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:

RPA and RAD51: fork reversal, fork protection, and genome stability

Nature Structural & Molecular Biology volume 25, pages 446–453 (2018)Cite this article

Subjects

Abstract

Replication protein A (RPA) and RAD51 are DNA-binding proteins that help maintain genome stability during DNA replication. These proteins regulate nucleases, helicases, DNA translocases, and signaling proteins to control replication, repair, recombination, and the DNA damage response. Their different DNA-binding mechanisms, enzymatic activities, and binding partners provide unique functionalities that cooperate to ensure that the appropriate activities are deployed at the right time to overcome replication challenges. Here we review and discuss the latest discoveries of the mechanisms by which these proteins work to preserve genome stability, with a focus on their actions in fork reversal and fork protection.

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

Access options

Buy this article

  • Purchase on Springer Link
  • Instant access to full article PDF
Buy now

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

Fig. 1: RPA and RAD51 have different biochemical characteristics.
Fig. 2: Replication-fork reversal is regulated by RPA and RAD51.
Fig. 3: Nascent strand degradation may be due to multiple rounds of fork reversal and nuclease action.
Fig. 4: Different functions of RAD51 may require different amounts of protein and ssDNA.

Similar content being viewed by others

References

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  2. Byun, T. S., Pacek, M., Yee, M. C., Walter, J. C. & Cimprich, K. A. Functional uncoupling of MCM helicase and DNA polymerase activities activates the ATR-dependent checkpoint. Genes Dev. 19, 1040–1052 (2005).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  3. García-Rodríguez, N., Morawska, M., Wong, R. P., Daigaku, Y. & Ulrich, H. D. Spatial separation between replisome- and template-induced replication stress signaling. EMBO J. 37, e98369 (2018).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  4. Takai, K. K., Kibe, T., Donigian, J. R., Frescas, D. & de Lange, T. Telomere protection by TPP1/POT1 requires tethering to TIN2. Mol. Cell 44, 647–659 (2011).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  5. Flynn, R. L. & Zou, L. Oligonucleotide/oligosaccharide-binding fold proteins: a growing family of genome guardians. Crit. Rev. Biochem. Mol. Biol. 45, 266–275 (2010).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  6. Kim, C., Paulus, B. F. & Wold, M. S. Interactions of human replication protein A with oligonucleotides. Biochemistry 33, 14197–14206 (1994).

    Article  PubMed  CAS  Google Scholar 

  7. Gibb, B. et al. Concentration-dependent exchange of replication protein A on single-stranded DNA revealed by single-molecule imaging. PLoS One 9, e87922 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  8. Arunkumar, A. I., Stauffer, M. E., Bochkareva, E., Bochkarev, A. & Chazin, W. J. Independent and coordinated functions of replication protein A tandem high affinity single-stranded DNA binding domains. J. Biol. Chem. 278, 41077–41082 (2003).

    Article  PubMed  CAS  Google Scholar 

  9. Bochkareva, E., Korolev, S., Lees-Miller, S. P. & Bochkarev, A. Structure of the RPA trimerization core and its role in the multistep DNA-binding mechanism of RPA. EMBO J. 21, 1855–1863 (2002).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  10. Fan, J. & Pavletich, N. P. Structure and conformational change of a replication protein A heterotrimer bound to ssDNA. Genes Dev. 26, 2337–2347 (2012).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  11. Nguyen, B. et al. Diffusion of human replication protein A along single-stranded DNA. J. Mol. Biol. 426, 3246–3261 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  12. Kemmerich, F. E. et al. Force regulated dynamics of RPA on a DNA fork. Nucleic Acids Res. 44, 5837–5848 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  13. Chen, R., Subramanyam, S., Elcock, A. H., Spies, M. & Wold, M. S. Dynamic binding of replication protein a is required for DNA repair. Nucleic Acids Res. 44, 5758–5772 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  14. 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).

    Article  PubMed  PubMed Central  Google Scholar 

  15. Bhat, K. P., Bétous, R. & Cortez, D. High-affinity DNA-binding domains of replication protein A (RPA) direct SMARCAL1-dependent replication fork remodeling. J. Biol. Chem. 290, 4110–4117 (2015).

    Article  PubMed  CAS  Google Scholar 

  16. Zou, L. & Elledge, S. J. Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 300, 1542–1548 (2003).

    Article  PubMed  CAS  Google Scholar 

  17. Ball, H. L., Myers, J. S. & Cortez, D. ATRIP binding to replication protein A-single-stranded DNA promotes ATR-ATRIP localization but is dispensable for Chk1 phosphorylation. Mol. Biol. Cell 16, 2372–2381 (2005).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  18. Xu, X. et al. The basic cleft of RPA70N binds multiple checkpoint proteins, including RAD9, to regulate ATR signaling. Mol. Cell. Biol. 28, 7345–7353 (2008).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  19. Bochkareva, E. et al. Single-stranded DNA mimicry in the p53 transactivation domain interaction with replication protein A. Proc. Natl Acad. Sci. USA 102, 15412–15417 (2005).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  21. Bass, T. E. et al. ETAA1 acts at stalled replication forks to maintain genome integrity. Nat. Cell Biol. 18, 1185–1195 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  22. Bansbach, C. E., Bétous, R., Lovejoy, C. A., Glick, G. G. & Cortez, D. The annealing helicase SMARCAL1 maintains genome integrity at stalled replication forks. Genes Dev. 23, 2405–2414 (2009).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  23. Ciccia, A. et al. The SIOD disorder protein SMARCAL1 is an RPA-interacting protein involved in replication fork restart. Genes Dev. 23, 2415–2425 (2009).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  24. Mer, G. et al. Structural basis for the recognition of DNA repair proteins UNG2, XPA, and RAD52 by replication factor RPA. Cell 103, 449–456 (2000).

    Article  PubMed  CAS  Google Scholar 

  25. Sugiyama, T., New, J. H. & Kowalczykowski, S. C. DNA annealing by RAD52 protein is stimulated by specific interaction with the complex of replication protein A and single-stranded DNA. Proc. Natl Acad. Sci. USA 95, 6049–6054 (1998).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  26. Melendy, T. & Stillman, B. An interaction between replication protein A and SV40 T antigen appears essential for primosome assembly during SV40 DNA replication. J. Biol. Chem. 268, 3389–3395 (1993).

    PubMed  CAS  Google Scholar 

  27. 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, 622–626 (2017).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  28. Toledo, L. I. et al. ATR prohibits replication catastrophe by preventing global exhaustion of RPA. Cell 155, 1088–1103 (2013).

    Article  PubMed  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  30. 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  PubMed  CAS  Google Scholar 

  31. Sarbajna, S. & West, S. C. Holliday junction processing enzymes as guardians of genome stability. Trends Biochem. Sci. 39, 409–419 (2014).

    Article  PubMed  CAS  Google Scholar 

  32. Bétous, R. et al. Substrate-selective repair and restart of replication forks by DNA translocases. Cell Rep. 3, 1958–1969 (2013). This paper showed that RPA directs the fork reversal activity of SMARCAL1, but not ZRANB3, to leading-strand gapped forks, suggesting that this family of SNF2 proteins may recognize different fork substrates.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  33. 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 

  34. Postow, L., Woo, E. M., Chait, B. T. & Funabiki, H. Identification of SMARCAL1 as a component of the DNA damage response. J. Biol. Chem. 284, 35951–35961 (2009).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  35. Yuan, J., Ghosal, G. & Chen, J. The annealing helicase HARP protects stalled replication forks. Genes Dev. 23, 2394–2399 (2009).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  37. Mason, A. C. et al. A structure-specific nucleic acid-binding domain conserved among DNA repair proteins. Proc. Natl Acad. Sci. USA 111, 7618–7623 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  38. Manosas, M. et al. RecG and UvsW catalyse robust DNA rewinding critical for stalled DNA replication fork rescue. Nat. Commun. 4, 2368 (2013).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  39. 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  PubMed  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  41. 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  PubMed  PubMed Central  CAS  Google Scholar 

  42. 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  PubMed  PubMed Central  CAS  Google Scholar 

  43. Poole, L. A. & Cortez, D. Functions of SMARCAL1, ZRANB3, and HLTF in maintaining genome stability. Crit. Rev. Biochem. Mol. Biol. 52, 696–714 (2017).

    Article  PubMed  CAS  Google Scholar 

  44. 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  PubMed  PubMed Central  CAS  Google Scholar 

  45. Shinohara, A., Ogawa, H. & Ogawa, T. Rad51 protein involved in repair and recombination in S. cerevisiae is a RecA-like protein. Cell 69, 457–470 (1992).

    Article  PubMed  CAS  Google Scholar 

  46. Miné, J. et al. Real-time measurements of the nucleation, growth and dissociation of single Rad51-DNA nucleoprotein filaments. Nucleic Acids Res. 35, 7171–7187 (2007).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  47. Hilario, J., Amitani, I., Baskin, R. J. & Kowalczykowski, S. C. Direct imaging of human Rad51 nucleoprotein dynamics on individual DNA molecules. Proc. Natl Acad. Sci. USA 106, 361–368 (2009).

    Article  PubMed  PubMed Central  Google Scholar 

  48. Benson, F. E., Stasiak, A. & West, S. C. Purification and characterization of the human Rad51 protein, an analogue of E. coli RecA. EMBO J. 13, 5764–5771 (1994).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  49. Sung, P. Catalysis of ATP-dependent homologous DNA pairing and strand exchange by yeast RAD51 protein. Science 265, 1241–1243 (1994).

    Article  PubMed  CAS  Google Scholar 

  50. Stasiak, A. & Egelman, E. H. Structure and function of RecA-DNA complexes. Experientia 50, 192–203 (1994).

    Article  PubMed  CAS  Google Scholar 

  51. Chi, P., Van Komen, S., Sehorn, M. G., Sigurdsson, S. & Sung, P. Roles of ATP binding and ATP hydrolysis in human Rad51 recombinase function. DNA Repair (Amst.) 5, 381–391 (2006).

    Article  CAS  Google Scholar 

  52. Bugreev, D. V. & Mazin, A. V. Ca2+ activates human homologous recombination protein Rad51 by modulating its ATPase activity. Proc. Natl Acad. Sci. USA 101, 9988–9993 (2004).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  53. 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 

  54. von Nicolai, C., Ehlén, Å., Martin, C., Zhang, X. & Carreira, A. A second DNA binding site in human BRCA2 promotes homologous recombination. Nat. Commun. 7, 12813 (2016).

    Article  CAS  Google Scholar 

  55. Carreira, A. et al. The BRC repeats of BRCA2 modulate the DNA-binding selectivity of RAD51. Cell 136, 1032–1043 (2009).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  56. Liu, J., Doty, T., Gibson, B. & Heyer, W. D. Human BRCA2 protein promotes RAD51 filament formation on RPA-covered single-stranded DNA. Nat. Struct. Mol. Biol. 17, 1260–1262 (2010).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  57. Jensen, R. B., Carreira, A. & Kowalczykowski, S. C. Purified human BRCA2 stimulates RAD51-mediated recombination. Nature 467, 678–683 (2010).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  58. McGlynn, P. & Lloyd, R. G. Rescue of stalled replication forks by RecG: simultaneous translocation on the leading and lagging strand templates supports an active DNA unwinding model of fork reversal and Holliday junction formation. Proc. Natl Acad. Sci. USA 98, 8227–8234 (2001).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  59. McGlynn, P. & Lloyd, R. G. Modulation of RNA polymerase by (p)ppGpp reveals a RecG-dependent mechanism for replication fork progression. Cell 101, 35–45 (2000).

    Article  PubMed  CAS  Google Scholar 

  60. Robu, M. E., Inman, R. B. & Cox, M. M. RecA protein promotes the regression of stalled replication forks in vitro. Proc. Natl Acad. Sci. USA 98, 8211–8218 (2001).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  61. Gupta, S., Yeeles, J. T. & Marians, K. J. Regression of replication forks stalled by leading-strand template damage: I. Both RecG and RuvAB catalyze regression, but RuvC cleaves the Holliday junctions formed by RecG preferentially. J. Biol. Chem. 289, 28376–28387 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  62. 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).The authors demonstrate that silencing RAD51 prevents fork reversal in response to a wide range of replication-stress agents.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  63. 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). This report finds that reversed forks generated by SMARCAL1 are degraded by the Mre11 nuclease if they cannot be stabilized by BRCA2 and RAD51.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  64. Mijic, S. et al. Replication fork reversal triggers fork degradation in BRCA2-defective cells. Nat. Commun. 8, 859 (2017). The authors report that BRCA2 is not needed for fork reversal but is important to stabilize RAD51 on reversed forks to prevent degradation.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  65. Reuter, M. et al. BRCA2 diffuses as oligomeric clusters with RAD51 and changes mobility after DNA damage in live cells. J. Cell Biol. 207, 599–613 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  PubMed  CAS  Google Scholar 

  67. Duro, E. et al. Identification of the MMS22L-TONSL complex that promotes homologous recombination. Mol. Cell 40, 632–644 (2010).

    Article  PubMed  CAS  Google Scholar 

  68. O’Donnell, L. et al. The MMS22L-TONSL complex mediates recovery from replication stress and homologous recombination. Mol. Cell 40, 619–631 (2010).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  69. Piwko, W. et al. The MMS22L-TONSL heterodimer directly promotes RAD51-dependent recombination upon replication stress. EMBO J. 35, 2584–2601 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  70. Courcelle, J., Donaldson, J. R., Chow, K. H. & Courcelle, C. T. DNA damage-induced replication fork regression and processing in Escherichia coli. Science 299, 1064–1067 (2003).

    Article  PubMed  CAS  Google Scholar 

  71. Defossez, P. A. et al. Elimination of replication block protein Fob1 extends the life span of yeast mother cells. Mol. Cell 3, 447–455 (1999).

    Article  PubMed  CAS  Google Scholar 

  72. Zou, H. & Rothstein, R. Holliday junctions accumulate in replication mutants via a RecA homolog-independent mechanism. Cell 90, 87–96 (1997).

    Article  PubMed  CAS  Google Scholar 

  73. Louarn, J., Cornet, F., François, V., Patte, J. & Louarn, J. M. Hyperrecombination in the terminus region of the Escherichia coli chromosome: possible relation to nucleoid organization. J. Bacteriol. 176, 7524–7531 (1994).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  74. Carr, A. M. & Lambert, S. Replication stress-induced genome instability: the dark side of replication maintenance by homologous recombination. J. Mol. Biol. 425, 4733–4744 (2013).

    Article  PubMed  CAS  Google Scholar 

  75. Margalef, P. et al. Stabilization of reversed replication forks by telomerase drives telomere catastrophe. Cell 172, 439–453.e14 (2018). This report provides an example of how fork reversal can be problematic in some circumstances because the reversed fork can be captured by proteins like telomerase.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  77. Hashimoto, Y., Ray Chaudhuri, A., 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  PubMed  PubMed Central  CAS  Google Scholar 

  78. 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  PubMed  PubMed Central  CAS  Google Scholar 

  79. Higgs, M. R. et al. BOD1L is required to suppress deleterious resection of stressed replication forks. Mol. Cell 59, 462–477 (2015).

    Article  PubMed  CAS  Google Scholar 

  80. 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  PubMed  CAS  Google Scholar 

  81. Iannascoli, C., Palermo, V., Murfuni, I., Franchitto, A. & Pichierri, P. The WRN exonuclease domain protects nascent strands from pathological MRE11/EXO1-dependent degradation. Nucleic Acids Res. 43, 9788–9803 (2015).

    PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  83. Lemaçon, 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). This paper reports that reversed forks are degraded by MRE11 in the absence of BRCA2 and that MUS81 acts in these cells to promote fork rescue.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  84. 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). The authors find that silencing any one of the SNF2 family fork remodeling enzymes SMARCAL1, HLTF, and ZRANB3 prevents nascent strand degradation, suggesting they may act sequentially to generate the substrate for MRE11 when BRCA2 is inactivated.

    Article  PubMed  CAS  PubMed Central  Google Scholar 

  85. Teixeira-Silva, A. et al. The end-joining factor Ku acts in the end-resection of double strand break-free arrested replication forks. Nat. Commun. 8, 1982 (2017).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  86. 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  PubMed  PubMed Central  CAS  Google Scholar 

  87. Dungrawala, H. et al. RADX promotes genome stability and modulates chemosensitivity by regulating RAD51 at replication forks. Mol. Cell 67, 374–386.e5 (2017). This paper identifies RADX as a new regulator of RAD51-dependent fork processing.

    Article  PubMed  CAS  PubMed Central  Google Scholar 

  88. Richardson, C., Stark, J. M., Ommundsen, M. & Jasin, M. Rad51 overexpression promotes alternative double-strand break repair pathways and genome instability. Oncogene 23, 546–553 (2004).

    Article  PubMed  CAS  Google Scholar 

  89. Klein, H. L. The consequences of Rad51 overexpression for normal and tumor cells. DNA Repair (Amst.) 7, 686–693 (2008).

    Article  CAS  Google Scholar 

  90. Tennstedt, P. et al. RAD51 overexpression is a negative prognostic marker for colorectal adenocarcinoma. Int. J. Cancer 132, 2118–2126 (2013).

    Article  PubMed  CAS  Google Scholar 

  91. Vispé, S., Cazaux, C., Lesca, C. & Defais, M. Overexpression of Rad51 protein stimulates homologous recombination and increases resistance of mammalian cells to ionizing radiation. Nucleic Acids Res. 26, 2859–2864 (1998).

    Article  PubMed  PubMed Central  Google Scholar 

  92. Hansen, L. T., Lundin, C., Spang-Thomsen, M., Petersen, L. N. & Helleday, T. The role of RAD51 in etoposide (VP16) resistance in small cell lung cancer. Int. J. Cancer 105, 472–479 (2003).

    Article  PubMed  CAS  Google Scholar 

  93. Drees, J. C., Lusetti, S. L., Chitteni-Pattu, S., Inman, R. B. & Cox, M. M. A RecA filament capping mechanism for RecX protein. Mol. Cell 15, 789–798 (2004).

    Article  PubMed  CAS  Google Scholar 

  94. Marians, K. J. Lesion bypass and the reactivation of stalled replication forks. Annu. Rev. Biochem. https://doi.org/10.1146/annurev-biochem-062917-011921 (2018).

    Article  PubMed  Google Scholar 

  95. 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  PubMed  CAS  PubMed Central  Google Scholar 

  96. Symington, L. S. End resection at double-strand breaks: mechanism and regulation. Cold Spring Harb. Perspect. Biol. 6, a016436 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  97. Kim, T. M. et al. RECQL5 and BLM exhibit divergent functions in cells defective for the Fanconi anemia pathway. Nucleic Acids Res. 43, 893–903 (2015).

    Article  PubMed  CAS  Google Scholar 

  98. Hartford, S. A. et al. Interaction with PALB2 is essential for maintenance of genomic integrity by BRCA2. PLoS Genet. 12, e1006236 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  99. Somyajit, K., Saxena, S., Babu, S., Mishra, A. & Nagaraju, G. Mammalian RAD51 paralogs protect nascent DNA at stalled forks and mediate replication restart. Nucleic Acids Res. 43, 9835–9855 (2015).

    PubMed  PubMed Central  CAS  Google Scholar 

  100. Patel, D. S., Misenko, S. M., Her, J. & Bunting, S. F. BLM helicase regulates DNA repair by counteracting RAD51 loading at DNA double-strand break sites. J. Cell Biol. 216, 3521–3534 (2017).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  103. 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  PubMed  PubMed Central  CAS  Google Scholar 

  104. Huh, M. S. et al. Stalled replication forks within heterochromatin require ATRX for protection. Cell Death Dis. 7, e2220 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  105. Ray Chaudhuri, A. et al. Replication fork stability confers chemoresistance in BRCA-deficient cells. Nature 535, 382–387 (2016). This is the first paper to show that inhibiting fork degradation in BRCA2-deficient cells can generate PARP-inhibitor resistance.

    Article  PubMed  CAS  Google Scholar 

  106. Mochizuki, A. L. et al. PARI regulates stalled replication fork processing to maintain genome stability upon replication stress in mice. Mol. Cell. Biol. 37, e00117–17 (2017).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  108. Yang, Y. et al. FANCD2 and REV1 cooperate in the protection of nascent DNA strands in response to replication stress. Nucleic Acids Res. 43, 8325–8339 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  109. 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  PubMed  PubMed Central  CAS  Google Scholar 

  110. 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 

  111. Villa, M., Bonetti, D., Carraro, M. & Longhese, M. P. Rad9/53BP1 protects stalled replication forks from degradation in Mec1/ATR-defective cells. EMBO Rep. 19, 351–367 (2018).

    Article  PubMed  CAS  Google Scholar 

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

    Article  PubMed  PubMed Central  Google Scholar 

Download references

Author information

Authors and Affiliations

  1. Department of Biochemistry, Vanderbilt University School of Medicine, Nashville, Tennessee, USA

    Kamakoti P. Bhat & David Cortez

Authors
  1. Kamakoti P. Bhat
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. David Cortez
    View author publications

    You can also search for this author in PubMed Google Scholar

Corresponding author

Correspondence to David Cortez.

Ethics declarations

Competing interests

The authors declare no competing financial interests.

Additional information

Publisher's note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Bhat, K.P., Cortez, D. RPA and RAD51: fork reversal, fork protection, and genome stability. Nat Struct Mol Biol 25, 446–453 (2018). https://doi.org/10.1038/s41594-018-0075-z

Download citation

  • Received:

  • Revised:

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41594-018-0075-z

This article is cited by

Search

Advanced 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