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:

Control of structure-specific endonucleases to maintain genome stability

Key Points

  • Structure-specific endonucleases (SSEs) process various types of DNA secondary structures that arise during DNA replication, repair, recombination and transcription, and are important for the maintenance of genome stability.

  • Elaborate regulatory mechanisms ensure that SSEs act in an efficient, specific and timely manner so that they do not themselves become a source of genome instability.

  • The control of SSEs relies on a combination of catalytic regulation; modulation of their cellular localization, which regulates their targeting to the appropriate substrate or instead ensures their sequestration to reduce the risk of uncontrolled DNA processing; and protein turnover.

  • SSEs can function genome-wide, such as during the repair of DNA adducts by nucleotide excision repair, or their function can be specific to genomic loci that are prone to the formation of DNA secondary structures that need to be processed during replication and/or transcription. Such loci include, for example, the ribosomal DNA or telomeres, which contain DNA repeats, or regions that replicate late in S phase and therefore might not be replicated on time before the onset of mitosis, such as common fragile sites.

  • The control of SSEs is critical during DNA replication, during which inappropriate SSE activity, especially of MUS81 nucleases, can have dire consequences for genome stability.

  • Nuclease scaffolds are particularly important for the regulation of SSEs, by helping in their efficient recruitment to DNA and coordination with other factors. Nuclease scaffolds can directly modulate the catalytic activity of SSEs and change their substrate specificity. Some scaffolds are dedicated to the control of a single SSE, whereas others control several SSEs, either independently or by coordinating their activity within the same pathway.

Abstract

Structure-specific endonucleases (SSEs) have key roles in DNA replication, recombination and repair, and emerging roles in transcription. These enzymes have specificity for DNA secondary structure rather than for sequence, and therefore their activity must be precisely controlled to ensure genome stability. In this Review, we discuss how SSEs are controlled as part of genome maintenance pathways in eukaryotes, with an emphasis on the elaborate mechanisms that regulate the members of the major SSE families — including the xeroderma pigmentosum group F-complementing protein (XPF) and MMS and UV-sensitive protein 81 (MUS81)-dependent nucleases, and the flap endonuclease 1 (FEN1), XPG and XPG-like endonuclease 1 (GEN1) enzymes — during processes such as DNA adduct repair, Holliday junction processing and replication stress. We also discuss newly characterized connections between SSEs and other classes of DNA-remodelling enzymes and cell cycle control machineries, which reveal the importance of SSE scaffolds such as the synthetic lethal of unknown function 4 (SLX4) tumour suppressor for the maintenance of genome stability.

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

Figure 1: DNA secondary structures processed by structure-specific endonucleases.
Figure 2: Control of structure-specific endonucleases during nucleotide excision repair.
Figure 3: Controlling the processing of Holliday junctions by structure-specific endonucleases.
Figure 4: Function of FEN1 in Okazaki fragment maturation.
Figure 5: Function of structure-specific endonucleases during replication stress.
Figure 6: Scaffold protein control of structure-specific endonucleases.

Similar content being viewed by others

References

  1. Lyamichev, V., Brow, M. A. & Dahlberg, J. E. Structure-specific endonucleolytic cleavage of nucleic acids by eubacterial DNA polymerases. Science 260, 778–783 (1993).

    Article  CAS  PubMed  Google Scholar 

  2. Robins, P., Pappin, D. J., Wood, R. D. & Lindahl, T. Structural and functional homology between mammalian DNase IV and the 5′-nuclease domain of Escherichia coli DNA polymerase I. J. Biol. Chem. 269, 28535–28538 (1994).

    CAS  PubMed  Google Scholar 

  3. Harrington, J. J. & Lieber, M. R. The characterization of a mammalian DNA structure-specific endonuclease. EMBO J. 13, 1235–1246 (1994).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. Harrington, J. J. & Lieber, M. R. Functional domains within FEN-1 and RAD2 define a family of structure-specific endonucleases: implications for nucleotide excision repair. Genes Dev. 8, 1344–1355 (1994).

    Article  CAS  PubMed  Google Scholar 

  5. O'Donovan, A., Davies, A. A., Moggs, J. G., West, S. C. & Wood, R. D. XPG endonuclease makes the 3′ incision in human DNA nucleotide excision repair. Nature 371, 432–435 (1994).

    Article  CAS  PubMed  Google Scholar 

  6. Davies, A. A., Friedberg, E. C., Tomkinson, A. E., Wood, R. D. & West, S. C. Role of the Rad1 and Rad10 proteins in nucleotide excision repair and recombination. J. Biol. Chem. 270, 24638–24641 (1995).

    Article  CAS  PubMed  Google Scholar 

  7. Sijbers, A. M. et al. Xeroderma pigmentosum group F caused by a defect in a structure-specific DNA repair endonuclease. Nucleic Acids Res. 86, 811–822 (1996).

    CAS  Google Scholar 

  8. Brookman, K. W. et al. ERCC4 (XPF) encodes a human nucleotide excision repair protein with eukaryotic recombination homologs. Mol. Cell. Biol. 16, 6553–6562 (1996).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  9. Mizuuchi, K., Kemper, B., Hays, J. & Weisberg, R. A. T4 endonuclease VII cleaves holliday structures. Cell 29, 357–365 (1982).

    Article  CAS  PubMed  Google Scholar 

  10. Dunderdale, H. J. et al. Formation and resolution of recombination intermediates by E. coli RecA and RuvC proteins. Nature 354, 506–510 (1991).

    Article  CAS  PubMed  Google Scholar 

  11. Iwasaki, H., Takahagi, M., Shiba, T., Nakata, A. & Shinagawa, H. Escherichia coli RuvC protein is an endonuclease that resolves the Holliday structure. EMBO J. 10, 4381–4389 (1991).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. Boddy, M. N. et al. Mus81–Eme1 are essential components of a Holliday junction resolvase. Cell 107, 537–548 (2001).

    Article  CAS  PubMed  Google Scholar 

  13. Chen, X. B. et al. Human Mus81-associated endonuclease cleaves Holliday junctions in vitro. Mol. Cell 8, 1117–1127 (2001).

    Article  CAS  PubMed  Google Scholar 

  14. Ip, S. C. Y. et al. Identification of Holliday junction resolvases from humans and yeast. Nature 456, 357–361 (2008).

    Article  CAS  PubMed  Google Scholar 

  15. Fekairi, S. et al. Human SLX4 is a Holliday junction resolvase subunit that binds multiple DNA repair/recombination endonucleases. Cell 138, 78–89 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  16. Munoz, I. M. et al. Coordination of structure-specific nucleases by human SLX4/BTBD12 is required for DNA repair. Mol. Cell 35, 116–127 (2009).

    Article  CAS  PubMed  Google Scholar 

  17. Svendsen, J. M. et al. Mammalian BTBD12/SLX4 assembles a Holliday junction resolvase and is required for DNA repair. Cell 138, 63–77 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  18. Andersen, S. L. et al. Drosophila MUS312 and the vertebrate ortholog BTBD12 interact with DNA structure-specific endonucleases in DNA repair and recombination. Mol. Cell 35, 128–135 (2009). References 15–18 describe the identification of human SLX4 and how it binds multiple proteins and/or complexes that are involved in genome maintenance, which include XPF–ERCC1, SLX1 and MUS81–EME1.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Rass, U. et al. Mechanism of Holliday junction resolution by the human GEN1 protein. Genes Dev. 24, 1559–1569 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Manandhar, M., Boulware, K. S. & Wood, R. D. The ERCC1 and ERCC4 (XPF) genes and gene products. Gene 569, 153–161 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  21. Tsutakawa, S. E., Lafrance-Vanasse, J. & Tainer, J. A. The cutting edges in DNA repair, licensing, and fidelity: DNA and RNA repair nucleases sculpt DNA to measure twice, cut once. DNA Repair (Amsterdam) 19, 95–107 (2014).

    Article  CAS  Google Scholar 

  22. Finger, L. D. et al. The wonders of flap endonucleases: structure, function, mechanism and regulation. Subcell. Biochem. 62, 301–326 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  23. Marteijn, J. A., Lans, H., Vermeulen, W. & Hoeijmakers, J. H. J. Understanding nucleotide excision repair and its roles in cancer and ageing. Nat. Rev. Mol. Cell Biol. 15, 465–481 (2014).

    Article  CAS  PubMed  Google Scholar 

  24. de Laat, W. L. et al. DNA-binding polarity of human replication protein A positions nucleases in nucleotide excision repair. Genes Dev. 12, 2598–2609 (1998). Describes how RPA binds to single-stranded DNA with a defined polarity and contributes to the control of XPF–ERCC1 and XPG at sites of DNA damage.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Staresincic, L. et al. Coordination of dual incision and repair synthesis in human nucleotide excision repair. EMBO J. 28, 1111–1120 (2009). Describes how the first incision in NER is made by XPF–ERCC1, which is necessary for the second incision that is carried out by XPG.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  26. Tripsianes, K. et al. Analysis of the XPA and ssDNA-binding surfaces on the central domain of human ERCC1 reveals evidence for subfunctionalization. Nucleic Acids Res. 35, 5789–5798 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  27. Tsodikov, O. V. et al. Structural basis for the recruitment of ERCC1–XPF to nucleotide excision repair complexes by XPA. EMBO J. 26, 4768–4776 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  28. Orelli, B. et al. The XPA-binding domain of ERCC1 is required for nucleotide excision repair but not other DNA repair pathways. J. Biol. Chem. 285, 3705–3712 (2010).

    Article  CAS  PubMed  Google Scholar 

  29. Sollier, J. et al. Transcription-coupled nucleotide excision repair factors promote R-loop-induced genome stability. Mol. Cell 56, 1–9 (2014).

    Article  CAS  Google Scholar 

  30. Fagbemi, A. F., Orelli, B. & Schärer, O. D. Regulation of endonuclease activity in human nucleotide excision repair. DNA Repair (Amsterdam) 10, 722–729 (2011).

    Article  CAS  Google Scholar 

  31. Zhang, J. & Walter, J. C. Mechanism and regulation of incisions during DNA interstrand cross-link repair. DNA Repair (Amsterdam) 19, 1–8 (2014).

    Article  CAS  Google Scholar 

  32. Lopez-Martinez, D., Liang, C.-C. & Cohn, M. A. Cellular response to DNA interstrand crosslinks: the Fanconi anemia pathway. Cell. Mol. Life Sci. 73, 1–18 (2016).

    Article  CAS  Google Scholar 

  33. Hodskinson, M. R. G. et al. Mouse SLX4 is a tumor suppressor that stimulates the activity of the nuclease XPF–ERCC1 in DNA crosslink repair. Mol. Cell 54, 472–484 (2014). Demonstrates that SLX4 is a tumour suppressor in mice, together with detailed biochemical analyses of the stimulation of XPF–ERCC1 by SLX4 on ICL repair intermediates.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Douwel, D. K. et al. XPF–ERCC1 acts in unhooking DNA interstrand crosslinks in cooperation with FANCD2 and FANCP/SLX4. Mol. Cell 54, 1–12 (2014). Describes the use of Xenopus laevis oocyte extracts to analyse the timely recruitment of FANCD2, SLX4 and XPF–ERCC1 to DNA during ICL repair.

    Article  CAS  Google Scholar 

  35. Lachaud, C. et al. Ubiquitinated Fancd2 recruits Fan1 to stalled replication forks to prevent genome instability. Science 351, 846–849 (2016). Shows that recruitment of FAN1 to ubiquitylated FANCD2 through its UBZ motif is crucial during replication inhibition, but is dispensable for ICL repair. A FAN1 variant from high-risk pancreatic cancer is not recruited to ubiquitylated FANCD2.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  36. Wang, R. et al. DNA repair. Mechanism of DNA interstrand cross-link processing by repair nuclease FAN1. Science 346, 1127–1130 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Liang, C.-C. et al. UHRF1 is a sensor for DNA interstrand crosslinks and recruits FANCD2 to initiate the Fanconi anemia pathway. Cell Rep. 10, 1947–1956 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  38. Tian, Y. et al. UHRF1 contributes to DNA damage repair as a lesion recognition factor and nuclease scaffold. Cell Rep. 10, 1957–1966 (2015). References 37 and 38 describe UHRF1 as a scaffold that is important in ICL repair and that recruits FANCD2, XPF–ERCC1 and MUS81–EME1.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Matos, J. & West, S. C. Holliday junction resolution: regulation in space and time. DNA Repair (Amsterdam) 19, 1–6 (2014).

    Article  CAS  Google Scholar 

  40. Blanco, M. G. & Matos, J. Hold your horSSEs: controlling structure-selective endonucleases MUS81 and Yen1/GEN1. Front. Genet. 6, 253 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  41. Matos, J., Blanco, M. G., Maslen, S., Skehel, J. M. & West, S. C. Regulatory control of the resolution of DNA recombination intermediates during meiosis and mitosis. Cell 147, 158–172 (2011). Seminal study about cell cycle control of the Holliday junction resolvases Mus81–Mms4 and Yen1 in meiosis and mitosis in S. cerevisiae , and MUS81–EME1 and GEN1 in mitotic human cells.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  42. Szakal, B. & Branzei, D. Premature Cdk1/Cdc5/Mus81 pathway activation induces aberrant replication and deleterious crossover. EMBO J. 32, 1155–1167 (2013). Describes the importance of the timely upregulation of Mus81–Mms4 in mitotic cells and the dire consequences if it is constitutively hyperactivate.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  43. Gallo-Fernandez, M. et al. Cell cycle-dependent regulation of the nuclease activity of Mus81–Eme1/Mms4. Nucleic Acids Res. 40, 8325–8335 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  44. Dehé, P.-M. 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 

  45. Wehrkamp-Richter, S., Hyppa, R. W., Prudden, J., Smith, G. R. & Boddy, M. N. Meiotic DNA joint molecule resolution depends on Nse5–Nse6 of the Smc5–Smc6 holocomplex. Nucleic Acids Res. 40, 9633–9646 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  46. Copsey, A. et al. Smc5/6 coordinates formation and resolution of joint molecules with chromosome morphology to ensure meiotic divisions. PLoS Genet. 9, e1004071 (2013).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  47. Sebesta, M. et al. Esc2 promotes Mus81 complex-activity via its SUMO-like and DNA binding domains. Nucleic Acids Res. http://dx.doi.org/10.1093/nar/gkw882 (2016).

  48. Wyatt, H. D. M., Sarbajna, S., Matos, J. & West, S. C. Coordinated actions of SLX1–SLX4 and MUS81–EME1 for Holliday junction resolution in human cells. Mol. Cell 52, 1–14 (2013).

    Article  CAS  Google Scholar 

  49. Castor, D. et al. Cooperative control of Holliday junction resolution and DNA repair by the SLX1 and MUS81–EME1 nucleases. Mol. Cell 52, 1–13 (2013). References 48 and 49 show that SLX4 controls and coordinates Holliday junction resolution by both SLX1 and MUS81–EME1.

    Article  CAS  Google Scholar 

  50. Eissler, C. L. et al. The Cdk/Cdc14 module controls activation of the Yen1 holliday junction resolvase to promote genome stability. Mol. Cell 54, 80–93 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  51. Blanco, M. G., Matos, J. & West, S. C. Dual control of Yen1 nuclease activity and cellular localization by Cdk and Cdc14 prevents genome instability. Mol. Cell 54, 94–106 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  52. García-Luis, J., Clemente-Blanco, A., Aragón, L. & Machin, F. Cdc14 targets the Holliday junction resolvase Yen1 to the nucleus in early anaphase. Cell Cycle 13, 1392–1399 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  53. Chan, Y. W. & West, S. C. Spatial control of the GEN1 Holliday junction resolvase ensures genome stability. Nat. Commun. 5, 4844–4811 (2014).

    Article  CAS  PubMed  Google Scholar 

  54. Bailly, A. P. et al. The Caenorhabditis elegans homolog of Gen1/Yen1 resolvases links DNA damage signaling to DNA double-strand break repair. PLoS Genet. 6, e1001025 (2010).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  55. Bellendir, S. P. & Sekelsky, J. An elegans solution for crossover formation. PLoS Genet. 9, e1003658 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  56. Andersen, S. L., Kuo, H. K., Savukoski, D., Brodsky, M. H. & Sekelsky, J. Three structure-selective endonucleases are essential in the absence of BLM helicase in Drosophila. PLoS Genet. 7, e1002315 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  57. Ölmezer, G. et al. Replication intermediates that escape Dna2 activity are processed by Holliday junction resolvase Yen1. Nat. Commun. 7, 13157 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  58. Kowalczykowski, S. C. An overview of the molecular mechanisms of recombinational DNA repair. Cold Spring Harb. Perspect. Biol. 7, a016410 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  59. Mazón, G. & Symington, L. S. Mph1 and Mus81-Mms4 prevent aberrant processing of mitotic recombination intermediates. Mol. Cell 52, 63–74 (2013).

    Article  PubMed  CAS  Google Scholar 

  60. Balakrishnan, L. & Bambara, R. A. Flap endonuclease 1. Annu. Rev. Biochem. 82, 119–138 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  61. Tsutakawa, S. E. et al. Human flap endonuclease structures, DNA double-base flipping, and a unified understanding of the FEN1 superfamily. Cell 145, 198–211 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  62. Tsutakawa, S. E. & Tainer, J. A. Double strand binding-single strand incision mechanism for human flap endonuclease: implications for the superfamily. Mech. Ageing Dev. 133, 195–202 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  63. Zheng, L. et al. Fen1 mutations result in autoimmunity, chronic inflammation and cancers. Nat. Med. 13, 812–819 (2007).

    Article  CAS  PubMed  Google Scholar 

  64. Li, X., Li, J., Harrington, J., Lieber, M. R. & Burgers, P. M. Lagging strand DNA synthesis at the eukaryotic replication fork involves binding and stimulation of FEN-1 by proliferating cell nuclear antigen. J. Biol. Chem. 270, 22109–22112 (1995).

    Article  CAS  PubMed  Google Scholar 

  65. Wu, X. et al. Processing of branched DNA intermediates by a complex of human FEN-1 and PCNA. Nucleic Acids Res. 24, 2036–2043 (1996).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  66. Henneke, G., Koundrioukoff, S. & Hubscher, U. Phosphorylation of human Fen1 by cyclin-dependent kinase modulates its role in replication fork regulation. Oncogene 22, 4301–4313 (2003).

    Article  CAS  PubMed  Google Scholar 

  67. Guo, Z. et al. Sequential posttranslational modifications program FEN1 degradation during cell-cycle progression. Mol. Cell 47, 444–456 (2012). Identifies a mechanism that controls the programmed degradation of FEN1 through sequential phosphorylation, sumoylation and ubiquitylation of FEN1.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  68. Guo, Z. et al. Methylation of FEN1 suppresses nearby phosphorylation and facilitates PCNA binding. Nat. Chem. Biol. 6, 766–773 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  69. Hasan, S. et al. Regulation of human flap endonuclease-1 activity by acetylation through the transcriptional coactivator p300. Mol. Cell 7, 1221–1231 (2001).

    Article  CAS  PubMed  Google Scholar 

  70. Choudhary, C. et al. Lysine acetylation targets protein complexes and co-regulates major cellular functions. Science 325, 834–840 (2009).

    Article  CAS  PubMed  Google Scholar 

  71. Balakrishnan, L., Stewart, J., Polaczek, P., Campbell, J. L. & Bambara, R. A. Acetylation of Dna2 endonuclease/helicase and flap endonuclease 1 by p300 promotes DNA stability by creating long flap intermediates. J. Biol. Chem. 285, 4398–4404 (2010).

    Article  CAS  PubMed  Google Scholar 

  72. Bae, S. H., Bae, K. H., Kim, J. A. & Seo, Y. S. RPA governs endonuclease switching during processing of Okazaki fragments in eukaryotes. Nature 412, 456–461 (2001).

    Article  CAS  PubMed  Google Scholar 

  73. Friedrich-Heineken, E. et al. The two DNA clamps Rad9/Rad1/Hus1 complex and proliferating cell nuclear antigen differentially regulate flap endonuclease 1 activity. J. Mol. Biol. 353, 980–989 (2005).

    Article  CAS  PubMed  Google Scholar 

  74. Sharma, S. et al. Stimulation of flap endonuclease-1 by the Bloom's syndrome protein. J. Biol. Chem. 279, 9847–9856 (2004).

    Article  CAS  PubMed  Google Scholar 

  75. Wang, W. & Bambara, R. A. Human Bloom protein stimulates flap endonuclease 1 activity by resolving DNA secondary structure. J. Biol. Chem. 280, 5391–5399 (2005).

    Article  CAS  PubMed  Google Scholar 

  76. Brosh, R. M. et al. Werner syndrome protein interacts with human flap endonuclease 1 and stimulates its cleavage activity. EMBO J. 20, 5791–5801 (2001).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  77. Zheng, L. et al. Novel function of the flap endonuclease 1 complex in processing stalled DNA replication forks. EMBO Rep. 6, 83–89 (2005).

    Article  CAS  PubMed  Google Scholar 

  78. Sharma, S. et al. The interaction site of Flap Endonuclease-1 with WRN helicase suggests a coordination of WRN and PCNA. Nucleic Acids Res. 33, 6769–6781 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  79. Sami, F. et al. RECQ1 interacts with FEN-1 and promotes binding of FEN-1 to telomeric chromatin. Biochem. J. 468, 227–244 (2015).

    Article  CAS  PubMed  Google Scholar 

  80. Schurman, S. H. et al. Direct and indirect roles of RECQL4 in modulating base excision repair capacity. Hum. Mol. Genet. 18, 3470–3483 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  81. Speina, E. et al. Human RECQL5beta stimulates flap endonuclease 1. Nucleic Acids Res. 38, 2904–2916 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  82. Magdalou, I., Lopez, B. S., Pasero, P. & Lambert, S. A. E. The causes of replication stress and their consequences on genome stability and cell fate. Semin. Cell Dev. Biol. 30, 154–164 (2014).

    Article  CAS  PubMed  Google Scholar 

  83. García-Muse, T. & Aguilera, A. Transcription–replication conflicts: how they occur and how they are resolved. Nat. Rev. Mol. Cell Biol. 17, 553–563 (2016).

    Article  PubMed  CAS  Google Scholar 

  84. 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 

  85. Guo, Z. et al. Nucleolar localization and dynamic roles of flap endonuclease 1 in ribosomal DNA replication and damage repair. Mol. Cell. Biol. 28, 4310–4319 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  86. Saharia, A. et al. FEN1 ensures telomere stability by facilitating replication fork re-initiation. J. Biol. Chem. 285, 27057–27066 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  87. Teasley, D. C. et al. Flap endonuclease 1 limits telomere fragility on the leading strand. J. Biol. Chem. 290, 15133–15145 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  88. Sharma, S. et al. WRN helicase and FEN-1 form a complex upon replication arrest and together process branchmigrating DNA structures associated with the replication fork. Mol. Biol. Cell 15, 734–750 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  89. Cheng, I-C. et al. Wuho is a new member in maintaining genome stability through its interaction with flap endonuclease 1. PLoS Biol. 14, e1002349 (2016). Identification of WUHO as a positive and negative regulator of the flap and gap endonuclease functions of FEN1, respectively.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  90. Chung, L. et al. The FEN1 E359K germline mutation disrupts the FEN1–WRN interaction and FEN1 GEN activity, causing aneuploidy-associated cancers. Oncogene 34, 902–911 (2015).

    Article  CAS  PubMed  Google Scholar 

  91. Shin, Y.-K., Amangyeld, T., Nguyen, T. A., Munashingha, P. R. & Seo, Y.-S. Human MUS81 complexes stimulate flap endonuclease 1. FEBS J. 279, 2412–2430 (2012).

    Article  CAS  PubMed  Google Scholar 

  92. Thu, H. P. T. et al. A physiological significance of the functional interaction between Mus81 and Rad27 in homologous recombination repair. Nucleic Acids Res. 43, 1684–1699 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  93. Kang, M.-J. et al. Genetic and functional interactions between Mus81–Mms4 and Rad27. Nucleic Acids Res. 38, 7611–7625 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  94. Chaudhury, I., Stroik, D. R. & Sobeck, A. FANCD2-controlled chromatin access of the Fanconi-associated nuclease FAN1 is crucial for the recovery of stalled replication forks. Mol. Cell. Biol. 34, 3939–3954 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  95. 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 

  96. Yuan, J., Ghosal, G. & Chen, J. The HARP-like domain-containing protein AH2/ZRANB3 binds to PCNA and participates in cellular response to replication stress. Mol. Cell 47, 410–421 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  97. Weston, R. et al. ZRANB3 is a structure-specific ATP-dependent endonuclease involved in replication stress response. Genes Dev. 26, 1558–1572 (2012). References 95–97 describe how the translocase ZRANB3 is an ATP-dependent SSE that is recruited to ubiquitylated PCNA.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  98. Badu-Nkansah, A., Mason, A. C., Eichman, B. F. & Cortez, D. Identification of a substrate recognition domain in the replication stress response protein zinc finger Ran-binding domain containing protein 3 (ZRANB3). J. Biol. Chem. 291 8251–8257 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  99. 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  PubMed  PubMed Central  Google Scholar 

  100. Sirbu, B. M. et al. Identification of proteins at active, stalled, and collapsed replication forks using isolation of proteins on nascent DNA (iPOND) coupled with mass spectrometry. J. Biol. Chem. 288, 31458–31467 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  101. Zeman, M. K. & Cimprich, K. A. Finally, polyubiquitinated PCNA gets recognized. Mol. Cell 47, 333–334 (2012).

    Article  CAS  PubMed  Google Scholar 

  102. Yao, Q. et al. Structure and specificity of the bacterial cysteine methyltransferase effector NleE suggests a novel substrate in human DNA repair pathway. PLoS Pathog. 10, e1004522 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  103. Doe, C. L., Ahn, J. S., Dixon, J. & Whitby, M. C. Mus81–Eme1 and Rqh1 involvement in processing stalled and collapsed replication forks. J. Biol. Chem. 277, 32753–32759 (2002).

    Article  CAS  PubMed  Google Scholar 

  104. Boddy, M. N. et al. Damage tolerance protein Mus81 associates with the FHA1 domain of checkpoint kinase Cds1. Mol. Cell. Biol. 20, 8758–8766 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  105. Interthal, H. & Heyer, W. D. MUS81 encodes a novel helix-hairpin-helix protein involved in the response to UV- and methylation-induced DNA damage in Saccharomyces cerevisiae. Mol. Gen. Genet. 263, 812–827 (2000).

    Article  CAS  PubMed  Google Scholar 

  106. Osman, F. & Whitby, M. Exploring the roles of Mus81–Eme1/Mms4 at perturbed replication forks. DNA Repair 6, 1004–1017 (2007).

    Article  CAS  PubMed  Google Scholar 

  107. Abraham, J. et al. Eme1 is involved in DNA damage processing and maintenance of genomic stability in mammalian cells. EMBO J. 22, 6137–6147 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  108. Dendouga, N. et al. Disruption of murine Mus81 increases genomic instability and DNA damage sensitivity but does not promote tumorigenesis. Mol. Cell. Biol. 25, 7569–7579 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  109. 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 

  110. Regairaz, M. et al. Mus81-mediated DNA cleavage resolves replication forks stalled by topoisomerase I–DNA complexes. J. Cell Biol. 195, 739–749 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  111. Franchitto, A. et al. Replication fork stalling in WRN-deficient cells is overcome by prompt activation of a MUS81-dependent pathway. J. Cell Biol. 183, 241–252 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  112. Noguchi, E., Noguchi, C., Du, L.-L. & Russell, P. Swi1 prevents replication fork collapse and controls checkpoint kinase Cds1. Mol. Cell. Biol. 23, 7861–7874 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  113. Noguchi, E., Noguchi, C., McDonald, W. H., Yates, J. R. & Russell, P. Swi1 and Swi3 are components of a replication fork protection complex in fission yeast. Mol. Cell. Biol. 24, 8342–8355 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  114. Bellaoui, M. et al. Elg1 forms an alternative RFC complex important for DNA replication and genome integrity. EMBO J. 22, 4304–4313 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  115. Pebernard, S., McDonald, W. H., Pavlova, Y., Yates, J. R. & Boddy, M. N. Nse1, Nse2, and a novel subunit of the Smc5–Smc6 complex, Nse3, play a crucial role in meiosis. Mol. Biol. Cell 15, 4866–4876 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  116. Boddy, M. N. et al. Replication checkpoint kinase Cds1 regulates recombinational repair protein Rad60. Mol. Cell. Biol. 23, 5939–5946 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  117. Irmisch, A., Ampatzidou, E., Mizuno, K., O'Connell, M. J. & Murray, J. M. Smc5/6 maintains stalled replication forks in a recombination-competent conformation. EMBO J. 28, 144–155 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  118. Torres-Rosell, J. et al. SMC5 and SMC6 genes are required for the segregation of repetitive chromosome regions. Nat. Cell Biol. 7, 412–419 (2005).

    Article  CAS  PubMed  Google Scholar 

  119. Mayle, R. et al. Mus81 and converging forks limit the mutagenicity of replication fork breakage. Science 349, 742–747 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  120. Gao, H., Chen, X.-B. & McGowan, C. H. Mus81 endonuclease localizes to nucleoli and to regions of DNA damage in human S-phase cells. Mol. Biol. Cell 14, 4826–4834 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  121. Shimura, T. et al. Bloom's syndrome helicase and Mus81 are required to induce transient double-strand DNA breaks in response to DNA replication stress. J. Mol. Biol. 375, 1152–1164 (2008).

    Article  CAS  PubMed  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

  124. Xing, M. et al. Acute MUS81 depletion leads to replication fork slowing and a constitutive DNA damage response. Oncotarget 6, 37638–37646 (2015).

    Article  PubMed  PubMed Central  Google Scholar 

  125. Zhang, R. et al. BLM helicase facilitates Mus81 endonuclease activity in human cells. Cancer Res. 65, 2526–2531 (2005).

    Article  CAS  PubMed  Google Scholar 

  126. Fadden, A. J. et al. A winged helix domain in human MUS81 binds DNA and modulates the endonuclease activity of MUS81 complexes. Nucleic Acids Res. 41, 9741–9752 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  127. Kikuchi, K. et al. Structure-specific endonucleases Xpf and Mus81 play overlapping but essential roles in DNA repair by homologous recombination. Cancer Res. 73, 4362–4371 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  128. Kai, M., Boddy, M. N., Russell, P. & Wang, T. S.-F. Replication checkpoint kinase Cds1 regulates Mus81 to preserve genome integrity during replication stress. Genes Dev. 19, 919–932 (2005). Provides evidence of the potentially deleterious effects of Mus81 during replication, especially when not controlled by Cds1.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  129. Froget, B., Blaisonneau, J., Lambert, S. & Baldacci, G. Cleavage of stalled forks by fission yeast Mus81/Eme1 in absence of DNA replication checkpoint. Mol. Biol. Cell 19, 445–456 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  130. Saugar, I. et al. Temporal regulation of the Mus81–Mms4 endonuclease ensures cell survival under conditions of DNA damage. Nucleic Acids Res. 41, 8943–8958 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  131. Schwartz, E. K. et al. Mus81–Mms4 function as a single heterodimer to cleave nicked intermediates in recombinational DNA repair. Mol. Cell. Biol. 32, 3065–3080 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  132. Gaillard, P.-H. L., Noguchi, E., Shanahan, P. & Russell, P. The endogenous Mus81–Eme1 complex resolves Holliday junctions by a nick and counternick mechanism. Mol. Cell 12, 747–759 (2003).

    Article  CAS  PubMed  Google Scholar 

  133. Domínguez-Kelly, R. et al. Wee1 controls genomic stability during replication by regulating the Mus81–Eme1 endonuclease. J. Cell Biol. 194, 567–579 (2011).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  134. 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  PubMed  PubMed Central  Google Scholar 

  135. Forment, J. V., Blasius, M., Guerini, I. & Jackson, S. P. Structure-specific DNA endonuclease Mus81/Eme1 generates DNA damage caused by Chk1 inactivation. PLoS ONE 6, e23517 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  136. Murfuni, I. et al. Survival of the replication checkpoint deficient cells requires MUS81–RAD52 function. PLoS Genet. 9, e1003910 (2013).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  137. Técher, H. et al. Signaling from Mus81–Eme2-dependent DNA damage elicited by Chk1 deficiency modulates replication fork speed and origin usage. Cell Rep. 14, 1114–1127 (2016).

    Article  PubMed  CAS  Google Scholar 

  138. Duda, H. et al. A mechanism for controlled breakage of under-replicated chromosomes during mitosis. Dev. Cell 1, 740–755 (2016).

    Article  CAS  Google Scholar 

  139. Ohouo, P. Y., Bastos de Oliveira, F. M., Almeida, B. S. & Smolka, M. B. DNA damage signaling recruits the Rtt107–Slx4 scaffolds via Dpb11 to mediate replication stress response. Mol. Cell 39, 300–306 (2010).

    Article  CAS  PubMed  Google Scholar 

  140. Ohouo, P. Y., Bastos de Oliveira, F. M., Liu, Y., Ma, C. J. & Smolka, M. B. DNA-repair scaffolds dampen checkpoint signalling by counteracting the adaptor Rad9. Nature 493, 120–124 (2013).

    Article  PubMed  CAS  Google Scholar 

  141. Princz, L. N., Gritenaite, D. & Pfander, B. The Slx4–Dpb11 scaffold complex: coordinating the response to replication fork stalling in S-phase and the subsequent mitosis. Cell Cycle 14, 488–494 (2015).

    Article  CAS  PubMed  Google Scholar 

  142. Balint, A. et al. Assembly of Slx4 signaling complexes behind DNA replication forks. EMBO J. 34, 2182–2197 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  143. Gritenaite, D. et al. A cell cycle-regulated Slx4–Dpb11 complex promotes the resolution of DNA repair intermediates linked to stalled replication. Genes Dev. 28, 1604–1619 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  144. Cussiol, J. R., Jablonowski, C. M., Yimit, A., Brown, G. W. & Smolka, M. B. Dampening DNA damage checkpoint signalling via coordinated BRCT domain interactions. EMBO J. 34, 1704–1717 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  145. Princz, L. N. et al. Dbf4-dependent kinase (DDK) and the Rtt107 scaffold promote Mus81–Mms4 resolvase activation during mitosis. EMBO J. http://dx.doi.org/10.15252/embj.201694831 (2017).

  146. 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 

  147. 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 

  148. Sogo, J. M., 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 

  149. Branzei, D. & Foiani, M. Maintaining genome stability at the replication fork. Nat. Rev. Mol. Cell Biol. 11, 208–219 (2010).

    Article  CAS  PubMed  Google Scholar 

  150. Ragland, R. L. et al. RNF4 and PLK1 are required for replication fork collapse in ATR-deficient cells. Genes Dev. 27, 2259–2273 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  151. Saito, T. T., Youds, J. L., Boulton, S. J. & Colaiácovo, M. P. Caenorhabditis elegans HIM-18/SLX-4 interacts with SLX-1 and XPF-1 and maintains genomic integrity in the germline by processing recombination intermediates. PLoS Genet. 5, e1000735 (2009).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  152. Fricke, W. M. & Brill, S. J. Slx1–Slx4 is a second structure-specific endonuclease functionally redundant with Sgs1–Top3. Genes Dev. 17, 1768–1778 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  153. Coulon, S. et al. Slx1–Slx4 are subunits of a structure-specific endonuclease that maintains ribosomal DNA in fission yeast. Mol. Biol. Cell 15, 71–80 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  154. Zakharyevich, K., Tang, S., Ma, Y. & Hunter, N. Delineation of joint molecule resolution pathways in meiosis identifies a crossover-specific resolvase. Cell 149, 334–347 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  155. Guervilly, J.-H. et al. The SLX4 complex is a SUMO E3 ligase that impacts on replication stress outcome and genome stability. Mol. Cell 57, 123–137 (2015).

    Article  CAS  PubMed  Google Scholar 

  156. Ouyang, J. et al. Noncovalent interactions with SUMO and ubiquitin orchestrate distinct functions of the SLX4 complex in genome maintenance. Mol. Cell 57, 108–122 (2015).

    Article  CAS  PubMed  Google Scholar 

  157. Minocherhomji, S. et al. Replication stress activates DNA repair synthesis in mitosis. Nature 528, 1–17 (2015).

    Article  CAS  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

  159. Naim, V., Wilhelm, T., Debatisse, M. & Rosselli, F. ERCC1 and MUS81–EME1 promote sister chromatid separation by processing late replication intermediates at common fragile sites during mitosis. Nat. Cell Biol. 15, 1008–1015 (2013).

    Article  CAS  PubMed  Google Scholar 

  160. Bergoglio, V. et al. DNA synthesis by Pol η promotes fragile site stability by preventing under-replicated DNA in mitosis. J. Cell Biol. 201, 395–408 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  161. Zhu, X.-D. et al. ERCC1/XPF removes the 3′ overhang from uncapped telomeres and represses formation of telomeric DNA-containing double minute chromosomes. Mol. Cell 12, 1489–1498 (2003).

    Article  CAS  PubMed  Google Scholar 

  162. Zeng, S. et al. Telomere recombination requires the MUS81 endonuclease. Nat. Cell Biol. 11, 616–623 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  163. Vannier, J.-B., Depeiges, A., White, C. & Gallego, M. E. ERCC1/XPF protects short telomeres from homologous recombination in Arabidopsis thaliana. PLoS Genet. 5, e1000380 (2009).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  164. Wan, B. et al. SLX4 assembles a telomere maintenance toolkit by bridging multiple endonucleases with telomeres. Cell Rep. 4, 861–869 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  165. Wilson, J. S. J. et al. Localization-dependent and -independent roles of SLX4 in regulating telomeres. Cell Rep. 4, 853–860 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  166. Sarkar, J. et al. SLX4 contributes to telomere preservation and regulated processing of telomeric joint molecule intermediates. Nucleic Acids Res. 43, 5912–5923 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  167. Vannier, J.-B., Pavicic-Kaltenbrunner, V., Petalcorin, M. I. R., Ding, H. & Boulton, S. J. RTEL1 dismantles T loops and counteracts telomeric G4-DNA to maintain telomere integrity. Cell 149, 795–806 (2012).

    Article  CAS  PubMed  Google Scholar 

  168. Saint-Léger, A. et al. The basic N-terminal domain of TRF2 limits recombination endonuclease action at human telomeres. Cell Cycle 13, 2469–2474 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  169. Poulet, A. et al. TRF2 promotes, remodels and protects telomeric Holliday junctions. EMBO J. 28, 641–651 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  170. Kim, Y. Nuclease delivery: versatile functions of SLX4/FANCP in genome maintenance. Mol. Cells 37, 569–574 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  171. Lachaud, C. et al. Distinct functional roles for the two SLX4 ubiquitin-binding UBZ domains mutated in Fanconi anemia. J. Cell Sci. 127, 2811–2817 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  172. Sarangi, P. et al. Sumoylation of the Rad1 nuclease promotes DNA repair and regulates its DNA association. Nucleic Acids Res. 42, 6393–6404 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  173. Flott, S. et al. Phosphorylation of Slx4 by Mec1 and Tel1 regulates the single-strand annealing mode of DNA repair in budding yeast. Mol. Cell. Biol. 27, 6433–6445 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  174. Sarangi, P. et al. A versatile scaffold contributes to damage survival via sumoylation and nuclease interactions. Cell Rep. 9, 143–152 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  175. Li, F. et al. Microarray-based genetic screen defines SAW1, a gene required for Rad1/Rad10-dependent processing of recombination intermediates. Mol. Cell 30, 325–335 (2008).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  176. Zhang, J.-M. et al. Fission yeast Pxd1 promotes proper DNA repair by activating Rad16XPF and inhibiting Dna2. PLoS Biol. 12, e1001946 (2014). Identifies Pxd1 as a novel nuclease scaffold in fission yeast that differentially regulates Rad16–Swi10 and Dna2 endonucleases.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  177. Bétous, 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  PubMed  PubMed Central  CAS  Google Scholar 

  178. Fugger, K. et al. FBH1 co-operates with MUS81 in inducing DNA double-strand breaks and cell death following replication stress. Nat. Commun. 4, 1423 (2013).

    Article  PubMed  CAS  Google Scholar 

  179. Burman, B., Zhang, Z. Z., Pegoraro, G., Lieb, J. D. & Misteli, T. Histone modifications predispose genome regions to breakage and translocation. Genes Dev. 29, 1393–1402 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  180. Le May, N. et al. NER factors are recruited to active promoters and facilitate chromatin modification for transcription in the absence of exogenous genotoxic attack. Mol. Cell 38, 54–66 (2010).

    Article  CAS  PubMed  Google Scholar 

  181. Trego, K. S. et al. Non-catalytic roles for XPG with BRCA1 and BRCA2 in homologous recombination and genome stability. Mol. Cell 61, 535–546 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  182. Laguette, N. et al. Premature activation of the SLX4 complex by Vpr promotes G2/M arrest and escape from innate immune sensing. Cell 156, 134–145 (2014).

    Article  CAS  PubMed  Google Scholar 

  183. Hartung, M. L. et al. H. pylori-induced DNA strand breaks are introduced by nucleotide excision repair endonucleases and promote NF-κB target gene expression. Cell Rep. 13, 70–79 (2015).

    Article  CAS  PubMed  Google Scholar 

  184. Ho, S. S. W. et al. The DNA structure-specific endonuclease MUS81 mediates DNA sensor STING-dependent host rejection of prostate cancer cells. Immunity 44, 1177–1189 (2016).

    Article  CAS  PubMed  Google Scholar 

  185. Nishino, T., Komori, K., Ishino, Y. & Morikawa, K. X-ray and biochemical anatomy of an archaeal XPF/Rad1/Mus81 family nuclease: similarity between its endonuclease domain and restriction enzymes. Structure 11, 445–457 (2003).

    Article  CAS  PubMed  Google Scholar 

  186. Ciccia, A., McDonald, N. & West, S. C. Structural and functional relationships of the XPF/MUS81 family of proteins. Annu. Rev. Biochem. 77, 259–287 (2008).

    Article  CAS  PubMed  Google Scholar 

  187. Sgouros, J., Gaillard, P. H. & Wood, R. D. A relationship betweena DNA-repair/recombination nuclease family and archaeal helicases. Trends Biochem. Sci. 24, 95–97 (1999).

    Article  CAS  PubMed  Google Scholar 

  188. Roberts, J. A., Bell, S. D. & White, M. F. An archaeal XPF repair endonuclease dependent on a heterotrimeric PCNA. Mol. Microbiol. 48, 361–371 (2003).

    Article  CAS  PubMed  Google Scholar 

  189. Gaillard, P. H. & Wood, R. D. Activity of individual ERCC1 and XPF subunits in DNA nucleotide excision repair. Nucleic Acids Res. 29, 872–879 (2001).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  190. Xue, X., Sung, P. & Zhao, X. Functions and regulation of the multitasking FANCM family of DNA motor proteins. Genes Dev. 29, 1777–1788 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  191. Smith, G. R., Boddy, M. N., Shanahan, P. & Russell, P. Fission yeast Mus81.Eme1 Holliday junction resolvase is required for meiotic crossing over but not for gene conversion. Genetics 165, 2289–2293 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  192. de los Santos, T., Loidl, J., Larkin, B. & Hollingsworth, N. M. A role for MMS4 in the processing of recombination intermediates during meiosis in Saccharomyces cerevisiae. Genetics 159, 1511–1525 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  193. Sekelsky, J. J., McKim, K. S., Chin, G. M. & Hawley, R. S. The Drosophila meiotic recombination gene mei-9 encodes a homologue of the yeast excision repair protein Rad1. Genetics 141, 619–627 (1995).

    CAS  PubMed  PubMed Central  Google Scholar 

  194. O'neil, N. J. et al. Joint molecule resolution requires the redundant activities of MUS-81 and XPF-1 during Caenorhabditis elegans meiosis. PLoS Genet. 9, e1003582 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  195. Makharashvili, N. et al. Catalytic and noncatalytic roles of the CtIP endonuclease in double-strand break end resection. Mol. Cell 54, 1022–1033 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  196. Wang, H. et al. CtIP maintains stability at common fragile sites and inverted repeats by end resection-independent endonuclease activity. Mol. Cell 54, 1012–1021 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  197. Matos, J., Blanco, M. G. & West, S. C. Cell-cycle kinases coordinate the resolution of recombination intermediates with chromosome segregation. Cell Rep. 4, 76–86 (2013).

    Article  CAS  PubMed  Google Scholar 

  198. 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 

  199. Kim, Y. et al. Mutations of the SLX4 gene in Fanconi anemia. Nat. Genet. 43, 142–146 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  200. Kaliraman, V. & Brill, S. J. Role of SGS1 and SLX4 in maintaining rDNA structure in Saccharomyces cerevisiae. Curr. Genet. 41, 389–400 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  201. Li, F. et al. Role of Saw1 in Rad1/Rad10 complex assembly at recombination intermediates in budding yeast. EMBO J. 32, 461–472 (2013). Demonstrates that the scaffold protein Saw1 is a structure-specific DNA-binding protein that targets and activates Rad1–Rad10 during SSA.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

Download references

Acknowledgements

The authors apologize for the many studies that were not cited owing to space limitations. The authors thank all members of their laboratory for stimulating discussions, especially S. Scaglione, J.-H. Guervilly and B. LLorente. The authors thank S. Scaglione and J.-H. Guervilly for their critical and careful reading of the manuscript. The authors also thank R. Wood, P. Russell, N. Boddy, P. McHugh and J. Matos for enlightening and enthusiastic discussions, with special thanks to R. Wood, P. Russell and N. Boddy for their feedback on the manuscript. Finally, the authors thank the referees for their thorough reviewing, which has undoubtedly helped improve the manuscript. This work was supported by grants from the Institut National du Cancer (PLBIO 2012–111), Agence Nationale de la Recherche (ANR Blanc EME1PHOSUMO) and the Siric-Cancéropôle PACA (AAP Projets émergents 2015).

Author information

Authors and Affiliations

Authors

Corresponding author

Correspondence to Pierre-Henri L. Gaillard.

Ethics declarations

Competing interests

The authors declare no competing financial interests.

Supplementary information

Supplementary information S1

Conserved families of structure-specific endonucleases (PDF 519 kb)

Supplementary information S2

Positive and negative regulation of SSEs (PDF 207 kb)

Supplementary information S3

Controlling the resolution of joint molecules by SSEs in meiosis. (PDF 783 kb)

Supplementary information S4

Structure-specific endonuclease scaffolds. (PDF 238 kb)

PowerPoint slides

Glossary

Single-stranded flaps

Single-stranded DNA protrusions from duplex DNA with either 5′ phosphate or 3′ hydroxyl ends.

Stem–loops

DNA structures that comprise two adjacent complementary sequences that form duplex DNA (the stem) with an intervening single-stranded closed loop.

Displacement loops

(D-loops). DNA structures that form early during homologous recombination by the invasion of duplex DNA by complementary single-stranded DNA.

Holliday junctions

Four-way DNA junction structures that form after complementary strand exchange between homologous sequences during DNA double-strand break repair by homologous recombination or during replication. Holliday junctions constitute covalent links between chromosomes.

DNA adducts

Chemical compounds or proteins that are covalently linked to DNA.

R-loops

Three-stranded nucleic acid structures that contain an RNA–DNA hybrid that results from the pairing of a nascent RNA with the DNA template during transcription.

Fanconi anaemia core complex

A multiprotein complex that resolves blocks to DNA replication such as DNA interstrand crosslinks. Mutations in the complex cause Fanconi anaemia, which is characterized mainly by haematological problems and cancer predisposition.

Trans-lesion DNA synthesis

Low-fidelity (and thereby mutagenic) DNA replication over damaged bases by specialized trans-lesion polymerases.

Ubiquitin-binding motif

A modular protein domain that mediates the non-covalent binding to monoubiquitin and/or polyubiquitin chains.

Double Holliday junctions

Structures that comprise two proximal Holliday junctions, which represent an intermediate stage of homologous recombination.

Holliday junction dissolution

A process by which double Holliday junctions are merged into a hemicatenane structure, in which one of the strands of one duplex passes between the strands of the other duplex. This structure is processed by a single-strand cut that releases the two duplexes from one another.

Bloom syndrome

A hereditary disorder that arises from mutations in the gene encoding the RecQ DNA helicase Bloom syndrome protein (BLM). It is characterized by growth deficiencies, sun sensitivity, immunodeficiency and predisposition to cancer.

Okazaki fragments

DNA fragments that are synthesized during replication of the lagging strand by DNA polymerase-α (Polα) and Polδ; the fragments are joined together by DNA ligase I to form the continuous lagging strand.

Common fragile sites

(CFSs). Chromosomal regions that are prone to replication stress. CFSs are a potential source of genome instability and are associated with chromosomal rearrangements in cancer.

NPL4 zinc finger domain

(NZF domain). A domain that mediates binding to monoubiquitin and polyubiquitin.

Winged-helix domain

A subclass of the helix-turn-helix DNA-binding domain that is found in DNA-processing enzymes and many transcription factors.

S phase checkpoint

A signalling pathway that is triggered by replication stress and that delays cell cycle progression and protects the integrity of replication forks.

Forkhead-associated domain

(FHA domain). A domain that mediates recognition of phosphopeptides.

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Dehé, PM., Gaillard, PH. Control of structure-specific endonucleases to maintain genome stability. Nat Rev Mol Cell Biol 18, 315–330 (2017). https://doi.org/10.1038/nrm.2016.177

Download citation

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/nrm.2016.177

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