoxicity-induced oncogenic mutations in hematopoietic stem cells. The extent to which TLS mediates mutagenic events leading to therapy-induced secondary neoplasms has not been determined.

TARGETING TLS FACTORS FOR THERAPY

There have been attempts to develop small molecules that target components of the TLS pathway (Table 1). The REV1 inhibitor JH-RE-06 was discovered by the Zhou and Hong laboratories in a screen for small molecule inhibitors that target the REV7-binding surface of the REV1 CTD and disrupt the REV1–REV7 interaction (241). Binding of JH-RE-06 induces REV1 dimerization, thereby blocking the REV1–REV7 interaction and preventing Polζ recruitment to the replisome. JH-RE-06 treatments induce all the hallmarks of TLS-deficiency including inhibition of mutagenic lesion bypass and enhancement of DNA damage-sensitivity in cultured human and mouse cell lines (241). Co-administration of JH-RE-06 with cisplatin suppresses the growth of human melanoma xenografts in vivo, thereby providing encouraging pre-clinical validation for TLS inhibitors as a novel class of chemosensitizers for cancer therapy. A different small molecule inhibitor of REV1 inhibitor that binds to the REV1-CTD and inhibits interaction with TLS insertion polymerases also sensitizes cultured murine and human cells to genotoxic exposures (242). Therefore, pharmacological disruption of associations between REV1 and its partner DNA polymerases is emerging as a feasible approach for inhibiting TLS and chemosensitizing cancer cells (243).

Table 1.

TLS targeting agents

Target	Inhibitor	Function	IC₅₀ or K_d	Structure of the Inhibitor-bound complex	In vitro & In vivo
REV1 (REV7 interface)	JH-RE-06	Blocks the REV1-REV7 interaction	IC₅₀ = 0.78 μM (AlphaScreen) K_d = 0.42 μM (ITC)	PDB: 6C8C (241)	In vitro: • Sensitize multiple tumor cell lines to cisplatin (241) • Selective killing of BRCA1-deficient breast and ovarian cancers (127) In vivo: • Synergize with cisplatin to suppress melanoma growth in mice (241); • Selective suppression of BRCA1-deficient breast tumorigenesis in mice (127); • Suppress lung tumorigenesis in mice (155).
REV1 (RIR interface)	Phenazopyridine (PAP) analog 1	Blocks the REV1-RIR interaction	K _d = 21 μM (MST)	PDB: 6WS5 (265)	In vitro & in vivo: • Synergize with Olaparib/ temozolomide and induces cell death in PDXres 1518–3 SCLC cells (258)
REV1 UBM2 (ubiquitin interface)	MLAF50	Blocks the REV1-ubiquitin interaction	K _d = 37 μM (SPR)	ND	In vitro: • Inhibit chromatin co-localization of REV1 with PCNA (259)
REV7	Compound 7	Binds REV7	IC₅₀ = 78 μM (FP)	ND	In vitro: • Sensitize HeLa cells to cisplatin (260)
Pol eta/REV1	PNR-7-02	Inhibits the catalytic activity of Pol eta and REV1 with similar efficiency	IC₅₀ = 8.0 μM (polymerase activity assay)	ND	In vitro: • Sensitize HAP-1 cells to cisplatin (261)
Pol kappa	IAG-10	Selectively inhibits human Pol Kappa	IC₅₀ = 0.7–7 μM (polymerase activity assay)	ND	In vitro: • Sensitize HAP-1 cells to TMZ (232)
PCNA	T2AA	Binds to PCNA in a bimolecular fashion; inhibits PNCA-interaction with Pol eta or REV1	IC₅₀ = 1 μM (FP)	PDB: 3WGW (262)	In vitro: • Sensitize HeLa and U2OS cells to cisplatin treatment (262,263)
MAGEA4 (Rad18 interface)	Cyclic and linear MTP peptides	Blocks MAGEA4/RAD18 interaction	IC₅₀: <1 nM to 10 μM (TR-FRET)	PDB: 7UOA (246)	In vitro: • Selectively pulldown MAGEA4 (246)

Open in a new tab

Given the central role of PCNA as a hub for inserter and extender polymerase engagement at the replisome, targeting the interaction between PCNA and TLS polymerases may be a feasible strategy for inhibiting TLS. Vanarotti and colleagues identified small molecules that bind to the UBM2 domain of REV1 and prevent its association with monoubiquitinated PCNA (244). The REV1–UBM2-binding compound prevents cisplatin-induced recruitment of REV1 to PCNA on chromatin, prevents UV-induced mutagenesis of the HPRT gene, and sensitizes cells to cyclophosphamide or cisplatin. Another study identified a PCNA-binding small molecule designated PCNA-I1S that stabilizes the PCNA trimer, and reduces levels of chromatin-associated PCNA (245). PCNA-I1S inhibits cell growth and enhances cytotoxic effects of UV radiation and cisplatin. However, it is unclear whether the cytotoxic effects of PCNA-I1S result from inhibition of TLS or other PCNA-mediated genome maintenance processes.

The most ideal therapeutic target pathways in cancer are ones that are critically required for neoplastic cell growth, yet are dispensable for viability of normal healthy cells. The TLS pathway meets this criterion in some contexts. For example, oncogene-expressing cells are more dependent on Rad18 and Polκ when compared with untransformed cells (123). Some cancer cells pathologically stabilize RAD18 via a mis-expressed Cancer/Testes Antigen (CTA), termed MAGEA4 (78). Moreover, MAGEA4-expressing cancer cells become critically dependent on MAGEA4 to maintain RAD18 expression, activate TLS, and tolerate DNA damage (78). Owing to its cancer cell-specific expression and requirement for DNA damage tolerance, the MAGEA4/RAD18 signaling axis is an appealing pathway for therapeutic inhibition. Fleming et al. identified small cyclic peptide inhibitors that bind directly to MAGEA4 and inhibit RAD18 binding with nanomolar potency (246). That study validates the chemical tractability of MAGEA4/RAD18 as a cancer-specific vulnerability and justifies further work to identify more drug-like pharmacological inhibitors of pathological TLS in cancer. RAD18 is also implicated in DSB repair (247,248,249) and ICL repair (250) independently of its roles in TLS. Inhibiting MAGEA4-dependent RAD18 activities in genome maintenance might also sensitize cancer cells to diverse classes of anti-cancer agents that induce replication fork stalling, DSB, and ICL.

CONCLUSIONS AND FUTURE DIRECTIONS

In the past decades our understanding of TLS as a PRR mechanism merely involving a PCNA-ubiquitination-mediated DNA polymerase switch has expanded considerably. We are now aware of multiple ubiquitination-dependent and independent TLS activation mechanisms that may operate both within and outside of S-phase, and that are intimately integrated with other components of the DDR. Future work will likely reveal additional complexity and fine-tuning mechanisms that regulate TLS polymerases and coordinate genome maintenance with other cellular processes. Matsuoka and Elledge showed that substrates of the proximal DDR mediators ATM and ATR are involved in diverse pathways including metabolism, chromatin packing and remodeling, trafficking, transcription, and translation (251). By analogy, integration of TLS with broad regulatory pathways in a cell is likely important for genome stability and DNA damage tolerance.

For many other genome maintenance mechanisms (notably DSB signaling and NER), chromatin architecture and histone modifications are critical determinants of DNA repair activity (252,253). Pharmacological modifiers of chromatin structure impact the nuclear mobility of TLS polymerases (254), yet little else is known regarding links between chromatin and TLS. More connections between chromatin modification and TLS are likely to be discovered. In the case of DSB repair, pathway choice between error-free HR and error-prone NHEJ/MMEJ is enormously consequential for genome stability (255,256). It is likely that mechanisms of pathway choice between error-prone TLS and error-free TS (or other processes) will similarly emerge as important determinants of mutagenesis and DNA damage tolerance.

Many questions remain regarding the relationship between TLS and cancer and the extent to which TLS sustains tumorigenic phenotypes such as mutagenesis and DNA damage tolerance. Whole genome sequencing is the new standard for evaluating roles of candidate genome maintenance pathways in cancer-associated mutation patterns. Genome sequencing approaches have provided evidence that TLS polymerases contribute to some of the cosmic mutation signatures in cancer. There is a need for more studies to model the effects of TLS-deficiencies on genome-wide mutation patterns and to determine whether TLS polymerases contribute to the mutation signatures of unknown etiology. The role of TLS in mediating therapy-induced mutations that drive disease recurrence and secondary malignancies is also unclear and must be addressed. Putative TLS-mediated mutational scars of cancer cells could provide biomarkers that are predictive or prognostic of sensitivity to specific therapeutic agents and that guide clinical decisions.

There is increasing evidence that TLS allows neoplastic cells to tolerate intrinsically-arising DNA damage. Such TLS-dependency could be a useful and actionable vulnerability that presents opportunities to target cancer cells using TLS inhibitors. It is important to define the pathological contexts in which tolerance of various types of intrinsic (and therapy-induced) DNA damage is TLS-dependent. For example ssDNA gap-suppression has been proposed as an important role of TLS in cancer cells (124), but not all cancer cells are sensitive to REV1-inhibition.

It is possible that different cancer cells harbor distinct forms of intrinsic DNA damage and also have differential TLS-dependencies based upon availability of other DDR pathways. It will also be valuable to identify synthetic lethal genetic interactions between TLS and other DNA repair pathways which could be exploited for therapy.

Genetically engineered mouse models (GEMM) have been invaluable for identifying genes that sustain multistep tumorigenesis. There exist sophisticated GEMM that recapitulate the genetics, disease progression and histopathological features of many human cancers. As yet however, there have been few studies to model impact of TLS perturbations on oncogene-driven cancers.

Therefore, GEMM would also be useful for defining the cancer settings in which TLS sustains tumor cells (and in which TLS inhibition would likely provide therapeutic benefit). GEMM would also be useful to determine whether the altered expression patterns of TLS polymerases and regulators observed in cancers (e.g. over-abundance of RAD18 or reduced REV1 expression compared with normal tissues) impact disease progression and shape mutation signatures. GEMM would also be extremely useful in preclinical studies to determine therapeutic efficacies of pharmacological TLS inhibitors.

For effective cancer therapy it is important to selectively discriminate between neoplastic cells and normal healthy cells. Representative untransformed cells and cancer cell lines might have different TLS-dependencies (123,257). However, cancer is a highly heterogeneous disease. Mechanisms of DNA damage tolerance and chemoresistance often differ tremendously between different patients, even those with the same cancers and cancer subtypes. Therefore, it may prove impossible to conclude that all cancers are globally dependent on TLS. Ultimately, precision medicine approaches that target individual tumors based on their unique disease characteristics (including driver oncogenes, intrinsic stresses and repertoire of available DDR pathways) might provide the best vehicle for TLS inhibitors as cancer therapies.

METHODS

Lung adenocarcinoma (LUAD), bladder urothelial carcinoma (BLCA), lung squamous cell carcinoma (LUSC), uterine corpus endometrial carcinoma (UCEC), pancreatic adenocarcinoma (PAAD), glioblastoma (GBM), were selected for evaluation of relationships between TLS/TS gene status and genome instability in human tumors. TCGA datasets containing RNA expression, mutation, genomic alteration and clinical information for these tumors were from the TCGA data portal (https://portal.gdc.cancer.gov) and were all downloaded in April 2022. Specific datasets used in this study include: (i) HTSeq-FPKM-UQ gene expression quantification (duplicated samples for the same patients were removed), (ii) somatic mutations aggregated and masked by Mutect2 and organized as MAF files, publicly available, (iii) smoking information for most subjects in LUAD, LUSC and BLCA patients. To avoid zeros when using a log scale, log2(FPKM + 1) was used to display gene expression data obtained by RNA-seq.

Statistical analysis and data presentation

R (version 4.1.0) was used for data analysis and presentation. Wilcoxon Rank Sum Test (wilcox.test, with option alterative = ‘two sided’ and paired = False) was used for comparison of gene expression between tumor and adjacent normal samples as well as the comparison of SNVs. R package ComplexHeatmap (version 2.8.0) was used to generate the heatmap illustrating relative expression levels of TLS and TS pathway genes in LUAD. Z-score transformed FPKM-UQ value was used as the gene expression value in the heatmap. Stage, p53 mutation, Replication stress (RS) and smoke annotations were derived from clinical files, Mutect2 MAF files and GISTIC2 copy number files.

DATA AVAILABILIY

The raw data in this study including RNAexpression, clinical information, mutation, and genomic alterations can be found at TCGA data portal (https://portal.gdc.cancer.gov).

Supplementary Material

zcad005_Supplemental_File

zcad005_supplemental_file.pdf^{(365.7KB, pdf)}

Contributor Information

Jay Anand, Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, 614 Brinkhous-Bullitt Building, Chapel Hill, NC 27599, USA.

Lilly Chiou, Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, 614 Brinkhous-Bullitt Building, Chapel Hill, NC 27599, USA; Curriculum in Genetics and Molecular Biology, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA.

Carly Sciandra, Department of Biochemistry, Duke University School of Medicine, Durham, NC 27710, USA.

Xingyuan Zhang, Department of Biostatistics, University of North Carolina at Chapel Hill, 135 Dauer Drive, 3101 McGavran-Greenberg Hall, Chapel Hill, NC 27599, USA.

Jiyong Hong, Department of Chemistry, Duke University, Durham, NC 27708, USA.

Di Wu, Department of Biostatistics, University of North Carolina at Chapel Hill, 135 Dauer Drive, 3101 McGavran-Greenberg Hall, Chapel Hill, NC 27599, USA.

Pei Zhou, Department of Biochemistry, Duke University School of Medicine, Durham, NC 27710, USA.

Cyrus Vaziri, Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, 614 Brinkhous-Bullitt Building, Chapel Hill, NC 27599, USA.

SUPPLEMENTARY DATA

Supplementary Data are available at NAR Cancer Online.

FUNDING

National Cancer Institute [R01 CA215347, R01 CA229530 to C.V., CA191448 to P.Z., J.H., in part]; National Institute of Environmental Health Sciences [R01 ES029079 to C.V.]; Duke Clinical and Translational Science Institute (CTSI) [UL1TR002553 to P.Z., J.H.]; developmental funds of the Duke Cancer Institute (to P.Z.) as part of the P30 Cancer Center Support Grant [NIH CA014236]; L.C. was supported in part by a grant from the National Institute of General Medical Sciences [5T32GM135128].

Conflict of interest statement. None declared.

REFERENCES

1. Hanahan D., Weinberg R.A. The hallmarks of cancer. Cell. 2000; 100:57–70. [DOI] [PubMed] [Google Scholar]
2. Hanahan D., Weinberg R.A. Hallmarks of cancer: the next generation. Cell. 2011; 144:646–674. [DOI] [PubMed] [Google Scholar]
3. Baynton K., Fuchs R.P. Lesions in DNA: hurdles for polymerases. Trends Biochem. Sci. 2000; 25:74–79. [DOI] [PubMed] [Google Scholar]
4. Ogi T., Lehmann A.R. the Y-family DNA polymerase kappa (pol kappa) functions in mammalian nucleotide-excision repair. Nat. Cell Biol. 2006; 8:640–642. [DOI] [PubMed] [Google Scholar]
5. Ogi T., Limsirichaikul S., Overmeer R.M., Volker M., Takenaka K., Cloney R., Nakazawa Y., Niimi A., Miki Y., Jaspers N.G. et al. Three DNA polymerases, recruited by different mechanisms, carry out NER repair synthesis in human cells. Mol. Cell. 2010; 37:714–727. [DOI] [PubMed] [Google Scholar]
6. Yang Y., Durando M., Smith-Roe S.L., Sproul C., Greenwalt A.M., Kaufmann W., Oh S., Hendrickson E.A., Vaziri C. Cell cycle stage-specific roles of Rad18 in tolerance and repair of oxidative DNA damage. Nucleic Acids Res. 2013; 41:2296–2312. [DOI] [PMC free article] [PubMed] [Google Scholar]
7. Zlatanou A., Despras E., Braz-Petta T., Boubakour-Azzouz I., Pouvelle C., Stewart G.S., Nakajima S., Yasui A., Ishchenko A.A., Kannouche P.L. The hMsh2-hMsh6 complex acts in concert with monoubiquitinated PCNA and Pol eta in response to oxidative DNA damage in human cells. Mol. Cell. 2011; 43:649–662. [DOI] [PubMed] [Google Scholar]
8. Tirman S., Quinet A., Wood M., Meroni A., Cybulla E., Jackson J., Pegoraro S., Simoneau A., Zou L., Vindigni A. Temporally distinct post-replicative repair mechanisms fill PRIMPOL-dependent ssDNA gaps in human cells. Mol. Cell. 2021; 81:4026–4040. [DOI] [PMC free article] [PubMed] [Google Scholar]
9. Karras G.I., Jentsch S. the RAD6 DNA damage tolerance pathway operates uncoupled from the replication fork and is functional beyond S phase. Cell. 2010; 141:255–267. [DOI] [PubMed] [Google Scholar]
10. Daigaku Y., Davies A.A., Ulrich H.D. Ubiquitin-dependent DNA damage bypass is separable from genome replication. Nature. 2010; 465:951–955. [DOI] [PMC free article] [PubMed] [Google Scholar]
11. Masutani C., Araki M., Yamada A., Kusumoto R., Nogimori T., Maekawa T., Iwai S., Hanaoka F. Xeroderma pigmentosum variant (XP-V) correcting protein from HeLa cells has a thymine dimer bypass DNA polymerase activity. EMBO J. 1999; 18:3491–3501. [DOI] [PMC free article] [PubMed] [Google Scholar]
12. Masutani C., Kusumoto R., Yamada A., Dohmae N., Yokoi M., Yuasa M., Araki M., Iwai S., Takio K., Hanaoka F. The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase eta. Nature. 1999; 399:700–704. [DOI] [PubMed] [Google Scholar]
13. Friedberg E.C. How nucleotide excision repair protects against cancer. Nat. Rev. Cancer. 2001; 1:22–33. [DOI] [PubMed] [Google Scholar]
14. Pfeifer G.P. Formation and processing of UV photoproducts: effects of DNA sequence and chromatin environment. Photochem. Photobiol. 1997; 65:270–283. [DOI] [PubMed] [Google Scholar]
15. Dupuy A., Sarasin A. DNA damage and gene therapy of xeroderma pigmentosum, a human DNA repair-deficient disease. Mutat. Res. 2015; 776:2–8. [DOI] [PubMed] [Google Scholar]
16. Di Lucca J., Guedj M., Lacapere J.J., Fargnoli M.C., Bourillon A., Dieude P., Dupin N., Wolkenstein P., Aegerter P., Saiag P. et al. Variants of the xeroderma pigmentosum variant gene (POLH) are associated with melanoma risk. Eur. J. Cancer. 2009; 45:3228–3236. [DOI] [PubMed] [Google Scholar]
17. Castro L.P., Batista-Vieira D., de Souza T.A., Timoteo A.R.S., Coutinho J.D.L., Pinheiro de Almeida I.C., Henriques S.R.M., de Azevedo F.M., Rosa R.C.A., Kannouche P.L. et al. XPC and POLH/XPV genes mutated in a genetic cluster of xeroderma pigmentosum patients in northeast Brazil. Front. Genet. 2021; 12:784963. [DOI] [PMC free article] [PubMed] [Google Scholar]
18. Ziv O., Geacintov N., Nakajima S., Yasui A., Livneh Z. DNA polymerase zeta cooperates with polymerases kappa and iota in translesion DNA synthesis across pyrimidine photodimers in cells from XPV patients. Proc. Natl. Acad. Sci. U.S.A. 2009; 106:11552–11557. [DOI] [PMC free article] [PubMed] [Google Scholar]
19. Shachar S., Ziv O., Avkin S., Adar S., Wittschieben J., Reissner T., Chaney S., Friedberg E.C., Wang Z., Carell T. et al. Two-polymerase mechanisms dictate error-free and error-prone translesion DNA synthesis in mammals. EMBO J. 2009; 28:383–393. [DOI] [PMC free article] [PubMed] [Google Scholar]
20. Limoli C.L., Giedzinski E., Morgan W.F., Cleaver J.E. 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. U.S.A. 2000; 97:7939–7946. [DOI] [PMC free article] [PubMed] [Google Scholar]
21. Vaziri C., Rogozin I.B., Gu Q., Wu D., Day T.A. Unravelling roles of error-prone DNA polymerases in shaping cancer genomes. Oncogene. 2021; 40:6549–6565. [DOI] [PMC free article] [PubMed] [Google Scholar]
22. Ceccaldi R., Liu J.C., Amunugama R., Hajdu I., Primack B., Petalcorin M.I., O’Connor K.W., Konstantinopoulos P.A., Elledge S.J., Boulton S.J. et al. Homologous-recombination-deficient tumours are dependent on Poltheta-mediated repair. Nature. 2015; 518:258–262. [DOI] [PMC free article] [PubMed] [Google Scholar]
23. Schimmel J., van Schendel R., den Dunnen J.T., Tijsterman M. Templated insertions: a smoking gun for polymerase theta-mediated end joining. Trends Genet. 2019; 35:632–644. [DOI] [PubMed] [Google Scholar]
24. Hwang T., Reh S., Dunbayev Y., Zhong Y., Takata Y., Shen J., McBride K.M., Murnane J.P., Bhak J., Lee S. et al. Defining the mutation signatures of DNA polymerase theta in cancer genomes. NAR Cancer. 2020; 2:zcaa017. [DOI] [PMC free article] [PubMed] [Google Scholar]
25. Prakash S., Johnson R.E., Prakash L. Eukaryotic translesion synthesis DNA polymerases: specificity of structure and function. Annu. Rev. Biochem. 2005; 74:317–353. [DOI] [PubMed] [Google Scholar]
26. Waters L.S., Minesinger B.K., Wiltrout M.E., D'Souza S., Woodruff R.V., Walker G.C Eukaryotic translesion polymerases and their roles and regulation in DNA damage tolerance. Microbiol. Mol. Biol. Rev. 2009; 73:134–154. [DOI] [PMC free article] [PubMed] [Google Scholar]
27. Ulrich H.D., Jentsch S. Two RING finger proteins mediate cooperation between ubiquitin-conjugating enzymes in DNA repair. EMBO J. 2000; 19:3388–3397. [DOI] [PMC free article] [PubMed] [Google Scholar]
28. Kannouche P.L., Wing J., Lehmann A.R. Interaction of human DNA polymerase eta with monoubiquitinated PCNA: a possible mechanism for the polymerase switch in response to DNA damage. Mol. Cell. 2004; 14:491–500. [DOI] [PubMed] [Google Scholar]
29. Davies A.A., Huttner D., Daigaku Y., Chen S., Ulrich H.D. Activation of ubiquitin-dependent DNA damage bypass is mediated by replication protein a. Mol. Cell. 2008; 29:625–636. [DOI] [PMC free article] [PubMed] [Google Scholar]
30. Zou L., Elledge S.J. Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science. 2003; 300:1542–1548. [DOI] [PubMed] [Google Scholar]
31. Pena-Diaz J., Bregenhorn S., Ghodgaonkar M., Follonier C., Artola-Boran M., Castor D., Lopes M., Sartori A.A., Jiricny J. Noncanonical mismatch repair as a source of genomic instability in human cells. Mol. Cell. 2012; 47:669–680. [DOI] [PubMed] [Google Scholar]
32. Ray A., Blevins C., Wani G., Wani A.A. ATR- and ATM-mediated DNA damage response is dependent on excision repair assembly during G1 but Not in S phase of cell cycle. PLoS One. 2016; 11:e0159344. [DOI] [PMC free article] [PubMed] [Google Scholar]
33. Bienko M., Green C.M., Crosetto N., Rudolf F., Zapart G., Coull B., Kannouche P., Wider G., Peter M., Lehmann A.R. et al. Ubiquitin-binding domains in Y-family polymerases regulate translesion synthesis. Science. 2005; 310:1821–1824. [DOI] [PubMed] [Google Scholar]
34. Masuda Y., Kanao R., Kaji K., Ohmori H., Hanaoka F., Masutani C. Different types of interaction between PCNA and PIP boxes contribute to distinct cellular functions of Y-family DNA polymerases. Nucleic Acids Res. 2015; 43:7898–7910. [DOI] [PMC free article] [PubMed] [Google Scholar]
35. Guo C., Sonoda E., Tang T.S., Parker J.L., Bielen A.B., Takeda S., Ulrich H.D., Friedberg E.C. REV1 protein interacts with PCNA: significance of the REV1 BRCT domain in vitro and in vivo. Mol. Cell. 2006; 23:265–271. [DOI] [PubMed] [Google Scholar]
36. Pustovalova Y., Maciejewski M.W., Korzhnev D.M. NMR mapping of PCNA interaction with translesion synthesis DNA polymerase Rev1 mediated by Rev1-BRCT domain. J. Mol. Biol. 2013; 425:3091–3105. [DOI] [PubMed] [Google Scholar]
37. Sharma N.M., Kochenova O.V., Shcherbakova P.V. the non-canonical protein binding site at the monomer-monomer interface of yeast proliferating cell nuclear antigen (PCNA) regulates the Rev1-PCNA interaction and Polzeta/Rev1-dependent translesion DNA synthesis. J. Biol. Chem. 2011; 286:33557–33566. [DOI] [PMC free article] [PubMed] [Google Scholar]
38. Durando M., Tateishi S., Vaziri C. a non-catalytic role of DNA polymerase η in recruiting Rad18 and promoting PCNA monoubiquitination at stalled replication forks. Nucleic Acids Res. 2013; 41:3079–3093. [DOI] [PMC free article] [PubMed] [Google Scholar]
39. Watanabe K., Tateishi S., Kawasuji M., Tsurimoto T., Inoue H., Yamaizumi M. Rad18 guides poleta to replication stalling sites through physical interaction and PCNA monoubiquitination. EMBO J. 2004; 23:3886–3896. [DOI] [PMC free article] [PubMed] [Google Scholar]
40. Masutani C., Kusumoto R., Iwai S., Hanaoka F. Mechanisms of accurate translesion synthesis by human DNA polymerase eta. EMBO J. 2000; 19:3100–3109. [DOI] [PMC free article] [PubMed] [Google Scholar]
41. Choi J.H., Pfeifer G.P. the role of DNA polymerase eta in UV mutational spectra. DNA Repair (Amst.). 2005; 4:211–220. [DOI] [PubMed] [Google Scholar]
42. Choi J.Y., Angel K.C., Guengerich F.P. Translesion synthesis across bulky N2-alkyl guanine DNA adducts by human DNA polymerase kappa. J. Biol. Chem. 2006; 281:21062–21072. [DOI] [PubMed] [Google Scholar]
43. Ogi T., Shinkai Y., Tanaka K., Ohmori H. Polkappa protects mammalian cells against the lethal and mutagenic effects of benzo[a]pyrene. Proc. Natl. Acad. Sci. U.S.A. 2002; 99:15548–15553. [DOI] [PMC free article] [PubMed] [Google Scholar]
44. Lin W., Xin H., Zhang Y., Wu X., Yuan F., Wang Z. the human REV1 gene codes for a DNA template-dependent dCMP transferase. Nucleic Acids Res. 1999; 27:4468–4475. [DOI] [PMC free article] [PubMed] [Google Scholar]
45. Hishiki A., Hashimoto H., Hanafusa T., Kamei K., Ohashi E., Shimizu T., Ohmori H., Sato M. Structural basis for novel interactions between human translesion synthesis polymerases and proliferating cell nuclear antigen. J. Biol. Chem. 2009; 284:10552–10560. [DOI] [PMC free article] [PubMed] [Google Scholar]
46. McCulloch S.D., Kokoska R.J., Masutani C., Iwai S., Hanaoka F., Kunkel T.A. Preferential cis-syn thymine dimer bypass by DNA polymerase eta occurs with biased fidelity. Nature. 2004; 428:97–100. [DOI] [PubMed] [Google Scholar]
47. Kusumoto R., Masutani C., Shimmyo S., Iwai S., Hanaoka F. DNA binding properties of human DNA polymerase eta: implications for fidelity and polymerase switching of translesion synthesis. Genes Cells. 2004; 9:1139–1150. [DOI] [PubMed] [Google Scholar]
48. Ohmori H., Friedberg E.C., Fuchs R.P., Goodman M.F., Hanaoka F., Hinkle D., Kunkel T.A., Lawrence C.W., Livneh Z., Nohmi T. et al. the Y-family of DNA polymerases. Mol. Cell. 2001; 8:7–8. [DOI] [PubMed] [Google Scholar]
49. Prakash S., Johnson R.E., Prakash L. Eukaryotic translesion synthesis DNA polymerases: specificity of structure and function. Annu. Rev. Biochem. 2005; 74:317–353. [DOI] [PubMed] [Google Scholar]
50. Johnson R.E., Yu S.-L., Prakash S., Prakash L. Yeast DNA polymerase zeta (zeta) is essential for error-free replication past thymine glycol. Genes Dev. 2003; 17:77–87. [DOI] [PMC free article] [PubMed] [Google Scholar]
51. Ross A.-L., Simpson L.J., Sale J.E. Vertebrate DNA damage tolerance requires the C-terminus but not BRCT or transferase domains of REV1. Nucleic Acids Res. 2005; 33:1280–1289. [DOI] [PMC free article] [PubMed] [Google Scholar]
52. Edmunds C.E., Simpson L.J., Sale J.E. PCNA ubiquitination and REV1 define temporally distinct mechanisms for controlling translesion synthesis in the avian cell line DT40. Mol. Cell. 2008; 30:519–529. [DOI] [PubMed] [Google Scholar]
53. Ohashi E., Murakumo Y., Kanjo N., Akagi J.-I., Masutani C., Hanaoka F., Ohmori H. Interaction of hREV1 with three human Y-family DNA polymerases. Genes Cells. 2004; 9:523–531. [DOI] [PubMed] [Google Scholar]
54. Ohashi E., Hanafusa T., Kamei K., Song I., Tomida J., Hashimoto H., Vaziri C., Ohmori H. Identification of a novel REV1-interacting motif necessary for DNA polymerase kappa function. Genes Cells. 2009; 14:101–111. [DOI] [PMC free article] [PubMed] [Google Scholar]
55. Pustovalova Y., Bezsonova I., Korzhnev D.M. the C-terminal domain of human Rev1 contains independent binding sites for DNA polymerase η and Rev7 subunit of polymerase ζ. FEBS Lett. 2012; 586:3051–3056. [DOI] [PMC free article] [PubMed] [Google Scholar]
56. Pustovalova Y., Magalhaes M.T., D'Souza S., Rizzo A.A., Korza G., Walker G.C., Korzhnev D.M Interaction between the Rev1 C-terminal domain and the PolD3 subunit of Polzeta suggests a mechanism of polymerase exchange upon Rev1/Polzeta-dependent translesion synthesis. Biochemistry. 2016; 55:2043–2053. [DOI] [PMC free article] [PubMed] [Google Scholar]
57. D'Souza S., Walker G.C. Novel role for the C terminus of Saccharomyces cerevisiae Rev1 in mediating protein-protein interactions. Mol. Cell. Biol. 2006; 26:8173–8182. [DOI] [PMC free article] [PubMed] [Google Scholar]
58. Guo C., Sonoda E., Tang T.-S., Parker J.L., Bielen A.B., Takeda S., Ulrich H.D., Friedberg E.C. REV1 protein interacts with PCNA: significance of the REV1 BRCT domain in vitro and in vivo. Mol. Cell. 2006; 23:265–271. [DOI] [PubMed] [Google Scholar]
59. Terai K., Abbas T., Jazaeri A.A., Dutta A. CRL4(Cdt2) E3 ubiquitin ligase monoubiquitinates PCNA to promote translesion DNA synthesis. Mol. Cell. 2010; 37:143–149. [DOI] [PMC free article] [PubMed] [Google Scholar]
60. Lin J.R., Zeman M.K., Chen J.Y., Yee M.C., Cimprich K.A. SHPRH and HLTF act in a damage-specific manner to coordinate different forms of postreplication repair and prevent mutagenesis. Mol. Cell. 2011; 42:237–249. [DOI] [PMC free article] [PubMed] [Google Scholar]
61. Haracska L., Unk I., Prakash L., Prakash S. Ubiquitylation of yeast proliferating cell nuclear antigen and its implications for translesion DNA synthesis. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:6477–6482. [DOI] [PMC free article] [PubMed] [Google Scholar]
62. Acharya N., Yoon J.H., Hurwitz J., Prakash L., Prakash S. DNA polymerase eta lacking the ubiquitin-binding domain promotes replicative lesion bypass in humans cells. Proc. Natl. Acad. Sci. U.S.A. 2010; 107:10401–10405. [DOI] [PMC free article] [PubMed] [Google Scholar]
63. Hendel A., Krijger P.H.L., Diamant N., Goren Z., Langerak P., Kim J., Reissner T., Lee K.-y., Geacintov N.E., Carell T. et al. PCNA ubiquitination is important, but not essential for translesion DNA synthesis in mammalian cells. PLos Genet. 2011; 7:e1002262. [DOI] [PMC free article] [PubMed] [Google Scholar]
64. Wit N., Buoninfante O.A., van den Berk P.C.M., Jansen J.G., Hogenbirk M.A., de Wind N., Jacobs H. Roles of PCNA ubiquitination and TLS polymerases κ and η in the bypass of methyl methanesulfonate-induced DNA damage. Nucleic Acids Res. 2015; 43:282–294. [DOI] [PMC free article] [PubMed] [Google Scholar]
65. Krijger P.H.L., van den Berk P.C.M., Wit N., Langerak P., Jansen J.G., Reynaud C.-A., de Wind N., Jacobs H. PCNA ubiquitination-independent activation of polymerase η during somatic hypermutation and DNA damage tolerance. DNA Repair (Amst.). 2011; 10:1051–1059. [DOI] [PubMed] [Google Scholar]
66. Huertas P., Cortes-Ledesma F., Sartori A.A., Aguilera A., Jackson S.P. CDK targets Sae2 to control DNA-end resection and homologous recombination. Nature. 2008; 455:689–692. [DOI] [PMC free article] [PubMed] [Google Scholar]
67. Ira G., Pellicioli A., Balijja A., Wang X., Fiorani S., Carotenuto W., Liberi G., Bressan D., Wan L., Hollingsworth N.M. et al. DNA end resection, homologous recombination and DNA damage checkpoint activation require CDK1. Nature. 2004; 431:1011–1017. [DOI] [PMC free article] [PubMed] [Google Scholar]
68. Aylon Y., Liefshitz B., Kupiec M. the CDK regulates repair of double-strand breaks by homologous recombination during the cell cycle. EMBO J. 2004; 23:4868–4875. [DOI] [PMC free article] [PubMed] [Google Scholar]
69. Yun M.H., Hiom K. CtIP-BRCA1 modulates the choice of DNA double-strand-break repair pathway throughout the cell cycle. Nature. 2009; 459:460–463. [DOI] [PMC free article] [PubMed] [Google Scholar]
70. Tomimatsu N., Mukherjee B., Catherine Hardebeck M., Ilcheva M., Vanessa Camacho C., Louise Harris J., Porteus M., Llorente B., Khanna K.K., Burma S. Phosphorylation of EXO1 by CDKs 1 and 2 regulates DNA end resection and repair pathway choice. Nat. Commun. 2014; 5:3561. [DOI] [PMC free article] [PubMed] [Google Scholar]
71. Falck J., Forment J.V., Coates J., Mistrik M., Lukas J., Bartek J., Jackson S.P. CDK targeting of NBS1 promotes DNA-end resection, replication restart and homologous recombination. EMBO Rep. 2012; 13:561–568. [DOI] [PMC free article] [PubMed] [Google Scholar]
72. Ferretti L.P., Lafranchi L., Sartori A.A. Controlling DNA-end resection: a new task for CDKs. Front. Genet. 2013; 4:99. [DOI] [PMC free article] [PubMed] [Google Scholar]
73. Luo K., Deng M., Li Y., Wu C., Xu Z., Yuan J., Lou Z. CDK-mediated RNF4 phosphorylation regulates homologous recombination in S-phase. Nucleic Acids Res. 2015; 43:5465–5475. [DOI] [PMC free article] [PubMed] [Google Scholar]
74. Swift M.L., Beishline K., Flashner S., Azizkhan-Clifford J. DSB repair pathway choice is regulated by recruitment of 53BP1 through cell cycle-dependent regulation of Sp1. Cell Rep. 2021; 34:108840. [DOI] [PubMed] [Google Scholar]
75. Uchiyama M., Terunuma J., Hanaoka F. The protein level of Rev1, a TLS polymerase in fission yeast, is strictly regulated during the cell cycle and after DNA damage. PLoS One. 2015; 10:e0130000. [DOI] [PMC free article] [PubMed] [Google Scholar]
76. Waters L.S., Walker G.C. the critical mutagenic translesion DNA polymerase Rev1 is highly expressed during G(2)/M phase rather than S phase. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:8971–8976. [DOI] [PMC free article] [PubMed] [Google Scholar]
77. Varanasi L., Do P.M., Goluszko E., Martinez L.A. Rad18 is a transcriptional target of E2F3. Cell Cycle. 2012; 11:1131–1141. [DOI] [PubMed] [Google Scholar]
78. Gao Y., Mutter-Rottmayer E., Greenwalt A.M., Goldfarb D., Yan F., Yang Y., Martinez-Chacin R.C., Pearce K.H., Tateishi S., Major M.B. et al. A neomorphic cancer cell-specific role of MAGE-A4 in trans-lesion synthesis. Nat. Commun. 2016; 7:12105. [DOI] [PMC free article] [PubMed] [Google Scholar]
79. Bi X., Barkley L.R., Slater D.M., Tateishi S., Yamaizumi M., Ohmori H., Vaziri C. Rad18 regulates DNA polymerase kappa and is required for recovery from S-phase checkpoint-mediated arrest. Mol. Cell. Biol. 2006; 26:3527–3540. [DOI] [PMC free article] [PubMed] [Google Scholar]
80. Song I.Y., Palle K., Gurkar A., Tateishi S., Kupfer G.M., Vaziri C. Rad18-mediated translesion synthesis of bulky DNA adducts is coupled to activation of the Fanconi anemia DNA repair pathway. J. Biol. Chem. 2010; 285:31525–31536. [DOI] [PMC free article] [PubMed] [Google Scholar]
81. Liu G., Chen X. DNA polymerase eta, the product of the xeroderma pigmentosum variant gene and a target of p53, modulates the DNA damage checkpoint and p53 activation. Mol. Cell. Biol. 2006; 26:1398–1413. [DOI] [PMC free article] [PubMed] [Google Scholar]
82. Tomicic M.T., Aasland D., Naumann S.C., Meise R., Barckhausen C., Kaina B., Christmann M. Translesion polymerase eta is upregulated by cancer therapeutics and confers anticancer drug resistance. Cancer Res. 2014; 74:5585–5596. [DOI] [PubMed] [Google Scholar]
83. Lerner L.K., Francisco G., Soltys D.T., Rocha C.R., Quinet A., Vessoni A.T., Castro L.P., David T.I., Bustos S.O., Strauss B.E. et al. Predominant role of DNA polymerase eta and p53-dependent translesion synthesis in the survival of ultraviolet-irradiated human cells. Nucleic Acids Res. 2017; 45:1270–1280. [DOI] [PMC free article] [PubMed] [Google Scholar]
84. Day T.A., Palle K., Barkley L.R., Kakusho N., Zou Y., Tateishi S., Verreault A., Masai H., Vaziri C. Phosphorylated Rad18 directs DNA polymerase η to sites of stalled replication. J. Cell Biol. 2010; 191:953–966. [DOI] [PMC free article] [PubMed] [Google Scholar]
85. Yamada M., Masai H., Bartek J. Regulation and roles of Cdc7 kinase under replication stress. Cell Cycle. 2014; 13:1859–1866. [DOI] [PMC free article] [PubMed] [Google Scholar]
86. Liu R., Huang Y. CDC7 as a novel biomarker and druggable target in cancer. Clin. Transl. Oncol. 2022; 24:1856–1864. [DOI] [PubMed] [Google Scholar]
87. Barkley L.R., Palle K., Durando M., Day T.A., Gurkar A., Kakusho N., Li J., Masai H., Vaziri C. c-Jun N-terminal kinase-mediated Rad18 phosphorylation facilitates Poleta recruitment to stalled replication forks. Mol. Biol. Cell. 2012; 23:1943–1954. [DOI] [PMC free article] [PubMed] [Google Scholar]
88. Hotamisligil G.S., Davis R.J. Cell signaling and stress responses. Cold Spring Harb. Perspect. Biol. 2016; 8:a006072. [DOI] [PMC free article] [PubMed] [Google Scholar]
89. Sarcevic B., Mawson A., Baker R.T., Sutherland R.L. Regulation of the ubiquitin-conjugating enzyme hHR6A by CDK-mediated phosphorylation. EMBO J. 2002; 21:2009–2018. [DOI] [PMC free article] [PubMed] [Google Scholar]
90. Chen Y.W., Cleaver J.E., Hatahet Z., Honkanen R.E., Chang J.Y., Yen Y., Chou K.M. Human DNA polymerase eta activity and translocation is regulated by phosphorylation. Proc. Natl. Acad. Sci. U.S.A. 2008; 105:16578–16583. [DOI] [PMC free article] [PubMed] [Google Scholar]
91. Gohler T., Sabbioneda S., Green C.M., Lehmann A.R. ATR-mediated phosphorylation of DNA polymerase eta is needed for efficient recovery from UV damage. J. Cell Biol. 2011; 192:219–227. [DOI] [PMC free article] [PubMed] [Google Scholar]
92. Bertoletti F., Cea V., Liang C.C., Lanati T., Maffia A., Avarello M.D.M., Cipolla L., Lehmann A.R., Cohn M.A., Sabbioneda S. Phosphorylation regulates human poleta stability and damage bypass throughout the cell cycle. Nucleic Acids Res. 2017; 45:9441–9454. [DOI] [PMC free article] [PubMed] [Google Scholar]
93. Lehmann A.R., Kirk-Bell S., Arlett C.F., Paterson M.C., Lohman P.H., de Weerd-Kastelein E.A., Bootsma D Xeroderma pigmentosum cells with normal levels of excision repair have a defect in DNA synthesis after UV-irradiation. Proc. Natl. Acad. Sci. U.S.A. 1975; 72:219–223. [DOI] [PMC free article] [PubMed] [Google Scholar]
94. Cleaver J.E., Thomas G.H., Park S.D. Xeroderma pigmentosum variants have a slow recovery of DNA synthesis after irradiation with ultraviolet light. Biochim. Biophys. Acta. 1979; 564:122–131. [DOI] [PubMed] [Google Scholar]
95. Quinet A., Martins D.J., Vessoni A.T., Biard D., Sarasin A., Stary A., Menck C.F. Translesion synthesis mechanisms depend on the nature of DNA damage in UV-irradiated human cells. Nucleic Acids Res. 2016; 44:5717–5731. [DOI] [PMC free article] [PubMed] [Google Scholar]
96. Vallerga M.B., Mansilla S.F., Federico M.B., Bertolin A.P., Gottifredi V. Rad51 recombinase prevents Mre11 nuclease-dependent degradation and excessive PrimPol-mediated elongation of nascent DNA after UV irradiation. Proc. Natl. Acad. Sci. U.S.A. 2015; 112:E6624–6633. [DOI] [PMC free article] [PubMed] [Google Scholar]
97. Sokol A.M., Cruet-Hennequart S., Pasero P., Carty M.P. DNA polymerase eta modulates replication fork progression and DNA damage responses in platinum-treated human cells. Sci. Rep. 2013; 3:3277. [DOI] [PMC free article] [PubMed] [Google Scholar]
98. Despras E., Sittewelle M., Pouvelle C., Delrieu N., Cordonnier A.M., Kannouche P.L. Rad18-dependent SUMOylation of human specialized DNA polymerase eta is required to prevent under-replicated DNA. Nat. Commun. 2016; 7:13326. [DOI] [PMC free article] [PubMed] [Google Scholar]
99. Benureau Y., Pouvelle C., Dupaigne P., Baconnais S., Moreira Tavares E., Mazon G., Despras E., Le Cam E., Kannouche P.L. Changes in the architecture and abundance of replication intermediates delineate the chronology of DNA damage tolerance pathways at UV-stalled replication forks in human cells. Nucleic Acids Res. 2022; 50:9909–9929. [DOI] [PMC free article] [PubMed] [Google Scholar]
100. Lehmann A.R. Post-replication repair of DNA in ultraviolet-irradiated mammalian cells. No gaps in DNA synthesized late after ultraviolet irradiation. Eur. J. Biochem. 1972; 31:438–445. [DOI] [PubMed] [Google Scholar]
101. Meneghini R. Gaps in DNA synthesized by ultraviolet light-irradiated WI38 human cells. Biochim. Biophys. Acta. 1976; 425:419–427. [DOI] [PubMed] [Google Scholar]
102. Quinet A., Lerner L.K., Martins D.J., Menck C.F.M. Filling gaps in translesion DNA synthesis in human cells. Mutat. Res. Genet. Toxicol. Environ. Mutagen. 2018; 836:127–142. [DOI] [PubMed] [Google Scholar]
103. Quinet A., Tirman S., Cybulla E., Meroni A., Vindigni A. to skip or not to skip: choosing repriming to tolerate DNA damage. Mol. Cell. 2021; 81:649–658. [DOI] [PMC free article] [PubMed] [Google Scholar]
104. Lopes M., Foiani M., Sogo J.M. Multiple mechanisms control chromosome integrity after replication fork uncoupling and restart at irreparable UV lesions. Mol. Cell. 2006; 21:15–27. [DOI] [PubMed] [Google Scholar]
105. Diamant N., Hendel A., Vered I., Carell T., Reissner T., de Wind N., Geacinov N., Livneh Z. DNA damage bypass operates in the S and G2 phases of the cell cycle and exhibits differential mutagenicity. Nucleic Acids Res. 2012; 40:170–180. [DOI] [PMC free article] [PubMed] [Google Scholar]
106. Elvers I., Johansson F., Groth P., Erixon K., Helleday T. UV stalled replication forks restart by re-priming in human fibroblasts. Nucleic Acids Res. 2011; 39:7049–7057. [DOI] [PMC free article] [PubMed] [Google Scholar]
107. Zellweger R., Dalcher D., Mutreja K., Berti M., Schmid J.A., Herrador R., Vindigni A., Lopes M. Rad51-mediated replication fork reversal is a global response to genotoxic treatments in human cells. J. Cell Biol. 2015; 208:563–579. [DOI] [PMC free article] [PubMed] [Google Scholar]
108. Bianchi J., Rudd S.G., Jozwiakowski S.K., Bailey L.J., Soura V., Taylor E., Stevanovic I., Green A.J., Stracker T.H., Lindsay H.D. et al. PrimPol bypasses UV photoproducts during eukaryotic chromosomal DNA replication. Mol. Cell. 2013; 52:566–573. [DOI] [PMC free article] [PubMed] [Google Scholar]
109. Mourón S., Rodriguez-Acebes S., Martínez-Jiménez M.I., García-Gómez S., Chocrón S., Blanco L., Méndez J. Repriming of DNA synthesis at stalled replication forks by human PrimPol. Nat. Struct. Mol. Biol. 2013; 20:1383–1389. [DOI] [PubMed] [Google Scholar]
110. Guilliam T.A., Jozwiakowski S.K., Ehlinger A., Barnes R.P., Rudd S.G., Bailey L.J., Skehel J.M., Eckert K.A., Chazin W.J., Doherty A.J. Human PrimPol is a highly error-prone polymerase regulated by single-stranded DNA binding proteins. Nucleic Acids Res. 2015; 43:1056–1068. [DOI] [PMC free article] [PubMed] [Google Scholar]
111. Guilliam T.A., Brissett N.C., Ehlinger A., Keen B.A., Kolesar P., Taylor E.M., Bailey L.J., Lindsay H.D., Chazin W.J., Doherty A.J. Molecular basis for PrimPol recruitment to replication forks by RPA. Nat. Commun. 2017; 8:15222. [DOI] [PMC free article] [PubMed] [Google Scholar]
112. Bainbridge L.J., Teague R., Doherty A.J. Repriming DNA synthesis: an intrinsic restart pathway that maintains efficient genome replication. Nucleic Acids Res. 2021; 49:4831–4847. [DOI] [PMC free article] [PubMed] [Google Scholar]
113. Broomfield S., Chow B.L., Xiao W. MMS2, encoding a ubiquitin-conjugating-enzyme-like protein, is a member of the yeast error-free postreplication repair pathway. Proc. Natl. Acad. Sci. U.S.A. 1998; 95:5678–5683. [DOI] [PMC free article] [PubMed] [Google Scholar]
114. Hofmann R.M., Pickart C.M. Noncanonical MMS2-encoded ubiquitin-conjugating enzyme functions in assembly of novel polyubiquitin chains for DNA repair. Cell. 1999; 96:645–653. [DOI] [PubMed] [Google Scholar]
115. Johnson R.E., Henderson S.T., Petes T.D., Prakash S., Bankmann M., Prakash L. Saccharomyces cerevisiae RAD5-encoded DNA repair protein contains DNA helicase and zinc-binding sequence motifs and affects the stability of simple repetitive sequences in the genome. Mol. Cell. Biol. 1992; 12:3807–3818. [DOI] [PMC free article] [PubMed] [Google Scholar]
116. Elserafy M., Abugable A.A., Atteya R., El-Khamisy S.F. Rad5, HLTF, and SHPRH: a Fresh View of an Old Story. Trends Genet. 2018; 34:574–577. [DOI] [PMC free article] [PubMed] [Google Scholar]
117. Unk I., Hajdu I., Fatyol K., Szakal B., Blastyak A., Bermudez V., Hurwitz J., Prakash L., Prakash S., Haracska L. Human SHPRH is a ubiquitin ligase for Mms2-Ubc13-dependent polyubiquitylation of proliferating cell nuclear antigen. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:18107–18112. [DOI] [PMC free article] [PubMed] [Google Scholar]
118. Masuda Y., Suzuki M., Kawai H., Hishiki A., Hashimoto H., Masutani C., Hishida T., Suzuki F., Kamiya K. En bloc transfer of polyubiquitin chains to PCNA in vitro is mediated by two different human E2-E3 pairs. Nucleic Acids Res. 2012; 40:10394–10407. [DOI] [PMC free article] [PubMed] [Google Scholar]
119. Motegi A., Sood R., Moinova H., Markowitz S.D., Liu P.P., Myung K. Human SHPRH suppresses genomic instability through proliferating cell nuclear antigen polyubiquitination. J. Cell Biol. 2006; 175:703–708. [DOI] [PMC free article] [PubMed] [Google Scholar]
120. Unk I., Hajdú I., Fátyol K., Hurwitz J., Yoon J.-H., Prakash L., Prakash S., Haracska L. Human HLTF functions as a ubiquitin ligase for proliferating cell nuclear antigen polyubiquitination. Proc. Natl. Acad. Sci. U.S.A. 2008; 105:3768–3773. [DOI] [PMC free article] [PubMed] [Google Scholar]
121. Ciccia A., Nimonkar A.V., Hu Y., Hajdu I., Achar Y.J., Izhar L., Petit S.A., Adamson B., Yoon J.C., Kowalczykowski S.C. et al. Polyubiquitinated PCNA recruits the ZRANB3 translocase to maintain genomic integrity after replication stress. Mol. Cell. 2012; 47:396–409. [DOI] [PMC free article] [PubMed] [Google Scholar]
122. Hishiki A., Hara K., Ikegaya Y., Yokoyama H., Shimizu T., Sato M., Hashimoto H. Structure of a Novel DNA-binding Domain of Helicase-like Transcription Factor (HLTF) and Its Functional Implication in DNA Damage Tolerance. J. Biol. Chem. 2015; 290:13215–13223. [DOI] [PMC free article] [PubMed] [Google Scholar]
123. Yang Y., Gao Y., Mutter-Rottmayer L., Zlatanou A., Durando M., Ding W., Wyatt D., Ramsden D., Tanoue Y., Tateishi S. et al. DNA repair factor RAD18 and DNA polymerase Polkappa confer tolerance of oncogenic DNA replication stress. J. Cell Biol. 2017; 216:3097–3115. [DOI] [PMC free article] [PubMed] [Google Scholar]
124. Nayak S., Calvo J.A., Cong K., Peng M., Berthiaume E., Jackson J., Zaino A.M., Vindigni A., Hadden M.K., Cantor S.B. Inhibition of the translesion synthesis polymerase REV1 exploits replication gaps as a cancer vulnerability. Sci. Adv. 2020; 6:eaaz7808. [DOI] [PMC free article] [PubMed] [Google Scholar]
125. Cong K., Peng M., Kousholt A.N., Lee W.T.C., Lee S., Nayak S., Krais J., VanderVere-Carozza P.S., Pawelczak K.S., Calvo J. et al. Replication gaps are a key determinant of PARP inhibitor synthetic lethality with BRCA deficiency. Mol. Cell. 2021; 81:3128–3144. [DOI] [PMC free article] [PubMed] [Google Scholar]
126. Cong K., Cantor S.B. Exploiting replication gaps for cancer therapy. Mol. Cell. 2022; 82:2363–2369. [DOI] [PMC free article] [PubMed] [Google Scholar]
127. Taglialatela A., Leuzzi G., Sannino V., Cuella-Martin R., Huang J.-W., Wu-Baer F., Baer R., Costanzo V., Ciccia A. REV1-Polζ maintains the viability of homologous recombination-deficient cancer cells through mutagenic repair of PRIMPOL-dependent ssDNA gaps. Mol. Cell. 2021; 81:4008–4025. [DOI] [PMC free article] [PubMed] [Google Scholar]
128. Alexandrov L.B., Kim J., Haradhvala N.J., Huang M.N., Tian Ng A.W., Wu Y., Boot A., Covington K.R., Gordenin D.A., Bergstrom E.N. et al. The repertoire of mutational signatures in human cancer. Nature. 2020; 578:94–101. [DOI] [PMC free article] [PubMed] [Google Scholar]
129. Roberts S.A., Sterling J., Thompson C., Harris S., Mav D., Shah R., Klimczak L.J., Kryukov G.V., Malc E., Mieczkowski P.A. et al. Clustered mutations in yeast and in human cancers can arise from damaged long single-strand DNA regions. Mol. Cell. 2012; 46:424–435. [DOI] [PMC free article] [PubMed] [Google Scholar]
130. Morganella S., Alexandrov L.B., Glodzik D., Zou X., Davies H., Staaf J., Sieuwerts A.M., Brinkman A.B., Martin S., Ramakrishna M. et al. The topography of mutational processes in breast cancer genomes. Nat. Commun. 2016; 7:11383. [DOI] [PMC free article] [PubMed] [Google Scholar]
131. Petljak M., Dananberg A., Chu K., Bergstrom E.N., Striepen J., von Morgen P., Chen Y., Shah H., Sale J.E., Alexandrov L.B. et al. Mechanisms of APOBEC3 mutagenesis in human cancer cells. Nature. 2022; 607:799–807. [DOI] [PMC free article] [PubMed] [Google Scholar]
132. Delbos F., De Smet A., Faili A., Aoufouchi S., Weill J.C., Reynaud C.A. Contribution of DNA polymerase eta to immunoglobulin gene hypermutation in the mouse. J. Exp. Med. 2005; 201:1191–1196. [DOI] [PMC free article] [PubMed] [Google Scholar]
133. Supek F., Lehner B. Clustered mutation signatures reveal that error-prone DNA repair targets mutations to active genes. Cell. 2017; 170:534–547. [DOI] [PubMed] [Google Scholar]
134. Xu H.L., Gao X.R., Zhang W., Cheng J.R., Tan Y.T., Zheng W., Shu X.O., Xiang Y.B. Effects of polymorphisms in translesion DNA synthesis genes on lung cancer risk and prognosis in Chinese men. Cancer Epidemiol. 2013; 37:917–922. [DOI] [PMC free article] [PubMed] [Google Scholar]
135. Dai Z.J., Liu X.H., Ma Y.F., Kang H.F., Jin T.B., Dai Z.M., Guan H.T., Wang M., Liu K., Dai C. et al. Association between single nucleotide polymorphisms in DNA polymerase kappa gene and breast cancer risk in chinese Han population: a STROBE-Compliant Observational Study. Medicine (Baltimore). 2016; 95:e2466. [DOI] [PMC free article] [PubMed] [Google Scholar]
136. Sakiyama T., Kohno T., Mimaki S., Ohta T., Yanagitani N., Sobue T., Kunitoh H., Saito R., Shimizu K., Hirama C. et al. Association of amino acid substitution polymorphisms in DNA repair genes TP53, POLI, REV1 and LIG4 with lung cancer risk. Int. J. Cancer. 2005; 114:730–737. [DOI] [PubMed] [Google Scholar]
137. Yeom M., Kim I.H., Kim J.K., Kang K., Eoff R.L., Guengerich F.P., Choi J.Y. Effects of twelve germline missense variations on DNA lesion and G-quadruplex bypass activities of human DNA polymerase REV1. Chem. Res. Toxicol. 2016; 29:367–379. [DOI] [PMC free article] [PubMed] [Google Scholar]
138. Zhao G., Gleave E.S., Lamers M.H. Single-molecule studies contrast ordered DNA replication with stochastic translesion synthesis. Elife. 2017; 6:e3217. [DOI] [PMC free article] [PubMed] [Google Scholar]
139. Sasatani M., Xi Y., Kajimura J., Kawamura T., Piao J., Masuda Y., Honda H., Kubo K., Mikamoto T., Watanabe H. et al. Overexpression of Rev1 promotes the development of carcinogen-induced intestinal adenomas via accumulation of point mutation and suppression of apoptosis proportionally to the Rev1 expression level. Carcinogenesis. 2017; 38:570–578. [DOI] [PMC free article] [PubMed] [Google Scholar]
140. Jung Y.S., Qian Y., Chen X. DNA polymerase eta is targeted by Mdm2 for polyubiquitination and proteasomal degradation in response to ultraviolet irradiation. DNA Repair (Amst.). 2012; 11:177–184. [DOI] [PMC free article] [PubMed] [Google Scholar]
141. Ziv O., Zeisel A., Mirlas-Neisberg N., Swain U., Nevo R., Ben-Chetrit N., Martelli M.P., Rossi R., Schiesser S., Canman C.E. et al. Identification of novel DNA-damage tolerance genes reveals regulation of translesion DNA synthesis by nucleophosmin. Nat. Commun. 2014; 5:5437. [DOI] [PMC free article] [PubMed] [Google Scholar]
142. Sitz J., Blanchet S.A., Gameiro S.F., Biquand E., Morgan T.M., Galloy M., Dessapt J., Lavoie E.G., Blondeau A., Smith B.C. et al. Humanpapillomavirus E7 oncoprotein targets RNF168 to hijack the host DNA damageresponse. Proc.Natl. Acad. Sci. U.S.A. 2019; 24:19552–19562. [DOI] [PMC free article] [PubMed] [Google Scholar]
143. Cipolla L., Bertoletti F., Maffia A., Liang C.C., Lehmann A.R., Cohn M.A., Sabbioneda S. UBR5 interacts with the replication fork and protects DNA replication from DNA polymerase eta toxicity. Nucleic Acids Res. 2019; 47:11268–11283. [DOI] [PMC free article] [PubMed] [Google Scholar]
144. Wittschieben J.P., Patil V., Glushets V., Robinson L.J., Kusewitt D.F., Wood R.D. Loss of DNA polymerase zeta enhances spontaneous tumorigenesis. Cancer Res. 2010; 70:2770–2778. [DOI] [PMC free article] [PubMed] [Google Scholar]
145. O-Wang J., Kawamura K., Tada Y., Ohmori H., Kimura H., Sakiyama S., Tagawa M. DNA polymerase kappa, implicated in spontaneous and DNA damage-induced mutagenesis, is overexpressed in lung cancer. Cancer Res. 2001; 61:5366–5369. [PubMed] [Google Scholar]
146. Wang Y., Seimiya M., Kawamura K., Yu L., Ogi T., Takenaga K., Shishikura T., Nakagawara A., Sakiyama S., Tagawa M. et al. Elevated expression of DNA polymerase kappa in human lung cancer is associated with p53 inactivation: negative regulation of POLK promoter activity by p53. Int. J. Oncol. 2004; 25:161–165. [PubMed] [Google Scholar]
147. Zhou J., Zhang S., Xie L., Liu P., Xie F., Wu J., Cao J., Ding W.Q. Overexpression of DNA polymerase iota (Poliota) in esophageal squamous cell carcinoma. Cancer Sci. 2012; 103:1574–1579. [DOI] [PMC free article] [PubMed] [Google Scholar]
148. Rimkus C., Friederichs J., Rosenberg R., Holzmann B., Siewert J.R., Janssen K.P. Expression of the mitotic checkpoint gene MAD2L2 has prognostic significance in colon cancer. Int. J. Cancer. 2007; 120:207–211. [DOI] [PubMed] [Google Scholar]
149. Moinova H.R., Chen W.D., Shen L., Smiraglia D., Olechnowicz J., Ravi L., Kasturi L., Myeroff L., Plass C., Parsons R. et al. HLTF gene silencing in human colon cancer. Proc. Natl. Acad. Sci. U.S.A. 2002; 99:4562–4567. [DOI] [PMC free article] [PubMed] [Google Scholar]
150. Yang Y., Gao Y., Zlatanou A., Tateishi S., Yurchenko V., Rogozin I.B., Vaziri C. Diverse roles of RAD18 and Y-family DNA polymerases in tumorigenesis. Cell Cycle. 2018; 17:833–843. [DOI] [PMC free article] [PubMed] [Google Scholar]
151. Kotsantis P., Petermann E., Boulton S.J. Mechanisms of oncogene-induced replication stress: jigsaw falling into place. Cancer Discov. 2018; 8:537–555. [DOI] [PMC free article] [PubMed] [Google Scholar]
152. Primo L.M.F., Teixeira L.K. DNA replication stress: oncogenes in the spotlight. Genet Mol Biol. 2019; 43:e20190138. [DOI] [PMC free article] [PubMed] [Google Scholar]
153. Kotsantis P., Silva L.M., Irmscher S., Jones R.M., Folkes L., Gromak N., Petermann E. Increased global transcription activity as a mechanism of replication stress in cancer. Nat. Commun. 2016; 7:13087. [DOI] [PMC free article] [PubMed] [Google Scholar]
154. Chang L., Ruiz P., Ito T., Sellers W.R. Targeting pan-essential genes in cancer: challenges and opportunities. Cancer Cell. 2021; 39:466–479. [DOI] [PMC free article] [PubMed] [Google Scholar]
155. Chen Y., Jie X., Xing B., Wu Z., Yang X., Rao X., Xu Y., Zhou D., Dong X., Zhang T. et al. REV1 promotes lung tumorigenesis by activating the Rad18/SERTAD2 axis. Cell Death. Dis. 2022; 13:110. [DOI] [PMC free article] [PubMed] [Google Scholar]
156. Zou S., Yang J., Guo J., Su Y., He C., Wu J., Yu L., Ding W.Q., Zhou J. RAD18 promotes the migration and invasion of esophageal squamous cell cancer via the JNK-MMPs pathway. Cancer Lett. 2018; 417:65–74. [DOI] [PubMed] [Google Scholar]
157. Yan X., He Y., Yang S., Zeng T., Hua Y., Bao S., Yang F., Duan N., Sun C., Liang Y. et al. a positive feedback loop: rAD18-YAP-TGF-beta between triple-negative breast cancer and macrophages regulates cancer stemness and progression. Cell Death Discov. 2022; 8:196. [DOI] [PMC free article] [PubMed] [Google Scholar]
158. Panzarino N.J., Krais J.J., Cong K., Peng M., Mosqueda M., Nayak S.U., Bond S.M., Calvo J.A., Doshi M.B., Bere M. et al. Replication gaps underlie BRCA deficiency and therapy response. Cancer Res. 2021; 81:1388–1397. [DOI] [PMC free article] [PubMed] [Google Scholar]
159. Tarantino D., Walker C., Weekes D., Pemberton H., Davidson K., Torga G., Frankum J., Mendes-Pereira A.M., Prince C., Ferro R. et al. Functional screening reveals HORMAD1-driven gene dependencies associated with translesion synthesis and replication stress tolerance. Oncogene. 2022; 41:3969–3977. [DOI] [PMC free article] [PubMed] [Google Scholar]
160. Watkins J., Weekes D., Shah V., Gazinska P., Joshi S., Sidhu B., Gillett C., Pinder S., Vanoli F., Jasin M. et al. Genomic complexity profiling reveals that HORMAD1 overexpression contributes to homologous recombination deficiency in triple-negative breast cancers. Cancer Discov. 2015; 5:488–505. [DOI] [PMC free article] [PubMed] [Google Scholar]
161. Gao Y., Kardos J., Yang Y., Tamir T.Y., Mutter-Rottmayer E., Weissman B., Major M.B., Kim W.Y., Vaziri C. The cancer/testes (CT) antigen HORMAD1 promotes homologous recombinational DNA repair and radioresistance in lung adenocarcinoma cells. Sci. Rep. 2018; 8:15304. [DOI] [PMC free article] [PubMed] [Google Scholar]
162. Nichols B.A., Oswald N.W., McMillan E.A., McGlynn K., Yan J., Kim M.S., Saha J., Mallipeddi P.L., LaDuke S.A., Villalobos P.A. et al. HORMAD1 is a negative prognostic indicator in lung adenocarcinoma and specifies resistance to oxidative and genotoxic stress. Cancer Res. 2018; 78:6196–6208. [DOI] [PMC free article] [PubMed] [Google Scholar]
163. Shimizu T., Tateishi S., Tanoue Y., Azuma T., Ohmori H. Somatic hypermutation of immunoglobulin genes in Rad18 knockout mice. DNA Repair (Amst.). 2017; 50:54–60. [DOI] [PubMed] [Google Scholar]
164. Lou J., Yang Y., Gu Q., Price B.A., Qiu Y., Fedoriw Y., Desai S., Mose L.E., Chen B., Tateishi S. et al. Rad18 mediates specific mutational signatures and shapes the genomic landscape of carcinogen-induced tumors in vivo. NAR Cancer. 2021; 3:zcaa037. [DOI] [PMC free article] [PubMed] [Google Scholar]
165. Stancel J.N., McDaniel L.D., Velasco S., Richardson J., Guo C., Friedberg E.C. Polk mutant mice have a spontaneous mutator phenotype. DNA Repair (Amst.). 2009; 8:1355–1362. [DOI] [PMC free article] [PubMed] [Google Scholar]
166. Jansen J.G., Langerak P., Tsaalbi-Shtylik A., van den Berk P., Jacobs H., de Wind N. Strand-biased defect in C/G transversions in hypermutating immunoglobulin genes in Rev1-deficient mice. J. Exp. Med. 2006; 203:319–323. [DOI] [PMC free article] [PubMed] [Google Scholar]
167. Fakouri N.B., Durhuus J.A., Regnell C.E., Angleys M., Desler C., Hasan-Olive M.M., Martin-Pardillos A., Tsaalbi-Shtylik A., Thomsen K., Lauritzen M. et al. Rev1 contributes to proper mitochondrial function via the PARP-NAD(+)-SIRT1-PGC1alpha axis. Sci. Rep. 2017; 7:12480. [DOI] [PMC free article] [PubMed] [Google Scholar]
168. Dumstorf C.A., Clark A.B., Lin Q., Kissling G.E., Yuan T., Kucherlapati R., McGregor W.G., Kunkel T.A. Participation of mouse DNA polymerase iota in strand-biased mutagenic bypass of UV photoproducts and suppression of skin cancer. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:18083–18088. [DOI] [PMC free article] [PubMed] [Google Scholar]
169. Ohkumo T., Kondo Y., Yokoi M., Tsukamoto T., Yamada A., Sugimoto T., Kanao R., Higashi Y., Kondoh H., Tatematsu M. et al. 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. 2006; 26:7696–7706. [DOI] [PMC free article] [PubMed] [Google Scholar]
170. Watanabe N., Mii S., Asai N., Asai M., Niimi K., Ushida K., Kato T., Enomoto A., Ishii H., Takahashi M. et al. the REV7 subunit of DNA polymerase zeta is essential for primordial germ cell maintenance in the mouse. J. Biol. Chem. 2013; 288:10459–10471. [DOI] [PMC free article] [PubMed] [Google Scholar]
171. Bemark M., Khamlichi A.A., Davies S.L., Neuberger M.S. Disruption of mouse polymerase zeta (Rev3) leads to embryonic lethality and impairs blastocyst development in vitro. Curr. Biol. 2000; 10:1213–1216. [DOI] [PubMed] [Google Scholar]
172. Esposito G., Godindagger I., Klein U., Yaspo M.L., Cumano A., Rajewsky K. Disruption of the Rev3l-encoded catalytic subunit of polymerase zeta in mice results in early embryonic lethality. Curr. Biol. 2000; 10:1221–1224. [DOI] [PubMed] [Google Scholar]
173. Wittschieben J., Shivji M.K., Lalani E., Jacobs M.A., Marini F., Gearhart P.J., Rosewell I., Stamp G., Wood R.D. Disruption of the developmentally regulated Rev3l gene causes embryonic lethality. Curr. Biol. 2000; 10:1217–1220. [DOI] [PubMed] [Google Scholar]
174. Lange S.S., Bedford E., Reh S., Wittschieben J.P., Carbajal S., Kusewitt D.F., DiGiovanni J., Wood R.D. Dual role for mammalian DNA polymerase zeta in maintaining genome stability and proliferative responses. Proc. Natl. Acad. Sci. U.S.A. 2013; 110:E687–696. [DOI] [PMC free article] [PubMed] [Google Scholar]
175. Lin Q., Clark A.B., McCulloch S.D., Yuan T., Bronson R.T., Kunkel T.A., Kucherlapati R. Increased susceptibility to UV-induced skin carcinogenesis in polymerase eta-deficient mice. Cancer Res. 2006; 66:87–94. [DOI] [PubMed] [Google Scholar]
176. Tanoue Y., Toyoda T., Sun J., Mustofa M.K., Tateishi C., Endo S., Motoyama N., Araki K., Wu D., Okuno Y. et al. Differential roles of Rad18 and Chk2 in genome maintenance and skin carcinogenesis following UV exposure. J. Invest. Dermatol. 2018; 138:2550–2557. [DOI] [PubMed] [Google Scholar]
177. Kelland L. the resurgence of platinum-based cancer chemotherapy. Nat. Rev. Cancer. 2007; 7:573–584. [DOI] [PubMed] [Google Scholar]
178. Kartalou M., Essigmann J.M. Mechanisms of resistance to cisplatin. Mutat. Res. 2001; 478:23–43. [DOI] [PubMed] [Google Scholar]
179. Galluzzi L., Senovilla L., Vitale I., Michels J., Martins I., Kepp O., Castedo M., Kroemer G. Molecular mechanisms of cisplatin resistance. Oncogene. 2012; 31:1869–1883. [DOI] [PubMed] [Google Scholar]
180. Park S.B., Krishnan A.V., Lin C.S., Goldstein D., Friedlander M., Kiernan M.C. Mechanisms underlying chemotherapy-induced neurotoxicity and the potential for neuroprotective strategies. Curr. Med. Chem. 2008; 15:3081–3094. [DOI] [PubMed] [Google Scholar]
181. Sioka C., Kyritsis A.P. Central and peripheral nervous system toxicity of common chemotherapeutic agents. Cancer Chemother. Pharmacol. 2009; 63:761–767. [DOI] [PubMed] [Google Scholar]
182. Lord C.J., Ashworth A. the DNA damage response and cancer therapy. Nature. 2012; 481:287–294. [DOI] [PubMed] [Google Scholar]
183. Mamenta E.L., Poma E.E., Kaufmann W.K., Delmastro D.A., Grady H.L., Chaney S.G. Enhanced replicative bypass of platinum-DNA adducts in cisplatin-resistant human ovarian carcinoma cell lines. Cancer Res. 1994; 54:3500–3505. [PubMed] [Google Scholar]
184. Bassett E., King N.M., Bryant M.F., Hector S., Pendyala L., Chaney S.G., Cordeiro-Stone M. the role of DNA polymerase eta in translesion synthesis past platinum-DNA adducts in human fibroblasts. Cancer Res. 2004; 64:6469–6475. [DOI] [PubMed] [Google Scholar]
185. Ummat A., Rechkoblit O., Jain R., Roy Choudhury J., Johnson R.E., Silverstein T.D., Buku A., Lone S., Prakash L., Prakash S. et al. Structural basis for cisplatin DNA damage tolerance by human polymerase eta during cancer chemotherapy. Nat. Struct. Mol. Biol. 2012; 19:628–632. [DOI] [PMC free article] [PubMed] [Google Scholar]
186. Zhao Y., Biertumpfel C., Gregory M.T., Hua Y.J., Hanaoka F., Yang W. Structural basis of human DNA polymerase eta-mediated chemoresistance to cisplatin. Proc. Natl. Acad. Sci. U.S.A. 2012; 109:7269–7274. [DOI] [PMC free article] [PubMed] [Google Scholar]
187. Chen Y.W., Cleaver J.E., Hanaoka F., Chang C.F., Chou K.M. a novel role of DNA polymerase eta in modulating cellular sensitivity to chemotherapeutic agents. Mol. Cancer Res. 2006; 4:257–265. [DOI] [PubMed] [Google Scholar]
188. Albertella M.R., Green C.M., Lehmann A.R., O’Connor M.J a role for polymerase eta in the cellular tolerance to cisplatin-induced damage. Cancer Res. 2005; 65:9799–9806. [DOI] [PubMed] [Google Scholar]
189. Alt A., Lammens K., Chiocchini C., Lammens A., Pieck J.C., Kuch D., Hopfner K.P., Carell T. Bypass of DNA lesions generated during anticancer treatment with cisplatin by DNA polymerase eta. Science. 2007; 318:967–970. [DOI] [PubMed] [Google Scholar]
190. Wagner J.M., Karnitz L.M. Cisplatin-induced DNA damage activates replication checkpoint signaling components that differentially affect tumor cell survival. Mol. Pharmacol. 2009; 76:208–214. [DOI] [PMC free article] [PubMed] [Google Scholar]
191. Yamashita Y.M., Okada T., Matsusaka T., Sonoda E., Zhao G.Y., Araki K., Tateishi S., Yamaizumi M., Takeda S. RAD18 and RAD54 cooperatively contribute to maintenance of genomic stability in vertebrate cells. EMBO J. 2002; 21:5558–5566. [DOI] [PMC free article] [PubMed] [Google Scholar]
192. Ceppi P., Novello S., Cambieri A., Longo M., Monica V., Lo Iacono M., Giaj-Levra M., Saviozzi S., Volante M., Papotti M. et al. Polymerase eta mRNA expression predicts survival of non-small cell lung cancer patients treated with platinum-based chemotherapy. Clin. Cancer Res. 2009; 15:1039–1045. [DOI] [PubMed] [Google Scholar]
193. Teng K.Y., Qiu M.Z., Li Z.H., Luo H.Y., Zeng Z.L., Luo R.Z., Zhang H.Z., Wang Z.Q., Li Y.H., Xu R.H. DNA polymerase eta protein expression predicts treatment response and survival of metastatic gastric adenocarcinoma patients treated with oxaliplatin-based chemotherapy. J. Transl. Med. 2010; 8:126. [DOI] [PMC free article] [PubMed] [Google Scholar]
194. Srivastava A.K., Han C., Zhao R., Cui T., Dai Y., Mao C., Zhao W., Zhang X., Yu J., Wang Q.E. Enhanced expression of DNA polymerase eta contributes to cisplatin resistance of ovarian cancer stem cells. Proc. Natl. Acad. Sci. U.S.A. 2015; 112:4411–4416. [DOI] [PMC free article] [PubMed] [Google Scholar]
195. Thul P.J., Akesson L., Wiking M., Mahdessian D., Geladaki A., Ait Blal H., Alm T., Asplund A., Bjork L., Breckels L.M. et al. A subcellular map of the human proteome. Science. 2017; 356:eaal3321. [DOI] [PubMed] [Google Scholar]
196. Uhlen M., Fagerberg L., Hallstrom B.M., Lindskog C., Oksvold P., Mardinoglu A., Sivertsson A., Kampf C., Sjostedt E., Asplund A. et al. Proteomics. Tissue-based map of the human proteome. Science. 2015; 347:1260419. [DOI] [PubMed] [Google Scholar]
197. Uhlen M., Zhang C., Lee S., Sjostedt E., Fagerberg L., Bidkhori G., Benfeitas R., Arif M., Liu Z., Edfors F. et al. a pathology atlas of the human cancer transcriptome. Science. 2017; 357:eaan2507. [DOI] [PubMed] [Google Scholar]
198. Doles J., Oliver T.G., Cameron E.R., Hsu G., Jacks T., Walker G.C., Hemann M.T. Suppression of Rev3, the catalytic subunit of Pol{zeta}, sensitizes drug-resistant lung tumors to chemotherapy. Proc. Natl. Acad. Sci. U.S.A. 2010; 107:20786–20791. [DOI] [PMC free article] [PubMed] [Google Scholar]
199. Xie K., Doles J., Hemann M.T., Walker G.C. Error-prone translesion synthesis mediates acquired chemoresistance. Proc. Nat. Acad. Sci. U.S.A. 2010; 107:20792–20797. [DOI] [PMC free article] [PubMed] [Google Scholar]
200. Xu X., Xie K., Zhang X.Q., Pridgen E.M., Park G.Y., Cui D.S., Shi J., Wu J., Kantoff P.W., Lippard S.J. et al. Enhancing tumor cell response to chemotherapy through nanoparticle-mediated codelivery of siRNA and cisplatin prodrug. Proc. Natl. Acad. Sci. U.S.A. 2013; 110:18638–18643. [DOI] [PMC free article] [PubMed] [Google Scholar]
201. Vassel F.M., Bian K., Walker G.C., Hemann M.T. Rev7 loss alters cisplatin response and increases drug efficacy in chemotherapy-resistant lung cancer. Proc. Natl. Acad. Sci. U.S.A. 2020; 117:28922–28924. [DOI] [PMC free article] [PubMed] [Google Scholar]
202. Ciccia A., Elledge S.J. the DNA damage response: making it safe to play with knives. Mol. Cell. 2010; 40:179–204. [DOI] [PMC free article] [PubMed] [Google Scholar]
203. Baillie K.E., Stirling P.C. Beyond kinases: targeting replication stress proteins in cancer therapy. Trends Cancer. 2021; 7:430–446. [DOI] [PubMed] [Google Scholar]
204. Smith J., Tho L.M., Xu N., Gillespie D.A. the ATM-Chk2 and ATR-Chk1 pathways in DNA damage signaling and cancer. Adv. Cancer. Res. 2010; 108:73–112. [DOI] [PubMed] [Google Scholar]
205. Yazinski S.A., Zou L. Functions, regulation, and therapeutic implications of the ATR checkpoint pathway. Annu. Rev. Genet. 2016; 50:155–173. [DOI] [PubMed] [Google Scholar]
206. Pilie P.G., Tang C., Mills G.B., Yap T.A. State-of-the-art strategies for targeting the DNA damage response in cancer. Nat. Rev. Clin. Oncol. 2019; 16:81–104. [DOI] [PMC free article] [PubMed] [Google Scholar]
207. Menendez D., Anand J.R., Murphy C.C., Bell W.J., Fu J., Slepushkina N., Buehler E., Martin S.E., Lal-Nag M., Nitiss J.L. et al. Etoposide-induced DNA damage is increased in p53 mutants: identification of ATR and other genes that influence effects of p53 mutations on Top2-induced cytotoxicity. OncoTargets Ther. 2022; 13:332–346. [DOI] [PMC free article] [PubMed] [Google Scholar]
208. Karnitz L.M., Zou L. Molecular pathways: targeting ATR in cancer therapy. Clin. Cancer Res. 2015; 21:4780–4785. [DOI] [PMC free article] [PubMed] [Google Scholar]
209. Despras E., Daboussi F., Hyrien O., Marheineke K., Kannouche P.L. ATR/Chk1 pathway is essential for resumption of DNA synthesis and cell survival in UV-irradiated XP variant cells. Hum. Mol. Genet. 2010; 19:1690–1701. [DOI] [PubMed] [Google Scholar]
210. Maher V.M., Ouellette L.M., Mittlestat M., McCormick J.J. Synergistic effect of caffeine on the cytotoxicity of ultraviolet irradiation and of hydrocarbon epoxides in strains of Xeroderma pigmentosum. Nature. 1975; 258:760–763. [DOI] [PubMed] [Google Scholar]
211. Elbaek C.R., Petrosius V., Sorensen C.S. WEE1 kinase limits CDK activities to safeguard DNA replication and mitotic entry. Mutat. Res. 2020; 819-820:111694. [DOI] [PubMed] [Google Scholar]
212. Puigvert J.C., Sanjiv K., Helleday T. Targeting DNA repair, DNA metabolism and replication stress as anti-cancer strategies. FEBS J. 2016; 283:232–245. [DOI] [PubMed] [Google Scholar]
213. Matheson C.J., Backos D.S., Reigan P. Targeting WEE1 Kinase in Cancer. Trends Pharmacol. Sci. 2016; 37:872–881. [DOI] [PubMed] [Google Scholar]
214. Lheureux S., Cristea M.C., Bruce J.P., Garg S., Cabanero M., Mantia-Smaldone G., Olawaiye A.B., Ellard S.L., Weberpals J.I., Wahner Hendrickson A.E. et al. Adavosertib plus gemcitabine for platinum-resistant or platinum-refractory recurrent ovarian cancer: a double-blind, randomised, placebo-controlled, phase 2 trial. Lancet North Am. Ed. 2021; 397:281–292. [DOI] [PMC free article] [PubMed] [Google Scholar]
215. Nayak S., Calvo J.A., Cantor S.B. Targeting translesion synthesis (TLS) to expose replication gaps, a unique cancer vulnerability. Expert Opin. Ther. Targets. 2021; 25:27–36. [DOI] [PMC free article] [PubMed] [Google Scholar]
216. Satoh M.S., Lindahl T. Role of poly(ADP-ribose) formation in DNA repair. Nature. 1992; 356:356–358. [DOI] [PubMed] [Google Scholar]
217. Krishnakumar R., Kraus W.L. The PARP side of the nucleus: molecular actions, physiological outcomes, and clinical targets. Mol. Cell. 2010; 39:8–24. [DOI] [PMC free article] [PubMed] [Google Scholar]
218. Bryant H.E., Schultz N., Thomas H.D., Parker K.M., Flower D., Lopez E., Kyle S., Meuth M., Curtin N.J., Helleday T. Specific killing of BRCA2-deficient tumours with inhibitors of poly(ADP-ribose) polymerase. Nature. 2005; 434:913–917. [DOI] [PubMed] [Google Scholar]
219. Farmer H., McCabe N., Lord C.J., Tutt A.N., Johnson D.A., Richardson T.B., Santarosa M., Dillon K.J., Hickson I., Knights C. et al. Targeting the DNA repair defect in BRCA mutant cells as a therapeutic strategy. Nature. 2005; 434:917–921. [DOI] [PubMed] [Google Scholar]
220. Lord C.J., Ashworth A. PARP inhibitors: synthetic lethality in the clinic. Science. 2017; 355:1152–1158. [DOI] [PMC free article] [PubMed] [Google Scholar]
221. Noordermeer S.M., van Attikum H. PARP inhibitor resistance: a tug-of-war in BRCA-mutated cells. Trends Cell Biol. 2019; 29:820–834. [DOI] [PubMed] [Google Scholar]
222. Helleday T. The underlying mechanism for the PARP and BRCA synthetic lethality: clearing up the misunderstandings. Mol Oncol. 2011; 5:387–393. [DOI] [PMC free article] [PubMed] [Google Scholar]
223. Paes Dias M., Tripathi V., van der Heijden I., Cong K., Manolika E.M., Bhin J., Gogola E., Galanos P., Annunziato S., Lieftink C. et al. Loss of nuclear DNA ligase III reverts PARP inhibitor resistance in BRCA1/53BP1 double-deficient cells by exposing ssDNA gaps. Mol. Cell. 2021; 81:4692–4708. [DOI] [PMC free article] [PubMed] [Google Scholar]
224. Johnson D.R., O’Neill B.P Glioblastoma survival in the United States before and during the temozolomide era. J. Neurooncol. 2012; 107:359–364. [DOI] [PubMed] [Google Scholar]
225. Lonardi S., Tosoni A., Brandes A.A. Adjuvant chemotherapy in the treatment of high grade gliomas. Cancer Treat. Rev. 2005; 31:79–89. [DOI] [PubMed] [Google Scholar]
226. Barthel F.P., Johnson K.C., Varn F.S., Moskalik A.D., Tanner G., Kocakavuk E., Anderson K.J., Abiola O., Aldape K., Alfaro K.D. et al. Longitudinal molecular trajectories of diffuse glioma in adults. Nature. 2019; 576:112–120. [DOI] [PMC free article] [PubMed] [Google Scholar]
227. Brennan C.W., Verhaak R.G., McKenna A., Campos B., Noushmehr H., Salama S.R., Zheng S., Chakravarty D., Sanborn J.Z., Berman S.H. et al. The somatic genomic landscape of glioblastoma. Cell. 2013; 155:462–477. [DOI] [PMC free article] [PubMed] [Google Scholar]
228. Wang J., Cazzato E., Ladewig E., Frattini V., Rosenbloom D.I., Zairis S., Abate F., Liu Z., Elliott O., Shin Y.J. et al. Clonal evolution of glioblastoma under therapy. Nat. Genet. 2016; 48:768–776. [DOI] [PMC free article] [PubMed] [Google Scholar]
229. Alexandrov L.B., Nik-Zainal S., Wedge D.C., Aparicio S.A., Behjati S., Biankin A.V., Bignell G.R., Bolli N., Borg A., Borresen-Dale A.L. et al. Signatures of mutational processes in human cancer. Nature. 2013; 500:415–421. [DOI] [PMC free article] [PubMed] [Google Scholar]
230. Touat M., Li Y.Y., Boynton A.N., Spurr L.F., Iorgulescu J.B., Bohrson C.L., Cortes-Ciriano I., Birzu C., Geduldig J.E., Pelton K. et al. Mechanisms and therapeutic implications of hypermutation in gliomas. Nature. 2020; 580:517–523. [DOI] [PMC free article] [PubMed] [Google Scholar]
231. Latancia M.T., Moreno N.C., Leandro G.S., Ribeiro V.C., de Souza I., Vieira W.K.M., Bastos A.U., Hoch N.C., Rocha C.R.R., Menck C.F.M. DNA polymerase eta protects human cells against DNA damage induced by the tumor chemotherapeutic temozolomide. Mutat. Res. Genet. Toxicol. Environ. Mutagen. 2022; 878:503498. [DOI] [PubMed] [Google Scholar]
232. Ketkar A., Maddukuri L., Penthala N.R., Reed M.R., Zafar M.K., Crooks P.A., Eoff R.L. Inhibition of human DNA polymerases eta and kappa by indole-derived molecules occurs through distinct mechanisms. ACS Chem. Biol. 2019; 14:1337–1351. [DOI] [PMC free article] [PubMed] [Google Scholar]
233. Rocha C.R.R., Reily Rocha A., Molina Silva M., Rodrigues Gomes L., Teatin Latancia M., Andrade Tomaz M., de Souza I., Karolynne Seregni Monteiro L., Menck C.F.M. Revealing temozolomide resistance mechanisms via genome-wide CRISPR libraries. Cells. 2020; 9:2573. [DOI] [PMC free article] [PubMed] [Google Scholar]
234. Peng C., Chen Z., Wang S., Wang H.W., Qiu W., Zhao L., Xu R., Luo H., Chen Y., Chen D. et al. The error-prone DNA polymerase kappa promotes temozolomide resistance in glioblastoma through Rad17-dependent activation of ATR-Chk1 signaling. Cancer Res. 2016; 76:2340–2353. [DOI] [PubMed] [Google Scholar]
235. Wang H., Wu W., Wang H.W., Wang S., Chen Y., Zhang X., Yang J., Zhao S., Ding H.F., Lu D Analysis of specialized DNA polymerases expression in human gliomas: association with prognostic significance. Neuro. Oncol. 2010; 12:679–686. [DOI] [PMC free article] [PubMed] [Google Scholar]
236. Szikriszt B., Poti A., Pipek O., Krzystanek M., Kanu N., Molnar J., Ribli D., Szeltner Z., Tusnady G.E., Csabai I. et al. a comprehensive survey of the mutagenic impact of common cancer cytotoxics. Genome Biol. 2016; 17:99. [DOI] [PMC free article] [PubMed] [Google Scholar]
237. Lee Y.S., Gregory M.T., Yang W. Human Pol zeta purified with accessory subunits is active in translesion DNA synthesis and complements Pol eta in cisplatin bypass. Proc. Natl. Acad. Sci. U.S.A. 2014; 111:2954–2959. [DOI] [PMC free article] [PubMed] [Google Scholar]
238. Brady S.W., Gout A.M., Zhang J. Therapeutic and prognostic insights from the analysis of cancer mutational signatures. Trends Genet. 2022; 38:194–208. [DOI] [PMC free article] [PubMed] [Google Scholar]
239. Kaldor J.M., Day N.E., Pettersson F., Clarke E.A., Pedersen D., Mehnert W., Bell J., Host H., Prior P., Karjalainen S. et al. Leukemia following chemotherapy for ovarian cancer. N. Engl. J. Med. 1990; 322:1–6. [DOI] [PubMed] [Google Scholar]
240. Travis L.B., Holowaty E.J., Bergfeldt K., Lynch C.F., Kohler B.A., Wiklund T., Curtis R.E., Hall P., Andersson M., Pukkala E. et al. Risk of leukemia after platinum-based chemotherapy for ovarian cancer. N. Engl. J. Med. 1999; 340:351–357. [DOI] [PubMed] [Google Scholar]
241. Wojtaszek J.L., Chatterjee N., Najeeb J., Ramos A., Lee M., Bian K., Xue J.Y., Fenton B.A., Park H., Li D. et al. A small molecule targeting mutagenic translesion synthesis improves chemotherapy. Cell. 2019; 178:152–159. [DOI] [PMC free article] [PubMed] [Google Scholar]
242. Sail V., Rizzo A.A., Chatterjee N., Dash R.C., Ozen Z., Walker G.C., Korzhnev D.M., Hadden M.K. Identification of small molecule translesion synthesis inhibitors that target the Rev1-CT/RIR protein−protein interaction. ACS Chem. Biol. 2017; 12:1903–1912. [DOI] [PMC free article] [PubMed] [Google Scholar]
243. Zhou J., Gelot C., Pantelidou C., Li A., Yucel H., Davis R.E., Farkkila A., Kochupurakkal B., Syed A., Shapiro G.I. et al. A first-in-class polymerase theta inhibitor selectively targets homologous-recombination-deficient tumors. Nat Cancer. 2021; 2:598–610. [DOI] [PMC free article] [PubMed] [Google Scholar]
244. Vanarotti M., Evison B.J., Actis M.L., Inoue A., McDonald E.T., Shao Y., Heath R.J., Fujii N. Small-molecules that bind to the ubiquitin-binding motif of REV1 inhibit REV1 interaction with K164-monoubiquitinated PCNA and suppress DNA damage tolerance. Bioorg. Med. Chem. 2018; 26:2345–2353. [DOI] [PMC free article] [PubMed] [Google Scholar]
245. Lu S., Dong Z. Additive effects of a small molecular PCNA inhibitor PCNA-I1S and DNA damaging agents on growth inhibition and DNA damage in prostate and lung cancer cells. PLoS One. 2019; 14:e0223894. [DOI] [PMC free article] [PubMed] [Google Scholar]
246. Fleming M.C., Chiou L.F., Tumbale P.P., Droby G.N., Lim J., Norris-Drouin J.L., Williams J.G., Pearce K.H., Williams R.S., Vaziri C. et al. Discovery and structural basis of the selectivity of potent cyclic peptide inhibitors of MAGE-A4. J. Med. Chem. 2022; 65:7231–7245. [DOI] [PMC free article] [PubMed] [Google Scholar]
247. Du M., Gu J., Liu C., Liu N., Yu Z., Zhou C., Heng W., Cao Z., Wei F., Zhu K. et al. Genome-wide CRISPR screen identified Rad18 as a determinant of doxorubicin sensitivity in osteosarcoma. J. Exp. Clin. Cancer Res. 2022; 41:154. [DOI] [PMC free article] [PubMed] [Google Scholar]
248. Huang J., Huen M.S.Y., Kim H., Leung C.C.Y., Glover J.N.M., Yu X., Chen J. RAD18 transmits DNA damage signalling to elicit homologous recombination repair. Nat. Cell Biol. 2009; 11:592–603. [DOI] [PMC free article] [PubMed] [Google Scholar]
249. Palle K., Vaziri C. Rad18 E3 ubiquitin ligase activity mediates Fanconi anemia pathway activation and cell survival following DNA Topoisomerase 1 inhibition. Cell Cycle. 2011; 10:1625–1638. [DOI] [PMC free article] [PubMed] [Google Scholar]
250. Räschle M., Smeenk G., Hansen R.K., Temu T., Oka Y., Hein M.Y., Nagaraj N., Long D.T., Walter J.C., Hofmann K. et al. DNA repair. Proteomics reveals dynamic assembly of repair complexes during bypass of DNA cross-links. Science. 2015; 348:1253671. [DOI] [PMC free article] [PubMed] [Google Scholar]
251. Matsuoka S., Ballif B.A., Smogorzewska A., McDonald E.R., Hurov K.E., Luo J., Bakalarski C.E., Zhao Z., Solimini N., Lerenthal Y. et al. ATM and ATR substrate analysis reveals extensive protein networks responsive to DNA damage. Science. 2007; 316:1160–1166. [DOI] [PubMed] [Google Scholar]
252. Adar S., Hu J., Lieb J.D., Sancar A. Genome-wide kinetics of DNA excision repair in relation to chromatin state and mutagenesis. Proc. Natl. Acad. Sci. U.S.A. 2016; 113:E2124–2133. [DOI] [PMC free article] [PubMed] [Google Scholar]
253. Sanchez A., Lee D., Kim D.I., Miller K.M. Making connections: integrative signaling mechanisms coordinate DNA break repair in chromatin. Front. Genet. 2021; 12:747734. [DOI] [PMC free article] [PubMed] [Google Scholar]
254. Sabbioneda S., Gourdin A.M., Green C.M., Zotter A., Giglia-Mari G., Houtsmuller A., Vermeulen W., Lehmann A.R. Effect of proliferating cell nuclear antigen ubiquitination and chromatin structure on the dynamic properties of the Y-family DNA polymerases. Mol. Biol. Cell. 2008; 19:5193–5202. [DOI] [PMC free article] [PubMed] [Google Scholar]
255. Clairmont C.S., D’Andrea A.D REV7 directs DNA repair pathway choice. Trends Cell Biol. 2021; 31:965–978. [DOI] [PMC free article] [PubMed] [Google Scholar]
256. Scully R., Panday A., Elango R., Willis N.A. DNA double-strand break repair-pathway choice in somatic mammalian cells. Nat. Rev. Mol. Cell Biol. 2019; 20:698–714. [DOI] [PMC free article] [PubMed] [Google Scholar]
257. Yoon J.H., Johnson R.E., Prakash L., Prakash S. Implications of inhibition of Rev1 interaction with Y family DNA polymerases for cisplatin chemotherapy. Genes Dev. 2021; 35:1256–1270. [DOI] [PMC free article] [PubMed] [Google Scholar]
258. Stanzione M., Zhong J., Wong E., LaSalle T.J., Wise J.F., Simoneau A., Myers D.T., Phat S., Sade-Feldman M., Lawrence M.S. et al. Translesion DNA synthesis mediates acquired resistance to olaparib plus temozolomide in small cell lung cancer. Sci. Adv. 2022; 8:eabn1229. [DOI] [PMC free article] [PubMed] [Google Scholar]
259. Vanarotti M., Grace C.R., Miller D.J., Actis M.L., Inoue A., Evison B.J., Vaithiyalingam S., Singh A.P., McDonald E.T., Fujii N. Structures of REV1 UBM2 domain complex with ubiquitin and with a small-molecule that inhibits the REV1 UBM2-ubiquitin interaction. J. Mol. Biol. 2018; 430:2857–2872. [DOI] [PMC free article] [PubMed] [Google Scholar]
260. Actis M.L., Ambaye N.D., Evison B.J., Shao Y., Vanarotti M., Inoue A., McDonald E.T., Kikuchi S., Heath R., Hara K. et al. Identification of the first small-molecule inhibitor of the REV7 DNA repair protein interaction. Bioorg. Med. Chem. 2016; 24:4339–4346. [DOI] [PMC free article] [PubMed] [Google Scholar]
261. Zafar M.K., Maddukuri L., Ketkar A., Penthala N.R., Reed M.R., Eddy S., Crooks P.A., Eoff R.L. A small-molecule inhibitor of human DNA polymerase eta potentiates the effects of cisplatin in tumor cells. Biochemistry. 2018; 57:1262–1273. [DOI] [PMC free article] [PubMed] [Google Scholar]
262. Punchihewa C., Inoue A., Hishiki A., Fujikawa Y., Connelly M., Evison B., Shao Y., Heath R., Kuraoka I., Rodrigues P. et al. Identification of small molecule proliferating cell nuclear antigen (PCNA) inhibitor that disrupts interactions with PIP-box proteins and inhibits DNA replication. J. Biol. Chem. 2012; 287:14289–14300. [DOI] [PMC free article] [PubMed] [Google Scholar]
263. Inoue A., Kikuchi S., Hishiki A., Shao Y., Heath R., Evison B.J., Actis M., Canman C.E., Hashimoto H., Fujii N. A small molecule inhibitor of monoubiquitinated proliferating cell nuclear antigen (PCNA) inhibits repair of interstrand DNA cross-link, enhances DNA double strand break, and sensitizes cancer cells to cisplatin. J. Biol. Chem. 2014; 289:7109–7120. [DOI] [PMC free article] [PubMed] [Google Scholar]
264. Konstantinopoulos P.A., da Costa A., Gulhan D., Lee E.K., Cheng S.C., Hendrickson A.E.W., Kochupurakkal B., Kolin D.L., Kohn E.C., Liu J.F. et al. a Replication stress biomarker is associated with response to gemcitabine versus combined gemcitabine and ATR inhibitor therapy in ovarian cancer. Nat. Commun. 2021; 12:5574. [DOI] [PMC free article] [PubMed] [Google Scholar]
265. McPherson K.S., Zaino A.M., Dash R.C., Rizzo A.A., Li Y., Hao B., Bezsonova I., Hadden M.K., Korzhnev D.M. Structure-Based Drug Design of Phenazopyridine Derivatives as Inhibitors of Rev1 Interactions in Translesion Synthesis. ChemMedChem. 2021; 16:1126–1132. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

zcad005_Supplemental_File

zcad005_supplemental_file.pdf^{(365.7KB, pdf)}

[B1] 1. Hanahan D., Weinberg R.A. The hallmarks of cancer. Cell. 2000; 100:57–70. [DOI] [PubMed] [Google Scholar]

[B2] 2. Hanahan D., Weinberg R.A. Hallmarks of cancer: the next generation. Cell. 2011; 144:646–674. [DOI] [PubMed] [Google Scholar]

[B3] 3. Baynton K., Fuchs R.P. Lesions in DNA: hurdles for polymerases. Trends Biochem. Sci. 2000; 25:74–79. [DOI] [PubMed] [Google Scholar]

[B4] 4. Ogi T., Lehmann A.R. the Y-family DNA polymerase kappa (pol kappa) functions in mammalian nucleotide-excision repair. Nat. Cell Biol. 2006; 8:640–642. [DOI] [PubMed] [Google Scholar]

[B5] 5. Ogi T., Limsirichaikul S., Overmeer R.M., Volker M., Takenaka K., Cloney R., Nakazawa Y., Niimi A., Miki Y., Jaspers N.G. et al. Three DNA polymerases, recruited by different mechanisms, carry out NER repair synthesis in human cells. Mol. Cell. 2010; 37:714–727. [DOI] [PubMed] [Google Scholar]

[B6] 6. Yang Y., Durando M., Smith-Roe S.L., Sproul C., Greenwalt A.M., Kaufmann W., Oh S., Hendrickson E.A., Vaziri C. Cell cycle stage-specific roles of Rad18 in tolerance and repair of oxidative DNA damage. Nucleic Acids Res. 2013; 41:2296–2312. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B7] 7. Zlatanou A., Despras E., Braz-Petta T., Boubakour-Azzouz I., Pouvelle C., Stewart G.S., Nakajima S., Yasui A., Ishchenko A.A., Kannouche P.L. The hMsh2-hMsh6 complex acts in concert with monoubiquitinated PCNA and Pol eta in response to oxidative DNA damage in human cells. Mol. Cell. 2011; 43:649–662. [DOI] [PubMed] [Google Scholar]

[B8] 8. Tirman S., Quinet A., Wood M., Meroni A., Cybulla E., Jackson J., Pegoraro S., Simoneau A., Zou L., Vindigni A. Temporally distinct post-replicative repair mechanisms fill PRIMPOL-dependent ssDNA gaps in human cells. Mol. Cell. 2021; 81:4026–4040. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B9] 9. Karras G.I., Jentsch S. the RAD6 DNA damage tolerance pathway operates uncoupled from the replication fork and is functional beyond S phase. Cell. 2010; 141:255–267. [DOI] [PubMed] [Google Scholar]

[B10] 10. Daigaku Y., Davies A.A., Ulrich H.D. Ubiquitin-dependent DNA damage bypass is separable from genome replication. Nature. 2010; 465:951–955. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B11] 11. Masutani C., Araki M., Yamada A., Kusumoto R., Nogimori T., Maekawa T., Iwai S., Hanaoka F. Xeroderma pigmentosum variant (XP-V) correcting protein from HeLa cells has a thymine dimer bypass DNA polymerase activity. EMBO J. 1999; 18:3491–3501. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B12] 12. Masutani C., Kusumoto R., Yamada A., Dohmae N., Yokoi M., Yuasa M., Araki M., Iwai S., Takio K., Hanaoka F. The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase eta. Nature. 1999; 399:700–704. [DOI] [PubMed] [Google Scholar]

[B13] 13. Friedberg E.C. How nucleotide excision repair protects against cancer. Nat. Rev. Cancer. 2001; 1:22–33. [DOI] [PubMed] [Google Scholar]

[B14] 14. Pfeifer G.P. Formation and processing of UV photoproducts: effects of DNA sequence and chromatin environment. Photochem. Photobiol. 1997; 65:270–283. [DOI] [PubMed] [Google Scholar]

[B15] 15. Dupuy A., Sarasin A. DNA damage and gene therapy of xeroderma pigmentosum, a human DNA repair-deficient disease. Mutat. Res. 2015; 776:2–8. [DOI] [PubMed] [Google Scholar]

[B16] 16. Di Lucca J., Guedj M., Lacapere J.J., Fargnoli M.C., Bourillon A., Dieude P., Dupin N., Wolkenstein P., Aegerter P., Saiag P. et al. Variants of the xeroderma pigmentosum variant gene (POLH) are associated with melanoma risk. Eur. J. Cancer. 2009; 45:3228–3236. [DOI] [PubMed] [Google Scholar]

[B17] 17. Castro L.P., Batista-Vieira D., de Souza T.A., Timoteo A.R.S., Coutinho J.D.L., Pinheiro de Almeida I.C., Henriques S.R.M., de Azevedo F.M., Rosa R.C.A., Kannouche P.L. et al. XPC and POLH/XPV genes mutated in a genetic cluster of xeroderma pigmentosum patients in northeast Brazil. Front. Genet. 2021; 12:784963. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B18] 18. Ziv O., Geacintov N., Nakajima S., Yasui A., Livneh Z. DNA polymerase zeta cooperates with polymerases kappa and iota in translesion DNA synthesis across pyrimidine photodimers in cells from XPV patients. Proc. Natl. Acad. Sci. U.S.A. 2009; 106:11552–11557. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B19] 19. Shachar S., Ziv O., Avkin S., Adar S., Wittschieben J., Reissner T., Chaney S., Friedberg E.C., Wang Z., Carell T. et al. Two-polymerase mechanisms dictate error-free and error-prone translesion DNA synthesis in mammals. EMBO J. 2009; 28:383–393. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B20] 20. Limoli C.L., Giedzinski E., Morgan W.F., Cleaver J.E. 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. U.S.A. 2000; 97:7939–7946. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B21] 21. Vaziri C., Rogozin I.B., Gu Q., Wu D., Day T.A. Unravelling roles of error-prone DNA polymerases in shaping cancer genomes. Oncogene. 2021; 40:6549–6565. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B22] 22. Ceccaldi R., Liu J.C., Amunugama R., Hajdu I., Primack B., Petalcorin M.I., O’Connor K.W., Konstantinopoulos P.A., Elledge S.J., Boulton S.J. et al. Homologous-recombination-deficient tumours are dependent on Poltheta-mediated repair. Nature. 2015; 518:258–262. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B23] 23. Schimmel J., van Schendel R., den Dunnen J.T., Tijsterman M. Templated insertions: a smoking gun for polymerase theta-mediated end joining. Trends Genet. 2019; 35:632–644. [DOI] [PubMed] [Google Scholar]

[B24] 24. Hwang T., Reh S., Dunbayev Y., Zhong Y., Takata Y., Shen J., McBride K.M., Murnane J.P., Bhak J., Lee S. et al. Defining the mutation signatures of DNA polymerase theta in cancer genomes. NAR Cancer. 2020; 2:zcaa017. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B25] 25. Prakash S., Johnson R.E., Prakash L. Eukaryotic translesion synthesis DNA polymerases: specificity of structure and function. Annu. Rev. Biochem. 2005; 74:317–353. [DOI] [PubMed] [Google Scholar]

[B26] 26. Waters L.S., Minesinger B.K., Wiltrout M.E., D'Souza S., Woodruff R.V., Walker G.C Eukaryotic translesion polymerases and their roles and regulation in DNA damage tolerance. Microbiol. Mol. Biol. Rev. 2009; 73:134–154. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B27] 27. Ulrich H.D., Jentsch S. Two RING finger proteins mediate cooperation between ubiquitin-conjugating enzymes in DNA repair. EMBO J. 2000; 19:3388–3397. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B28] 28. Kannouche P.L., Wing J., Lehmann A.R. Interaction of human DNA polymerase eta with monoubiquitinated PCNA: a possible mechanism for the polymerase switch in response to DNA damage. Mol. Cell. 2004; 14:491–500. [DOI] [PubMed] [Google Scholar]

[B29] 29. Davies A.A., Huttner D., Daigaku Y., Chen S., Ulrich H.D. Activation of ubiquitin-dependent DNA damage bypass is mediated by replication protein a. Mol. Cell. 2008; 29:625–636. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B30] 30. Zou L., Elledge S.J. Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science. 2003; 300:1542–1548. [DOI] [PubMed] [Google Scholar]

[B31] 31. Pena-Diaz J., Bregenhorn S., Ghodgaonkar M., Follonier C., Artola-Boran M., Castor D., Lopes M., Sartori A.A., Jiricny J. Noncanonical mismatch repair as a source of genomic instability in human cells. Mol. Cell. 2012; 47:669–680. [DOI] [PubMed] [Google Scholar]

[B32] 32. Ray A., Blevins C., Wani G., Wani A.A. ATR- and ATM-mediated DNA damage response is dependent on excision repair assembly during G1 but Not in S phase of cell cycle. PLoS One. 2016; 11:e0159344. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B33] 33. Bienko M., Green C.M., Crosetto N., Rudolf F., Zapart G., Coull B., Kannouche P., Wider G., Peter M., Lehmann A.R. et al. Ubiquitin-binding domains in Y-family polymerases regulate translesion synthesis. Science. 2005; 310:1821–1824. [DOI] [PubMed] [Google Scholar]

[B34] 34. Masuda Y., Kanao R., Kaji K., Ohmori H., Hanaoka F., Masutani C. Different types of interaction between PCNA and PIP boxes contribute to distinct cellular functions of Y-family DNA polymerases. Nucleic Acids Res. 2015; 43:7898–7910. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B35] 35. Guo C., Sonoda E., Tang T.S., Parker J.L., Bielen A.B., Takeda S., Ulrich H.D., Friedberg E.C. REV1 protein interacts with PCNA: significance of the REV1 BRCT domain in vitro and in vivo. Mol. Cell. 2006; 23:265–271. [DOI] [PubMed] [Google Scholar]

[B36] 36. Pustovalova Y., Maciejewski M.W., Korzhnev D.M. NMR mapping of PCNA interaction with translesion synthesis DNA polymerase Rev1 mediated by Rev1-BRCT domain. J. Mol. Biol. 2013; 425:3091–3105. [DOI] [PubMed] [Google Scholar]

[B37] 37. Sharma N.M., Kochenova O.V., Shcherbakova P.V. the non-canonical protein binding site at the monomer-monomer interface of yeast proliferating cell nuclear antigen (PCNA) regulates the Rev1-PCNA interaction and Polzeta/Rev1-dependent translesion DNA synthesis. J. Biol. Chem. 2011; 286:33557–33566. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B38] 38. Durando M., Tateishi S., Vaziri C. a non-catalytic role of DNA polymerase η in recruiting Rad18 and promoting PCNA monoubiquitination at stalled replication forks. Nucleic Acids Res. 2013; 41:3079–3093. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B39] 39. Watanabe K., Tateishi S., Kawasuji M., Tsurimoto T., Inoue H., Yamaizumi M. Rad18 guides poleta to replication stalling sites through physical interaction and PCNA monoubiquitination. EMBO J. 2004; 23:3886–3896. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B40] 40. Masutani C., Kusumoto R., Iwai S., Hanaoka F. Mechanisms of accurate translesion synthesis by human DNA polymerase eta. EMBO J. 2000; 19:3100–3109. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B41] 41. Choi J.H., Pfeifer G.P. the role of DNA polymerase eta in UV mutational spectra. DNA Repair (Amst.). 2005; 4:211–220. [DOI] [PubMed] [Google Scholar]

[B42] 42. Choi J.Y., Angel K.C., Guengerich F.P. Translesion synthesis across bulky N2-alkyl guanine DNA adducts by human DNA polymerase kappa. J. Biol. Chem. 2006; 281:21062–21072. [DOI] [PubMed] [Google Scholar]

[B43] 43. Ogi T., Shinkai Y., Tanaka K., Ohmori H. Polkappa protects mammalian cells against the lethal and mutagenic effects of benzo[a]pyrene. Proc. Natl. Acad. Sci. U.S.A. 2002; 99:15548–15553. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B44] 44. Lin W., Xin H., Zhang Y., Wu X., Yuan F., Wang Z. the human REV1 gene codes for a DNA template-dependent dCMP transferase. Nucleic Acids Res. 1999; 27:4468–4475. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B45] 45. Hishiki A., Hashimoto H., Hanafusa T., Kamei K., Ohashi E., Shimizu T., Ohmori H., Sato M. Structural basis for novel interactions between human translesion synthesis polymerases and proliferating cell nuclear antigen. J. Biol. Chem. 2009; 284:10552–10560. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B46] 46. McCulloch S.D., Kokoska R.J., Masutani C., Iwai S., Hanaoka F., Kunkel T.A. Preferential cis-syn thymine dimer bypass by DNA polymerase eta occurs with biased fidelity. Nature. 2004; 428:97–100. [DOI] [PubMed] [Google Scholar]

[B47] 47. Kusumoto R., Masutani C., Shimmyo S., Iwai S., Hanaoka F. DNA binding properties of human DNA polymerase eta: implications for fidelity and polymerase switching of translesion synthesis. Genes Cells. 2004; 9:1139–1150. [DOI] [PubMed] [Google Scholar]

[B48] 48. Ohmori H., Friedberg E.C., Fuchs R.P., Goodman M.F., Hanaoka F., Hinkle D., Kunkel T.A., Lawrence C.W., Livneh Z., Nohmi T. et al. the Y-family of DNA polymerases. Mol. Cell. 2001; 8:7–8. [DOI] [PubMed] [Google Scholar]

[B49] 49. Prakash S., Johnson R.E., Prakash L. Eukaryotic translesion synthesis DNA polymerases: specificity of structure and function. Annu. Rev. Biochem. 2005; 74:317–353. [DOI] [PubMed] [Google Scholar]

[B50] 50. Johnson R.E., Yu S.-L., Prakash S., Prakash L. Yeast DNA polymerase zeta (zeta) is essential for error-free replication past thymine glycol. Genes Dev. 2003; 17:77–87. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B51] 51. Ross A.-L., Simpson L.J., Sale J.E. Vertebrate DNA damage tolerance requires the C-terminus but not BRCT or transferase domains of REV1. Nucleic Acids Res. 2005; 33:1280–1289. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B52] 52. Edmunds C.E., Simpson L.J., Sale J.E. PCNA ubiquitination and REV1 define temporally distinct mechanisms for controlling translesion synthesis in the avian cell line DT40. Mol. Cell. 2008; 30:519–529. [DOI] [PubMed] [Google Scholar]

[B53] 53. Ohashi E., Murakumo Y., Kanjo N., Akagi J.-I., Masutani C., Hanaoka F., Ohmori H. Interaction of hREV1 with three human Y-family DNA polymerases. Genes Cells. 2004; 9:523–531. [DOI] [PubMed] [Google Scholar]

[B54] 54. Ohashi E., Hanafusa T., Kamei K., Song I., Tomida J., Hashimoto H., Vaziri C., Ohmori H. Identification of a novel REV1-interacting motif necessary for DNA polymerase kappa function. Genes Cells. 2009; 14:101–111. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B55] 55. Pustovalova Y., Bezsonova I., Korzhnev D.M. the C-terminal domain of human Rev1 contains independent binding sites for DNA polymerase η and Rev7 subunit of polymerase ζ. FEBS Lett. 2012; 586:3051–3056. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B56] 56. Pustovalova Y., Magalhaes M.T., D'Souza S., Rizzo A.A., Korza G., Walker G.C., Korzhnev D.M Interaction between the Rev1 C-terminal domain and the PolD3 subunit of Polzeta suggests a mechanism of polymerase exchange upon Rev1/Polzeta-dependent translesion synthesis. Biochemistry. 2016; 55:2043–2053. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B57] 57. D'Souza S., Walker G.C. Novel role for the C terminus of Saccharomyces cerevisiae Rev1 in mediating protein-protein interactions. Mol. Cell. Biol. 2006; 26:8173–8182. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B58] 58. Guo C., Sonoda E., Tang T.-S., Parker J.L., Bielen A.B., Takeda S., Ulrich H.D., Friedberg E.C. REV1 protein interacts with PCNA: significance of the REV1 BRCT domain in vitro and in vivo. Mol. Cell. 2006; 23:265–271. [DOI] [PubMed] [Google Scholar]

[B59] 59. Terai K., Abbas T., Jazaeri A.A., Dutta A. CRL4(Cdt2) E3 ubiquitin ligase monoubiquitinates PCNA to promote translesion DNA synthesis. Mol. Cell. 2010; 37:143–149. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B60] 60. Lin J.R., Zeman M.K., Chen J.Y., Yee M.C., Cimprich K.A. SHPRH and HLTF act in a damage-specific manner to coordinate different forms of postreplication repair and prevent mutagenesis. Mol. Cell. 2011; 42:237–249. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B61] 61. Haracska L., Unk I., Prakash L., Prakash S. Ubiquitylation of yeast proliferating cell nuclear antigen and its implications for translesion DNA synthesis. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:6477–6482. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B62] 62. Acharya N., Yoon J.H., Hurwitz J., Prakash L., Prakash S. DNA polymerase eta lacking the ubiquitin-binding domain promotes replicative lesion bypass in humans cells. Proc. Natl. Acad. Sci. U.S.A. 2010; 107:10401–10405. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B63] 63. Hendel A., Krijger P.H.L., Diamant N., Goren Z., Langerak P., Kim J., Reissner T., Lee K.-y., Geacintov N.E., Carell T. et al. PCNA ubiquitination is important, but not essential for translesion DNA synthesis in mammalian cells. PLos Genet. 2011; 7:e1002262. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B64] 64. Wit N., Buoninfante O.A., van den Berk P.C.M., Jansen J.G., Hogenbirk M.A., de Wind N., Jacobs H. Roles of PCNA ubiquitination and TLS polymerases κ and η in the bypass of methyl methanesulfonate-induced DNA damage. Nucleic Acids Res. 2015; 43:282–294. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B65] 65. Krijger P.H.L., van den Berk P.C.M., Wit N., Langerak P., Jansen J.G., Reynaud C.-A., de Wind N., Jacobs H. PCNA ubiquitination-independent activation of polymerase η during somatic hypermutation and DNA damage tolerance. DNA Repair (Amst.). 2011; 10:1051–1059. [DOI] [PubMed] [Google Scholar]

[B66] 66. Huertas P., Cortes-Ledesma F., Sartori A.A., Aguilera A., Jackson S.P. CDK targets Sae2 to control DNA-end resection and homologous recombination. Nature. 2008; 455:689–692. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B67] 67. Ira G., Pellicioli A., Balijja A., Wang X., Fiorani S., Carotenuto W., Liberi G., Bressan D., Wan L., Hollingsworth N.M. et al. DNA end resection, homologous recombination and DNA damage checkpoint activation require CDK1. Nature. 2004; 431:1011–1017. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B68] 68. Aylon Y., Liefshitz B., Kupiec M. the CDK regulates repair of double-strand breaks by homologous recombination during the cell cycle. EMBO J. 2004; 23:4868–4875. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B69] 69. Yun M.H., Hiom K. CtIP-BRCA1 modulates the choice of DNA double-strand-break repair pathway throughout the cell cycle. Nature. 2009; 459:460–463. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B70] 70. Tomimatsu N., Mukherjee B., Catherine Hardebeck M., Ilcheva M., Vanessa Camacho C., Louise Harris J., Porteus M., Llorente B., Khanna K.K., Burma S. Phosphorylation of EXO1 by CDKs 1 and 2 regulates DNA end resection and repair pathway choice. Nat. Commun. 2014; 5:3561. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B71] 71. Falck J., Forment J.V., Coates J., Mistrik M., Lukas J., Bartek J., Jackson S.P. CDK targeting of NBS1 promotes DNA-end resection, replication restart and homologous recombination. EMBO Rep. 2012; 13:561–568. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B72] 72. Ferretti L.P., Lafranchi L., Sartori A.A. Controlling DNA-end resection: a new task for CDKs. Front. Genet. 2013; 4:99. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B73] 73. Luo K., Deng M., Li Y., Wu C., Xu Z., Yuan J., Lou Z. CDK-mediated RNF4 phosphorylation regulates homologous recombination in S-phase. Nucleic Acids Res. 2015; 43:5465–5475. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B74] 74. Swift M.L., Beishline K., Flashner S., Azizkhan-Clifford J. DSB repair pathway choice is regulated by recruitment of 53BP1 through cell cycle-dependent regulation of Sp1. Cell Rep. 2021; 34:108840. [DOI] [PubMed] [Google Scholar]

[B75] 75. Uchiyama M., Terunuma J., Hanaoka F. The protein level of Rev1, a TLS polymerase in fission yeast, is strictly regulated during the cell cycle and after DNA damage. PLoS One. 2015; 10:e0130000. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B76] 76. Waters L.S., Walker G.C. the critical mutagenic translesion DNA polymerase Rev1 is highly expressed during G(2)/M phase rather than S phase. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:8971–8976. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B77] 77. Varanasi L., Do P.M., Goluszko E., Martinez L.A. Rad18 is a transcriptional target of E2F3. Cell Cycle. 2012; 11:1131–1141. [DOI] [PubMed] [Google Scholar]

[B78] 78. Gao Y., Mutter-Rottmayer E., Greenwalt A.M., Goldfarb D., Yan F., Yang Y., Martinez-Chacin R.C., Pearce K.H., Tateishi S., Major M.B. et al. A neomorphic cancer cell-specific role of MAGE-A4 in trans-lesion synthesis. Nat. Commun. 2016; 7:12105. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B79] 79. Bi X., Barkley L.R., Slater D.M., Tateishi S., Yamaizumi M., Ohmori H., Vaziri C. Rad18 regulates DNA polymerase kappa and is required for recovery from S-phase checkpoint-mediated arrest. Mol. Cell. Biol. 2006; 26:3527–3540. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B80] 80. Song I.Y., Palle K., Gurkar A., Tateishi S., Kupfer G.M., Vaziri C. Rad18-mediated translesion synthesis of bulky DNA adducts is coupled to activation of the Fanconi anemia DNA repair pathway. J. Biol. Chem. 2010; 285:31525–31536. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B81] 81. Liu G., Chen X. DNA polymerase eta, the product of the xeroderma pigmentosum variant gene and a target of p53, modulates the DNA damage checkpoint and p53 activation. Mol. Cell. Biol. 2006; 26:1398–1413. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B82] 82. Tomicic M.T., Aasland D., Naumann S.C., Meise R., Barckhausen C., Kaina B., Christmann M. Translesion polymerase eta is upregulated by cancer therapeutics and confers anticancer drug resistance. Cancer Res. 2014; 74:5585–5596. [DOI] [PubMed] [Google Scholar]

[B83] 83. Lerner L.K., Francisco G., Soltys D.T., Rocha C.R., Quinet A., Vessoni A.T., Castro L.P., David T.I., Bustos S.O., Strauss B.E. et al. Predominant role of DNA polymerase eta and p53-dependent translesion synthesis in the survival of ultraviolet-irradiated human cells. Nucleic Acids Res. 2017; 45:1270–1280. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B84] 84. Day T.A., Palle K., Barkley L.R., Kakusho N., Zou Y., Tateishi S., Verreault A., Masai H., Vaziri C. Phosphorylated Rad18 directs DNA polymerase η to sites of stalled replication. J. Cell Biol. 2010; 191:953–966. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B85] 85. Yamada M., Masai H., Bartek J. Regulation and roles of Cdc7 kinase under replication stress. Cell Cycle. 2014; 13:1859–1866. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B86] 86. Liu R., Huang Y. CDC7 as a novel biomarker and druggable target in cancer. Clin. Transl. Oncol. 2022; 24:1856–1864. [DOI] [PubMed] [Google Scholar]

[B87] 87. Barkley L.R., Palle K., Durando M., Day T.A., Gurkar A., Kakusho N., Li J., Masai H., Vaziri C. c-Jun N-terminal kinase-mediated Rad18 phosphorylation facilitates Poleta recruitment to stalled replication forks. Mol. Biol. Cell. 2012; 23:1943–1954. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B88] 88. Hotamisligil G.S., Davis R.J. Cell signaling and stress responses. Cold Spring Harb. Perspect. Biol. 2016; 8:a006072. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B89] 89. Sarcevic B., Mawson A., Baker R.T., Sutherland R.L. Regulation of the ubiquitin-conjugating enzyme hHR6A by CDK-mediated phosphorylation. EMBO J. 2002; 21:2009–2018. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B90] 90. Chen Y.W., Cleaver J.E., Hatahet Z., Honkanen R.E., Chang J.Y., Yen Y., Chou K.M. Human DNA polymerase eta activity and translocation is regulated by phosphorylation. Proc. Natl. Acad. Sci. U.S.A. 2008; 105:16578–16583. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B91] 91. Gohler T., Sabbioneda S., Green C.M., Lehmann A.R. ATR-mediated phosphorylation of DNA polymerase eta is needed for efficient recovery from UV damage. J. Cell Biol. 2011; 192:219–227. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B92] 92. Bertoletti F., Cea V., Liang C.C., Lanati T., Maffia A., Avarello M.D.M., Cipolla L., Lehmann A.R., Cohn M.A., Sabbioneda S. Phosphorylation regulates human poleta stability and damage bypass throughout the cell cycle. Nucleic Acids Res. 2017; 45:9441–9454. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B93] 93. Lehmann A.R., Kirk-Bell S., Arlett C.F., Paterson M.C., Lohman P.H., de Weerd-Kastelein E.A., Bootsma D Xeroderma pigmentosum cells with normal levels of excision repair have a defect in DNA synthesis after UV-irradiation. Proc. Natl. Acad. Sci. U.S.A. 1975; 72:219–223. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B94] 94. Cleaver J.E., Thomas G.H., Park S.D. Xeroderma pigmentosum variants have a slow recovery of DNA synthesis after irradiation with ultraviolet light. Biochim. Biophys. Acta. 1979; 564:122–131. [DOI] [PubMed] [Google Scholar]

[B95] 95. Quinet A., Martins D.J., Vessoni A.T., Biard D., Sarasin A., Stary A., Menck C.F. Translesion synthesis mechanisms depend on the nature of DNA damage in UV-irradiated human cells. Nucleic Acids Res. 2016; 44:5717–5731. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B96] 96. Vallerga M.B., Mansilla S.F., Federico M.B., Bertolin A.P., Gottifredi V. Rad51 recombinase prevents Mre11 nuclease-dependent degradation and excessive PrimPol-mediated elongation of nascent DNA after UV irradiation. Proc. Natl. Acad. Sci. U.S.A. 2015; 112:E6624–6633. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B97] 97. Sokol A.M., Cruet-Hennequart S., Pasero P., Carty M.P. DNA polymerase eta modulates replication fork progression and DNA damage responses in platinum-treated human cells. Sci. Rep. 2013; 3:3277. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B98] 98. Despras E., Sittewelle M., Pouvelle C., Delrieu N., Cordonnier A.M., Kannouche P.L. Rad18-dependent SUMOylation of human specialized DNA polymerase eta is required to prevent under-replicated DNA. Nat. Commun. 2016; 7:13326. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B99] 99. Benureau Y., Pouvelle C., Dupaigne P., Baconnais S., Moreira Tavares E., Mazon G., Despras E., Le Cam E., Kannouche P.L. Changes in the architecture and abundance of replication intermediates delineate the chronology of DNA damage tolerance pathways at UV-stalled replication forks in human cells. Nucleic Acids Res. 2022; 50:9909–9929. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B100] 100. Lehmann A.R. Post-replication repair of DNA in ultraviolet-irradiated mammalian cells. No gaps in DNA synthesized late after ultraviolet irradiation. Eur. J. Biochem. 1972; 31:438–445. [DOI] [PubMed] [Google Scholar]

[B101] 101. Meneghini R. Gaps in DNA synthesized by ultraviolet light-irradiated WI38 human cells. Biochim. Biophys. Acta. 1976; 425:419–427. [DOI] [PubMed] [Google Scholar]

[B102] 102. Quinet A., Lerner L.K., Martins D.J., Menck C.F.M. Filling gaps in translesion DNA synthesis in human cells. Mutat. Res. Genet. Toxicol. Environ. Mutagen. 2018; 836:127–142. [DOI] [PubMed] [Google Scholar]

[B103] 103. Quinet A., Tirman S., Cybulla E., Meroni A., Vindigni A. to skip or not to skip: choosing repriming to tolerate DNA damage. Mol. Cell. 2021; 81:649–658. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B104] 104. Lopes M., Foiani M., Sogo J.M. Multiple mechanisms control chromosome integrity after replication fork uncoupling and restart at irreparable UV lesions. Mol. Cell. 2006; 21:15–27. [DOI] [PubMed] [Google Scholar]

[B105] 105. Diamant N., Hendel A., Vered I., Carell T., Reissner T., de Wind N., Geacinov N., Livneh Z. DNA damage bypass operates in the S and G2 phases of the cell cycle and exhibits differential mutagenicity. Nucleic Acids Res. 2012; 40:170–180. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B106] 106. Elvers I., Johansson F., Groth P., Erixon K., Helleday T. UV stalled replication forks restart by re-priming in human fibroblasts. Nucleic Acids Res. 2011; 39:7049–7057. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B107] 107. Zellweger R., Dalcher D., Mutreja K., Berti M., Schmid J.A., Herrador R., Vindigni A., Lopes M. Rad51-mediated replication fork reversal is a global response to genotoxic treatments in human cells. J. Cell Biol. 2015; 208:563–579. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B108] 108. Bianchi J., Rudd S.G., Jozwiakowski S.K., Bailey L.J., Soura V., Taylor E., Stevanovic I., Green A.J., Stracker T.H., Lindsay H.D. et al. PrimPol bypasses UV photoproducts during eukaryotic chromosomal DNA replication. Mol. Cell. 2013; 52:566–573. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B109] 109. Mourón S., Rodriguez-Acebes S., Martínez-Jiménez M.I., García-Gómez S., Chocrón S., Blanco L., Méndez J. Repriming of DNA synthesis at stalled replication forks by human PrimPol. Nat. Struct. Mol. Biol. 2013; 20:1383–1389. [DOI] [PubMed] [Google Scholar]

[B110] 110. Guilliam T.A., Jozwiakowski S.K., Ehlinger A., Barnes R.P., Rudd S.G., Bailey L.J., Skehel J.M., Eckert K.A., Chazin W.J., Doherty A.J. Human PrimPol is a highly error-prone polymerase regulated by single-stranded DNA binding proteins. Nucleic Acids Res. 2015; 43:1056–1068. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B111] 111. Guilliam T.A., Brissett N.C., Ehlinger A., Keen B.A., Kolesar P., Taylor E.M., Bailey L.J., Lindsay H.D., Chazin W.J., Doherty A.J. Molecular basis for PrimPol recruitment to replication forks by RPA. Nat. Commun. 2017; 8:15222. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B112] 112. Bainbridge L.J., Teague R., Doherty A.J. Repriming DNA synthesis: an intrinsic restart pathway that maintains efficient genome replication. Nucleic Acids Res. 2021; 49:4831–4847. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B113] 113. Broomfield S., Chow B.L., Xiao W. MMS2, encoding a ubiquitin-conjugating-enzyme-like protein, is a member of the yeast error-free postreplication repair pathway. Proc. Natl. Acad. Sci. U.S.A. 1998; 95:5678–5683. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B114] 114. Hofmann R.M., Pickart C.M. Noncanonical MMS2-encoded ubiquitin-conjugating enzyme functions in assembly of novel polyubiquitin chains for DNA repair. Cell. 1999; 96:645–653. [DOI] [PubMed] [Google Scholar]

[B115] 115. Johnson R.E., Henderson S.T., Petes T.D., Prakash S., Bankmann M., Prakash L. Saccharomyces cerevisiae RAD5-encoded DNA repair protein contains DNA helicase and zinc-binding sequence motifs and affects the stability of simple repetitive sequences in the genome. Mol. Cell. Biol. 1992; 12:3807–3818. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B116] 116. Elserafy M., Abugable A.A., Atteya R., El-Khamisy S.F. Rad5, HLTF, and SHPRH: a Fresh View of an Old Story. Trends Genet. 2018; 34:574–577. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B117] 117. Unk I., Hajdu I., Fatyol K., Szakal B., Blastyak A., Bermudez V., Hurwitz J., Prakash L., Prakash S., Haracska L. Human SHPRH is a ubiquitin ligase for Mms2-Ubc13-dependent polyubiquitylation of proliferating cell nuclear antigen. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:18107–18112. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B118] 118. Masuda Y., Suzuki M., Kawai H., Hishiki A., Hashimoto H., Masutani C., Hishida T., Suzuki F., Kamiya K. En bloc transfer of polyubiquitin chains to PCNA in vitro is mediated by two different human E2-E3 pairs. Nucleic Acids Res. 2012; 40:10394–10407. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B119] 119. Motegi A., Sood R., Moinova H., Markowitz S.D., Liu P.P., Myung K. Human SHPRH suppresses genomic instability through proliferating cell nuclear antigen polyubiquitination. J. Cell Biol. 2006; 175:703–708. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B120] 120. Unk I., Hajdú I., Fátyol K., Hurwitz J., Yoon J.-H., Prakash L., Prakash S., Haracska L. Human HLTF functions as a ubiquitin ligase for proliferating cell nuclear antigen polyubiquitination. Proc. Natl. Acad. Sci. U.S.A. 2008; 105:3768–3773. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B121] 121. Ciccia A., Nimonkar A.V., Hu Y., Hajdu I., Achar Y.J., Izhar L., Petit S.A., Adamson B., Yoon J.C., Kowalczykowski S.C. et al. Polyubiquitinated PCNA recruits the ZRANB3 translocase to maintain genomic integrity after replication stress. Mol. Cell. 2012; 47:396–409. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B122] 122. Hishiki A., Hara K., Ikegaya Y., Yokoyama H., Shimizu T., Sato M., Hashimoto H. Structure of a Novel DNA-binding Domain of Helicase-like Transcription Factor (HLTF) and Its Functional Implication in DNA Damage Tolerance. J. Biol. Chem. 2015; 290:13215–13223. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B123] 123. Yang Y., Gao Y., Mutter-Rottmayer L., Zlatanou A., Durando M., Ding W., Wyatt D., Ramsden D., Tanoue Y., Tateishi S. et al. DNA repair factor RAD18 and DNA polymerase Polkappa confer tolerance of oncogenic DNA replication stress. J. Cell Biol. 2017; 216:3097–3115. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B124] 124. Nayak S., Calvo J.A., Cong K., Peng M., Berthiaume E., Jackson J., Zaino A.M., Vindigni A., Hadden M.K., Cantor S.B. Inhibition of the translesion synthesis polymerase REV1 exploits replication gaps as a cancer vulnerability. Sci. Adv. 2020; 6:eaaz7808. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B125] 125. Cong K., Peng M., Kousholt A.N., Lee W.T.C., Lee S., Nayak S., Krais J., VanderVere-Carozza P.S., Pawelczak K.S., Calvo J. et al. Replication gaps are a key determinant of PARP inhibitor synthetic lethality with BRCA deficiency. Mol. Cell. 2021; 81:3128–3144. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B126] 126. Cong K., Cantor S.B. Exploiting replication gaps for cancer therapy. Mol. Cell. 2022; 82:2363–2369. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B127] 127. Taglialatela A., Leuzzi G., Sannino V., Cuella-Martin R., Huang J.-W., Wu-Baer F., Baer R., Costanzo V., Ciccia A. REV1-Polζ maintains the viability of homologous recombination-deficient cancer cells through mutagenic repair of PRIMPOL-dependent ssDNA gaps. Mol. Cell. 2021; 81:4008–4025. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B128] 128. Alexandrov L.B., Kim J., Haradhvala N.J., Huang M.N., Tian Ng A.W., Wu Y., Boot A., Covington K.R., Gordenin D.A., Bergstrom E.N. et al. The repertoire of mutational signatures in human cancer. Nature. 2020; 578:94–101. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B129] 129. Roberts S.A., Sterling J., Thompson C., Harris S., Mav D., Shah R., Klimczak L.J., Kryukov G.V., Malc E., Mieczkowski P.A. et al. Clustered mutations in yeast and in human cancers can arise from damaged long single-strand DNA regions. Mol. Cell. 2012; 46:424–435. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B130] 130. Morganella S., Alexandrov L.B., Glodzik D., Zou X., Davies H., Staaf J., Sieuwerts A.M., Brinkman A.B., Martin S., Ramakrishna M. et al. The topography of mutational processes in breast cancer genomes. Nat. Commun. 2016; 7:11383. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B131] 131. Petljak M., Dananberg A., Chu K., Bergstrom E.N., Striepen J., von Morgen P., Chen Y., Shah H., Sale J.E., Alexandrov L.B. et al. Mechanisms of APOBEC3 mutagenesis in human cancer cells. Nature. 2022; 607:799–807. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B132] 132. Delbos F., De Smet A., Faili A., Aoufouchi S., Weill J.C., Reynaud C.A. Contribution of DNA polymerase eta to immunoglobulin gene hypermutation in the mouse. J. Exp. Med. 2005; 201:1191–1196. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B133] 133. Supek F., Lehner B. Clustered mutation signatures reveal that error-prone DNA repair targets mutations to active genes. Cell. 2017; 170:534–547. [DOI] [PubMed] [Google Scholar]

[B134] 134. Xu H.L., Gao X.R., Zhang W., Cheng J.R., Tan Y.T., Zheng W., Shu X.O., Xiang Y.B. Effects of polymorphisms in translesion DNA synthesis genes on lung cancer risk and prognosis in Chinese men. Cancer Epidemiol. 2013; 37:917–922. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B135] 135. Dai Z.J., Liu X.H., Ma Y.F., Kang H.F., Jin T.B., Dai Z.M., Guan H.T., Wang M., Liu K., Dai C. et al. Association between single nucleotide polymorphisms in DNA polymerase kappa gene and breast cancer risk in chinese Han population: a STROBE-Compliant Observational Study. Medicine (Baltimore). 2016; 95:e2466. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B136] 136. Sakiyama T., Kohno T., Mimaki S., Ohta T., Yanagitani N., Sobue T., Kunitoh H., Saito R., Shimizu K., Hirama C. et al. Association of amino acid substitution polymorphisms in DNA repair genes TP53, POLI, REV1 and LIG4 with lung cancer risk. Int. J. Cancer. 2005; 114:730–737. [DOI] [PubMed] [Google Scholar]

[B137] 137. Yeom M., Kim I.H., Kim J.K., Kang K., Eoff R.L., Guengerich F.P., Choi J.Y. Effects of twelve germline missense variations on DNA lesion and G-quadruplex bypass activities of human DNA polymerase REV1. Chem. Res. Toxicol. 2016; 29:367–379. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B138] 138. Zhao G., Gleave E.S., Lamers M.H. Single-molecule studies contrast ordered DNA replication with stochastic translesion synthesis. Elife. 2017; 6:e3217. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B139] 139. Sasatani M., Xi Y., Kajimura J., Kawamura T., Piao J., Masuda Y., Honda H., Kubo K., Mikamoto T., Watanabe H. et al. Overexpression of Rev1 promotes the development of carcinogen-induced intestinal adenomas via accumulation of point mutation and suppression of apoptosis proportionally to the Rev1 expression level. Carcinogenesis. 2017; 38:570–578. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B140] 140. Jung Y.S., Qian Y., Chen X. DNA polymerase eta is targeted by Mdm2 for polyubiquitination and proteasomal degradation in response to ultraviolet irradiation. DNA Repair (Amst.). 2012; 11:177–184. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B141] 141. Ziv O., Zeisel A., Mirlas-Neisberg N., Swain U., Nevo R., Ben-Chetrit N., Martelli M.P., Rossi R., Schiesser S., Canman C.E. et al. Identification of novel DNA-damage tolerance genes reveals regulation of translesion DNA synthesis by nucleophosmin. Nat. Commun. 2014; 5:5437. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B142] 142. Sitz J., Blanchet S.A., Gameiro S.F., Biquand E., Morgan T.M., Galloy M., Dessapt J., Lavoie E.G., Blondeau A., Smith B.C. et al. Humanpapillomavirus E7 oncoprotein targets RNF168 to hijack the host DNA damageresponse. Proc.Natl. Acad. Sci. U.S.A. 2019; 24:19552–19562. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B143] 143. Cipolla L., Bertoletti F., Maffia A., Liang C.C., Lehmann A.R., Cohn M.A., Sabbioneda S. UBR5 interacts with the replication fork and protects DNA replication from DNA polymerase eta toxicity. Nucleic Acids Res. 2019; 47:11268–11283. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B144] 144. Wittschieben J.P., Patil V., Glushets V., Robinson L.J., Kusewitt D.F., Wood R.D. Loss of DNA polymerase zeta enhances spontaneous tumorigenesis. Cancer Res. 2010; 70:2770–2778. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B145] 145. O-Wang J., Kawamura K., Tada Y., Ohmori H., Kimura H., Sakiyama S., Tagawa M. DNA polymerase kappa, implicated in spontaneous and DNA damage-induced mutagenesis, is overexpressed in lung cancer. Cancer Res. 2001; 61:5366–5369. [PubMed] [Google Scholar]

[B146] 146. Wang Y., Seimiya M., Kawamura K., Yu L., Ogi T., Takenaga K., Shishikura T., Nakagawara A., Sakiyama S., Tagawa M. et al. Elevated expression of DNA polymerase kappa in human lung cancer is associated with p53 inactivation: negative regulation of POLK promoter activity by p53. Int. J. Oncol. 2004; 25:161–165. [PubMed] [Google Scholar]

[B147] 147. Zhou J., Zhang S., Xie L., Liu P., Xie F., Wu J., Cao J., Ding W.Q. Overexpression of DNA polymerase iota (Poliota) in esophageal squamous cell carcinoma. Cancer Sci. 2012; 103:1574–1579. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B148] 148. Rimkus C., Friederichs J., Rosenberg R., Holzmann B., Siewert J.R., Janssen K.P. Expression of the mitotic checkpoint gene MAD2L2 has prognostic significance in colon cancer. Int. J. Cancer. 2007; 120:207–211. [DOI] [PubMed] [Google Scholar]

[B149] 149. Moinova H.R., Chen W.D., Shen L., Smiraglia D., Olechnowicz J., Ravi L., Kasturi L., Myeroff L., Plass C., Parsons R. et al. HLTF gene silencing in human colon cancer. Proc. Natl. Acad. Sci. U.S.A. 2002; 99:4562–4567. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B150] 150. Yang Y., Gao Y., Zlatanou A., Tateishi S., Yurchenko V., Rogozin I.B., Vaziri C. Diverse roles of RAD18 and Y-family DNA polymerases in tumorigenesis. Cell Cycle. 2018; 17:833–843. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B151] 151. Kotsantis P., Petermann E., Boulton S.J. Mechanisms of oncogene-induced replication stress: jigsaw falling into place. Cancer Discov. 2018; 8:537–555. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B152] 152. Primo L.M.F., Teixeira L.K. DNA replication stress: oncogenes in the spotlight. Genet Mol Biol. 2019; 43:e20190138. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B153] 153. Kotsantis P., Silva L.M., Irmscher S., Jones R.M., Folkes L., Gromak N., Petermann E. Increased global transcription activity as a mechanism of replication stress in cancer. Nat. Commun. 2016; 7:13087. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B154] 154. Chang L., Ruiz P., Ito T., Sellers W.R. Targeting pan-essential genes in cancer: challenges and opportunities. Cancer Cell. 2021; 39:466–479. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B155] 155. Chen Y., Jie X., Xing B., Wu Z., Yang X., Rao X., Xu Y., Zhou D., Dong X., Zhang T. et al. REV1 promotes lung tumorigenesis by activating the Rad18/SERTAD2 axis. Cell Death. Dis. 2022; 13:110. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B156] 156. Zou S., Yang J., Guo J., Su Y., He C., Wu J., Yu L., Ding W.Q., Zhou J. RAD18 promotes the migration and invasion of esophageal squamous cell cancer via the JNK-MMPs pathway. Cancer Lett. 2018; 417:65–74. [DOI] [PubMed] [Google Scholar]

[B157] 157. Yan X., He Y., Yang S., Zeng T., Hua Y., Bao S., Yang F., Duan N., Sun C., Liang Y. et al. a positive feedback loop: rAD18-YAP-TGF-beta between triple-negative breast cancer and macrophages regulates cancer stemness and progression. Cell Death Discov. 2022; 8:196. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B158] 158. Panzarino N.J., Krais J.J., Cong K., Peng M., Mosqueda M., Nayak S.U., Bond S.M., Calvo J.A., Doshi M.B., Bere M. et al. Replication gaps underlie BRCA deficiency and therapy response. Cancer Res. 2021; 81:1388–1397. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B159] 159. Tarantino D., Walker C., Weekes D., Pemberton H., Davidson K., Torga G., Frankum J., Mendes-Pereira A.M., Prince C., Ferro R. et al. Functional screening reveals HORMAD1-driven gene dependencies associated with translesion synthesis and replication stress tolerance. Oncogene. 2022; 41:3969–3977. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B160] 160. Watkins J., Weekes D., Shah V., Gazinska P., Joshi S., Sidhu B., Gillett C., Pinder S., Vanoli F., Jasin M. et al. Genomic complexity profiling reveals that HORMAD1 overexpression contributes to homologous recombination deficiency in triple-negative breast cancers. Cancer Discov. 2015; 5:488–505. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B161] 161. Gao Y., Kardos J., Yang Y., Tamir T.Y., Mutter-Rottmayer E., Weissman B., Major M.B., Kim W.Y., Vaziri C. The cancer/testes (CT) antigen HORMAD1 promotes homologous recombinational DNA repair and radioresistance in lung adenocarcinoma cells. Sci. Rep. 2018; 8:15304. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B162] 162. Nichols B.A., Oswald N.W., McMillan E.A., McGlynn K., Yan J., Kim M.S., Saha J., Mallipeddi P.L., LaDuke S.A., Villalobos P.A. et al. HORMAD1 is a negative prognostic indicator in lung adenocarcinoma and specifies resistance to oxidative and genotoxic stress. Cancer Res. 2018; 78:6196–6208. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B163] 163. Shimizu T., Tateishi S., Tanoue Y., Azuma T., Ohmori H. Somatic hypermutation of immunoglobulin genes in Rad18 knockout mice. DNA Repair (Amst.). 2017; 50:54–60. [DOI] [PubMed] [Google Scholar]

[B164] 164. Lou J., Yang Y., Gu Q., Price B.A., Qiu Y., Fedoriw Y., Desai S., Mose L.E., Chen B., Tateishi S. et al. Rad18 mediates specific mutational signatures and shapes the genomic landscape of carcinogen-induced tumors in vivo. NAR Cancer. 2021; 3:zcaa037. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B165] 165. Stancel J.N., McDaniel L.D., Velasco S., Richardson J., Guo C., Friedberg E.C. Polk mutant mice have a spontaneous mutator phenotype. DNA Repair (Amst.). 2009; 8:1355–1362. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B166] 166. Jansen J.G., Langerak P., Tsaalbi-Shtylik A., van den Berk P., Jacobs H., de Wind N. Strand-biased defect in C/G transversions in hypermutating immunoglobulin genes in Rev1-deficient mice. J. Exp. Med. 2006; 203:319–323. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B167] 167. Fakouri N.B., Durhuus J.A., Regnell C.E., Angleys M., Desler C., Hasan-Olive M.M., Martin-Pardillos A., Tsaalbi-Shtylik A., Thomsen K., Lauritzen M. et al. Rev1 contributes to proper mitochondrial function via the PARP-NAD(+)-SIRT1-PGC1alpha axis. Sci. Rep. 2017; 7:12480. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B168] 168. Dumstorf C.A., Clark A.B., Lin Q., Kissling G.E., Yuan T., Kucherlapati R., McGregor W.G., Kunkel T.A. Participation of mouse DNA polymerase iota in strand-biased mutagenic bypass of UV photoproducts and suppression of skin cancer. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:18083–18088. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B169] 169. Ohkumo T., Kondo Y., Yokoi M., Tsukamoto T., Yamada A., Sugimoto T., Kanao R., Higashi Y., Kondoh H., Tatematsu M. et al. 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. 2006; 26:7696–7706. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B170] 170. Watanabe N., Mii S., Asai N., Asai M., Niimi K., Ushida K., Kato T., Enomoto A., Ishii H., Takahashi M. et al. the REV7 subunit of DNA polymerase zeta is essential for primordial germ cell maintenance in the mouse. J. Biol. Chem. 2013; 288:10459–10471. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B171] 171. Bemark M., Khamlichi A.A., Davies S.L., Neuberger M.S. Disruption of mouse polymerase zeta (Rev3) leads to embryonic lethality and impairs blastocyst development in vitro. Curr. Biol. 2000; 10:1213–1216. [DOI] [PubMed] [Google Scholar]

[B172] 172. Esposito G., Godindagger I., Klein U., Yaspo M.L., Cumano A., Rajewsky K. Disruption of the Rev3l-encoded catalytic subunit of polymerase zeta in mice results in early embryonic lethality. Curr. Biol. 2000; 10:1221–1224. [DOI] [PubMed] [Google Scholar]

[B173] 173. Wittschieben J., Shivji M.K., Lalani E., Jacobs M.A., Marini F., Gearhart P.J., Rosewell I., Stamp G., Wood R.D. Disruption of the developmentally regulated Rev3l gene causes embryonic lethality. Curr. Biol. 2000; 10:1217–1220. [DOI] [PubMed] [Google Scholar]

[B174] 174. Lange S.S., Bedford E., Reh S., Wittschieben J.P., Carbajal S., Kusewitt D.F., DiGiovanni J., Wood R.D. Dual role for mammalian DNA polymerase zeta in maintaining genome stability and proliferative responses. Proc. Natl. Acad. Sci. U.S.A. 2013; 110:E687–696. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B175] 175. Lin Q., Clark A.B., McCulloch S.D., Yuan T., Bronson R.T., Kunkel T.A., Kucherlapati R. Increased susceptibility to UV-induced skin carcinogenesis in polymerase eta-deficient mice. Cancer Res. 2006; 66:87–94. [DOI] [PubMed] [Google Scholar]

[B176] 176. Tanoue Y., Toyoda T., Sun J., Mustofa M.K., Tateishi C., Endo S., Motoyama N., Araki K., Wu D., Okuno Y. et al. Differential roles of Rad18 and Chk2 in genome maintenance and skin carcinogenesis following UV exposure. J. Invest. Dermatol. 2018; 138:2550–2557. [DOI] [PubMed] [Google Scholar]

[B177] 177. Kelland L. the resurgence of platinum-based cancer chemotherapy. Nat. Rev. Cancer. 2007; 7:573–584. [DOI] [PubMed] [Google Scholar]

[B178] 178. Kartalou M., Essigmann J.M. Mechanisms of resistance to cisplatin. Mutat. Res. 2001; 478:23–43. [DOI] [PubMed] [Google Scholar]

[B179] 179. Galluzzi L., Senovilla L., Vitale I., Michels J., Martins I., Kepp O., Castedo M., Kroemer G. Molecular mechanisms of cisplatin resistance. Oncogene. 2012; 31:1869–1883. [DOI] [PubMed] [Google Scholar]

[B180] 180. Park S.B., Krishnan A.V., Lin C.S., Goldstein D., Friedlander M., Kiernan M.C. Mechanisms underlying chemotherapy-induced neurotoxicity and the potential for neuroprotective strategies. Curr. Med. Chem. 2008; 15:3081–3094. [DOI] [PubMed] [Google Scholar]

[B181] 181. Sioka C., Kyritsis A.P. Central and peripheral nervous system toxicity of common chemotherapeutic agents. Cancer Chemother. Pharmacol. 2009; 63:761–767. [DOI] [PubMed] [Google Scholar]

[B182] 182. Lord C.J., Ashworth A. the DNA damage response and cancer therapy. Nature. 2012; 481:287–294. [DOI] [PubMed] [Google Scholar]

[B183] 183. Mamenta E.L., Poma E.E., Kaufmann W.K., Delmastro D.A., Grady H.L., Chaney S.G. Enhanced replicative bypass of platinum-DNA adducts in cisplatin-resistant human ovarian carcinoma cell lines. Cancer Res. 1994; 54:3500–3505. [PubMed] [Google Scholar]

[B184] 184. Bassett E., King N.M., Bryant M.F., Hector S., Pendyala L., Chaney S.G., Cordeiro-Stone M. the role of DNA polymerase eta in translesion synthesis past platinum-DNA adducts in human fibroblasts. Cancer Res. 2004; 64:6469–6475. [DOI] [PubMed] [Google Scholar]

[B185] 185. Ummat A., Rechkoblit O., Jain R., Roy Choudhury J., Johnson R.E., Silverstein T.D., Buku A., Lone S., Prakash L., Prakash S. et al. Structural basis for cisplatin DNA damage tolerance by human polymerase eta during cancer chemotherapy. Nat. Struct. Mol. Biol. 2012; 19:628–632. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B186] 186. Zhao Y., Biertumpfel C., Gregory M.T., Hua Y.J., Hanaoka F., Yang W. Structural basis of human DNA polymerase eta-mediated chemoresistance to cisplatin. Proc. Natl. Acad. Sci. U.S.A. 2012; 109:7269–7274. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B187] 187. Chen Y.W., Cleaver J.E., Hanaoka F., Chang C.F., Chou K.M. a novel role of DNA polymerase eta in modulating cellular sensitivity to chemotherapeutic agents. Mol. Cancer Res. 2006; 4:257–265. [DOI] [PubMed] [Google Scholar]

[B188] 188. Albertella M.R., Green C.M., Lehmann A.R., O’Connor M.J a role for polymerase eta in the cellular tolerance to cisplatin-induced damage. Cancer Res. 2005; 65:9799–9806. [DOI] [PubMed] [Google Scholar]

[B189] 189. Alt A., Lammens K., Chiocchini C., Lammens A., Pieck J.C., Kuch D., Hopfner K.P., Carell T. Bypass of DNA lesions generated during anticancer treatment with cisplatin by DNA polymerase eta. Science. 2007; 318:967–970. [DOI] [PubMed] [Google Scholar]

[B190] 190. Wagner J.M., Karnitz L.M. Cisplatin-induced DNA damage activates replication checkpoint signaling components that differentially affect tumor cell survival. Mol. Pharmacol. 2009; 76:208–214. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B191] 191. Yamashita Y.M., Okada T., Matsusaka T., Sonoda E., Zhao G.Y., Araki K., Tateishi S., Yamaizumi M., Takeda S. RAD18 and RAD54 cooperatively contribute to maintenance of genomic stability in vertebrate cells. EMBO J. 2002; 21:5558–5566. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B192] 192. Ceppi P., Novello S., Cambieri A., Longo M., Monica V., Lo Iacono M., Giaj-Levra M., Saviozzi S., Volante M., Papotti M. et al. Polymerase eta mRNA expression predicts survival of non-small cell lung cancer patients treated with platinum-based chemotherapy. Clin. Cancer Res. 2009; 15:1039–1045. [DOI] [PubMed] [Google Scholar]

[B193] 193. Teng K.Y., Qiu M.Z., Li Z.H., Luo H.Y., Zeng Z.L., Luo R.Z., Zhang H.Z., Wang Z.Q., Li Y.H., Xu R.H. DNA polymerase eta protein expression predicts treatment response and survival of metastatic gastric adenocarcinoma patients treated with oxaliplatin-based chemotherapy. J. Transl. Med. 2010; 8:126. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B194] 194. Srivastava A.K., Han C., Zhao R., Cui T., Dai Y., Mao C., Zhao W., Zhang X., Yu J., Wang Q.E. Enhanced expression of DNA polymerase eta contributes to cisplatin resistance of ovarian cancer stem cells. Proc. Natl. Acad. Sci. U.S.A. 2015; 112:4411–4416. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B195] 195. Thul P.J., Akesson L., Wiking M., Mahdessian D., Geladaki A., Ait Blal H., Alm T., Asplund A., Bjork L., Breckels L.M. et al. A subcellular map of the human proteome. Science. 2017; 356:eaal3321. [DOI] [PubMed] [Google Scholar]

[B196] 196. Uhlen M., Fagerberg L., Hallstrom B.M., Lindskog C., Oksvold P., Mardinoglu A., Sivertsson A., Kampf C., Sjostedt E., Asplund A. et al. Proteomics. Tissue-based map of the human proteome. Science. 2015; 347:1260419. [DOI] [PubMed] [Google Scholar]

[B197] 197. Uhlen M., Zhang C., Lee S., Sjostedt E., Fagerberg L., Bidkhori G., Benfeitas R., Arif M., Liu Z., Edfors F. et al. a pathology atlas of the human cancer transcriptome. Science. 2017; 357:eaan2507. [DOI] [PubMed] [Google Scholar]

[B198] 198. Doles J., Oliver T.G., Cameron E.R., Hsu G., Jacks T., Walker G.C., Hemann M.T. Suppression of Rev3, the catalytic subunit of Pol{zeta}, sensitizes drug-resistant lung tumors to chemotherapy. Proc. Natl. Acad. Sci. U.S.A. 2010; 107:20786–20791. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B199] 199. Xie K., Doles J., Hemann M.T., Walker G.C. Error-prone translesion synthesis mediates acquired chemoresistance. Proc. Nat. Acad. Sci. U.S.A. 2010; 107:20792–20797. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B200] 200. Xu X., Xie K., Zhang X.Q., Pridgen E.M., Park G.Y., Cui D.S., Shi J., Wu J., Kantoff P.W., Lippard S.J. et al. Enhancing tumor cell response to chemotherapy through nanoparticle-mediated codelivery of siRNA and cisplatin prodrug. Proc. Natl. Acad. Sci. U.S.A. 2013; 110:18638–18643. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B201] 201. Vassel F.M., Bian K., Walker G.C., Hemann M.T. Rev7 loss alters cisplatin response and increases drug efficacy in chemotherapy-resistant lung cancer. Proc. Natl. Acad. Sci. U.S.A. 2020; 117:28922–28924. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B202] 202. Ciccia A., Elledge S.J. the DNA damage response: making it safe to play with knives. Mol. Cell. 2010; 40:179–204. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B203] 203. Baillie K.E., Stirling P.C. Beyond kinases: targeting replication stress proteins in cancer therapy. Trends Cancer. 2021; 7:430–446. [DOI] [PubMed] [Google Scholar]

[B204] 204. Smith J., Tho L.M., Xu N., Gillespie D.A. the ATM-Chk2 and ATR-Chk1 pathways in DNA damage signaling and cancer. Adv. Cancer. Res. 2010; 108:73–112. [DOI] [PubMed] [Google Scholar]

[B205] 205. Yazinski S.A., Zou L. Functions, regulation, and therapeutic implications of the ATR checkpoint pathway. Annu. Rev. Genet. 2016; 50:155–173. [DOI] [PubMed] [Google Scholar]

[B206] 206. Pilie P.G., Tang C., Mills G.B., Yap T.A. State-of-the-art strategies for targeting the DNA damage response in cancer. Nat. Rev. Clin. Oncol. 2019; 16:81–104. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B207] 207. Menendez D., Anand J.R., Murphy C.C., Bell W.J., Fu J., Slepushkina N., Buehler E., Martin S.E., Lal-Nag M., Nitiss J.L. et al. Etoposide-induced DNA damage is increased in p53 mutants: identification of ATR and other genes that influence effects of p53 mutations on Top2-induced cytotoxicity. OncoTargets Ther. 2022; 13:332–346. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B208] 208. Karnitz L.M., Zou L. Molecular pathways: targeting ATR in cancer therapy. Clin. Cancer Res. 2015; 21:4780–4785. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B209] 209. Despras E., Daboussi F., Hyrien O., Marheineke K., Kannouche P.L. ATR/Chk1 pathway is essential for resumption of DNA synthesis and cell survival in UV-irradiated XP variant cells. Hum. Mol. Genet. 2010; 19:1690–1701. [DOI] [PubMed] [Google Scholar]

[B210] 210. Maher V.M., Ouellette L.M., Mittlestat M., McCormick J.J. Synergistic effect of caffeine on the cytotoxicity of ultraviolet irradiation and of hydrocarbon epoxides in strains of Xeroderma pigmentosum. Nature. 1975; 258:760–763. [DOI] [PubMed] [Google Scholar]

[B211] 211. Elbaek C.R., Petrosius V., Sorensen C.S. WEE1 kinase limits CDK activities to safeguard DNA replication and mitotic entry. Mutat. Res. 2020; 819-820:111694. [DOI] [PubMed] [Google Scholar]

[B212] 212. Puigvert J.C., Sanjiv K., Helleday T. Targeting DNA repair, DNA metabolism and replication stress as anti-cancer strategies. FEBS J. 2016; 283:232–245. [DOI] [PubMed] [Google Scholar]

[B213] 213. Matheson C.J., Backos D.S., Reigan P. Targeting WEE1 Kinase in Cancer. Trends Pharmacol. Sci. 2016; 37:872–881. [DOI] [PubMed] [Google Scholar]

[B214] 214. Lheureux S., Cristea M.C., Bruce J.P., Garg S., Cabanero M., Mantia-Smaldone G., Olawaiye A.B., Ellard S.L., Weberpals J.I., Wahner Hendrickson A.E. et al. Adavosertib plus gemcitabine for platinum-resistant or platinum-refractory recurrent ovarian cancer: a double-blind, randomised, placebo-controlled, phase 2 trial. Lancet North Am. Ed. 2021; 397:281–292. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B215] 215. Nayak S., Calvo J.A., Cantor S.B. Targeting translesion synthesis (TLS) to expose replication gaps, a unique cancer vulnerability. Expert Opin. Ther. Targets. 2021; 25:27–36. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B216] 216. Satoh M.S., Lindahl T. Role of poly(ADP-ribose) formation in DNA repair. Nature. 1992; 356:356–358. [DOI] [PubMed] [Google Scholar]

[B217] 217. Krishnakumar R., Kraus W.L. The PARP side of the nucleus: molecular actions, physiological outcomes, and clinical targets. Mol. Cell. 2010; 39:8–24. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B218] 218. Bryant H.E., Schultz N., Thomas H.D., Parker K.M., Flower D., Lopez E., Kyle S., Meuth M., Curtin N.J., Helleday T. Specific killing of BRCA2-deficient tumours with inhibitors of poly(ADP-ribose) polymerase. Nature. 2005; 434:913–917. [DOI] [PubMed] [Google Scholar]

[B219] 219. Farmer H., McCabe N., Lord C.J., Tutt A.N., Johnson D.A., Richardson T.B., Santarosa M., Dillon K.J., Hickson I., Knights C. et al. Targeting the DNA repair defect in BRCA mutant cells as a therapeutic strategy. Nature. 2005; 434:917–921. [DOI] [PubMed] [Google Scholar]

[B220] 220. Lord C.J., Ashworth A. PARP inhibitors: synthetic lethality in the clinic. Science. 2017; 355:1152–1158. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B221] 221. Noordermeer S.M., van Attikum H. PARP inhibitor resistance: a tug-of-war in BRCA-mutated cells. Trends Cell Biol. 2019; 29:820–834. [DOI] [PubMed] [Google Scholar]

[B222] 222. Helleday T. The underlying mechanism for the PARP and BRCA synthetic lethality: clearing up the misunderstandings. Mol Oncol. 2011; 5:387–393. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B223] 223. Paes Dias M., Tripathi V., van der Heijden I., Cong K., Manolika E.M., Bhin J., Gogola E., Galanos P., Annunziato S., Lieftink C. et al. Loss of nuclear DNA ligase III reverts PARP inhibitor resistance in BRCA1/53BP1 double-deficient cells by exposing ssDNA gaps. Mol. Cell. 2021; 81:4692–4708. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B224] 224. Johnson D.R., O’Neill B.P Glioblastoma survival in the United States before and during the temozolomide era. J. Neurooncol. 2012; 107:359–364. [DOI] [PubMed] [Google Scholar]

[B225] 225. Lonardi S., Tosoni A., Brandes A.A. Adjuvant chemotherapy in the treatment of high grade gliomas. Cancer Treat. Rev. 2005; 31:79–89. [DOI] [PubMed] [Google Scholar]

[B226] 226. Barthel F.P., Johnson K.C., Varn F.S., Moskalik A.D., Tanner G., Kocakavuk E., Anderson K.J., Abiola O., Aldape K., Alfaro K.D. et al. Longitudinal molecular trajectories of diffuse glioma in adults. Nature. 2019; 576:112–120. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B227] 227. Brennan C.W., Verhaak R.G., McKenna A., Campos B., Noushmehr H., Salama S.R., Zheng S., Chakravarty D., Sanborn J.Z., Berman S.H. et al. The somatic genomic landscape of glioblastoma. Cell. 2013; 155:462–477. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B228] 228. Wang J., Cazzato E., Ladewig E., Frattini V., Rosenbloom D.I., Zairis S., Abate F., Liu Z., Elliott O., Shin Y.J. et al. Clonal evolution of glioblastoma under therapy. Nat. Genet. 2016; 48:768–776. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B229] 229. Alexandrov L.B., Nik-Zainal S., Wedge D.C., Aparicio S.A., Behjati S., Biankin A.V., Bignell G.R., Bolli N., Borg A., Borresen-Dale A.L. et al. Signatures of mutational processes in human cancer. Nature. 2013; 500:415–421. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B230] 230. Touat M., Li Y.Y., Boynton A.N., Spurr L.F., Iorgulescu J.B., Bohrson C.L., Cortes-Ciriano I., Birzu C., Geduldig J.E., Pelton K. et al. Mechanisms and therapeutic implications of hypermutation in gliomas. Nature. 2020; 580:517–523. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B231] 231. Latancia M.T., Moreno N.C., Leandro G.S., Ribeiro V.C., de Souza I., Vieira W.K.M., Bastos A.U., Hoch N.C., Rocha C.R.R., Menck C.F.M. DNA polymerase eta protects human cells against DNA damage induced by the tumor chemotherapeutic temozolomide. Mutat. Res. Genet. Toxicol. Environ. Mutagen. 2022; 878:503498. [DOI] [PubMed] [Google Scholar]

[B232] 232. Ketkar A., Maddukuri L., Penthala N.R., Reed M.R., Zafar M.K., Crooks P.A., Eoff R.L. Inhibition of human DNA polymerases eta and kappa by indole-derived molecules occurs through distinct mechanisms. ACS Chem. Biol. 2019; 14:1337–1351. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B233] 233. Rocha C.R.R., Reily Rocha A., Molina Silva M., Rodrigues Gomes L., Teatin Latancia M., Andrade Tomaz M., de Souza I., Karolynne Seregni Monteiro L., Menck C.F.M. Revealing temozolomide resistance mechanisms via genome-wide CRISPR libraries. Cells. 2020; 9:2573. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B234] 234. Peng C., Chen Z., Wang S., Wang H.W., Qiu W., Zhao L., Xu R., Luo H., Chen Y., Chen D. et al. The error-prone DNA polymerase kappa promotes temozolomide resistance in glioblastoma through Rad17-dependent activation of ATR-Chk1 signaling. Cancer Res. 2016; 76:2340–2353. [DOI] [PubMed] [Google Scholar]

[B235] 235. Wang H., Wu W., Wang H.W., Wang S., Chen Y., Zhang X., Yang J., Zhao S., Ding H.F., Lu D Analysis of specialized DNA polymerases expression in human gliomas: association with prognostic significance. Neuro. Oncol. 2010; 12:679–686. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B236] 236. Szikriszt B., Poti A., Pipek O., Krzystanek M., Kanu N., Molnar J., Ribli D., Szeltner Z., Tusnady G.E., Csabai I. et al. a comprehensive survey of the mutagenic impact of common cancer cytotoxics. Genome Biol. 2016; 17:99. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B237] 237. Lee Y.S., Gregory M.T., Yang W. Human Pol zeta purified with accessory subunits is active in translesion DNA synthesis and complements Pol eta in cisplatin bypass. Proc. Natl. Acad. Sci. U.S.A. 2014; 111:2954–2959. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B238] 238. Brady S.W., Gout A.M., Zhang J. Therapeutic and prognostic insights from the analysis of cancer mutational signatures. Trends Genet. 2022; 38:194–208. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B239] 239. Kaldor J.M., Day N.E., Pettersson F., Clarke E.A., Pedersen D., Mehnert W., Bell J., Host H., Prior P., Karjalainen S. et al. Leukemia following chemotherapy for ovarian cancer. N. Engl. J. Med. 1990; 322:1–6. [DOI] [PubMed] [Google Scholar]

[B240] 240. Travis L.B., Holowaty E.J., Bergfeldt K., Lynch C.F., Kohler B.A., Wiklund T., Curtis R.E., Hall P., Andersson M., Pukkala E. et al. Risk of leukemia after platinum-based chemotherapy for ovarian cancer. N. Engl. J. Med. 1999; 340:351–357. [DOI] [PubMed] [Google Scholar]

[B241] 241. Wojtaszek J.L., Chatterjee N., Najeeb J., Ramos A., Lee M., Bian K., Xue J.Y., Fenton B.A., Park H., Li D. et al. A small molecule targeting mutagenic translesion synthesis improves chemotherapy. Cell. 2019; 178:152–159. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B242] 242. Sail V., Rizzo A.A., Chatterjee N., Dash R.C., Ozen Z., Walker G.C., Korzhnev D.M., Hadden M.K. Identification of small molecule translesion synthesis inhibitors that target the Rev1-CT/RIR protein−protein interaction. ACS Chem. Biol. 2017; 12:1903–1912. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B243] 243. Zhou J., Gelot C., Pantelidou C., Li A., Yucel H., Davis R.E., Farkkila A., Kochupurakkal B., Syed A., Shapiro G.I. et al. A first-in-class polymerase theta inhibitor selectively targets homologous-recombination-deficient tumors. Nat Cancer. 2021; 2:598–610. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B244] 244. Vanarotti M., Evison B.J., Actis M.L., Inoue A., McDonald E.T., Shao Y., Heath R.J., Fujii N. Small-molecules that bind to the ubiquitin-binding motif of REV1 inhibit REV1 interaction with K164-monoubiquitinated PCNA and suppress DNA damage tolerance. Bioorg. Med. Chem. 2018; 26:2345–2353. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B245] 245. Lu S., Dong Z. Additive effects of a small molecular PCNA inhibitor PCNA-I1S and DNA damaging agents on growth inhibition and DNA damage in prostate and lung cancer cells. PLoS One. 2019; 14:e0223894. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B246] 246. Fleming M.C., Chiou L.F., Tumbale P.P., Droby G.N., Lim J., Norris-Drouin J.L., Williams J.G., Pearce K.H., Williams R.S., Vaziri C. et al. Discovery and structural basis of the selectivity of potent cyclic peptide inhibitors of MAGE-A4. J. Med. Chem. 2022; 65:7231–7245. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B247] 247. Du M., Gu J., Liu C., Liu N., Yu Z., Zhou C., Heng W., Cao Z., Wei F., Zhu K. et al. Genome-wide CRISPR screen identified Rad18 as a determinant of doxorubicin sensitivity in osteosarcoma. J. Exp. Clin. Cancer Res. 2022; 41:154. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B248] 248. Huang J., Huen M.S.Y., Kim H., Leung C.C.Y., Glover J.N.M., Yu X., Chen J. RAD18 transmits DNA damage signalling to elicit homologous recombination repair. Nat. Cell Biol. 2009; 11:592–603. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B249] 249. Palle K., Vaziri C. Rad18 E3 ubiquitin ligase activity mediates Fanconi anemia pathway activation and cell survival following DNA Topoisomerase 1 inhibition. Cell Cycle. 2011; 10:1625–1638. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B250] 250. Räschle M., Smeenk G., Hansen R.K., Temu T., Oka Y., Hein M.Y., Nagaraj N., Long D.T., Walter J.C., Hofmann K. et al. DNA repair. Proteomics reveals dynamic assembly of repair complexes during bypass of DNA cross-links. Science. 2015; 348:1253671. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B251] 251. Matsuoka S., Ballif B.A., Smogorzewska A., McDonald E.R., Hurov K.E., Luo J., Bakalarski C.E., Zhao Z., Solimini N., Lerenthal Y. et al. ATM and ATR substrate analysis reveals extensive protein networks responsive to DNA damage. Science. 2007; 316:1160–1166. [DOI] [PubMed] [Google Scholar]

[B252] 252. Adar S., Hu J., Lieb J.D., Sancar A. Genome-wide kinetics of DNA excision repair in relation to chromatin state and mutagenesis. Proc. Natl. Acad. Sci. U.S.A. 2016; 113:E2124–2133. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B253] 253. Sanchez A., Lee D., Kim D.I., Miller K.M. Making connections: integrative signaling mechanisms coordinate DNA break repair in chromatin. Front. Genet. 2021; 12:747734. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B254] 254. Sabbioneda S., Gourdin A.M., Green C.M., Zotter A., Giglia-Mari G., Houtsmuller A., Vermeulen W., Lehmann A.R. Effect of proliferating cell nuclear antigen ubiquitination and chromatin structure on the dynamic properties of the Y-family DNA polymerases. Mol. Biol. Cell. 2008; 19:5193–5202. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B255] 255. Clairmont C.S., D’Andrea A.D REV7 directs DNA repair pathway choice. Trends Cell Biol. 2021; 31:965–978. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B256] 256. Scully R., Panday A., Elango R., Willis N.A. DNA double-strand break repair-pathway choice in somatic mammalian cells. Nat. Rev. Mol. Cell Biol. 2019; 20:698–714. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B257] 257. Yoon J.H., Johnson R.E., Prakash L., Prakash S. Implications of inhibition of Rev1 interaction with Y family DNA polymerases for cisplatin chemotherapy. Genes Dev. 2021; 35:1256–1270. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B258] 258. Stanzione M., Zhong J., Wong E., LaSalle T.J., Wise J.F., Simoneau A., Myers D.T., Phat S., Sade-Feldman M., Lawrence M.S. et al. Translesion DNA synthesis mediates acquired resistance to olaparib plus temozolomide in small cell lung cancer. Sci. Adv. 2022; 8:eabn1229. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B259] 259. Vanarotti M., Grace C.R., Miller D.J., Actis M.L., Inoue A., Evison B.J., Vaithiyalingam S., Singh A.P., McDonald E.T., Fujii N. Structures of REV1 UBM2 domain complex with ubiquitin and with a small-molecule that inhibits the REV1 UBM2-ubiquitin interaction. J. Mol. Biol. 2018; 430:2857–2872. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B260] 260. Actis M.L., Ambaye N.D., Evison B.J., Shao Y., Vanarotti M., Inoue A., McDonald E.T., Kikuchi S., Heath R., Hara K. et al. Identification of the first small-molecule inhibitor of the REV7 DNA repair protein interaction. Bioorg. Med. Chem. 2016; 24:4339–4346. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B261] 261. Zafar M.K., Maddukuri L., Ketkar A., Penthala N.R., Reed M.R., Eddy S., Crooks P.A., Eoff R.L. A small-molecule inhibitor of human DNA polymerase eta potentiates the effects of cisplatin in tumor cells. Biochemistry. 2018; 57:1262–1273. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B262] 262. Punchihewa C., Inoue A., Hishiki A., Fujikawa Y., Connelly M., Evison B., Shao Y., Heath R., Kuraoka I., Rodrigues P. et al. Identification of small molecule proliferating cell nuclear antigen (PCNA) inhibitor that disrupts interactions with PIP-box proteins and inhibits DNA replication. J. Biol. Chem. 2012; 287:14289–14300. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B263] 263. Inoue A., Kikuchi S., Hishiki A., Shao Y., Heath R., Evison B.J., Actis M., Canman C.E., Hashimoto H., Fujii N. A small molecule inhibitor of monoubiquitinated proliferating cell nuclear antigen (PCNA) inhibits repair of interstrand DNA cross-link, enhances DNA double strand break, and sensitizes cancer cells to cisplatin. J. Biol. Chem. 2014; 289:7109–7120. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B264] 264. Konstantinopoulos P.A., da Costa A., Gulhan D., Lee E.K., Cheng S.C., Hendrickson A.E.W., Kochupurakkal B., Kolin D.L., Kohn E.C., Liu J.F. et al. a Replication stress biomarker is associated with response to gemcitabine versus combined gemcitabine and ATR inhibitor therapy in ovarian cancer. Nat. Commun. 2021; 12:5574. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B265] 265. McPherson K.S., Zaino A.M., Dash R.C., Rizzo A.A., Li Y., Hao B., Bezsonova I., Hadden M.K., Korzhnev D.M. Structure-Based Drug Design of Phenazopyridine Derivatives as Inhibitors of Rev1 Interactions in Translesion Synthesis. ChemMedChem. 2021; 16:1126–1132. [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

Roles of trans-lesion synthesis (TLS) DNA polymerases in tumorigenesis and cancer therapy

Jay Anand

Lilly Chiou

Carly Sciandra

Xingyuan Zhang

Jiyong Hong

Di Wu

Pei Zhou

Cyrus Vaziri

Abstract

INTRODUCTION

TLS PATHWAY IMBALANCE DUE TO POLη-DEFICIENCY CAUSES SKIN CANCER

Figure 1.

THE TLS POLYMERASE SWITCH: ROLES OF RAD18-MEDIATED UBIQUITIN SIGNALING

RAD18-INDEPENDENT MECHANISMS OF TLS PATHWAY ACTIVATION

REGULATED EXPRESSION AND ACTIVITY OF TLS FACTORS

TLS OCCURS BOTH AT STALLED LEADING DNA REPLICATION FORK AND POST-REPLICATIVELY

TLS on-the-fly

TLS at post-replicative ssDNA gaps

Figure 2.

EVIDENCE THAT TLS POLYMERASES CONTRIBUTE TO CANCER

Contributions of TLS to cancer based on analysis of mutation signatures

Altered expression of TLS polymerases and their regulators in cancer

Figure 3.

TLS is a dependency of some cancer cells

Contributions of TLS to cancer based on mouse models

Spontaneous tumorigenesis

Oncogene-induced tumorigenesis

Orthotopic tumor models

Carcinogen-induced tumorigenesis

HOW DOES TLS AFFECT RESPONSE TO CANCER THERAPY?

Platinum drugs

Checkpoint kinase inhibitors

Poly(ADP-ribose) polymerase (PARP) inhibitors

ROLE OF TLS IN THERAPY-INDUCED MUTAGENESIS

TARGETING TLS FACTORS FOR THERAPY

Table 1.

CONCLUSIONS AND FUTURE DIRECTIONS

METHODS

Statistical analysis and data presentation

DATA AVAILABILIY

Supplementary Material

Contributor Information

SUPPLEMENTARY DATA

FUNDING

REFERENCES

Associated Data

Supplementary Materials

ACTIONS

PERMALINK

RESOURCES

Similar articles

Cited by other articles

Links to NCBI Databases