Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Review Article
  • Published:

The plasticity of DNA replication forks in response to clinically relevant genotoxic stress

Abstract

Complete and accurate DNA replication requires the progression of replication forks through DNA damage, actively transcribed regions, structured DNA and compact chromatin. Recent studies have revealed a remarkable plasticity of the replication process in dealing with these obstacles, which includes modulation of replication origin firing, of the architecture of replication forks, and of the functional organization of the replication machinery in response to replication stress. However, these specialized mechanisms also expose cells to potentially dangerous transactions while replicating DNA. In this Review, we discuss how replication forks are actively stalled, remodelled, processed, protected and restarted in response to specific types of stress. We also discuss adaptations of the replication machinery and the role of chromatin modifications during these transactions. Finally, we discuss interesting recent data on the relevance of replication fork plasticity to human health, covering its role in tumorigenesis, its crosstalk with innate immunity responses and its potential as an effective cancer therapy target.

This is a preview of subscription content, access via your institution

Access options

Buy this article

Prices may be subject to local taxes which are calculated during checkout

Fig. 1: Overview of the mechanisms underlying replication fork plasticity.
Fig. 2: Replication fork remodelling and restart during replication stress.
Fig. 3: Nucleolytic processing and restart of stalled replication forks.
Fig. 4: Replisome plasticity under replication stress.
Fig. 5: Crosstalk between replication fork protection and chromatin modification.
Fig. 6: DNA replication and repair can generate cytoplasmic DNA that activates an innate immunity response.

Similar content being viewed by others

References

  1. Saldivar, J. C., Cortez, D. & Cimprich, K. A. The essential kinase ATR: ensuring faithful duplication of a challenging genome. Nat. Rev. Mol. Cell Biol. 18, 622–636 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  2. Berti, M. & Vindigni, A. Replication stress: getting back on track. Nat. Struct. Mol. Biol. 23, 103–109 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Cortez, D. Replication-coupled DNA repair. Mol. Cell 74, 866–876 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. Rickman, K. & Smogorzewska, A. Advances in understanding DNA processing and protection at stalled replication forks. J. Cell Biol. 218, 1096–1107 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  5. 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  PubMed  PubMed Central  Google Scholar 

  6. 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  PubMed  Google Scholar 

  7. 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). This extensive electron microscopy analysis of replication intermediates shows that RAD51-dependent replication fork slowing and reversal is a general response to sublethal genotoxic treatments.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  8. Higgins, N. P., Kato, K. & Strauss, B. A model for replication repair in mammalian cells. J. Mol. Biol. 101, 417–425 (1976).

    Article  CAS  PubMed  Google Scholar 

  9. Atkinson, J. & McGlynn, P. Replication fork reversal and the maintenance of genome stability. Nucleic Acids Res. 37, 3475–3492 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  10. Bermejo, R. et al. The replication checkpoint protects fork stability by releasing transcribed genes from nuclear pores. Cell 146, 233–246 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  11. Sogo, J., Lopes, M. & Foiani, M. Fork reversal and ssDNA accumulation at stalled replication forks owing to checkpoint defects. Science 297, 599–602 (2002).

    Article  CAS  PubMed  Google Scholar 

  12. Ray Chaudhuri, A. et al. Topoisomerase I poisoning results in PARP-mediated replication fork reversal. Nat. Struct. Mol. Biol. 19, 417–423 (2012).

    Article  CAS  PubMed  Google Scholar 

  13. 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). A combination of single-molecule and biochemical approaches show a crucial role of the RECQ1 helicase in restarting reversed forks under the negative regulation of PARP1-mediated parylation.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Quinet, A., Lemacon, D. & Vindigni, A. Replication fork reversal: players and guardians. Mol. Cell 68, 830–833 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  15. Neelsen, K. J. et al. Deregulated origin licensing leads to chromosomal breaks by rereplication of a gapped DNA template. Genes Dev. 27, 2537–2542 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  16. Follonier, C., Oehler, J., Herrador, R. & Lopes, M. Friedreich’s ataxia-associated GAA repeats induce replication-fork reversal and unusual molecular junctions. Nat. Struct. Mol. Biol. 20, 486–494 (2013).

    Article  CAS  PubMed  Google Scholar 

  17. Schmid, J. A. et al. Histone ubiquitination by the DNA damage response is required for efficient DNA replication in unperturbed S phase. Mol. Cell 71, 897–910.e8 (2018). This study identifies canonical DDR factors that are essential mediators of efficient DNA replication in unperturbed S phase, through the modulation of resection and restart of chromatinized reversed forks.

    Article  CAS  PubMed  Google Scholar 

  18. Kile, A. C. et al. HLTF’s ancient HIRAN domain binds 3´ DNA ends to drive replication fork reversal. Mol. Cell 58, 1090–1100 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. 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  PubMed  PubMed Central  Google Scholar 

  20. 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. 20, 698–714 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  21. Bhat, K. P. & Cortez, D. RPA and RAD51: fork reversal, fork protection, and genome stability. Nat. Struct. Mol. Biol. 25, 446–453 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  22. 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  PubMed  PubMed Central  CAS  Google Scholar 

  23. Mason, J. M., Chan, Y.-L., Weichselbaum, R. W. & Bishop, D. K. Non-enzymatic roles of human RAD51 at stalled replication forks. Nat. Commun. 10, 4410–4411 (2019).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  24. Mijic, S. et al. Replication fork reversal triggers fork degradation in BRCA2-defective cells. Nat. Commun. 8, 859 (2017).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

  26. 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  PubMed  PubMed Central  Google Scholar 

  27. Bhat, K. P. et al. RADX modulates RAD51 activity to control replication fork protection. Cell Rep. 24, 538–545 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  28. Schubert, L. et al. RADX interacts with single-stranded DNA to promote replication fork stability. EMBO Rep. 18, 1991–2003 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  29. Chappidi, N. et al. Fork cleavage-religation cycle and active transcription mediate replication restart after fork stalling at Co-transcriptional R-loops. Mol. Cell 77, 528–541.e8 (2020). This report uncovers fork cleavage–re-ligation cycles as a mechanism of fork restart following drug-induced transcription–replication conflicts; these cycles coordinate transcription restart with the resumption of semiconservative DNA synthesis.

    Article  CAS  PubMed  Google Scholar 

  30. García-Rodríguez, N., Wong, R. P. & Ulrich, H. D. Functions of ubiquitin and SUMO in DNA replication and replication stress. Front. Genet. 7, 87 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  31. Ciccia, A. et al. Polyubiquitinated PCNA recruits the ZRANB3 translocase to maintain genomic integrity after replication stress. Mol. Cell 47, 396–409 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Weston, R., Peeters, H. & Ahel, D. ZRANB3 is a structure-specific ATP-dependent endonuclease involved in replication stress response. Genes Dev. 26, 1558–1572 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Betous, R. et al. Substrate-selective repair and restart of replication forks by DNA translocases. Cell Rep. 3, 1958–1969 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  35. 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  PubMed  PubMed Central  Google Scholar 

  36. 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  PubMed  PubMed Central  Google Scholar 

  37. Blastyák, 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  PubMed  CAS  Google Scholar 

  38. Bai, G. et al. HLTF promotes fork reversal, limiting replication stress resistance and preventing multiple mechanisms of unrestrained DNA synthesis. Mol. Cell https://doi.org/10.1016/j.molcel.2020.04.031 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  39. 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  PubMed  PubMed Central  Google Scholar 

  40. Couch, F. B. & Cortez, D. Fork reversal, too much of a good thing. Cell Cycle 13, 1049–1050 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. Quinet, A. et al. PRIMPOL-mediated adaptive response suppresses replication fork reversal in BRCA-deficient cells. Mol. Cell 77, 461–474.e9 (2020). This study reports that PrimPol-mediated fork repriming can compete with fork reversal in response to prolonged genotoxic treatments, thereby preventing fork degradation and providing a clinically relevant mechanistic understanding of chemoresistance in BRCA-defective tumours.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  42. Goodman, M. F. & Woodgate, R. Translesion DNA polymerases. Cold Spring Harb. Perspect. Biol. 5, a010363–a010363 (2013).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  43. Ait Saada, A., Lambert, S. A. E. & Carr, A. M. Preserving replication fork integrity and competence via the homologous recombination pathway. DNA Repair 71, 135–147 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  44. 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  PubMed  PubMed Central  Google Scholar 

  45. 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  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  PubMed  CAS  Google Scholar 

  49. Guilliam, T. A. et al. Molecular basis for PrimPol recruitment to replication forks by RPA. Nat. Commun. 8, 15222 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  50. Branzei, D. & Psakhye, I. DNA damage tolerance. Curr. Opin. Cell Biol. 40, 137–144 (2016).

    Article  CAS  PubMed  Google Scholar 

  51. Giannattasio, M. et al. Visualization of recombination-mediated damage bypass by template switching. Nat. Struct. Mol. Biol. 21, 884–892 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  52. 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.USA 112, E6624–E6633 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  53. 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). Using DNA fibre spreading, this report is the first to show a crucial role for BRCA2 and RAD51 in protecting stalled forks from nucleolytic degradation, uncoupled from their canonical DNA repair roles.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  54. Schlacher, K., Wu, H. & Jasin, M. A distinct replication fork protection pathway connects Fanconi anemia tumor suppressors to RAD51–BRCA1/2. Cancer Cell 22, 106–116 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  55. Reynolds, J. J. et al. Mutations in DONSON disrupt replication fork stability and cause microcephalic dwarfism. Nat. Genet. 49, 537–549 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  56. Przetocka, S. et al. CtIP-mediated fork protection synergizes with BRCA1 to suppress genomic instability upon DNA replication stress. Mol. Cell 72, 568–582.e6 (2018).

    Article  CAS  PubMed  Google Scholar 

  57. Leuzzi, G., Marabitti, V., Pichierri, P. & Franchitto, A. WRNIP1 protects stalled forks from degradation and promotes fork restart after replication stress. EMBO J. 35, 1437–1451 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  58. Porebski, B. et al. WRNIP1 protects reversed DNA replication forks from SLX4-dependent nucleolytic cleavage. iScience 21, 31–41 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  59. Garzón, J., Ursich, S., Lopes, M., Hiraga, S. & Donaldson, A. D. Human RIF1-protein phosphatase 1 prevents degradation and breakage of nascent DNA on replication stalling. Cell Rep. 27, 2558–2566.e4 (2019).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  60. Mukherjee, C. et al. RIF1 promotes replication fork protection and efficient restart to maintain genome stability. Nat. Commun. 10, 3287 (2019).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  61. 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  PubMed  PubMed Central  Google Scholar 

  62. Xu, S. et al. Abro1 maintains genome stability and limits replication stress by protecting replication fork stability. Genes Dev. 31, 1469–1482 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  63. Coquel, F. et al. SAMHD1 acts at stalled replication forks to prevent interferon induction. Nature 557, 57–61 (2018).

    Article  CAS  PubMed  Google Scholar 

  64. Rainey, M. D. et al. CDC7 kinase promotes MRE11 fork processing, modulating fork speed and chromosomal breakage. EMBO Rep. https://doi.org/10.15252/embr.201948920 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  65. 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  PubMed  PubMed Central  Google Scholar 

  66. Taglialatela, A. et al. Restoration of replication fork stability in BRCA1- and BRCA2-deficient cells by inactivation of SNF2-family fork remodelers. Mol. Cell 68, 414–430.e8 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  67. Hashimoto, Y., Chaudhuri, A. R., Lopes, M. & Costanzo, V. Rad51 protects nascent DNA from Mre11-dependent degradation and promotes continuous DNA synthesis. Nat. Struct. Mol. Biol. 17, 1305–1311 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  68. Dehé, P. et al. Regulation of Mus81–Eme1 Holliday junction resolvase in response to DNA damage. Nat. Struct. Mol. Biol. 20, 598–603 (2013).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  69. Hanada, K. et al. The structure-specific endonuclease Mus81–Eme1 promotes conversion of interstrand DNA crosslinks into double-strands breaks. EMBO J. 25, 4921–4932 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  70. Hanada, K. et al. The structure-specific endonuclease Mus81 contributes to replication restart by generating double-strand DNA breaks. Nat. Struct. Mol. Biol. 14, 1096–1104 (2007).

    Article  CAS  PubMed  Google Scholar 

  71. Ying, S. et al. MUS81 promotes common fragile site expression. Nat. Cell Biol. 15, 1001–1007 (2013).

    Article  CAS  PubMed  Google Scholar 

  72. Di Marco, S. et al. RECQ5 helicase cooperates with MUS81 endonuclease in processing stalled replication forks at common fragile sites during mitosis. Mol. Cell 66, 658–671.e8 (2017).

    Article  PubMed  CAS  Google Scholar 

  73. Minocherhomji, S. et al. Replication stress activates DNA repair synthesis in mitosis. Nature 528, 286–290 (2015).

    Article  CAS  PubMed  Google Scholar 

  74. Neelsen, K. J., Zanini, I. M. Y., Herrador, R. & Lopes, M. Oncogenes induce genotoxic stress by mitotic processing of unusual replication intermediates. J. Cell Biol. 200, 699–708 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  75. Pepe, A. & West, S. C. MUS81–EME2 promotes replication fork restart. Cell Rep. 7, 1048–1055 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  76. Fu, H. et al. The DNA repair endonuclease Mus81 facilitates fast DNA replication in the absence of exogenous damage. Nat. Commun. 6, 6714–6746 (2015).

    Article  CAS  Google Scholar 

  77. Lai, X. et al. MUS81 nuclease activity is essential for replication stress tolerance and chromosome segregation in BRCA2-deficient cells. Nat. Commun. 8, 15913–15983 (2017).

    Article  CAS  Google Scholar 

  78. Matos, D. A. et al. ATR protects the genome against R loops through a MUS81-triggered feedback loop. Mol. Cell 77, 514–527.e4 (2020).

    Article  CAS  PubMed  Google Scholar 

  79. Kramara, J., Osia, B. & Malkova, A. Break-induced replication: the where, the why, and the how. Trends Genet. 34, 518–531 (2018).

    Article  CAS  PubMed  Google Scholar 

  80. Lydeard, J. R., Jain, S., Yamaguchi, M. & Haber, J. E. Break-induced replication and telomerase-independent telomere maintenance require Pol32. Nature 448, 820–823 (2007).

    Article  CAS  PubMed  Google Scholar 

  81. Wilson, M. A. et al. Pif1 helicase and Polδ promote recombination-coupled DNA synthesis via bubble migration. Nature 502, 393–396 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  82. Costantino, L. et al. Break-induced replication repair of damaged forks induces genomic duplications in human cells. Science 343, 88–91 (2014).

    Article  CAS  PubMed  Google Scholar 

  83. Bhowmick, R., Minocherhomji, S. & Hickson, I. D. RAD52 facilitates mitotic DNA synthesis following replication stress. Mol. Cell 64, 1117–1126 (2016).

    Article  CAS  PubMed  Google Scholar 

  84. Dilley, R. L. et al. Break-induced telomere synthesis underlies alternative telomere maintenance. Nature 539, 54–58 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  85. Sotiriou, S. K. et al. Mammalian RAD52 functions in break-induced replication repair of collapsed DNA replication forks. Mol. Cell 64, 1127–1134 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  86. Piazza, A., Wright, W. D. & Heyer, W.-D. Multi-invasions are recombination byproducts that induce chromosomal rearrangements. Cell 170, 760–773.e15 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  87. Lee, J. A., Carvalho, C. M. B. & Lupski, J. R. A DNA replication mechanism for generating nonrecurrent rearrangements associated with genomic disorders. Cell 131, 1235–1247 (2007).

    Article  CAS  PubMed  Google Scholar 

  88. Zhang, C.-Z., Leibowitz, M. L. & Pellman, D. Chromothripsis and beyond: rapid genome evolution from complex chromosomal rearrangements. Genes Dev. 27, 2513–2530 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  89. Willis, N. A., Rass, E. & Scully, R. Deciphering the code of the cancer genome: mechanisms of chromosome rearrangement. Trends Cancer 1, 217–230 (2015).

    Article  PubMed  PubMed Central  Google Scholar 

  90. Amunugama, R. et al. Replication fork reversal during DNA interstrand crosslink repair requires CMG unloading. Cell Rep. 23, 3419–3428 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  91. Kang, S., Kang, M. S., Ryu, E. & Myung, K. Eukaryotic DNA replication: orchestrated action of multi-subunit protein complexes. Mutat. Res. 809, 58–69 (2018).

    Article  CAS  PubMed  Google Scholar 

  92. Dewar, J. M. & Walter, J. C. Mechanisms of DNA replication termination. Nat. Rev. Mol. Cell Biol. 18, 507–516 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  93. Berens, T. J. & Toczyski, D. P. Keeping it together in times of stress: checkpoint function at stalled replication forks. Mol. Cell 45, 585–586 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  94. Dungrawala, H. et al. The replication checkpoint prevents two types of fork collapse without regulating replisome stability. Mol. Cell 59, 998–1010 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  95. Petermann, E., Orta, M. L., Issaeva, N., Schultz, N. & Helleday, T. Hydroxyurea-stalled replication forks become progressively inactivated and require two different RAD51-mediated pathways for restart and repair. Mol. Cell 37, 492–502 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  96. Harley, M. E. et al. TRAIP promotes DNA damage response during genome replication and is mutated in primordial dwarfism. Nat. Genet. 48, 36–43 (2016).

    Article  CAS  PubMed  Google Scholar 

  97. Wu, R. A. et al. TRAIP is a master regulator of DNA interstrand crosslink repair. Nature 567, 267–272 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  98. Larsen, N. B. et al. Replication-coupled DNA-protein crosslink repair by SPRTN and the proteasome in Xenopus egg extracts. Mol. Cell 73, 574–588.e7 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  99. Deng, L. et al. Mitotic CDK promotes replisome disassembly, fork breakage, and complex DNA rearrangements. Mol. Cell 73, 915–929.e6 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  100. Trakselis, M. A., Seidman, M. M. & Brosh, R. M. Mechanistic insights into how CMG helicase facilitates replication past DNA roadblocks. DNA Repair 55, 76–82 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  101. Huang, J. et al. The DNA translocase FANCM/MHF promotes replication traverse of DNA interstrand crosslinks. Mol. Cell 52, 434–446 (2013). Combining DNA fibre spreading with direct detection of interstrand crosslinks (ICLs), this report surprisingly shows that replication forks efficiently traverse ICLs, which had long been assumed to block replication fork progression.

    Article  CAS  PubMed  Google Scholar 

  102. González-Acosta, D. et al. PrimPol primase mediates replication traverse of DNA interstrand crosslinks. Preprint at https://doi.org/10.1101/2020.05.19.104729 (2020).

  103. Huang, J. et al. Remodeling of interstrand crosslink proximal replisomes is dependent on ATR, FANCM, and FANCD2. Cell Rep. 27, 1794–1808.e5 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  104. Sparks, J. L. et al. The CMG helicase bypasses DNA–protein cross-links to facilitate their repair. Cell 176, 167–181.e21 (2019).

    Article  CAS  PubMed  Google Scholar 

  105. Georgescu, R. et al. Structure of eukaryotic CMG helicase at a replication fork and implications to replisome architecture and origin initiation. Proc. Natl Acad. Sci. USA 114, E697–E706 (2017). This study of the structure of CMG helicase in complex with a replication fork shows a bipartite, polar organization, in which the C-tier motor lies behind and threads ssDNA through the N-tier ring, thereby separating the DNA strands.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  106. Douglas, M. E., Ali, F. A., Costa, A. & Diffley, J. F. X. The mechanism of eukaryotic CMG helicase activation. Nature 555, 265–268 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  107. Aria, V. & Yeeles, J. T. P. Mechanism of bidirectional leading-strand synthesis establishment at eukaryotic DNA replication origins. Mol. Cell 73, 199–211.e10 (2019).

    Article  CAS  PubMed Central  Google Scholar 

  108. Sun, J. et al. The architecture of a eukaryotic replisome. Nat. Struct. Mol. Biol. 22, 976–982 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  109. Wasserman, M. R., Schauer, G. D., O’Donnell, M. E. & Liu, S. Replication fork activation is enabled by a single-stranded DNA gate in CMG helicase. Cell 178, 600–611.e16 (2019). This report shows that CMG actively translocating on ssDNA can transit to an inactive, dsDNA-binding conformation when uncoupled from polymerases, and that it can revert to ssDNA as a means to renucleate a functional replisome with the assistance of MCM10 and a yet-unidentified ssDNA gate in the CMG structure.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  110. Thu, Y. M. & Bielinsky, A.-K. MCM10: one tool for all — integrity, maintenance and damage control. Semin. Cell Dev. Biol. 30, 121–130 (2014).

    Article  CAS  PubMed  Google Scholar 

  111. Samel, S. A. et al. A unique DNA entry gate serves for regulated loading of the eukaryotic replicative helicase MCM2–7 onto DNA. Genes Dev. 28, 1653–1666 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  112. Frigola, J. et al. Cdt1 stabilizes an open MCM ring for helicase loading. Nat. Commun. 8, 15720 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  113. Costa, A. et al. The structural basis for MCM2–7 helicase activation by GINS and Cdc45. Nat. Struct. Mol. Biol. 18, 471–477 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  114. Goswami, P. et al. Structure of DNA–CMG–Pol epsilon elucidates the roles of the non-catalytic polymerase modules in the eukaryotic replisome. Nat. Commun. 9, 5061 (2018).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  115. Manosas, M., Perumal, S. K., Croquette, V. & Benkovic, S. J. Direct observation of stalled fork restart via fork regression in the T4 replication system. Science 338, 1217–1220 (2012).

    Article  CAS  PubMed  Google Scholar 

  116. Mayle, R. et al. Mcm10 has potent strand-annealing activity and limits translocase-mediated fork regression. Proc. Natl Acad. Sci.USA 116, 798–803 (2019).

    Article  CAS  PubMed  Google Scholar 

  117. Graham, J. E., Marians, K. J. & Kowalczykowski, S. C. Independent and stochastic action of DNA polymerases in the replisome. Cell 169, 1201–1213.e17 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  118. Yeeles, J. T. P., Janska, A., Early, A. & Diffley, J. F. X. How the eukaryotic replisome achieves rapid and efficient DNA replication. Mol. Cell 65, 105–116 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  119. Hashimoto, Y., Puddu, F. & Costanzo, V. RAD51- and MRE11-dependent reassembly of uncoupled CMG helicase complex at collapsed replication forks. Nat. Struct. Mol. Biol. 19, 17–24 (2012).

    Article  CAS  Google Scholar 

  120. Petojevic, T. et al. Cdc45 (cell division cycle protein 45) guards the gate of the eukaryote replisome helicase stabilizing leading strand engagement. Proc. Natl Acad. Sci. USA 112, E249–E258 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  121. Gambus, A. et al. GINS maintains association of Cdc45 with MCM in replisome progression complexes at eukaryotic DNA replication forks. Nat. Cell Biol. 8, 358–366 (2006).

    Article  CAS  PubMed  Google Scholar 

  122. Yu, C. et al. Strand-specific analysis shows protein binding at replication forks and PCNA unloading from lagging strands when forks stall. Mol. Cell 56, 551–563 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  123. Baretic, D. et al. Cryo-EM structure of the fork protection complex bound to CMG at a replication fork. Mol. Cell 78, 926–940 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  124. Somyajit, K. et al. Redox-sensitive alteration of replisome architecture safeguards genome integrity. Science 358, 797–802 (2017).

    Article  CAS  PubMed  Google Scholar 

  125. Lukas, J., Lukas, C. & Bartek, J. More than just a focus: the chromatin response to DNA damage and its role in genome integrity maintenance. Nat. Cell Biol. 13, 1161–1169 (2011).

    Article  CAS  PubMed  Google Scholar 

  126. Alabert, C. & Groth, A. Chromatin replication and epigenome maintenance. Nat. Rev. Mol. Cell Biol. 13, 153–167 (2012).

    Article  CAS  PubMed  Google Scholar 

  127. Hammond, C. M., Strømme, C. B., Huang, H., Patel, D. J. & Groth, A. Histone chaperone networks shaping chromatin function. Nat. Rev. Mol. Cell Biol. 18, 141–158 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  128. Saredi, G. et al. H4K20me0 marks post-replicative chromatin and recruits the TONSL–MMS22L DNA repair complex. Nature 534, 714–718 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  129. Pellegrino, S., Michelena, J., Teloni, F., Imhof, R. & Altmeyer, M. Replication-coupled dilution of H4K20me2 guides 53BP1 to pre-replicative chromatin. Cell Rep. 19, 1819–1831 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  130. Nakamura, K. et al. H4K20me0 recognition by BRCA1–BARD1 directs homologous recombination to sister chromatids. Nat. Cell Biol. 21, 311–318 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  131. Xu, Y. et al. 53BP1 and BRCA1 control pathway choice for stalled replication restart. eLife 6, e30523 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  132. Chen, B.-R. et al. XLF and H2AX function in series to promote replication fork stability. J. Cell Biol. 218, 2113–2123 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  133. Her, J., Ray, C., Altshuler, J., Zheng, H. & Bunting, S. F. 53BP1 mediates ATR–Chk1 signaling and protects replication forks under conditions of replication stress. Mol. Cell. Biol. 38, e00472-17 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  134. Higgs, M. R. et al. Histone methylation by SETD1A protects nascent DNA through the nucleosome chaperone activity of FANCD2. Mol. Cell 71, 25–41.e6 (2018). This study reports that H3K4 methylation by SETD1A–BOD1L is crucial to protecting stalled replication forks from nucleolytic degradation, by stimulating FANCD2 histone chaperone activity and promoting the stability of RAD51 filaments.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  135. Ray Chaudhuri, A. et al. Replication fork stability confers chemoresistance in BRCA-deficient cells. Nature 535, 382–387 (2016). This report shows that the loss of MLL3–MLL4–PTIP-mediated H3K4 methylation prevents MRE11 recruitment to stalled forks and restores fork stability in BRCA1- or BRCA2-deficient cells, thereby promoting chemoresistance despite the persistence of a DSB repair defect.

    Article  PubMed  CAS  Google Scholar 

  136. 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  PubMed  PubMed Central  Google Scholar 

  137. Alsulami, M. et al. SETD1A methyltransferase is physically and functionally linked to the DNA damage repair protein RAD18. Mol. Cell. Proteom. 18, 1428–1436 (2019).

    Article  CAS  Google Scholar 

  138. 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  PubMed  Google Scholar 

  139. Jasencakova, Z. et al. Replication stress interferes with histone recycling and predeposition marking of new histones. Mol. Cell 37, 736–743 (2010).

    Article  CAS  PubMed  Google Scholar 

  140. Feng, G. et al. Replication fork stalling elicits chromatin compaction for the stability of stalling replication forks. Proc. Natl Acad. Sci. USA 116, 14563–14572 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  141. Reverón-Gómez, N. et al. Accurate recycling of parental histones reproduces the histone modification landscape during DNA replication. Mol. Cell 72, 239–249.e5 (2018). This report of the development of ChOR-seq, to monitor chromatin modifications before and after replication, shows that histone modifications are re-established across the cell cycle, with modification- and locus-specific kinetics.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  142. Talbert, P. B. & Henikoff, S. Histone variants on the move: substrates for chromatin dynamics. Nat. Rev. Mol. Cell Biol. 18, 115–126 (2017).

    Article  CAS  PubMed  Google Scholar 

  143. Piquet, S. et al. The histone chaperone FACT coordinates H2A.X-dependent signaling and repair of DNA damage. Mol. Cell 72, 888–901.e7 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  144. Kim, J. et al. Replication stress shapes a protective chromatin environment across fragile genomic regions. Mol. Cell 69, 36–47.e7 (2018). In cells experiencing replication stress, FACT-dependent deposition of macroH2A1.2 at common fragile sites promotes BRCA1 recruitment and fork protection and prevents genome instability.

    Article  CAS  PubMed  Google Scholar 

  145. Rogakou, E. P., Boon, C., Redon, C. & Bonner, W. M. Megabase chromatin domains involved in DNA double-strand breaks in vivo. J. Cell Biol. 146, 905–916 (1999).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  146. Courbet, S. et al. Replication fork movement sets chromatin loop size and origin choice in mammalian cells. Nature 455, 557–560 (2008).

    Article  CAS  PubMed  Google Scholar 

  147. Pope, B. D. et al. Topologically associating domains are stable units of replication-timing regulation. Nature 515, 402–405 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  148. Yamazaki, S., Hayano, M. & Masai, H. Replication timing regulation of eukaryotic replicons: Rif1 as a global regulator of replication timing. Trends Genet. 29, 449–460 (2013).

    Article  CAS  PubMed  Google Scholar 

  149. Sarkies, P., Reams, C., Simpson, L. J. & Sale, J. E. Epigenetic instability due to defective replication of structured DNA. Mol. Cell 40, 703–713 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  150. Taneja, N. et al. SNF2 family protein Fft3 suppresses nucleosome turnover to promote epigenetic inheritance and proper replication. Mol. Cell 66, 50–62.e6 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  151. Jasencakova, Z. & Groth, A. Replication stress, a source of epigenetic aberrations in cancer? BioEssays 32, 847–855 (2010).

    Article  CAS  PubMed  Google Scholar 

  152. Wang, A. T. et al. A dominant mutation in human RAD51 reveals its function in DNA interstrand crosslink repair independent of homologous recombination. Mol. Cell 59, 478–490 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  153. Rickman, K. A. et al. Distinct roles of BRCA2 in replication fork protection in response to hydroxyurea and DNA interstrand crosslinks. Genes Dev. 34, 832–846 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  154. Billing, D. et al. The BRCT domains of the BRCA1 and BARD1 tumor suppressors differentially regulate homology-directed repair and stalled fork protection. Mol. Cell 72, 127–139.e8 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  155. Feng, W. & Jasin, M. BRCA2 suppresses replication stress-induced mitotic and G1 abnormalities through homologous recombination. Nat. Commun. 8, 525 (2017).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  156. Pathania, S. et al. BRCA1 haploinsufficiency for replication stress suppression in primary cells. Nat. Commun. 5, 5496 (2014).

    Article  PubMed  Google Scholar 

  157. Venkitaraman, A. R. How do mutations affecting the breast cancer genes BRCA1 and BRCA2 cause cancer susceptibility? DNA Repair 81, 102668 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  158. Tan, S. L. W. et al. A class of environmental and endogenous toxins induces BRCA2 haploinsufficiency and genome instability. Cell 169, 1105–1118.e15 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  159. 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  PubMed  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

  161. Forment, J. V. & O’Connor, M. J. Targeting the replication stress response in cancer. Pharmacol. Ther. 188, 155–167 (2018).

    Article  CAS  PubMed  Google Scholar 

  162. 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).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  163. Ding, X. et al. Synthetic viability by BRCA2 and PARP1/ARTD1 deficiencies. Nat. Commun. 7, 12425 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  164. Daza-Martin, M. et al. Isomerization of BRCA1–BARD1 promotes replication fork protection. Nature 571, 521–527 (2019). This study reports that the BRCA1–BARD1 complex has a genetically HR-independent role in fork protection that is regulated by prolyl isomerization and that contributes to the complex’s tumour suppressor function.

    Article  CAS  PubMed  Google Scholar 

  165. Sakai, W. et al. Secondary mutations as a mechanism of cisplatin resistance in BRCA2-mutated cancers. Nature 451, 1116–1120 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  166. Swisher, E. M. et al. Secondary BRCA1 mutations in BRCA1-mutated ovarian carcinomas with platinum resistance. Cancer Res. 68, 2581–2586 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  167. Edwards, S. L. et al. Resistance to therapy caused by intragenic deletion in BRCA2. Nature 451, 1111–1115 (2008).

    Article  CAS  PubMed  Google Scholar 

  168. Patch, A.-M. et al. Whole–genome characterization of chemoresistant ovarian cancer. Nature 521, 489–494 (2015).

    Article  CAS  PubMed  Google Scholar 

  169. Christie, E. L. et al. Reversion of BRCA1/2 germline mutations detected in circulating tumor DNA from patients with high-grade serous ovarian cancer. J. Clin. Oncol. 35, 1274–1280 (2017).

    Article  CAS  PubMed  Google Scholar 

  170. Goodall, J. et al. Circulating cell-free DNA to guide prostate cancer treatment with PARP inhibition. Cancer Discov. 7, 1006–1017 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  171. Quigley, D. et al. Analysis of circulating cell-free DNA identifies multiclonal heterogeneity of BRCA2 Reversion mutations associated with resistance to PARP inhibitors. Cancer Discov. 7, 999–1005 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  172. Pilié, 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. 16, 81–104 (2019).

    Article  PubMed  CAS  PubMed Central  Google Scholar 

  173. Mohni, K. N. et al. A synthetic lethal screen identifies DNA repair pathways that sensitize cancer cells to combined ATR inhibition and cisplatin treatments. PLoS ONE 10, e0125482 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  174. Xu, G. et al. REV7 counteracts DNA double-strand break resection and affects PARP inhibition. Nature 521, 541–544 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  175. Tkáč, J. et al. HELB is a feedback inhibitor of DNA end resection. Mol. Cell 61, 405–418 (2016).

    Article  PubMed  CAS  Google Scholar 

  176. 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  PubMed  PubMed Central  Google Scholar 

  177. Brown, J. S., Sundar, R. & Lopez, J. Combining DNA damaging therapeutics with immunotherapy: more haste, less speed. Br. J. Cancer 118, 312–324 (2018).

    Article  CAS  PubMed  Google Scholar 

  178. Rizvi, N. A. et al. Mutational landscape determines sensitivity to PD-1 blockade in non–small cell lung cancer. Science 348, 124–128 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  179. Le, D. T. et al. PD-1 blockade in tumors with mismatch-repair deficiency. N. Engl. J. Med. 372, 2509–2520 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  180. Hugo, W. et al. Genomic and transcriptomic features of response to anti-PD-1 therapy in metastatic melanoma. Cell 165, 35–44 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  181. Nolan, E. et al. Combined immune checkpoint blockade as a therapeutic strategy for BRCA1-mutated breast cancer. Sci. Transl. Med. 9, eaal4922 (2017).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  182. Pfirschke, C. et al. Immunogenic chemotherapy sensitizes tumors to checkpoint blockade therapy. Immunity 44, 343–354 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  183. Gandhi, L. et al. Pembrolizumab plus chemotherapy in metastatic non-small-cell lung cancer. N. Engl. J. Med. 378, 2078–2092 (2018).

    Article  CAS  PubMed  Google Scholar 

  184. Motwani, M., Pesiridis, S. & Fitzgerald, K. A. DNA sensing by the cGAS–STING pathway in health and disease. Nat. Rev. Genet. 20, 657–674 (2019).

    Article  CAS  PubMed  Google Scholar 

  185. Li, T. & Chen, Z. J. The cGAS–cGAMP–STING pathway connects DNA damage to inflammation, senescence, and cancer. J. Exp. Med. 215, 1287–1299 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  186. Chabanon, R. M. et al. PARP inhibition enhances tumor cell-intrinsic immunity in ERCC1-deficient non-small cell lung cancer. J. Clin. Invest. 129, 1211–1228 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  187. Heijink, A. M. et al. BRCA2 deficiency instigates cGAS-mediated inflammatory signaling and confers sensitivity to tumor necrosis factor-alpha-mediated cytotoxicity. Nat. Commun. 10, 100 (2019).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  188. Reisländer, T. et al. BRCA2 abrogation triggers innate immune responses potentiated by treatment with PARP inhibitors. Nat. Commun. 10, 3143 (2019).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  189. Shen, J. et al. PARPi triggers the STING-dependent immune response and enhances the therapeutic efficacy of immune checkpoint blockade independent of brcaness. Cancer Res. 79, 311–319 (2019).

    Article  CAS  PubMed  Google Scholar 

  190. Mackenzie, K. J. et al. Ribonuclease H2 mutations induce a cGAS/STING-dependent innate immune response. EMBO J. 35, 831–844 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  191. Gratia, M. et al. Bloom syndrome protein restrains innate immune sensing of micronuclei by cGAS. J. Exp. Med. 216, 1199–1213 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  192. Bartsch, K. et al. Absence of RNase H2 triggers generation of immunogenic micronuclei removed by autophagy. Hum. Mol. Genet. 26, 3960–3972 (2017).

    Article  CAS  PubMed  Google Scholar 

  193. Martin, S. K., Tomida, J. & Wood, R. D. Deficiency in translesion DNA polymerase ζ induces an innate immune response. Preprint at https://doi.org/10.1101/2020.03.02.972513 (2020).

  194. Yang, Y.-G., Lindahl, T. & Barnes, D. E. Trex1 exonuclease degrades ssDNA to prevent chronic checkpoint activation and autoimmune disease. Cell 131, 873–886 (2007).

    Article  CAS  PubMed  Google Scholar 

  195. Wolf, C. et al. RPA and Rad51 constitute a cell intrinsic mechanism to protect the cytosol from self DNA. Nat. Commun. 7, 11752 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  196. Bhattacharya, S. et al. RAD51 interconnects between DNA replication, DNA repair and immunity. Nucleic Acids Res. 45, 4590–4605 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  197. Pantelidou, C. et al. PARP inhibitor efficacy depends on CD8+ T-cell recruitment via intratumoral STING pathway activation in BRCA-deficient models of triple-negative breast cancer. Cancer Discov. 9, 722–737 (2019). This study suggests that the efficacy of PARP inhibitors in BRCA-deficient triple-negative breast cancers partly depends on the activation of the cGAS–STING signalling pathway.

    Article  PubMed  PubMed Central  Google Scholar 

  198. Raso, M. C. et al. High levels of ISG15 promotes high DNA replication speed and chromosomal breakage via deregulated fork restart. J. Cell Biol. https://doi.org/10.1083/jcb.202002175 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  199. Dehé, P.-M. & Gaillard, P.-H. L. Control of structure-specific endonucleases to maintain genome stability. Nat. Rev. Mol. Cell Biol. 18, 315–330 (2017).

    Article  PubMed  CAS  Google Scholar 

  200. O’Driscoll, M., Ruiz-Perez, V. L., Woods, C. G., Jeggo, P. A. & Goodship, J. A. A splicing mutation affecting expression of ataxia-telangiectasia and Rad3-related protein (ATR) results in Seckel syndrome. Nat. Genet. 33, 497–501 (2003).

    Article  PubMed  CAS  Google Scholar 

  201. Ellis, N. A. et al. The Bloom’s syndrome gene product is homologous to RecQ helicases. Cell 83, 655–666 (1995).

    Article  CAS  PubMed  Google Scholar 

  202. Taylor, A. M. R. et al. Chromosome instability syndromes. Nat. Rev. Dis. Prim. 5, 64 (2019).

    Article  PubMed  Google Scholar 

  203. Bicknell, L. S. et al. Mutations in the pre-replication complex cause Meier–Gorlin syndrome. Nat. Genet. 43, 356–359 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  204. Bicknell, L. S. et al. Mutations in ORC1, encoding the largest subunit of the origin recognition complex, cause microcephalic primordial dwarfism resembling Meier–Gorlin syndrome. Nat. Genet. 43, 350–355 (2011).

    Article  CAS  PubMed  Google Scholar 

  205. Guernsey, D. L. et al. Mutations in origin recognition complex gene ORC4 cause Meier–Gorlin syndrome. Nat. Genet. 43, 360–364 (2011).

    Article  CAS  PubMed  Google Scholar 

  206. Logan, C. V. et al. DNA polymerase epsilon deficiency causes IMAGe syndrome with variable immunodeficiency. Am. J. Hum. Genet. 103, 1038–1044 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  207. Kitao, S. et al. Mutations in RECQL4 cause a subset of cases of Rothmund–Thomson syndrome. Nat. Genet. 22, 82–84 (1999).

    Article  CAS  PubMed  Google Scholar 

  208. Boerkoel, C. F. et al. Mutant chromatin remodelling protein SMARCAL1 causes Schimke immuno-osseous dysplasia. Nat. Genet. 30, 215–220 (2002).

    Article  CAS  PubMed  Google Scholar 

  209. Yu, C.-E. et al. Positional cloning of the Werner’s syndrome gene. Science 272, 258–262 (1996).

    Article  CAS  PubMed  Google Scholar 

Download references

Acknowledgements

The authors apologize to all those researchers whose relevant work could not be cited because of space limitations. This work was supported by a Marie Skłodowska-Curie Fellowship (704817) to M.B., by a US National Institutes of Health grant (R01GM116616) to D.C., and by grants from the Swiss National Science Foundation (31003A_169959 and 310030_189206), Swiss Cancer League (KFS-3967-08-2016) and European Research Council (Consolidator Grant 617102) to M.L.

Author information

Authors and Affiliations

Authors

Contributions

All authors contributed to reviewing the literature, discussing the manuscript content, writing the article and editing it before publication.

Corresponding author

Correspondence to Massimo Lopes.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Peer review information

Nature Reviews Molecular Cell Biology thanks Jiri Bartek and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Glossary

Repriming

Synthesis of short RNA sequences required for the restart of DNA synthesis after transient DNA polymerase blocking.

Replisome

A large protein complex that carries out DNA replication through its multiple enzymatic activities and that includes helicase, primase and DNA polymerases.

Fork uncoupling

Here, mainly used to describe uncoupling between helicase translocation and interrupted leading-strand polymerization.

Fork reversal

Conversion of standard three-way replication forks into four-way junctions, via re-annealing of the unwound parental duplex, followed by backtracking of the replication fork and annealing of the newly synthesized strands.

Template switching

DNA-damage tolerance pathway based on the use of a newly synthesized strand as an alternative template for DNA synthesis when the parental DNA is damaged.

Branch migration

Consecutive exchange of base pairings on homologous DNA strands at reversed forks or Holliday junctions, which moves the branch point up or down the DNA sequence.

Primase activity

Synthesis of short RNA sequences that are required for priming the activity of DNA polymerases.

MCM2–5 gate

An interface between the eponymous subunits of the MCM helicase. Opening of the gate is required in order to transition the helicase from dsDNA binding at replication origins to encircling the ssDNA template for leading-strand synthesis.

Lock–washer conformation

A transiently open conformation of the MCM helicase that allows the transition from dsDNA binding to ssDNA binding during the activation of DNA synthesis at replication origins.

Topologically associated domains

A 3D genome organization feature identified by chromosome conformation capture techniques, referring to genomic regions in which DNA sequences preferentially interact with each other rather than with other genomic sequences.

Microsatellite instability-high cancers

Tumours (typically arising in the colon) that display genetic hypermutability, usually owing to impaired DNA mismatch repair.

Anti-PD-1 therapy

Cancer treatment using drugs that block the interaction of specific cancer ligands with the immune checkpoint receptor PD-1 present on the surface of T cells, thereby preventing cancer cell evasion from the immune system.

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Berti, M., Cortez, D. & Lopes, M. The plasticity of DNA replication forks in response to clinically relevant genotoxic stress. Nat Rev Mol Cell Biol 21, 633–651 (2020). https://doi.org/10.1038/s41580-020-0257-5

Download citation

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41580-020-0257-5

This article is cited by

Search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing