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Leveraging the replication stress response to optimize cancer therapy

Nature Reviews Cancer volume 23, pages 6–24 (2023)Cite this article

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Abstract

High-fidelity DNA replication is critical for the faithful transmission of genetic information to daughter cells. Following genotoxic stress, specialized DNA damage tolerance pathways are activated to ensure replication fork progression. These pathways include translesion DNA synthesis, template switching and repriming. In this Review, we describe how DNA damage tolerance pathways impact genome stability, their connection with tumorigenesis and their effects on cancer therapy response. We discuss recent findings that single-strand DNA gap accumulation impacts chemoresponse and explore a growing body of evidence that suggests that different DNA damage tolerance factors, including translesion synthesis polymerases, template switching proteins and enzymes affecting single-stranded DNA gaps, represent useful cancer targets. We further outline how the consequences of DNA damage tolerance mechanisms could inform the discovery of new biomarkers to refine cancer therapies.

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Fig. 1: Major mediators of the replication stress response.
Fig. 2: The replication stress response in cancer and emerging therapeutic targets.
Fig. 3: Chemotherapy and PARPi in BRCA-deficient cancers.
Fig. 4: The replication stress response in cancer and implications for cancer biomarkers.

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References

  1. Zeman, M. K. & Cimprich, K. A. Causes and consequences of replication stress. Nat. Cell Biol. 16, 2–9 (2014).

    Article  CAS  Google Scholar 

  2. Hamperl, S., Bocek, M. J., Saldivar, J. C., Swigut, T. & Cimprich, K. A. Transcription-replication conflict orientation modulates R-loop levels and activates distinct DNA damage responses. Cell 170, 774–786.e19 (2017).

    Article  CAS  Google Scholar 

  3. Genois, M. M. et al. CARM1 regulates replication fork speed and stress response by stimulating PARP1. Mol. Cell 81, 784–800.e8 (2021).

    Article  CAS  Google Scholar 

  4. Quinet, A. et al. PRIMPOL-mediated adaptive response suppresses replication fork reversal in BRCA-deficient cells. Mol. Cell 77, 461–474.e9 (2020). This paper demonstrates how the balance between repriming and fork reversal governs the adaptive response of BRCA1-deficient cells to cisplatin.

    Article  CAS  Google Scholar 

  5. Hanzlikova, H. et al. The importance of poly(ADP-ribose) polymerase as a sensor of unligated Okazaki fragments during DNA replication. Mol. Cell 71, 319–331.e3 (2018).

    Article  CAS  Google Scholar 

  6. Vaitsiankova, A. et al. PARP inhibition impedes the maturation of nascent DNA strands during DNA replication. Nat. Struct. Mol. Biol. 29, 329–338 (2022). This work uncovers the mechanism by which PARPi promotes the formation of ssDNA gaps, which are an emerging determinant of PARPi response in BRCA-deficient cancers.

    Article  CAS  Google Scholar 

  7. van Wietmarschen, N. & Nussenzweig, A. Mechanism for synthetic lethality in BRCA-deficient cancers: no longer lagging behind. Mol. Cell 71, 877–878 (2018).

    Article  Google Scholar 

  8. Cong, K. et al. Replication gaps are a key determinant of PARP inhibitor synthetic lethality with BRCA deficiency. Mol. Cell 81, 3227 (2021). This publication links ssDNA gap accumulation and defective Okazaki fragment processing with PARPi sensitivity in BRCA1-deficient cells, revealing new potential targets in BRCA-deficient cancers.

    Article  CAS  Google Scholar 

  9. Donne, R. et al. Replication stress triggered by nucleotide pool imbalance drives DNA damage and cGAS-STING pathway activation in NAFLD. Dev. Cell 57, 1728–1741.e6 (2022).

    Article  CAS  Google Scholar 

  10. Koç, A., Wheeler, L. J., Mathews, C. K. & Merrill, G. F. Hydroxyurea arrests DNA replication by a mechanism that preserves basal dNTP pools. J. Biol. Chem. 279, 223–230 (2004).

    Article  Google Scholar 

  11. Flach, J. et al. Replication stress is a potent driver of functional decline in ageing haematopoietic stem cells. Nature 512, 198–202 (2014).

    Article  CAS  Google Scholar 

  12. Ubhi, T. & Brown, G. W. Exploiting DNA replication stress for cancer treatment. Cancer Res. 79, 1730–1739 (2019).

    Article  CAS  Google Scholar 

  13. Bartkova, J. et al. Oncogene-induced senescence is part of the tumorigenesis barrier imposed by DNA damage checkpoints. Nature 444, 633–637 (2006).

    Article  CAS  Google Scholar 

  14. Gorgoulis, V. G. et al. Activation of the DNA damage checkpoint and genomic instability in human precancerous lesions. Nature 434, 907–913 (2005).

    Article  CAS  Google Scholar 

  15. Maya-Mendoza, A. et al. High speed of fork progression induces DNA replication stress and genomic instability. Nature 559, 279–284 (2018).

    Article  CAS  Google Scholar 

  16. Quinet, A. & Vindigni, A. Superfast DNA replication causes damage in cancer cells. Nature 559, 186–187 (2018).

    Article  CAS  Google Scholar 

  17. Sale, J. E. Competition, collaboration and coordination–determining how cells bypass DNA damage. J. Cell Sci. 125, 1633–1643 (2012).

    CAS  Google Scholar 

  18. Sale, J. E. Translesion DNA synthesis and mutagenesis in eukaryotes. Cold Spring Harb. Perspect. Biol. 5, a012708 (2013).

    Article  Google Scholar 

  19. Branzei, D. & Szakal, B. DNA damage tolerance by recombination: molecular pathways and DNA structures. DNA Repair 44, 68–75 (2016).

    Article  CAS  Google Scholar 

  20. Quinet, A., Tirman, S., Cybulla, E., Meroni, A. & Vindigni, A. To skip or not to skip: choosing repriming to tolerate DNA damage. Mol. Cell 81, 649–658 (2021).

    Article  CAS  Google Scholar 

  21. Waters, L. S. et al. Eukaryotic translesion polymerases and their roles and regulation in DNA damage tolerance. Microbiol. Mol. Biol. Rev. 73, 134–154 (2009).

    Article  CAS  Google Scholar 

  22. Branzei, D. Ubiquitin family modifications and template switching. FEBS Lett. 585, 2810–2817 (2011).

    Article  CAS  Google Scholar 

  23. Neelsen, K. J. & Lopes, M. Replication fork reversal in eukaryotes: from dead end to dynamic response. Nat. Rev. Mol. Cell Biol. 16, 207–220 (2015).

    Article  CAS  Google Scholar 

  24. Zellweger, R. et al. Rad51-mediated replication fork reversal is a global response to genotoxic treatments in human cells. J. Cell Biol. 208, 563–579 (2015).

    Article  CAS  Google Scholar 

  25. Bianchi, J. et al. PrimPol bypasses UV photoproducts during eukaryotic chromosomal DNA replication. Mol. Cell 52, 566–573 (2013).

    Article  CAS  Google Scholar 

  26. Garcia-Gomez, S. et al. PrimPol, an archaic primase/polymerase operating in human cells. Mol. Cell 52, 541–553 (2013).

    Article  CAS  Google Scholar 

  27. Mouron, S. et al. Repriming of DNA synthesis at stalled replication forks by human PrimPol. Nat. Struct. Mol. Biol. 20, 1383–1389 (2013).

    Article  CAS  Google Scholar 

  28. Wan, L. et al. hPrimpol1/CCDC111 is a human DNA primase-polymerase required for the maintenance of genome integrity. EMBO Rep. 14, 1104–1112 (2013).

    Article  CAS  Google Scholar 

  29. Adar, S., Izhar, L., Hendel, A., Geacintov, N. & Livneh, Z. Repair of gaps opposite lesions by homologous recombination in mammalian cells. Nucleic Acids Res. 37, 5737–5748 (2009).

    Article  CAS  Google Scholar 

  30. 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 30, 519–529 (2008).

    Article  CAS  Google Scholar 

  31. Jansen, J. G. et al. Mammalian polymerase zeta is essential for post-replication repair of UV-induced DNA lesions. DNA Repair 8, 1444–1451 (2009).

    Article  CAS  Google Scholar 

  32. Tirman, S. et al. Temporally distinct post-replicative repair mechanisms fill PRIMPOL-dependent ssDNA gaps in human cells. Mol. Cell 81, 4026–4040.e8 (2021). This study describes previously uncharacterized ssDNA gap-filling pathways in human cells and shows that these pathways can be targeted to increase cancer cell genomic instability and sensitivity to PARPi and cisplatin treatment.

    Article  CAS  Google Scholar 

  33. Sale, J. E., Lehmann, A. R. & Woodgate, R. Y-family DNA polymerases and their role in tolerance of cellular DNA damage. Nat. Rev. Mol. Cell Biol. 13, 141–152 (2012).

    Article  CAS  Google Scholar 

  34. Gan, G. N., Wittschieben, J. P., Wittschieben, B. O. & Wood, R. D. DNA polymerase zeta (pol zeta) in higher eukaryotes. Cell Res. 18, 174–183 (2008).

    Article  CAS  Google Scholar 

  35. McCulloch, S. D. & Kunkel, T. A. The fidelity of DNA synthesis by eukaryotic replicative and translesion synthesis polymerases. Cell Res. 18, 148–161 (2008).

    Article  CAS  Google Scholar 

  36. Yang, Y. et al. Diverse roles of RAD18 and Y-family DNA polymerases in tumorigenesis. Cell Cycle 17, 833–843 (2018).

    Article  CAS  Google Scholar 

  37. Yamanaka, K., Chatterjee, N., Hemann, M. T. & Walker, G. C. Inhibition of mutagenic translesion synthesis: a possible strategy for improving chemotherapy? PLoS Genet. 13, e1006842 (2017).

    Article  Google Scholar 

  38. Watanabe, K. et al. Rad18 guides poleta to replication stalling sites through physical interaction and PCNA monoubiquitination. EMBO J. 23, 3886–3896 (2004).

    Article  CAS  Google Scholar 

  39. Tateishi, S. et al. Enhanced genomic instability and defective postreplication repair in RAD18 knockout mouse embryonic stem cells. Mol. Cell Biol. 23, 474–481 (2003).

    Article  CAS  Google Scholar 

  40. Yoon, J. H., Prakash, S. & Prakash, L. Requirement of Rad18 protein for replication through DNA lesions in mouse and human cells. Proc. Natl Acad. Sci. USA 109, 7799–7804 (2012).

    Article  CAS  Google Scholar 

  41. Lou, J. et al. Rad18 mediates specific mutational signatures and shapes the genomic landscape of carcinogen-induced tumors in vivo. NAR Cancer 3, zcaa037 (2021).

    Article  Google Scholar 

  42. The Human Protein Atlas. RAD18. Protein Atlas https://www.proteinatlas.org/ENSG00000070950-RAD18 (2022).

  43. Baatar, S. et al. High RAD18 expression is associated with disease progression and poor prognosis in patients with gastric cancer. Ann. Surg. Oncol. 27, 4360–4368 (2020).

    Article  Google Scholar 

  44. Wu, B. et al. High expression of RAD18 in glioma induces radiotherapy resistance via down-regulating P53 expression. Biomed. Pharmacother. 112, 108555 (2019).

    Article  CAS  Google Scholar 

  45. Kikuchi, S., Hara, K., Shimizu, T., Sato, M. & Hashimoto, H. Structural basis of recruitment of DNA polymerase zeta by interaction between REV1 and REV7 proteins. J. Biol. Chem. 287, 33847–33852 (2012).

    Article  CAS  Google Scholar 

  46. In Het Panhuis, W. et al. Rev1 deficiency induces replication stress to cause metabolic dysfunction differently in males and females. Am. J. Physiol. Endocrinol. Metab. 322, E319–E329 (2022).

    Article  Google Scholar 

  47. Jansen, J. G. et al. Strand-biased defect in C/G transversions in hypermutating immunoglobulin genes in Rev1-deficient mice. J. Exp. Med. 203, 319–323 (2006).

    Article  Google Scholar 

  48. Roberts, S. A. & Gordenin, D. A. Hypermutation in human cancer genomes: footprints and mechanisms. Nat. Rev. Cancer 14, 786–800 (2014).

    Article  CAS  Google Scholar 

  49. Dumstorf, C. A., Mukhopadhyay, S., Krishnan, E., Haribabu, B. & McGregor, W. G. REV1 is implicated in the development of carcinogen-induced lung cancer. Mol. Cancer Res. 7, 247–254 (2009).

    Article  CAS  Google Scholar 

  50. Wittschieben, J. P., Reshmi, S. C., Gollin, S. M. & Wood, R. D. Loss of DNA polymerase zeta causes chromosomal instability in mammalian cells. Cancer Res. 66, 134–142 (2006).

    Article  CAS  Google Scholar 

  51. Wittschieben, J. P. et al. Loss of DNA polymerase zeta enhances spontaneous tumorigenesis. Cancer Res. 70, 2770–2778 (2010).

    Article  CAS  Google Scholar 

  52. Martin, S. K., Tomida, J. & Wood, R. D. Disruption of DNA polymerase zeta engages an innate immune response. Cell Rep. 34, 108775 (2021).

    Article  CAS  Google Scholar 

  53. Seplyarskiy, V. B., Bazykin, G. A. & Soldatov, R. A. Polymerase zeta activity is linked to replication timing in humans: evidence from mutational signatures. Mol. Biol. Evol. 32, 3158–3172 (2015).

    CAS  Google Scholar 

  54. Zhang, S. et al. REV3L 3’UTR 460T>C polymorphism in microRNA target sites contributes to lung cancer susceptibility. Oncogene 32, 242–250 (2013).

    Article  CAS  Google Scholar 

  55. Wang, H. et al. REV3L confers chemoresistance to cisplatin in human gliomas: the potential of its RNAi for synergistic therapy. Neuro-Oncol. 11, 790–802 (2009).

    Article  Google Scholar 

  56. Zhou, J. et al. Overexpression of DNA polymerase iota (Poliota) in esophageal squamous cell carcinoma. Cancer Sci. 103, 1574–1579 (2012).

    Article  CAS  Google Scholar 

  57. Zhu, X. et al. REV3L, the catalytic subunit of DNA polymerase zeta, is involved in the progression and chemoresistance of esophageal squamous cell carcinoma. Oncol. Rep. 35, 1664–1670 (2016).

    Article  CAS  Google Scholar 

  58. Brondello, J. M. et al. Novel evidences for a tumor suppressor role of Rev3, the catalytic subunit of Pol zeta. Oncogene 27, 6093–6101 (2008).

    Article  CAS  Google Scholar 

  59. Pan, Q., Fang, Y., Xu, Y., Zhang, K. & Hu, X. Down-regulation of DNA polymerases kappa, eta, iota, and zeta in human lung, stomach, and colorectal cancers. Cancer Lett. 217, 139–147 (2005).

    Article  CAS  Google Scholar 

  60. Yoon, J. H., Prakash, L. & Prakash, S. Highly error-free role of DNA polymerase eta in the replicative bypass of UV-induced pyrimidine dimers in mouse and human cells. Proc. Natl Acad. Sci. USA 106, 18219–18224 (2009).

    Article  CAS  Google Scholar 

  61. Masutani, C. et al. The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase eta. Nature 399, 700–704 (1999).

    Article  CAS  Google Scholar 

  62. Srivastava, A. K. et al. Enhanced expression of DNA polymerase eta contributes to cisplatin resistance of ovarian cancer stem cells. Proc. Natl Acad. Sci. USA 112, 4411–4416 (2015).

    Article  CAS  Google Scholar 

  63. Zhou, W. et al. Expression of DNA translesion synthesis polymerase eta in head and neck squamous cell cancer predicts resistance to gemcitabine and cisplatin-based chemotherapy. PLoS ONE 8, e83978 (2013).

    Article  Google Scholar 

  64. Alexandrov, L. B. & Stratton, M. R. Mutational signatures: the patterns of somatic mutations hidden in cancer genomes. Curr. Opin. Genet. Dev. 24, 52–60 (2014).

    Article  CAS  Google Scholar 

  65. Rogozin, I. B. et al. DNA polymerase eta mutational signatures are found in a variety of different types of cancer. Cell Cycle 17, 348–355 (2018).

    Article  CAS  Google Scholar 

  66. Jansen, J. G. et al. Redundancy of mammalian Y family DNA polymerases in cellular responses to genomic DNA lesions induced by ultraviolet light. Nucleic Acids Res. 42, 11071–11082 (2014).

    Article  CAS  Google Scholar 

  67. Sertic, S. et al. Coordinated activity of Y family TLS polymerases and EXO1 protects non-S phase cells from UV-induced cytotoxic lesions. Mol. Cell 70, 34–47.e4 (2018).

    Article  CAS  Google Scholar 

  68. McIntyre, J. Polymerase iota - an odd sibling among Y family polymerases. DNA Repair 86, 102753 (2020).

    Article  CAS  Google Scholar 

  69. Luedeke, M. et al. Predisposition for TMPRSS2-ERG fusion in prostate cancer by variants in DNA repair genes. Cancer Epidemiol. Biomark. Prev. 18, 3030–3035 (2009).

    Article  CAS  Google Scholar 

  70. Silvestrov, P., Maier, S. J., Fang, M. & Cisneros, G. A. DNArCdb: a database of cancer biomarkers in DNA repair genes that includes variants related to multiple cancer phenotypes. DNA Repair 70, 10–17 (2018).

    Article  CAS  Google Scholar 

  71. Sakiyama, T. et al. Association of amino acid substitution polymorphisms in DNA repair genes TP53, POLI, REV1 and LIG4 with lung cancer risk. Int. J. Cancer 114, 730–737 (2005).

    Article  CAS  Google Scholar 

  72. Yang, J., Chen, Z., Liu, Y., Hickey, R. J. & Malkas, L. H. Altered DNA polymerase iota expression in breast cancer cells leads to a reduction in DNA replication fidelity and a higher rate of mutagenesis. Cancer Res. 64, 5597–5607 (2004).

    Article  CAS  Google Scholar 

  73. Yuan, F. et al. Overexpressed DNA polymerase iota regulated by JNK/c-Jun contributes to hypermutagenesis in bladder cancer. PLoS ONE 8, e69317 (2013).

    Article  CAS  Google Scholar 

  74. Zou, S. et al. DNA polymerase iota (Pol iota) promotes invasion and metastasis of esophageal squamous cell carcinoma. Oncotarget 7, 32274–32285 (2016).

    Article  Google Scholar 

  75. Twayana, S. et al. Translesion polymerase eta both facilitates DNA replication and promotes increased human genetic variation at common fragile sites. Proc. Natl Acad. Sci. USA 118, e2106477118 (2021).

    Article  CAS  Google Scholar 

  76. Tonzi, P., Yin, Y., Lee, C. W. T., Rothenberg, E. & Huang, T. T. Translesion polymerase kappa-dependent DNA synthesis underlies replication fork recovery. eLife 7, e41426 (2018).

    Article  Google Scholar 

  77. Hile, S. E. & Eckert, K. A. DNA polymerase kappa produces interrupted mutations and displays polar pausing within mononucleotide microsatellite sequences. Nucleic Acids Res. 36, 688–696 (2008).

    Article  CAS  Google Scholar 

  78. Tubbs, A. et al. Dual roles of poly(dA:dT) tracts in replication initiation and fork collapse. Cell 174, 1127–1142.e19 (2018).

    Article  CAS  Google Scholar 

  79. Ma, K. et al. Common fragile sites: genomic hotspots of DNA damage and carcinogenesis. Int. J. Mol. Sci. 13, 11974–11999 (2012).

    Article  CAS  Google Scholar 

  80. Stern, H. R., Sefcikova, J., Chaparro, V. E. & Beuning, P. J. Mammalian DNA polymerase kappa activity and specificity. Molecules 24, 2805 (2019).

    Article  Google Scholar 

  81. Q-Wang, J. et al. DNA polymerase kappa, implicated in spontaneous and DNA damage-induced mutagenesis, is overexpressed in lung cancer. Cancer Res. 61, 5366–5369 (2001).

    Google Scholar 

  82. Lovett, S. T. Template-switching during replication fork repair in bacteria. DNA Repair 56, 118–128 (2017).

    Article  CAS  Google Scholar 

  83. Ulrich, H. D. Timing and spacing of ubiquitin-dependent DNA damage bypass. FEBS Lett. 585, 2861–2867 (2011).

    Article  CAS  Google Scholar 

  84. Branzei, D., Seki, M. & Enomoto, T. Rad18/Rad5/Mms2-mediated polyubiquitination of PCNA is implicated in replication completion during replication stress. Genes Cell 9, 1031–1042 (2004).

    Article  CAS  Google Scholar 

  85. Takahashi, T. S., Wollscheid, H. P., Lowther, J. & Ulrich, H. D. Effects of chain length and geometry on the activation of DNA damage bypass by polyubiquitylated PCNA. Nucleic Acids Res. 48, 3042–3052 (2020).

    Article  CAS  Google Scholar 

  86. Chiu, R. K. et al. Lysine 63-polyubiquitination guards against translesion synthesis-induced mutations. PLoS Genet. 2, e116 (2006).

    Article  Google Scholar 

  87. The Human Protein Atlas. UBE2N. Protein Atlas https://www.proteinatlas.org/ENSG00000177889-UBE2N (2022).

  88. 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 42, 237–249 (2011).

    Article  CAS  Google Scholar 

  89. Motegi, A. et al. Polyubiquitination of proliferating cell nuclear antigen by HLTF and SHPRH prevents genomic instability from stalled replication forks. Proc. Natl Acad. Sci. USA 105, 12411–12416 (2008).

    Article  CAS  Google Scholar 

  90. The Human Protein Atlas. HLTF. Protein Atlas https://www.proteinatlas.org/ENSG00000071794-HLTF (2022).

  91. The Human Protein Atlas. SHPRH. Protein Atlas https://www.proteinatlas.org/ENSG00000146414-SHPRH (2022).

  92. Liu, L. et al. HLTF suppresses the migration and invasion of colorectal cancer cells via TGF‑β/SMAD signaling in vitro. Int. J. Oncol. 53, 2780–2788 (2018).

    CAS  Google Scholar 

  93. Moinova, H. R. et al. HLTF gene silencing in human colon cancer. Proc. Natl Acad. Sci. USA 99, 4562–4567 (2002).

    Article  CAS  Google Scholar 

  94. Sandhu, S. et al. Loss of HLTF function promotes intestinal carcinogenesis. Mol. Cancer 11, 18 (2012).

    Article  CAS  Google Scholar 

  95. Dhont, L., Mascaux, C. & Belayew, A. The helicase-like transcription factor (HLTF) in cancer: loss of function or oncomorphic conversion of a tumor suppressor? Cell Mol. Life Sci. 73, 129–147 (2016).

    Article  CAS  Google Scholar 

  96. Jasin, M. & Rothstein, R. Repair of strand breaks by homologous recombination. Cold Spring Harb. Perspect. Biol. 5, a012740 (2013).

    Article  Google Scholar 

  97. Piberger, A. L. et al. PrimPol-dependent single-stranded gap formation mediates homologous recombination at bulky DNA adducts. Nat. Commun. 11, 5863 (2020).

    Article  CAS  Google Scholar 

  98. Liberi, G. et al. Rad51-dependent DNA structures accumulate at damaged replication forks in sgs1 mutants defective in the yeast ortholog of BLM RecQ helicase. Genes Dev. 19, 339–350 (2005).

    Article  CAS  Google Scholar 

  99. Pietrobon, V. et al. The chromatin assembly factor 1 promotes Rad51-dependent template switches at replication forks by counteracting D-loop disassembly by the RecQ-type helicase Rqh1. PLoS Biol. 12, e1001968 (2014).

    Article  Google Scholar 

  100. The Human Protein Atlas. BLM. Protein Atlas https://www.proteinatlas.org/ENSG00000197299-BLM (2022).

  101. The Human Protein Atlas. NBN. Protein Atlas https://www.proteinatlas.org/ENSG00000104320-NBN (2022).

  102. Hasty, P. & Montagna, C. Chromosomal rearrangements in cancer: detection and potential causal mechanisms. Mol. Cell. Oncol. 1, e29904 (2014).

    Article  Google Scholar 

  103. Tanaka, H. & Watanabe, T. Mechanisms underlying recurrent genomic amplification in human cancers. Trends Cancer 6, 462–477 (2020).

    Article  CAS  Google Scholar 

  104. Vicario, R. et al. Patterns of HER2 gene amplification and response to anti-HER2 therapies. PLoS ONE 10, e0129876 (2015).

    Article  Google Scholar 

  105. Sakofsky, C. J. & Malkova, A. Break induced replication in eukaryotes: mechanisms, functions, and consequences. Crit. Rev. Biochem. Mol. Biol. 52, 395–413 (2017).

    Article  CAS  Google Scholar 

  106. Blastyak, A. et al. Yeast Rad5 protein required for postreplication repair has a DNA helicase activity specific for replication fork regression. Mol. Cell 28, 167–175 (2007).

    Article  CAS  Google Scholar 

  107. Bugreev, D. V., Rossi, M. J. & Mazin, A. V. Cooperation of RAD51 and RAD54 in regression of a model replication fork. Nucleic Acids Res. 39, 2153–2164 (2011).

    Article  CAS  Google Scholar 

  108. Betous, R. et al. SMARCAL1 catalyzes fork regression and Holliday junction migration to maintain genome stability during DNA replication. Genes Dev. 26, 151–162 (2012).

    Article  CAS  Google Scholar 

  109. Kolinjivadi, A. M. et al. Smarcal1-mediated fork reversal triggers Mre11-dependent degradation of nascent DNA in the absence of Brca2 and Stable Rad51 nucleofilaments. Mol. Cell 67, 867–881.e7 (2017).

    Article  CAS  Google Scholar 

  110. Gari, K., Decaillet, C., Stasiak, A. Z., Stasiak, A. & Constantinou, A. The Fanconi anemia protein FANCM can promote branch migration of Holliday junctions and replication forks. Mol. Cell 29, 141–148 (2008).

    Article  CAS  Google Scholar 

  111. Vujanovic, M. et al. Replication fork slowing and reversal upon DNA damage require PCNA polyubiquitination and ZRANB3 DNA translocase activity. Mol. Cell 67, 882–890.e5 (2017).

    Article  CAS  Google Scholar 

  112. Yusufzai, T. & Kadonaga, J. T. Annealing helicase 2 (AH2), a DNA-rewinding motor with an HNH motif. Proc. Natl Acad. Sci. USA 107, 20970–20973 (2010).

    Article  CAS  Google Scholar 

  113. Bai, G. et al. HLTF promotes fork reversal, limiting replication stress resistance and preventing multiple mechanisms of unrestrained DNA synthesis. Mol. Cell 78, 1237–1251.e7 (2020).

    Article  CAS  Google Scholar 

  114. Blastyak, A., Hajdu, I., Unk, I. & Haracska, L. Role of double-stranded DNA translocase activity of human HLTF in replication of damaged DNA. Mol. Cell Biol. 30, 684–693 (2010).

    Article  CAS  Google Scholar 

  115. Bansbach, C. E., Boerkoel, C. F. & Cortez, D. SMARCAL1 and replication stress: an explanation for SIOD? Nucleus 1, 245–248 (2010).

    Google Scholar 

  116. Elizondo, L. I. et al. Schimke immuno-osseous dysplasia: SMARCAL1 loss-of-function and phenotypic correlation. J. Med. Genet. 46, 49–59 (2009).

    Article  CAS  Google Scholar 

  117. The Human Protein Atlas. SMARCAL1. Protein Atlas https://www.proteinatlas.org/ENSG00000138375-SMARCAL1 (2022).

  118. Bogliolo, M. et al. Biallelic truncating FANCM mutations cause early-onset cancer but not Fanconi anemia. Genet. Med. 20, 458–463 (2018).

    Article  CAS  Google Scholar 

  119. Catucci, I. et al. Individuals with FANCM biallelic mutations do not develop Fanconi anemia, but show risk for breast cancer, chemotherapy toxicity and may display chromosome fragility. Genet. Med. 20, 452–457 (2018).

    Article  CAS  Google Scholar 

  120. Jones, S. et al. Genomic analyses of gynaecologic carcinosarcomas reveal frequent mutations in chromatin remodelling genes. Nat. Commun. 5, 5006 (2014).

    Article  CAS  Google Scholar 

  121. The Human Protein Atlas. ZRANB3. Protein Atlas https://www.proteinatlas.org/ENSG00000121988-ZRANB3 (2022).

  122. Hiramoto, T. et al. Mutations of a novel human RAD54 homologue, RAD54B, in primary cancer. Oncogene 18, 3422–3426 (1999).

    Article  CAS  Google Scholar 

  123. Smirnova, M., Van Komen, S., Sung, P. & Klein, H. L. Effects of tumor-associated mutations on Rad54 functions. J. Biol. Chem. 279, 24081–24088 (2004).

    Article  CAS  Google Scholar 

  124. Fugger, K. et al. FBH1 catalyzes regression of stalled replication forks. Cell Rep. 10, 1749–1757 (2015).

    Article  CAS  Google Scholar 

  125. Berti, M. et al. Sequential role of RAD51 paralog complexes in replication fork remodeling and restart. Nat. Commun. 11, 3531 (2020).

    Article  CAS  Google Scholar 

  126. Rein, H. L., Bernstein, K. A. & Baldock, R. A. RAD51 paralog function in replicative DNA damage and tolerance. Curr. Opin. Genet. Dev. 71, 86–91 (2021).

    Article  CAS  Google Scholar 

  127. Grundy, M. K., Buckanovich, R. J. & Bernstein, K. A. Regulation and pharmacological targeting of RAD51 in cancer. NAR Cancer 2, zcaa024 (2020).

    Article  Google Scholar 

  128. Cruz, C. et al. RAD51 foci as a functional biomarker of homologous recombination repair and PARP inhibitor resistance in germline BRCA-mutated breast cancer. Ann. Oncol. 29, 1203–1210 (2018).

    Article  CAS  Google Scholar 

  129. Maacke, H. et al. DNA repair and recombination factor Rad51 is over-expressed in human pancreatic adenocarcinoma. Oncogene 19, 2791–2795 (2000).

    Article  CAS  Google Scholar 

  130. Maacke, H. et al. Over-expression of wild-type Rad51 correlates with histological grading of invasive ductal breast cancer. Int. J. Cancer 88, 907–913 (2000).

    Article  CAS  Google Scholar 

  131. Raderschall, E. et al. Elevated levels of Rad51 recombination protein in tumor cells. Cancer Res. 62, 219–225 (2002).

    CAS  Google Scholar 

  132. Berti, M. et al. Human RECQ1 promotes restart of replication forks reversed by DNA topoisomerase I inhibition. Nat. Struct. Mol. Biol. 20, 347–354 (2013).

    Article  CAS  Google Scholar 

  133. Thangavel, S. et al. DNA2 drives processing and restart of reversed replication forks in human cells. J. Cell Biol. 208, 545–562 (2015).

    Article  CAS  Google Scholar 

  134. Chu, W. K. & Hickson, I. D. RecQ helicases: multifunctional genome caretakers. Nat. Rev. Cancer 9, 644–654 (2009).

    Article  CAS  Google Scholar 

  135. Abu-Libdeh, B. et al. RECON syndrome is a genome instability disorder caused by mutations in the DNA helicase RECQL1. J. Clin. Invest. 132, e147301 (2022).

    Article  CAS  Google Scholar 

  136. Cybulski, C. et al. Germline RECQL mutations are associated with breast cancer susceptibility. Nat. Genet. 47, 643–646 (2015).

    Article  CAS  Google Scholar 

  137. The Human Protein Atlas. DNA2. Protein Atlas https://www.proteinatlas.org/ENSG00000138346-DNA2 (2022).

  138. The Human Protein Atlas. WRN. Protein Atlas https://www.proteinatlas.org/ENSG00000165392-WRN (2022).

  139. The Human Protein Atlas. RECQL. Protein Atlas https://www.proteinatlas.org/ENSG00000004700-RECQL (2022).

  140. Ray Chaudhuri, A. et al. Replication fork stability confers chemoresistance in BRCA-deficient cells. Nature 535, 382–387 (2016). This publication was one of the first to explore how replication fork stability is associated with chemoresponse in BRCA-deficient cancer cells.

    Article  Google Scholar 

  141. Schlacher, K. et al. Double-strand break repair-independent role for BRCA2 in blocking stalled replication fork degradation by MRE11. Cell 145, 529–542 (2011).

    Article  CAS  Google Scholar 

  142. Ying, S., Hamdy, F. C. & Helleday, T. Mre11-dependent degradation of stalled DNA replication forks is prevented by BRCA2 and PARP1. Cancer Res. 72, 2814–2821 (2012).

    Article  CAS  Google Scholar 

  143. Dungrawala, H. et al. RADX promotes genome stability and modulates chemosensitivity by regulating RAD51 at replication forks. Mol. Cell 67, 374–386.e5 (2017).

    Article  CAS  Google Scholar 

  144. Guillemette, S. et al. Resistance to therapy in BRCA2 mutant cells due to loss of the nucleosome remodeling factor CHD4. Genes Dev. 29, 489–494 (2015).

    Article  CAS  Google Scholar 

  145. The Human Protein Atlas. MUS81. Protein Atlas https://www.proteinatlas.org/ENSG00000172732-MUS81 (2022).

  146. Lemacon, D. et al. MRE11 and EXO1 nucleases degrade reversed forks and elicit MUS81-dependent fork rescue in BRCA2-deficient cells. Nat. Commun. 8, 860 (2017).

    Article  Google Scholar 

  147. Rondinelli, B. et al. EZH2 promotes degradation of stalled replication forks by recruiting MUS81 through histone H3 trimethylation. Nat. Cell Biol. 19, 1371–1378 (2017).

    Article  CAS  Google Scholar 

  148. Krais, J. J. & Johnson, N. Ectopic RNF168 expression promotes break-induced replication-like DNA synthesis at stalled replication forks. Nucleic Acids Res. 48, 4298–4308 (2020).

    Article  CAS  Google Scholar 

  149. Zong, D. et al. BRCA1 haploinsufficiency is masked by RNF168-mediated chromatin ubiquitylation. Mol. Cell 73, 1267–1281.e7 (2019).

    Article  CAS  Google Scholar 

  150. Batenburg, N. L. et al. Cockayne syndrome group B protein regulates fork restart, fork progression and MRE11-dependent fork degradation in BRCA1/2-deficient cells. Nucleic Acids Res. 49, 12836–12854 (2021).

    Article  CAS  Google Scholar 

  151. The Human Protein Atlas. ERCC6. Protein Atlas https://www.proteinatlas.org/ENSG00000225830-ERCC6 (2022).

  152. Eckelmann, B. J. et al. XRCC1 promotes replication restart, nascent fork degradation and mutagenic DNA repair in BRCA2-deficient cells. NAR Cancer 2, zcaa013 (2020).

    Article  Google Scholar 

  153. The Human Protein Atlas. XRCC1. Protein Atlas https://www.proteinatlas.org/ENSG00000073050-XRCC1 (2022).

  154. Heller, R. C. & Marians, K. J. Replication fork reactivation downstream of a blocked nascent leading strand. Nature 439, 557–562 (2006).

    Article  CAS  Google Scholar 

  155. Fumasoni, M., Zwicky, K., Vanoli, F., Lopes, M. & Branzei, D. Error-free DNA damage tolerance and sister chromatid proximity during DNA replication rely on the Polα/Primase/Ctf4 complex. Mol. Cell 57, 812–823 (2015).

    Article  CAS  Google Scholar 

  156. Bailey, L. J., Bianchi, J. & Doherty, A. J. PrimPol is required for the maintenance of efficient nuclear and mitochondrial DNA replication in human cells. Nucleic Acids Res. 47, 4026–4038 (2019).

    Article  CAS  Google Scholar 

  157. González-Acosta, D. et al. PrimPol-mediated repriming facilitates replication traverse of DNA interstrand crosslinks. EMBO J. 40, e106355 (2021).

    Article  Google Scholar 

  158. Keen, B. A., Bailey, L. J., Jozwiakowski, S. K. & Doherty, A. J. Human PrimPol mutation associated with high myopia has a DNA replication defect. Nucleic Acids Res. 42, 12102–12111 (2014).

    Article  CAS  Google Scholar 

  159. Kobayashi, K. et al. Repriming by PrimPol is critical for DNA replication restart downstream of lesions and chain-terminating nucleosides. Cell Cycle 15, 1997–2008 (2016).

    Article  CAS  Google Scholar 

  160. Panzarino, N. J. et al. Replication gaps underlie BRCA deficiency and therapy response. Cancer Res. 81, 1388–1397 (2021). This work links replication-associated ssDNA gaps with therapy response in BRCA-deficient cancer cells and proposes a gap-centred framework for understanding ‘BRCAness’ phenotypes.

    Article  CAS  Google Scholar 

  161. Taglialatela, A. et al. REV1-Polζ maintains the viability of homologous recombination-deficient cancer cells through mutagenic repair of PRIMPOL-dependent ssDNA gaps. Mol. Cell 81, 4008–4025.e7 (2021). This study highlights the dependence of BRCA-deficient cells on ssDNA gap filling to promote cell survival, supporting the use of novel TLS inhibitors in combination with other cancer therapies.

    Article  CAS  Google Scholar 

  162. The Human Protein Atlas. PRIMPOL. Protein Atlas https://www.proteinatlas.org/ENSG00000164306-PRIMPOL (2022).

  163. Diaz-Talavera, A. et al. A cancer-associated point mutation disables the steric gate of human PrimPol. Sci. Rep. 9, 1121 (2019).

    Article  Google Scholar 

  164. Bamford, S. et al. The COSMIC (Catalogue of Somatic Mutations in Cancer) database and website. Br. J. Cancer 91, 355–358 (2004).

    Article  CAS  Google Scholar 

  165. Wong, R. P., Garcia-Rodriguez, N., Zilio, N., Hanulova, M. & Ulrich, H. D. Processing of DNA polymerase-blocking lesions during genome replication is spatially and temporally segregated from replication forks. Mol. Cell 77, 3–16.e14 (2020).

    Article  CAS  Google Scholar 

  166. Nayak, S. et al. Inhibition of the translesion synthesis polymerase REV1 exploits replication gaps as a cancer vulnerability. Sci. Adv. 6, eaaz7808 (2020).

    Article  CAS  Google Scholar 

  167. Thakar, T. et al. Ubiquitinated-PCNA protects replication forks from DNA2-mediated degradation by regulating Okazaki fragment maturation and chromatin assembly. Nat. Commun. 11, 2147 (2020).

    Article  CAS  Google Scholar 

  168. Thakar, T. et al. Lagging strand gap suppression connects BRCA-mediated fork protection to nucleosome assembly by ensuring PCNA-dependent CAF-1 recycling. Nat. Commun. 13, 5323 (2022).

    Article  CAS  Google Scholar 

  169. Liu, R. L., Dong, Y., Deng, Y. Z., Wang, W. J. & Li, W. D. Tumor suppressor miR-145 reverses drug resistance by directly targeting DNA damage-related gene RAD18 in colorectal cancer. Tumour Biol. 36, 5011–5019 (2015).

    Article  CAS  Google Scholar 

  170. Williams, S. A., Longerich, S., Sung, P., Vaziri, C. & Kupfer, G. M. The E3 ubiquitin ligase RAD18 regulates ubiquitylation and chromatin loading of FANCD2 and FANCI. Blood 117, 5078–5087 (2011).

    Article  CAS  Google Scholar 

  171. Geng, L., Huntoon, C. J. & Karnitz, L. M. RAD18-mediated ubiquitination of PCNA activates the Fanconi anemia DNA repair network. J. Cell Biol. 191, 249–257 (2010).

    Article  CAS  Google Scholar 

  172. Lopez-Martinez, D., Liang, C. C. & Cohn, M. A. Cellular response to DNA interstrand crosslinks: the Fanconi anemia pathway. Cell Mol. Life Sci. 73, 3097–3114 (2016).

    Article  CAS  Google Scholar 

  173. Kim, H. & D’Andrea, A. D. Regulation of DNA cross-link repair by the Fanconi anemia/BRCA pathway. Genes. Dev. 26, 1393–1408 (2012).

    Article  CAS  Google Scholar 

  174. Fenteany, G. et al. A series of xanthenes inhibiting Rad6 function and Rad6-Rad18 interaction in the PCNA ubiquitination cascade. iScience 25, 104053 (2022).

    Article  CAS  Google Scholar 

  175. Punchihewa, C. 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. 287, 14289–14300 (2012).

    Article  CAS  Google Scholar 

  176. Inoue, A. et al. 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. 289, 7109–7120 (2014).

    Article  CAS  Google Scholar 

  177. Qin, Z. et al. DNA-damage tolerance mediated by PCNA*Ub fusions in human cells is dependent on Rev1 but not Polη. Nucleic Acids Res. 41, 7356–7369 (2013).

    Article  CAS  Google Scholar 

  178. Coleman, K. E. et al. USP1-trapping lesions as a source of DNA replication stress and genomic instability. Nat. Commun. 13, 1740 (2022).

    Article  CAS  Google Scholar 

  179. Lim, K. S. et al. USP1 is required for replication fork protection in BRCA1-deficient tumors. Mol. Cell 72, 925–941.e4 (2018).

    Article  CAS  Google Scholar 

  180. Lin, X., Okuda, T., Trang, J. & Howell, S. B. Human REV1 modulates the cytotoxicity and mutagenicity of cisplatin in human ovarian carcinoma cells. Mol. Pharmacol. 69, 1748–1754 (2006).

    Article  CAS  Google Scholar 

  181. Okuda, T., Lin, X., Trang, J. & Howell, S. B. Suppression of hREV1 expression reduces the rate at which human ovarian carcinoma cells acquire resistance to cisplatin. Mol. Pharmacol. 67, 1852–1860 (2005).

    Article  CAS  Google Scholar 

  182. Wojtaszek, J. L. et al. A small molecule targeting mutagenic translesion synthesis improves chemotherapy. Cell 178, 152–159.e11 (2019).

    Article  CAS  Google Scholar 

  183. Ikeh, K. E. et al. REV1 inhibition enhances radioresistance and autophagy. Cancers 13, 5290 (2021).

    Article  CAS  Google Scholar 

  184. Mulcahy Levy, J. M. & Thorburn, A. Autophagy in cancer: moving from understanding mechanism to improving therapy responses in patients. Cell Death Differ. 27, 843–857 (2020).

    Article  Google Scholar 

  185. Ketkar, A. et al. Inhibition of human DNA polymerases eta and kappa by indole-derived molecules occurs through distinct mechanisms. ACS Chem. Biol. 14, 1337–1351 (2019).

    Article  CAS  Google Scholar 

  186. Coggins, G. E. et al. N-Aroyl indole thiobarbituric acids as inhibitors of DNA repair and replication stress response polymerases. ACS Chem. Biol. 8, 1722–1729 (2013).

    Article  CAS  Google Scholar 

  187. Zafar, M. K. et al. A small-molecule inhibitor of human DNA polymerase η potentiates the effects of cisplatin in tumor cells. Biochemistry 57, 1262–1273 (2018).

    Article  CAS  Google Scholar 

  188. Ceppi, P. et al. Polymerase eta mRNA expression predicts survival of non-small cell lung cancer patients treated with platinum-based chemotherapy. Clin. Cancer Res. 15, 1039–1045 (2009).

    Article  CAS  Google Scholar 

  189. Zhang, J. et al. A PolH transcript with a short 3’UTR enhances PolH expression and mediates cisplatin resistance. Cancer Res. 79, 3714–3724 (2019).

    Article  CAS  Google Scholar 

  190. Jha, V. & Ling, H. Structural basis for human DNA polymerase kappa to bypass cisplatin intrastrand cross-link (Pt-GG) lesion as an efficient and accurate extender. J. Mol. Biol. 430, 1577–1589 (2018).

    Article  CAS  Google Scholar 

  191. Yang, Y. et al. DNA repair factor RAD18 and DNA polymerase Polκ confer tolerance of oncogenic DNA replication stress. J. Cell Biol. 216, 3097–3115 (2017).

    Article  CAS  Google Scholar 

  192. Doles, J. et al. Suppression of Rev3, the catalytic subunit of Polζ, sensitizes drug-resistant lung tumors to chemotherapy. Proc. Natl Acad. Sci. USA 107, 20786–20791 (2010).

    Article  CAS  Google Scholar 

  193. 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. USA 117, 28922–28924 (2020).

    Article  CAS  Google Scholar 

  194. Okina, S. et al. High expression of REV7 is an independent prognostic indicator in patients with diffuse large B-cell lymphoma treated with rituximab. Int. J. Hematol. 102, 662–669 (2015).

    Article  CAS  Google Scholar 

  195. Niimi, K. et al. Suppression of REV7 enhances cisplatin sensitivity in ovarian clear cell carcinoma cells. Cancer Sci. 105, 545–552 (2014).

    Article  CAS  Google Scholar 

  196. Ghezraoui, H. et al. 53BP1 cooperation with the REV7-shieldin complex underpins DNA structure-specific NHEJ. Nature 560, 122–127 (2018).

    Article  CAS  Google Scholar 

  197. Xu, G. et al. REV7 counteracts DNA double-strand break resection and affects PARP inhibition. Nature 521, 541–544 (2015). This work shows that REV7 loss but not loss of the REV1 or REV3L TLS polymerases represents a BRCA1-independent mechanism of HR restoration that contributes to PARPi resistance.

    Article  CAS  Google Scholar 

  198. Yousefzadeh, M. J. & Wood, R. D. DNA polymerase POLQ and cellular defense against DNA damage. DNA Repair 12, 1–9 (2013).

    Article  CAS  Google Scholar 

  199. Ceccaldi, R. et al. Homologous-recombination-deficient tumours are dependent on Poltheta-mediated repair. Nature 518, 258–262 (2015).

    Article  CAS  Google Scholar 

  200. Mateos-Gomez, P. A. et al. Mammalian polymerase theta promotes alternative NHEJ and suppresses recombination. Nature 518, 254–257 (2015).

    Article  CAS  Google Scholar 

  201. Zhou, J. et al. A first-in-class polymerase theta inhibitor selectively targets homologous-recombination-deficient tumors. Nat. Cancer 2, 598–610 (2021). This study, along with Ceccaldi et al. (2015) and Mateos-Gomez et al. (2015), establishes Polθ as a target in BRCA-deficient cancers and provides critical mechanistic insight for utility of Polθ inhibitors, one of which is now being used in a clinical trial.

    Article  CAS  Google Scholar 

  202. Schrempf, A., Slyskova, J. & Loizou, J. I. Targeting the DNA repair enzyme polymerase theta in cancer therapy. Trends Cancer 7, 98–111 (2021).

    Article  CAS  Google Scholar 

  203. Alexandrov, L. B. et al. The repertoire of mutational signatures in human cancer. Nature 578, 94–101 (2020). This paper provides a comprehensive landscape of tumour mutational signatures and links replication stress response mechanisms with several of these mutational patterns in cancers.

    Article  CAS  Google Scholar 

  204. Higgins, G. S. et al. Overexpression of POLQ confers a poor prognosis in early breast cancer patients. Oncotarget 1, 175–184 (2010).

    Article  Google Scholar 

  205. Lemée, F. et al. DNA polymerase theta up-regulation is associated with poor survival in breast cancer, perturbs DNA replication, and promotes genetic instability. Proc. Natl Acad. Sci. USA 107, 13390–13395 (2010).

    Article  Google Scholar 

  206. Hodge, C. D., Spyracopoulos, L. & Glover, J. N. Ubc13: the Lys63 ubiquitin chain building machine. Oncotarget 7, 64471–64504 (2016).

    Article  Google Scholar 

  207. Wu, Z., Shen, S., Zhang, Z., Zhang, W. & Xiao, W. Ubiquitin-conjugating enzyme complex Uev1A-Ubc13 promotes breast cancer metastasis through nuclear factor-small ka, CyrillicB mediated matrix metalloproteinase-1 gene regulation. Breast Cancer Res. 16, R75 (2014).

    Article  Google Scholar 

  208. Wu, Z., Neufeld, H., Torlakovic, E. & Xiao, W. Uev1A-Ubc13 promotes colorectal cancer metastasis through regulating CXCL1 expression via NF-small ka, CyrillicB activation. Oncotarget 9, 15952–15967 (2018).

    Article  Google Scholar 

  209. Dikshit, A. et al. UBE2N promotes melanoma growth via MEK/FRA1/SOX10 signaling. Cancer Res. 78, 6462–6472 (2018).

    Article  CAS  Google Scholar 

  210. Zhang, X. et al. The inhibition of UBC13 expression and blockage of the DNMT1-CHFR-Aurora A pathway contribute to paclitaxel resistance in ovarian cancer. Cell Death Dis. 9, 93 (2018).

    Article  Google Scholar 

  211. Cheng, J. et al. A small-molecule inhibitor of UBE2N induces neuroblastoma cell death via activation of p53 and JNK pathways. Cell Death Dis. 5, e1079 (2014).

    Article  CAS  Google Scholar 

  212. Pulvino, M. et al. Inhibition of proliferation and survival of diffuse large B-cell lymphoma cells by a small-molecule inhibitor of the ubiquitin-conjugating enzyme Ubc13-Uev1A. Blood 120, 1668–1677 (2012).

    Article  CAS  Google Scholar 

  213. Lu, R. et al. MUS81 participates in the progression of serous ovarian cancer associated with dysfunctional DNA repair system. Front. Oncol. 9, 1189 (2019).

    Article  Google Scholar 

  214. Zhong, A. et al. MUS81 inhibition increases the sensitivity to therapy effect in epithelial ovarian cancer via regulating CyclinB pathway. J. Cancer 10, 2276–2287 (2019).

    Article  CAS  Google Scholar 

  215. Krais, J. J. et al. RNF168-mediated ubiquitin signaling inhibits the viability of BRCA1-null cancers. Cancer Res. 80, 2848–2860 (2020).

    Article  CAS  Google Scholar 

  216. Luijsterburg, M. S. et al. A PALB2-interacting domain in RNF168 couples homologous recombination to DNA break-induced chromatin ubiquitylation. eLife 6, e20922 (2017).

    Article  Google Scholar 

  217. Bryant, H. E. et al. Specific killing of BRCA2-deficient tumours with inhibitors of poly(ADP-ribose) polymerase. Nature 434, 913–917 (2005).

    Article  CAS  Google Scholar 

  218. Farmer, H. et al. Targeting the DNA repair defect in BRCA mutant cells as a therapeutic strategy. Nature 434, 917–921 (2005).

    Article  CAS  Google Scholar 

  219. Fong, P. C. et al. Inhibition of poly(ADP-ribose) polymerase in tumors from BRCA mutation carriers. N. Engl. J. Med. 361, 123–134 (2009).

    Article  CAS  Google Scholar 

  220. Turner, N., Tutt, A. & Ashworth, A. Targeting the DNA repair defect of BRCA tumours. Curr. Opin. Pharmacol. 5, 388–393 (2005).

    Article  CAS  Google Scholar 

  221. Zimmermann, M. et al. CRISPR screens identify genomic ribonucleotides as a source of PARP-trapping lesions. Nature 559, 285–289 (2018).

    Article  CAS  Google Scholar 

  222. D’Andrea, A. D. Mechanisms of PARP inhibitor sensitivity and resistance. DNA Repair 71, 172–176 (2018).

    Article  Google Scholar 

  223. Janysek, D. C., Kim, J., Duijf, P. H. G. & Dray, E. Clinical use and mechanisms of resistance for PARP inhibitors in homologous recombination-deficient cancers. Transl. Oncol. 14, 101012 (2021).

    Article  CAS  Google Scholar 

  224. Fugger, K., Hewitt, G., West, S. C. & Boulton, S. J. Tackling PARP inhibitor resistance. Trends Cancer 7, 1102–1118 (2021).

    Article  CAS  Google Scholar 

  225. Gatti, M., Imhof, R., Huang, Q., Baudis, M. & Altmeyer, M. The ubiquitin Ligase TRIP12 limits PARP1 trapping and constrains PARP inhibitor efficiency. Cell Rep. 32, 107985 (2020).

    Article  CAS  Google Scholar 

  226. Krastev, D. B. et al. The ubiquitin-dependent ATPase p97 removes cytotoxic trapped PARP1 from chromatin. Nat. Cell Biol. 24, 62–73 (2022).

    Article  CAS  Google Scholar 

  227. Noordermeer, S. M. & van Attikum, H. PARP inhibitor resistance: a tug-of-war in BRCA-mutated cells. Trends Cell Biol. 29, 820–834 (2019).

    Article  CAS  Google Scholar 

  228. Gogola, E. et al. Selective loss of PARG restores parylation and counteracts PARP inhibitor-mediated synthetic lethality. Cancer Cell 33, 1078–1093.e12 (2018).

    Article  CAS  Google Scholar 

  229. Yazinski, S. A. et al. ATR inhibition disrupts rewired homologous recombination and fork protection pathways in PARP inhibitor-resistant BRCA-deficient cancer cells. Genes Dev. 31, 318–332 (2017).

    Article  CAS  Google Scholar 

  230. Simoneau, A., Xiong, R. & Zou, L. The trans cell cycle effects of PARP inhibitors underlie their selectivity toward BRCA1/2-deficient cells. Genes Dev. 35, 1271–1289 (2021).

    Article  Google Scholar 

  231. Mehta, K. P. M. et al. CHK1 phosphorylates PRIMPOL to promote replication stress tolerance. Sci. Adv. 8, eabm0314 (2022).

    Article  CAS  Google Scholar 

  232. Guo, E. et al. FEN1 endonuclease as a therapeutic target for human cancers with defects in homologous recombination. Proc. Natl Acad. Sci. USA 117, 19415–19424 (2020).

    Article  CAS  Google Scholar 

  233. Paes Dias, M. et al. Loss of nuclear DNA ligase III reverts PARP inhibitor resistance in BRCA1/53BP1 double-deficient cells by exposing ssDNA gaps. Mol. Cell 81, 4692–4708.e9 (2021). This publication describes a mechanism of PARPi resistance mediated by LIG3 loss and accumulation of ssDNA in BRCA1/53BP1-deficient cells, pointing towards the clinical promise of targeting ssDNA gaps therapeutically to address chemoresistance.

    Article  CAS  Google Scholar 

  234. Cantor, S. B. Revisiting the BRCA-pathway through the lens of replication gap suppression: “Gaps determine therapy response in BRCA mutant cancer”. DNA Repair 107, 103209 (2021).

    Article  CAS  Google Scholar 

  235. Cong, K. & Cantor, S. B. Exploiting replication gaps for cancer therapy. Mol. Cell 82, 2363–2369 (2022).

    Article  CAS  Google Scholar 

  236. Dev, H. et al. Shieldin complex promotes DNA end-joining and counters homologous recombination in BRCA1-null cells. Nat. Cell Biol. 20, 954–965 (2018).

    Article  CAS  Google Scholar 

  237. Saldivar, J. C. et al. An intrinsic S/G. Science 361, 806–810 (2018).

    Article  CAS  Google Scholar 

  238. Couch, F. B. et al. ATR phosphorylates SMARCAL1 to prevent replication fork collapse. Genes Dev. 27, 1610–1623 (2013).

    Article  CAS  Google Scholar 

  239. Dibitetto, D. et al. Fork slowing and reversal as an adaptive response to chronic ATR inhibition. Preprint at bioRxriv https://doi.org/10.1101/2021.05.18.444697 (2021).

    Article  Google Scholar 

  240. Mutreja, K. et al. ATR-mediated global fork slowing and reversal assist fork traverse and prevent chromosomal breakage at DNA interstrand cross-links. Cell Rep. 24, 2629–2642.e5 (2018).

    Article  CAS  Google Scholar 

  241. Yazinski, S. A. & Zou, L. Functions, regulation, and therapeutic implications of the ATR checkpoint pathway. Annu. Rev. Genet. 50, 155–173 (2016).

    Article  CAS  Google Scholar 

  242. Parmar, K. et al. The CHK1 inhibitor prexasertib exhibits monotherapy activity in high-grade serous ovarian cancer models and sensitizes to PARP inhibition. Clin. Cancer Res. 25, 6127–6140 (2019).

    Article  CAS  Google Scholar 

  243. Chaikovsky, A. C. et al. The AMBRA1 E3 ligase adaptor regulates the stability of cyclin D. Nature 592, 794–798 (2021).

    Article  CAS  Google Scholar 

  244. Maiani, E. et al. AMBRA1 regulates cyclin D to guard S-phase entry and genomic integrity. Nature 592, 799–803 (2021).

    Article  CAS  Google Scholar 

  245. Simoneschi, D. et al. CRL4. Nature 592, 789–793 (2021).

    Article  CAS  Google Scholar 

  246. Young, L. A. et al. Differential activity of ATR and WEE1 inhibitors in a highly sensitive subpopulation of DLBCL linked to replication stress. Cancer Res. 79, 3762–3775 (2019).

    Article  CAS  Google Scholar 

  247. Ghelli Luserna Di Rora, A. et al. Synergism through WEE1 and CHK1 inhibition in acute lymphoblastic leukemia. Cancers 11, 1654 (2019).

    Article  Google Scholar 

  248. Chen, X. et al. Targeting replicative stress and DNA repair by combining PARP and Wee1 kinase inhibitors is synergistic in triple negative breast cancers with cyclin E or BRCA1 alteration. Cancers 13, 1656 (2021).

    Article  CAS  Google Scholar 

  249. Ha, D. H. et al. Antitumor effect of a WEE1 inhibitor and potentiation of olaparib sensitivity by DNA damage response modulation in triple-negative breast cancer. Sci. Rep. 10, 9930 (2020).

    Article  CAS  Google Scholar 

  250. Beck, H. et al. Cyclin-dependent kinase suppression by WEE1 kinase protects the genome through control of replication initiation and nucleotide consumption. Mol. Cell Biol. 32, 4226–4236 (2012).

    Article  CAS  Google Scholar 

  251. Elbaek, C. R. et al. WEE1 kinase protects the stability of stalled DNA replication forks by limiting CDK2 activity. Cell Rep. 38, 110261 (2022).

    Article  CAS  Google Scholar 

  252. Pillay, N. et al. DNA replication vulnerabilities render ovarian cancer cells sensitive to poly(ADP-Ribose) glycohydrolase inhibitors. Cancer Cell 35, 519–533.e8 (2019). This work is the first to address how PARG inhibition impacts replication stress across a variety of ovarian cancer models and to test combinatorial treatment of PARG inhibitors with those of CHK1 and WEE1 in preclinical models.

    Article  CAS  Google Scholar 

  253. Buisson, R., Boisvert, J. L., Benes, C. H. & Zou, L. Distinct but concerted roles of ATR, DNA-PK, and Chk1 in countering replication stress during S phase. Mol. Cell 59, 1011–1024 (2015).

    Article  CAS  Google Scholar 

  254. Balmus, G. et al. ATM orchestrates the DNA-damage response to counter toxic non-homologous end-joining at broken replication forks. Nat. Commun. 10, 87 (2019).

    Article  CAS  Google Scholar 

  255. Bartek, J. & Lukas, J. Chk1 and Chk2 kinases in checkpoint control and cancer. Cancer Cell 3, 421–429 (2003).

    Article  CAS  Google Scholar 

  256. Choi, M., Kipps, T. & Kurzrock, R. ATM mutations in cancer: therapeutic implications. Mol. Cancer Ther. 15, 1781–1791 (2016).

    Article  CAS  Google Scholar 

  257. Lavin, M. F. & Yeo, A. J. Clinical potential of ATM inhibitors. Mutat. Res. 821, 111695 (2020).

    Article  CAS  Google Scholar 

  258. Lin, Y. F. et al. PIDD mediates the association of DNA-PKcs and ATR at stalled replication forks to facilitate the ATR signaling pathway. Nucleic Acids Res. 46, 1847–1859 (2018).

    Article  CAS  Google Scholar 

  259. Hafsi, H. et al. Combined ATR and DNA-PK inhibition radiosensitizes tumor cells independently of their p53 status. Front. Oncol. 8, 245 (2018).

    Article  Google Scholar 

  260. Fok, J. H. L. et al. AZD7648 is a potent and selective DNA-PK inhibitor that enhances radiation, chemotherapy and olaparib activity. Nat. Commun. 10, 5065 (2019).

    Article  Google Scholar 

  261. Zhang, M. et al. CDK inhibitors in cancer therapy, an overview of recent development. Am. J. Cancer Res. 11, 1913–1935 (2021).

    CAS  Google Scholar 

  262. Alvarez-Fernandez, M. & Malumbres, M. Mechanisms of sensitivity and resistance to CDK4/6 inhibition. Cancer Cell 37, 514–529 (2020).

    Article  CAS  Google Scholar 

  263. Asghar, U. S., Kanani, R., Roylance, R. & Mittnacht, S. Systematic review of molecular biomarkers predictive of resistance to CDK4/6 inhibition in metastatic breast cancer. JCO Precis. Oncol. 6, e2100002 (2022).

    Article  Google Scholar 

  264. Dean, J. L., McClendon, A. K. & Knudsen, E. S. Modification of the DNA damage response by therapeutic CDK4/6 inhibition. J. Biol. Chem. 287, 29075–29087 (2012).

    Article  CAS  Google Scholar 

  265. Johnson, S. M. et al. Mitigation of hematologic radiation toxicity in mice through pharmacological quiescence induced by CDK4/6 inhibition. J. Clin. Invest. 120, 2528–2536 (2010).

    Article  CAS  Google Scholar 

  266. Roberts, P. J. et al. Multiple roles of cyclin-dependent kinase 4/6 inhibitors in cancer therapy. J. Natl Cancer Inst. 104, 476–487 (2012).

    Article  CAS  Google Scholar 

  267. Li, S. et al. Pan-cancer analysis reveals synergistic effects of CDK4/6i and PARPi combination treatment in RB-proficient and RB-deficient breast cancer cells. Cell Death Dis. 11, 219 (2020).

    Article  CAS  Google Scholar 

  268. Crozier, L. et al. CDK4/6 inhibitors induce replication stress to cause long-term cell cycle withdrawal. EMBO J. 41, e108599 (2022).

    Article  CAS  Google Scholar 

  269. Salvador-Barbero, B. et al. CDK4/6 inhibitors impair recovery from cytotoxic chemotherapy in pancreatic adenocarcinoma. Cancer Cell 38, 584 (2020). This study provides mechanistic evidence to support the emerging treatment paradigm of CDK4/6 inhibition, which is used clinically in combination with DNA-damaging therapies.

    Article  CAS  Google Scholar 

  270. Panagiotou, E., Gomatou, G., Trontzas, I. P., Syrigos, N. & Kotteas, E. Cyclin-dependent kinase (CDK) inhibitors in solid tumors: a review of clinical trials. Clin. Transl. Oncol. 24, 161–192 (2022).

    Article  CAS  Google Scholar 

  271. Coquel, F. et al. SAMHD1 acts at stalled replication forks to prevent interferon induction. Nature 557, 57–61 (2018). This work provides a mechanistic link between replication fork stress, nucleolytic processing and accumulation of cytosolic DNA that activates an innate immune-like response.

    Article  CAS  Google Scholar 

  272. Kreienkamp, R. et al. A cell-intrinsic interferon-like response links replication stress to cellular aging caused by progerin. Cell Rep. 22, 2006–2015 (2018).

    Article  CAS  Google Scholar 

  273. Orvain, C. et al. Hair follicle stem cell replication stress drives IFI16/STING-dependent inflammation in hidradenitis suppurativa. J. Clin. Invest. 130, 3777–3790 (2020).

    Article  CAS  Google Scholar 

  274. Shen, Y. J. et al. Genome-derived cytosolic DNA mediates type I interferon-dependent rejection of B cell lymphoma cells. Cell Rep. 11, 460–473 (2015).

    Article  CAS  Google Scholar 

  275. Técher, H. & Pasero, P. The replication stress response on a narrow path between genomic instability and inflammation. Front. Cell Dev. Biol. 9, 702584 (2021).

    Article  Google Scholar 

  276. Harding, S. M. et al. Mitotic progression following DNA damage enables pattern recognition within micronuclei. Nature 548, 466–470 (2017).

    Article  CAS  Google Scholar 

  277. Mackenzie, K. J. et al. cGAS surveillance of micronuclei links genome instability to innate immunity. Nature 548, 461–465 (2017).

    Article  CAS  Google Scholar 

  278. Ragu, S., Matos-Rodrigues, G. & Lopez, B. S. Replication stress, DNA damage, inflammatory cytokines and innate immune response. Genes 11, 409 (2020).

    Article  CAS  Google Scholar 

  279. Ni, Y. et al. High TGF-beta signature predicts immunotherapy resistance in gynecologic cancer patients treated with immune checkpoint inhibition. NPJ Precis. Oncol. 5, 101 (2021).

    Article  CAS  Google Scholar 

  280. Li, S. et al. Cancer immunotherapy via targeted TGF-beta signalling blockade in TH cells. Nature 587, 121–125 (2020).

    Article  CAS  Google Scholar 

  281. Jackson, L. M. et al. Loss of MED12 activates the TGFbeta pathway to promote chemoresistance and replication fork stability in BRCA-deficient cells. Nucleic Acids Res. 49, 12855–12869 (2021).

    Article  CAS  Google Scholar 

  282. Moskwa, P. et al. miR-182-mediated downregulation of BRCA1 impacts DNA repair and sensitivity to PARP inhibitors. Mol. Cell 41, 210–220 (2011).

    Article  CAS  Google Scholar 

  283. Martinez-Ruiz, H. et al. A TGFbeta-miR-182-BRCA1 axis controls the mammary differentiation hierarchy. Sci. Signal. 9, ra118 (2016).

    Article  Google Scholar 

  284. Kirshner, J. et al. Inhibition of transforming growth factor-beta1 signaling attenuates ataxia telangiectasia mutated activity in response to genotoxic stress. Cancer Res. 66, 10861–10869 (2006).

    Article  CAS  Google Scholar 

  285. Wang, M. et al. Novel Smad proteins localize to IR-induced double-strand breaks: interplay between TGFβ and ATM pathways. Nucleic Acids Res. 41, 933–942 (2013).

    Article  CAS  Google Scholar 

  286. Wiegman, E. M., Blaese, M. A., Loeffler, H., Coppes, R. P. & Rodemann, H. P. TGFbeta-1 dependent fast stimulation of ATM and p53 phosphorylation following exposure to ionizing radiation does not involve TGFbeta-receptor I signalling. Radiother. Oncol. 83, 289–295 (2007).

    Article  CAS  Google Scholar 

  287. Tang, B. et al. Transforming growth factor-beta can suppress tumorigenesis through effects on the putative cancer stem or early progenitor cell and committed progeny in a breast cancer xenograft model. Cancer Res. 67, 8643–8652 (2007).

    Article  CAS  Google Scholar 

  288. Wen, H. et al. Inhibiting of self-renewal, migration and invasion of ovarian cancer stem cells by blocking TGF-beta pathway. PLoS ONE 15, e0230230 (2020).

    Article  CAS  Google Scholar 

  289. Yan, G. et al. TGFbeta/cyclin D1/Smad-mediated inhibition of BMP4 promotes breast cancer stem cell self-renewal activity. Oncogenesis 10, 21 (2021).

    Article  CAS  Google Scholar 

  290. Chan, T. A. et al. Development of tumor mutation burden as an immunotherapy biomarker: utility for the oncology clinic. Ann. Oncol. 30, 44–56 (2019).

    Article  CAS  Google Scholar 

  291. Strickler, J. H., Hanks, B. A. & Khasraw, M. Tumor mutational burden as a predictor of immunotherapy response: is more always better? Clin. Cancer Res. 27, 1236–1241 (2021).

    Article  CAS  Google Scholar 

  292. Wang, S., Xie, K. & Liu, T. Cancer immunotherapies: from efficacy to resistance mechanisms - not only checkpoint matters. Front. Immunol. 12, 690112 (2021).

    Article  CAS  Google Scholar 

  293. Sim, M. J. W. & Sun, P. D. T cell recognition of tumor neoantigens and insights into T cell immunotherapy. Front. Immunol. 13, 833017 (2022).

    Article  CAS  Google Scholar 

  294. Salas-Benito, D. et al. Paradigms on immunotherapy combinations with chemotherapy. Cancer Discov. 11, 1353–1367 (2021).

    Article  CAS  Google Scholar 

  295. Zhang, J. Y., Yan, Y. Y., Li, J. J., Adhikari, R. & Fu, L. W. PD-1/PD-L1 based combinational cancer therapy: icing on the cake. Front. Pharmacol. 11, 722 (2020).

    Article  CAS  Google Scholar 

  296. McGrail, D. J. et al. Replication stress response defects are associated with response to immune checkpoint blockade in nonhypermutated cancers. Sci. Transl Med. 13, eabe6201 (2021).

    Article  CAS  Google Scholar 

  297. Zou, S. et al. RAD18 promotes the migration and invasion of esophageal squamous cell cancer via the JNK-MMPs pathway. Cancer Lett. 417, 65–74 (2018).

    Article  CAS  Google Scholar 

  298. Hill, S. J. et al. Prediction of DNA repair inhibitor response in short-term patient-derived ovarian cancer organoids. Cancer Discov. 8, 1404–1421 (2018). This work provides evidence that DNA replication fork dynamics could be used to predict therapy sensitivity in patient-derived tumour samples, building on previous findings that connected replication fork degradation phenotypes with drug response in cell-based models.

    Article  CAS  Google Scholar 

  299. Maréchal, A. & Zou, L. RPA-coated single-stranded DNA as a platform for post-translational modifications in the DNA damage response. Cell Res. 25, 9–23 (2015).

    Article  Google Scholar 

  300. Michelena, J. et al. Analysis of PARP inhibitor toxicity by multidimensional fluorescence microscopy reveals mechanisms of sensitivity and resistance. Nat. Commun. 9, 2678 (2018).

    Article  Google Scholar 

  301. Hussmann, J. A. et al. Mapping the genetic landscape of DNA double-strand break repair. Cell 184, 5653–5669.e25 (2021).

    Article  CAS  Google Scholar 

  302. Olivieri, M. et al. A genetic map of the response to DNA damage in human cells. Cell 182, 481–496.e21 (2020). This publication uses a powerful genome-wide screening to demonstrate the breadth of DNA damage response pathways essential for coping with a variety of replication stress inducers and DNA-damaging agents.

    Article  CAS  Google Scholar 

  303. Chen, M., Linstra, R. & van Vugt, M. A. T. M. Genomic instability, inflammatory signaling and response to cancer immunotherapy. Biochim. Biophys. Acta Rev. Cancer 1877, 188661 (2022).

    Article  CAS  Google Scholar 

  304. Chabanon, R. M. et al. Targeting the DNA damage response in immuno-oncology: developments and opportunities. Nat. Rev. Cancer 21, 701–717 (2021).

    Article  CAS  Google Scholar 

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Acknowledgements

The authors would like to thank L. Zou, P. Verma and J. Eissenberg for their careful reading of this manuscript and for their insightful feedback, and A. Meroni for comments on the figures. This work was supported by the National Cancer Institute (NCI) grants F30CA254215 to E.C. and R01CA237263 and R01CA248526 to A.V.; the US Department of Defense (DOD) Breast Cancer Research Program (BRCP) Expansion Award BC191374 to A.V.; the Alvin J. Siteman Cancer Center Siteman Investment Program (supported by The Foundation for Barnes-Jewish Hospital, Cancer Frontier Fund) to A.V.; and the Barnard Foundation to A.V.

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Authors and Affiliations

  1. Division of Oncology, Department of Medicine, Washington University in St. Louis, St. Louis, MO, USA

    Emily Cybulla & Alessandro Vindigni

  2. Edward A. Doisy Department of Biochemistry and Molecular Biology, Saint Louis University School of Medicine, St. Louis, MO, USA

    Emily Cybulla

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Glossary

DNA lesions

Modifications introduced on the DNA helix by different genotoxic agents.

Xeroderma pigmentosum

An autosomal recessive genetic disease caused by biallelic mutations of specific proteins that are involved in molecular mechanisms required to cope with UV-induced DNA lesions, including Polη.

Polar pausing

Transient pausing of the replication fork in response to a unidirectional barrier that only inhibits replication fork progression in one direction.

Schimke immuno-osseous dysplasia

A multi-system autosomal recessive genetic disease caused by inheritance of biallelic SMARCAL1 mutations, with renal disease being a major cause of mortality in patients with this disease.

RECON syndrome

An autosomal recessive genetic disease caused by biallelic mutations in the RECQL1 DNA helicase, which functions in the DNA damage response.

Microhomology-mediated end joining

(MMEJ). One of the DNA double-strand break repair pathways, along with homologous recombination and non-homologous end joining, which relies on microhomology sequences (1–16 nucleotides) to anneal and align double-strand break ends for repair.

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Cybulla, E., Vindigni, A. Leveraging the replication stress response to optimize cancer therapy. Nat Rev Cancer 23, 6–24 (2023). https://doi.org/10.1038/s41568-022-00518-6

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