The impact of replication stress on replication dynamics and DNA damage in vertebrate cells

Nature Reviews Genetics

Volumn 18, Issue 9, 2017, Pages 535-550

  Técher, Hervé ^a   Koundrioukoff, Stéphane ^b,c   Nicolas, Alain ^d   Debatisse, Michelle ^b,c  

^a FIRC INSTITUTE OF MOLECULAR ONCOLOGY   (Italy)

^b CNRS   (France)

^c SORBONNE UNIVERSITÉ   (France)

^d Institut Curie   (France)

Author keywords

[No Author keywords available]

Indexed keywords

ATR PROTEIN; CELL PROTEIN; CHECKPOINT KINASE 1; PARATHION; REPLICATION FACTOR A; UNCLASSIFIED DRUG; WEE1 PROTEIN; DEOXYRIBONUCLEOTIDE;

ADAPTATION; CELL CYCLE PROGRESSION; CELL LINE; DNA DAMAGE; DNA DAMAGE CHECKPOINT; DNA DAMAGE RESPONSE; DNA REPLICATION; DNA REPLICATION ORIGIN; DNA REPLICATION TIMING; DNA RNA HYBRIDIZATION; DNA SYNTHESIS INHIBITION; GENE ACTIVATION; GENETIC ORGANIZATION; GENETIC TRANSCRIPTION; GENOME ANALYSIS; GENOMIC INSTABILITY; HUMAN; NONHUMAN; ONCOGENE; OXIDATIVE STRESS; PRIORITY JOURNAL; REGULATORY MECHANISM; REPLICATION FORK PROGRESSION; REPLICATION FORK STALLING; REVIEW; S PHASE CELL CYCLE CHECKPOINT; SIGNAL TRANSDUCTION; VERTEBRATE; ANIMAL; CELL CYCLE; CELLS; CONFORMATION; METABOLISM;

ANIMALS; CELL CYCLE; CELLS; DEOXYRIBONUCLEOTIDES; DNA DAMAGE; DNA REPLICATION; HUMANS; NUCLEIC ACID CONFORMATION; TRANSCRIPTION, GENETIC;

EID: 85027511305     PISSN: 14710056     EISSN: 14710064     Source Type: Journal    
DOI: 10.1038/nrg.2017.46     Document Type: Review

References (233)

1. Tourriere, H. & Pasero, P. Maintenance of fork integrity at damaged DNA and natural pause sites. DNA Repair (Amst.) 6, 900-913 (2007).
2. Macheret, M. & Halazonetis, T. D. DNA replication stress as a hallmark of cancer. Annu. Rev. Pathol. 10, 425-448 (2015).
3. Zeman, M. K. & Cimprich, K. A. Causes and consequences of replication stress. Nat. Cell Biol. 16, 2-9 (2014).
4. Halazonetis, T. D., Gorgoulis, V. G. & Bartek, J. An oncogene-induced DNA damage model for cancer development. Science 319, 1352-1355 (2008). The authors propose for the first time that oncogene activation in precancerous lesions induces replication stress, resulting in the formation of DNA breaks and subsequent genomic instability. This ultimately selects for cells that are deficient in checkpoint surveillance.
5. Negrini, S., Gorgoulis, V. G. & Halazonetis, T. D. Genomic instability - An evolving hallmark of cancer. Nat. Rev. Mol. Cell Biol. 11, 220-228 (2010).
6. Dereli-Oz, A., Versini, G. & Halazonetis, T. D. Studies of genomic copy number changes in human cancers reveal signatures of DNA replication stress. Mol. Oncol. 5, 308-314 (2011).
7. Barlow, J. H. et al. Identification of early replicating fragile sites that contribute to genome instability. Cell 152, 620-632 (2013).
8. Le Tallec, B. et al. Common fragile site profiling in epithelial and erythroid cells reveals that most recurrent cancer deletions lie in fragile sites hosting large genes. Cell Rep. 4, 420-428 (2013).
9. Aladjem, M. I. & Redon, C. E. Order from clutter: Selective interactions at mammalian replication origins. Nat. Rev. Genet. 18, 101-116 (2017).
10. Sorensen, C. S. & Syljuasen, R. G. Safeguarding genome integrity: The checkpoint kinases ATR, CHK1 and WEE1 restrain CDK activity during normal DNA replication. Nucleic Acids Res. 40, 477-486 (2012).
11. Yekezare, M., Gomez-Gonzalez, B. & Diffley, J. F. Controlling DNA replication origins in response to DNA damage - inhibit globally, activate locally. J. Cell Sci. 126, 1297-1306 (2013).
12. Dehe, P. M. & Gaillard, P. H. Control of structure-specific endonucleases to maintain genome stability. Nat. Rev. Mol. Cell Biol. 18, 315-330 (2017).
13. Gilbert, D. M. et al. Space and time in the nucleus: Developmental control of replication timing and chromosome architecture. Cold Spring Harb. Symp. Quant. Biol. 75, 143-153 (2010).
14. Gilbert, D. M. Cell fate transitions and the replication timing decision point. J. Cell Biol. 191, 899-903 (2010).
15. Pope, B. D. et al. Replication-timing boundaries facilitate cell-type and species-specific regulation of a rearranged human chromosome in mouse. Hum. Mol. Genet. 21, 4162-4170 (2012).
16. Dileep, V., Rivera-Mulia, J. C., Sima, J. & Gilbert, D. M. Large-scale chromatin structure-function relationships during the cell cycle and development: Insights from replication timing. Cold Spring Harb. Symp. Quant. Biol. 80, 53-63 (2015).
17. Dileep, V. et al. Topologically associating domains and their long-range contacts are established during early G1 coincident with the establishment of the replication-timing program. Genome Res. 25, 1104-1113 (2015).
18. Sexton, T. & Cavalli, G. The role of chromosome domains in shaping the functional genome. Cell 160, 1049-1059 (2015).
19. Naumova, N. et al. Organization of the mitotic chromosome. Science 342, 948-953 (2013).
20. Chagin, V. O. et al. 4D visualization of replication foci in mammalian cells corresponding to individual replicons. Nat. Commun. 7, 11231 (2016).
21. Schepers, A. & Papior, P. Why are we where we are? Understanding replication origins and initiation sites in eukaryotes using ChIP-approaches. Chromosome Res. 18, 63-77 (2010).
22. Deegan, T. D. & Diffley, J. F. MCM: One ring to rule them all. Curr. Opin. Struct. Biol. 37, 145-151 (2016).
23. Gambus, A., Khoudoli, G. A., Jones, R. C. & Blow, J. J. MCM2-7 form double hexamers at licensed origins in Xenopus egg extract. J. Biol. Chem. 286, 11855-11864 (2011).
24. Evrin, C. et al. A double-hexameric MCM2-7 complex is loaded onto origin DNA during licensing of eukaryotic DNA replication. Proc. Natl Acad. Sci. USA 106, 20240-20245 (2009).
25. Siddiqui, K., On, K. F. & Diffley, J. F. Regulating DNA replication in eukarya. Cold Spring Harb. Perspect. Biol. 5, a012930 (2013).
26. Yeeles, J. T., Deegan, T. D., Janska, A., Early, A. & Diffley, J. F. Regulated eukaryotic DNA replication origin firing with purified proteins. Nature 519, 431-435 (2015). This pioneer work reports the purification and in vitro reconstitution of the different steps of pre-RC assembly and activation up to origin firing. The authors identified for the first time the minimal set of proteins essential for initiating replication in eukaryotes.
27. Hyrien, O. Peaks cloaked in the mist: The landscape of mammalian replication origins. J. Cell Biol. 208, 147-160 (2015).
28. Prioleau, M. N. & MacAlpine, D. M. DNA replication origins-where do we begin? Genes Dev. 30, 1683-1697 (2016).
29. Fragkos, M., Ganier, O., Coulombe, P. & Mechali, M. DNA replication origin activation in space and time. Nat. Rev. Mol. Cell Biol. 16, 360-374 (2015).
30. Lujan, S. A., Williams, J. S. & Kunkel, T. A. DNA polymerases divide the labor of genome replication. Trends Cell Biol. 26, 640-654 (2016).
31. Yeeles, J. T., Janska, A., Early, A. & Diffley, J. F. How the eukaryotic replisome achieves rapid and efficient DNA replication. Mol. Cell 65, 105-116 (2017).
32. Kurat, C. F., Yeeles, J. T., Patel, H., Early, A. & Diffley, J. F. Chromatin controls DNA replication origin selection, lagging-strand synthesis, and replication fork rates. Mol. Cell 65, 117-130 (2017).
33. Devbhandari, S., Jiang, J., Kumar, C., Whitehouse, I. & Remus, D. Chromatin constrains the initiation and elongation of DNA replication. Mol. Cell 65, 131-141 (2017).
34. Guilbaud, G. et al. Evidence for sequential and increasing activation of replication origins along replication timing gradients in the human genome. PLoS Comput. Biol. 7, e1002322 (2011).
35. Techer, H. et al. Replication dynamics: Biases and robustness of DNA fiber analysis. J. Mol. Biol. 425, 4845-4855 (2013).
36. Koundrioukoff, S. et al. Stepwise activation of the ATR signaling pathway upon increasing replication stress impacts fragile site integrity. PLoS Genet. 9, e1003643 (2013).
37. Speroni, J., Federico, M. B., Mansilla, S. F., Soria, G. & Gottifredi, V. Kinase-independent function of checkpoint kinase 1 (Chk1) in the replication of damaged DNA. Proc. Natl Acad. Sci. USA 109, 7344-7349 (2012).
38. Tuduri, S. et al. Topoisomerase I suppresses genomic instability by preventing interference between replication and transcription. Nat. Cell Biol. 11, 1315-1324 (2009). This study shows how topoisomerase I deficiency leads to genomic instability through enhanced conflicts between replication forks and the transcription machinery.
39. Poli, J. et al. dNTP pools determine fork progression and origin usage under replication stress. EMBO J. 31, 883-894 (2012).
40. Techer, 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). This study shows that Chk1 deficiency triggers nuclease-dependent DNA damage. Damage activates the ATM pathway and, in turn, induces fork slowing and subsequent firing of latent origins, helping replication to proceed along damaged templates.
41. Wilhelm, T. et al. Slow replication fork velocity of homologous recombination-defective cells results from endogenous oxidative stress. PLoS Genet. 12, e1006007 (2016).
42. Chabosseau, P. et al. Pyrimidine pool imbalance induced by BLM helicase deficiency contributes to genetic instability in Bloom syndrome. Nat. Commun. 2, 368 (2011).
43. Bester, A. C. et al. Nucleotide deficiency promotes genomic instability in early stages of cancer development. Cell 145, 435-446 (2011).
44. Gay, S. et al. Nucleotide supply, not local histone acetylation, sets replication origin usage in transcribed regions. EMBO Rep. 11, 698-704 (2010).
45. Groth, A. et al. Regulation of replication fork progression through histone supply and demand. Science 318, 1928-1931 (2007).
46. Mejlvang, J. et al. New histone supply regulates replication fork speed and PCNA unloading. J. Cell Biol. 204, 29-43 (2014).
47. Dungrawala, H. et al. The replication checkpoint prevents two types of fork collapse without regulating replisome stability. Mol. Cell 59, 998-1010 (2015). This study provides an exhaustive and quantitative analysis of the protein landscape at replication forks in a variety of experimental conditions, which reveals unexpected mechanisms by which the DDR preserves genome integrity.
48. Gordenin, D. A. & Resnick, M. A. Yeast ARMs (DNA at-risk motifs) can reveal sources of genome instability. Mutat. Res. 400, 45-58 (1998).
49. Liu, P., Carvalho, C. M., Hastings, P. J. & Lupski, J. R. Mechanisms for recurrent and complex human genomic rearrangements. Curr. Opin. Genet. Dev. 22, 211-220 (2012).
50. Sutherland, G. R., Baker, E. & Richards, R. I. Fragile sites still breaking. Trends Genet. 14, 501-506 (1998).
51. Mirkin, E. V. & Mirkin, S. M. Replication fork stalling at natural impediments. Microbiol. Mol. Biol. Rev. 71, 13-35 (2007).
52. Wang, G. & Vasquez, K. M. Impact of alternative DNA structures on DNA damage, DNA repair, and genetic instability. DNA Repair (Amst.) 19, 143-151 (2014).
53. Usdin, K., House, N. C. & Freudenreich, C. H. Repeat instability during DNA repair: Insights from model systems. Crit. Rev. Biochem. Mol. Biol. 50, 142-167 (2015).
54. Wickramasinghe, C. M., Arzouk, H., Frey, A., Maiter, A. & Sale, J. E. Contributions of the specialised DNA polymerases to replication of structured DNA. DNA Repair (Amst.) 29, 83-90 (2015).
55. Barthelemy, J., Hanenberg, H. & Leffak, M. FANCJ is essential to maintain microsatellite structure genome-wide during replication stress. Nucleic Acids Res. 44, 6803-6816 (2016).
56. Mendoza, O., Bourdoncle, A., Boule, J. B., Brosh, R. M. Jr & Mergny, J. L. G-Quadruplexes and helicases. Nucleic Acids Res. 44, 1989-2006 (2016).
57. Leon-Ortiz, A. M., Svendsen, J. & Boulton, S. J. Metabolism of DNA secondary structures at the eukaryotic replication fork. DNA Repair (Amst.) 19, 152-162 (2014).
58. Schmidt, M. H. & Pearson, C. E. Disease-associated repeat instability and mismatch repair. DNA Repair (Amst.) 38, 117-126 (2016).
59. Maizels, N. G4-associated human diseases. EMBO Rep. 16, 910-922 (2015).
60. Rhodes, D. & Lipps, H. J. G-Quadruplexes and their regulatory roles in biology. Nucleic Acids Res. 43, 8627-8637 (2015).
61. Sollier, J. & Cimprich, K. A. Breaking bad: R-loops and genome integrity. Trends Cell Biol. 25, 514-522 (2015).
62. Santos-Pereira, J. M. & Aguilera, A. R loops: New modulators of genome dynamics and function. Nat. Rev. Genet. 16, 583-597 (2015).
63. Garcia-Muse, T. & Aguilera, A. Transcription-replication conflicts: How they occur and how they are resolved. Nat. Rev. Mol. Cell Biol. 17, 553-563 (2016).
64. Sollier, J. et al. Transcription-coupled nucleotide excision repair factors promote R-loop-induced genome instability. Mol. Cell 56, 777-785 (2014). This study shows that R-loops are actively processed into DNA double-strand breaks by the NER endonucleases XPF-ERCC1 and XPG.
65. Stork, C. T. et al. Co-transcriptional R-loops are the main cause of estrogen-induced DNA damage. eLife 5, e17548 (2016).
66. Ginno, P. A., Lott, P. L., Christensen, H. C., Korf, I. & Chedin, F. R-Loop formation is a distinctive characteristic of unmethylated human CpG island promoters. Mol. Cell 45, 814-825 (2012). This study reports the development of DRIP (DNA- RNA hybrid immunoprecipitation), a technique that allows the genome-wide mapping of these hybrids. The authors show that long R-loop structures form preferentially upon the transcription of regions with high GC skew.
67. Ginno, P. A., Lim, Y. W., Lott, P. L., Korf, I. & Chedin, F. GC skew at the 5? and 3? ends of human genes links R-loop formation to epigenetic regulation and transcription termination. Genome Res. 23, 1590-1600 (2013).
68. Sanz, L. A. et al. Prevalent, dynamic, and conserved R-loop structures associate with specific epigenomic signatures in mammals. Mol. Cell 63, 167-178 (2016).
69. Kantidakis, T. et al. Mutation of cancer driver MLL2 results in transcription stress and genome instability. Genes Dev. 30, 408-420 (2016).
70. Pommier, Y., Sun, Y., Huang, S. N. & Nitiss, J. L. Roles of eukaryotic topoisomerases in transcription, replication and genomic stability. Nat. Rev. Mol. Cell Biol. 17, 703-721 (2016).
71. Bhatia, V. et al. BRCA2 prevents R-loop accumulation and associates with TREX-2 mRNA export factor PCID2. Nature 511, 362-365 (2014). This study shows that BRCA2 and FANC proteins protect genome stability by promoting the elimination of R-loops, which block replication fork progression.
72. Garcia-Rubio, M. L. et al. The Fanconi anemia pathway protects genome integrity from R-loops. PLoS Genet. 11, e1005674 (2015).
73. Yeo, C. Q. et al. p53 maintains genomic stability by preventing interference between transcription and replication. Cell Rep. 15, 132-146 (2016).
74. Herrera-Moyano, E., Mergui, X., Garcia-Rubio, M. L., Barroso, S. & Aguilera, A. The yeast and human FACT chromatin-reorganizing complexes solve R-loop-mediated transcription-replication conflicts. Genes Dev. 28, 735-748 (2014).
75. Gan, W. et al. R-Loop-mediated genomic instability is caused by impairment of replication fork progression. Genes Dev. 25, 2041-2056 (2011).
76. Durkin, S. G. & Glover, T. W. Chromosome fragile sites. Annu. Rev. Genet. 41, 169-192 (2007).
77. Debatisse, M., Le Tallec, B., Letessier, A., Dutrillaux, B. & Brison, O. Common fragile sites: Mechanisms of instability revisited. Trends Genet. 28, 22-32 (2012).
78. Helmrich, A., Ballarino, M. & Tora, L. Collisions between replication and transcription complexes cause common fragile site instability at the longest human genes. Mol. Cell 44, 966-977 (2011).
79. Wilson, T. E. et al. Large transcription units unify copy number variants and common fragile sites arising under replication stress. Genome Res. 25, 189-200 (2015).
80. Letessier, A. et al. Cell-type-specific replication initiation programs set fragility of the FRA3B fragile site. Nature 470, 120-123 (2011).
81. Le Tallec, B. et al. Molecular profiling of common fragile sites in human fibroblasts. Nat. Struct. Mol. Biol. 18, 1421-1423 (2011).
82. Gros, J. et al. Post-licensing specification of eukaryotic replication origins by facilitated Mcm2-7 sliding along DNA. Mol. Cell 60, 797-807 (2015).
83. Powell, S. K. et al. Dynamic loading and redistribution of the Mcm2-7 helicase complex through the cell cycle. EMBO J. 34, 531-543 (2015).
84. Chon, H. et al. RNase H2 roles in genome integrity revealed by unlinking its activities. Nucleic Acids Res. 41, 3130-3143 (2013).
85. Madireddy, A. et al. FANCD2 facilitates replication through common fragile sites. Mol. Cell 64, 388-404 (2016).
86. Saponaro, M. et al. RECQL5 controls transcript elongation and suppresses genome instability associated with transcription stress. Cell 157, 1037-1049 (2014).
87. Stillman, B. Deoxynucleoside triphosphate (dNTP) synthesis and destruction regulate the replication of both cell and virus genomes. Proc. Natl Acad. Sci. USA 110, 14120-14121 (2013).
88. Murthy, S. & Reddy, G. P. Replitase: Complete machinery for DNA synthesis. J. Cell. Physiol. 209, 711-717 (2006).
89. Mathews, C. K. Deoxyribonucleotide metabolism, mutagenesis and cancer. Nat. Rev. Cancer 15, 528-539 (2015).
90. Chabes, A. L., Pfleger, C. M., Kirschner, M. W. & Thelander, L. Mouse ribonucleotide reductase R2 protein: A new target for anaphase-promoting complex-Cdh1-mediated proteolysis. Proc. Natl Acad. Sci. USA 100, 3925-3929 (2003).
91. D'Angiolella, V. et al. Cyclin F-mediated degradation of ribonucleotide reductase M2 controls genome integrity and DNA repair. Cell 149, 1023-1034 (2012).
92. Hu, C. M. et al. Tumor cells require thymidylate kinase to prevent dUTP incorporation during DNA repair. Cancer Cell 22, 36-50 (2012).
93. Chen, C. W. et al. The impact of dUTPase on ribonucleotide reductase-induced genome instability in cancer cells. Cell Rep. 16, 1287-1299 (2016).
94. Hakansson, P., Hofer, A. & Thelander, L. Regulation of mammalian ribonucleotide reduction and dNTP pools after DNA damage and in resting cells. J. Biol. Chem. 281, 7834-7841 (2006).
95. Pontarin, G. et al. p53R2-dependent ribonucleotide reduction provides deoxyribonucleotides in quiescent human fibroblasts in the absence of induced DNA damage. J. Biol. Chem. 282, 16820-16828 (2007).
96. Tanaka, H. et al. A ribonucleotide reductase gene involved in a p53-dependent cell-cycle checkpoint for DNA damage. Nature 404, 42-49 (2000).
97. Chang, L. et al. ATM-mediated serine 72 phosphorylation stabilizes ribonucleotide reductase small subunit p53R2 protein against MDM2 to DNA damage. Proc. Natl Acad. Sci. USA 105, 18519-18524 (2008).
98. Lin, Z. P., Belcourt, M. F., Cory, J. G. & Sartorelli, A. C. Stable suppression of the R2 subunit of ribonucleotide reductase by R2-targeted short interference RNA sensitizes p53-/- HCT-116 colon cancer cells to DNA-damaging agents and ribonucleotide reductase inhibitors. J. Biol. Chem. 279, 27030-27038 (2004).
99. Reddy, G. P. & Mathews, C. K. Functional compartmentation of DNA precursors in T4 phage-infected bacteria. J. Biol. Chem. 253, 3461-3467 (1978).
100. Mathews, C. K. & Sinha, N. K. Are DNA precursors concentrated at replication sites? Proc. Natl Acad. Sci. USA 79, 302-306 (1982).
101. Franzolin, E. et al. The deoxynucleotide triphosphohydrolase SAMHD1 is a major regulator of DNA precursor pools in mammalian cells. Proc. Natl Acad. Sci. USA 110, 14272-14277 (2013).
102. Clifford, R. et al. SAMHD1 is mutated recurrently in chronic lymphocytic leukemia and is involved in response to DNA damage. Blood 123, 1021-1031 (2014).
103. Bessman, M. J. et al. Enzymatic synthesis of deoxyribonucleic acid. III. The incorporation of pyrimidine and purine analogues into deoxyribonucleic acid. Proc. Natl Acad. Sci. USA 44, 633-640 (1958).
104. Nick McElhinny, S. A. et al. Genome instability due to ribonucleotide incorporation into DNA. Nat. Chem. Biol. 6, 774-781 (2010).
105. Reijns, M. A. et al. Enzymatic removal of ribonucleotides from DNA is essential for mammalian genome integrity and development. Cell 149, 1008-1022 (2012).
106. Spadari, S. et al. Control of cell division by aphidicolin without adverse effects upon resting cells. Arzneimittelforschung 35, 1108-1116 (1985).
107. Dominguez-Kelly, R. et al. Wee1 controls genomic stability during replication by regulating the Mus81-Eme1 endonuclease. J. Cell Biol. 194, 567-579 (2011).
108. Wilhelm, T. et al. Spontaneous slow replication fork progression elicits mitosis alterations in homologous recombination-deficient mammalian cells. Proc. Natl Acad. Sci. USA 111, 763-768 (2014).
109. Lossaint, G. et al. FANCD2 binds MCM proteins and controls replisome function upon activation of s phase checkpoint signaling. Mol. Cell 51, 678-690 (2013). This study shows that the ATR-FANCD2-FANCI pathway actively regulates fork progression in response to replication stress through a mechanism involving the MCM complex.
110. 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).
111. Taylor, J. H. Increase in DNA replication sites in cells held at the beginning of S phase. Chromosoma 62, 291-300 (1977). This pioneer work shows that slowing the progression of replication forks triggers a compensatory increase in the density of initiation events.
112. Anglana, M., Apiou, F., Bensimon, A. & Debatisse, M. Dynamics of DNA replication in mammalian somatic cells: Nucleotide pool modulates origin choice and interorigin spacing. Cell 114, 385-394 (2003).
113. Marheineke, K. & Hyrien, O. Control of replication origin density and firing time in Xenopus egg extracts: Role of a caffeine-sensitive, ATR-dependent checkpoint. J. Biol. Chem. 279, 28071-28081 (2004).
114. Courbet, S. et al. Replication fork movement sets chromatin loop size and origin choice in mammalian cells. Nature 455, 557-560 (2008).
115. Ge, X. Q., Jackson, D. A. & Blow, J. J. Dormant origins licensed by excess Mcm2-7 are required for human cells to survive replicative stress. Genes Dev. 21, 3331-3341 (2007).
116. Ibarra, A., Schwob, E. & Mendez, J. Excess MCM proteins protect human cells from replicative stress by licensing backup origins of replication. Proc. Natl Acad. Sci. USA 105, 8956-8961 (2008).
117. Kohler, C. et al. Cdc45 is limiting for replication initiation in humans. Cell Cycle 15, 974-985 (2016).
118. Lachaud, C. et al. Ubiquitinated Fancd2 recruits Fan1 to stalled replication forks to prevent genome instability. Science 351, 846-849 (2016).
119. Chen, Y. H. et al. ATR-mediated phosphorylation of FANCI regulates dormant origin firing in response to replication stress. Mol. Cell 58, 323-338 (2015).
120. Sheu, Y. J., Kinney, J. B., Lengronne, A., Pasero, P. & Stillman, B. Domain within the helicase subunit Mcm4 integrates multiple kinase signals to control DNA replication initiation and fork progression. Proc. Natl Acad. Sci. USA 111, E1899-E1908 (2014).
121. Dimitrova, D. S. & Gilbert, D. M. Temporally coordinated assembly and disassembly of replication factories in the absence of DNA synthesis. Nat. Cell Biol. 2, 686-694 (2000).
122. Zachos, G., Rainey, M. D. & Gillespie, D. A. Chk1-deficient tumour cells are viable but exhibit multiple checkpoint and survival defects. EMBO J. 22, 713-723 (2003).
123. Zegerman, P. & Diffley, J. F. DNA replication as a target of the DNA damage checkpoint. DNA Repair (Amst.) 8, 1077-1088 (2009).
124. Ge, X. Q. & Blow, J. J. Chk1 inhibits replication factory activation but allows dormant origin firing in existing factories. J. Cell Biol. 191, 1285-1297 (2010).
125. Rybak, P., Waligorska, A., Bujnowicz, L., Hoang, A. & Dobrucki, J. W. Activation of new replication foci under conditions of replication stress. Cell Cycle 14, 2634-2647 (2015).
126. Eykelenboom, J. K. et al. ATR activates the S-M checkpoint during unperturbed growth to ensure sufficient replication prior to mitotic onset. Cell Rep. 5, 1095-1107 (2013).
127. Buisson, R., Boisvert, J. L., Benes, C. H. & Zou, L. Distinct but concerted roles of ATR, DNA-PK, and Chk1 in countering replication stress during S phase. Mol. Cell 59, 1011-1024 (2015).
128. Couch, F. B. et al. ATR phosphorylates SMARCAL1 to prevent replication fork collapse. Genes Dev. 27, 1610-1623 (2013).
129. Maya-Mendoza, A., Petermann, E., Gillespie, D. A., Caldecott, K. W. & Jackson, D. A. Chk1 regulates the density of active replication origins during the vertebrate S phase. EMBO J. 26, 2719-2731 (2007).
130. Petermann, E., Woodcock, M. & Helleday, T. Chk1 promotes replication fork progression by controlling replication initiation. Proc. Natl Acad. Sci. USA 107, 16090-16095 (2010).
131. Katsuno, Y. et al. Cyclin A-Cdk1 regulates the origin firing program in mammalian cells. Proc. Natl Acad. Sci. USA 106, 3184-3189 (2009).
132. 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).
133. Pfister, S. X. et al. Inhibiting WEE1 selectively kills histone H3K36me3-deficient cancers by dNTP starvation. Cancer Cell 28, 557-568 (2015).
134. Petermann, E., Helleday, T. & Caldecott, K. W. Claspin promotes normal replication fork rates in human cells. Mol. Biol. Cell 19, 2373-2378 (2008).
135. Miotto, B. et al. The RBBP6/ZBTB38/MCM10 axis regulates DNA replication and common fragile site stability. Cell Rep. 7, 575-587 (2014).
136. Choi, H. J. et al. NEK8 links the ATR-regulated replication stress response and S phase CDK activity to renal ciliopathies. Mol. Cell 51, 423-439 (2013).
137. Fu, H. et al. The DNA repair endonuclease Mus81 facilitates fast DNA replication in the absence of exogenous damage. Nat. Commun. 6, 6746 (2015).
138. Giunco, S. et al. Cross talk between EBV and telomerase: The role of TERT and NOTCH2 in the switch of latent/lytic cycle of the virus. Cell Death Dis. 6, e1774 (2015).
139. Park, Y. B. et al. Alterations in the INK4a/ARF locus and their effects on the growth of human osteosarcoma cell lines. Cancer Genet. Cytogenet. 133, 105-111 (2002).
140. Ahuja, A. K. et al. A short G1 phase imposes constitutive replication stress and fork remodelling in mouse embryonic stem cells. Nat. Commun. 7, 10660 (2016). This study shows the massive accumulation of ssDNA gaps, reduced fork speed and frequent fork reversal in mouse embryonic stem cells. These marks of replication stress can be suppressed by delaying the G1-S transition, showing that rapid cell cycle progression challenges genome integrity.
141. 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).
142. Murfuni, I. et al. Survival of the replication checkpoint deficient cells requires MUS81-RAD52 function. PLoS Genet. 9, e1003910 (2013).
143. Thompson, R., Montano, R. & Eastman, A. The Mre11 nuclease is critical for the sensitivity of cells to Chk1 inhibition. PLoS ONE 7, e44021 (2012).
144. Beck, H. et al. Regulators of cyclin-dependent kinases are crucial for maintaining genome integrity in S phase. J. Cell Biol. 188, 629-638 (2010).
145. Pepe, A. & West, S. C. MUS81-EME2 promotes replication fork restart. Cell Rep. 7, 1048-1055 (2014).
146. Buis, J., Stoneham, T., Spehalski, E. & Ferguson, D. O. Mre11 regulates CtIP-dependent double-strand break repair by interaction with CDK2. Nat. Struct. Mol. Biol. 19, 246-252 (2012).
147. Huertas, P. & Jackson, S. P. Human CtIP mediates cell cycle control of DNA end resection and double strand break repair. J. Biol. Chem. 284, 9558-9565 (2009).
148. Peterson, S. E. et al. Cdk1 uncouples CtIP-dependent resection and Rad51 filament formation during M-phase double-strand break repair. J. Cell Biol. 194, 705-720 (2011).
149. Dehe, P. M. et al. Regulation of Mus81-Eme1 Holliday junction resolvase in response to DNA damage. Nat. Struct. Mol. Biol. 20, 598-603 (2013).
150. Falck, J. et al. CDK targeting of NBS1 promotes DNA-end resection, replication restart and homologous recombination. EMBO Rep. 13, 561-568 (2012).
151. Shechter, D., Costanzo, V. & Gautier, J. Regulation of DNA replication by ATR: Signaling in response to DNA intermediates. DNA Repair (Amst.) 3, 901-908 (2004).
152. Syljuasen, R. G. et al. Inhibition of human Chk1 causes increased initiation of DNA replication, phosphorylation of ATR targets, and DNA breakage. Mol. Cell. Biol. 25, 3553-3562 (2005).
153. Niida, H. et al. Essential role of Tip60-dependent recruitment of ribonucleotide reductase at DNA damage sites in DNA repair during G1 phase. Genes Dev. 24, 333-338 (2010). This study shows that, during the G1 phase, RNR re-localizes to repair foci in a TIP60-dependent manner, suggesting that a local increase in dNTP pools favours repair synthesis.
154. 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).
155. Kolinjivadi, A. M. et al. Moonlighting at replication forks - A new life for homologous recombination proteins BRCA1, BRCA2 and RAD51. FEBS Lett. 591, 1083-1100 (2017).
156. Toledo, L. I. et al. ATR prohibits replication catastrophe by preventing global exhaustion of RPA. Cell 155, 1088-1103 (2013). This study shows that RPA is present in limiting amount in cells, which compromises ssDNA protection upon stringent replication stress, especially in cells deficient in ATR.
157. Hossain, M. & Stillman, B. Opposing roles for DNA replication initiator proteins ORC1 and CDC6 in control of Cyclin E gene transcription. eLife 5, e12785 (2016). This study shows that ORC1 and CDC6 have opposing effects in controlling the level of cyclin E expression. Cyclin E level then regulates cell passage through the restriction point and S phase onset, which ensures genome stability.
158. Sanjiv, K. et al. Cancer-specific synthetic lethality between ATR and CHK1 kinase activities. Cell Rep. 14, 298-309 (2016).
159. Mokrani-Benhelli, H. et al. Primary microcephaly, impaired DNA replication, and genomic instability caused by compound heterozygous ATR mutations. Hum. Mutat. 34, 374-384 (2013).
160. Murga, M. et al. A mouse model of ATR-Seckel shows embryonic replicative stress and accelerated aging. Nat. Genet. 41, 891-898 (2009).
161. Lopez-Contreras, A. J. et al. Increased Rrm2 gene dosage reduces fragile site breakage and prolongs survival of ATR mutant mice. Genes Dev. 29, 690-695 (2015). This study shows that extra-alleles of the RNR regulatory subunit RRM2 enhance RNR activity, resulting in reduced chromosomal breaks at fragile sites, and extending the lifespan of ATR-mutant mice.
162. Hills, S. A. & Diffley, J. F. DNA replication and oncogene-induced replicative stress. Curr. Biol. 24, R435-R444 (2014).
163. Neelsen, K. J., Zanini, I. M., Herrador, R. & Lopes, M. Oncogenes induce genotoxic stress by mitotic processing of unusual replication intermediates. J. Cell Biol. 200, 699-708 (2013).
164. Maya-Mendoza, A. et al. Myc and Ras oncogenes engage different energy metabolism programs and evoke distinct patterns of oxidative and DNA replication stress. Mol. Oncol. 9, 601-616 (2015).
165. Srinivasan, S. V., Dominguez-Sola, D., Wang, L. C., Hyrien, O. & Gautier, J. Cdc45 is a critical effector of myc-dependent DNA replication stress. Cell Rep. 3, 1629-1639 (2013).
166. Jones, R. M. et al. Increased replication initiation and conflicts with transcription underlie Cyclin E-induced replication stress. Oncogene 32, 3744-3753 (2013).
167. Dominguez-Sola, D. et al. Non-transcriptional control of DNA replication by c-Myc. Nature 448, 445-451 (2007).
168. Di Micco, R. et al. Oncogene-induced senescence is a DNA damage response triggered by DNA hyper-replication. Nature 444, 638-642 (2006).
169. Ohtsubo, M., Theodoras, A. M., Schumacher, J., Roberts, J. M. & Pagano, M. Human cyclin E, a nuclear protein essential for the G1-to-S phase transition. Mol. Cell. Biol. 15, 2612-2624 (1995).
170. Costantino, L. et al. Break-induced replication repair of damaged forks induces genomic duplications in human cells. Science 343, 88-91 (2014). This report shows that POLD3 is crucial for the survival of U2OS cells overexpressing cyclin E, suggesting that BIR is required to repair broken forks. Strikingly, POLD3-dependent repair generates gross chromosome rearrangements similar to those seen in human breast and ovarian cancers.
171. Bartkova, J. et al. Oncogene-induced senescence is part of the tumorigenesis barrier imposed by DNA damage checkpoints. Nature 444, 633-637 (2006).
172. Aird, K. M. et al. Suppression of nucleotide metabolism underlies the establishment and maintenance of oncogene-induced senescence. Cell Rep. 3, 1252-1265 (2013).
173. Miron, K., Golan-Lev, T., Dvir, R., Ben-David, E. & Kerem, B. Oncogenes create a unique landscape of fragile sites. Nat. Commun. 6, 7094 (2015). The authors show that the overexpression of RAS or cyclin E in the same cells induces a set of fragile sites specific to each oncogene. These sites nest within large genes and colocalize with cancer breakpoints.
174. Lee, A. C. et al. Ras proteins induce senescence by altering the intracellular levels of reactive oxygen species. J. Biol. Chem. 274, 7936-7940 (1999).
175. Vafa, O. et al. c-Myc can induce DNA damage, increase reactive oxygen species, and mitigate p53 function: A mechanism for oncogene-induced genetic instability. Mol. Cell 9, 1031-1044 (2002).
176. Weyemi, U. et al. ROS-generating NADPH oxidase NOX4 is a critical mediator in oncogenic H-Ras-induced DNA damage and subsequent senescence. Oncogene 31, 1117-1129 (2012).
177. Ogrunc, M. et al. Oncogene-induced reactive oxygen species fuel hyperproliferation and DNA damage response activation. Cell Death Differ. 21, 998-1012 (2014).
178. Guo, Z., Kozlov, S., Lavin, M. F., Person, M. D. & Paull, T. T. ATM activation by oxidative stress. Science 330, 517-521 (2010).
179. Torres-Rosell, J. et al. Anaphase onset before complete DNA replication with intact checkpoint responses. Science 315, 1411-1415 (2007).
180. Naim, V. & Rosselli, F. The FANC pathway and mitosis: A replication legacy. Cell Cycle 8, 2907-2911 (2009).
181. Lukas, C. et al. 53BP1 nuclear bodies form around DNA lesions generated by mitotic transmission of chromosomes under replication stress. Nat. Cell Biol. 13, 243-253 (2011).
182. Lemmens, B., van Schendel, R. & Tijsterman, M. Mutagenic consequences of a single G-quadruplex demonstrate mitotic inheritance of DNA replication fork barriers. Nat. Commun. 6, 8909 (2015).
183. Maciejowski, J., Li, Y., Bosco, N., Campbell, P. J. & de Lange, T. Chromothripsis and kataegis induced by telomere crisis. Cell 163, 1641-1654 (2015).
184. Crosetto, N. et al. Nucleotide-resolution DNA double-strand break mapping by next-generation sequencing. Nat. Methods 10, 361-365 (2013).
185. 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).
186. Ying, S. et al. MUS81 promotes common fragile site expression. Nat. Cell Biol. 15, 1001-1007 (2013). References 185 and 186 show that sequences remaining under-replicated in mitotic cells, notably at CFSs, are converted into DNA breaks by specialized nucleases such as MUS81-EME1.
187. 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).
188. Bergoglio, V. et al. DNA synthesis by Pol eta promotes fragile site stability by preventing under-replicated DNA in mitosis. J. Cell Biol. 201, 395-408 (2013).
189. Minocherhomji, S. et al. Replication stress activates DNA repair synthesis in mitosis. Nature 528, 286-290 (2015).
190. Despras, E. et al. Rad18-dependent SUMOylation of human specialized DNA polymerase eta is required to prevent under-replicated DNA. Nat. Commun. 7, 13326 (2016).
191. Fernandez-Vidal, A. et al. A role for DNA polymerase theta in the timing of DNA replication. Nat. Commun. 5, 4285 (2014).
192. Hayano, M. et al. Rif1 is a global regulator of timing of replication origin firing in fission yeast. Genes Dev. 26, 137-150 (2012).
193. Dave, A., Cooley, C., Garg, M. & Bianchi, A. Protein phosphatase 1 recruitment by Rif1 regulates DNA replication origin firing by counteracting DDK activity. Cell Rep. 7, 53-61 (2014).
194. Mattarocci, S. et al. Rif1 controls DNA replication timing in yeast through the PP1 phosphatase Glc7. Cell Rep. 7, 62-69 (2014).
195. Cornacchia, D. et al. Mouse Rif1 is a key regulator of the replication-timing programme in mammalian cells. EMBO J. 31, 3678-3690 (2012).
196. Yamazaki, S. et al. Rif1 regulates the replication timing domains on the human genome. EMBO J. 31, 3667-3677 (2012).
197. Foti, R. et al. Nuclear architecture organized by Rif1 underpins the replication-timing program. Mol. Cell 61, 260-273 (2016).
198. Hassan-Zadeh, V. et al. USF binding sequences from the HS4 insulator element impose early replication timing on a vertebrate replicator. PLoS Biol. 10, e1001277 (2012).
199. Therizols, P. et al. Chromatin decondensation is sufficient to alter nuclear organization in embryonic stem cells. Science 346, 1238-1242 (2014).
200. Hansel-Hertsch, R. et al. G-Quadruplex structures mark human regulatory chromatin. Nat. Genet. 48, 1267-1272 (2016).
201. Chambers, V. S. et al. High-throughput sequencing of DNA G-quadruplex structures in the human genome. Nat. Biotechnol. 33, 877-881 (2015).
202. Biffi, G., Tannahill, D., McCafferty, J. & Balasubramanian, S. Quantitative visualization of DNA G-quadruplex structures in human cells. Nat. Chem. 5, 182-186 (2013).
203. Henderson, A. et al. Detection of G-quadruplex DNA in mammalian cells. Nucleic Acids Res. 42, 860-869 (2014).
204. Castillo Bosch, P. et al. FANCJ promotes DNA synthesis through G-quadruplex structures. EMBO J. 33, 2521-2533 (2014).
205. Ribeyre, C. et al. The yeast Pif1 helicase prevents genomic instability caused by G-quadruplex-forming CEB1 sequences in vivo. PLoS Genet. 5, e1000475 (2009).
206. Piazza, A. et al. Genetic instability triggered by G-quadruplex interacting Phen-DC compounds in Saccharomyces cerevisiae. Nucleic Acids Res. 38, 4337-4348 (2010).
207. Piazza, A. et al. Stimulation of gross chromosomal rearrangements by the human CEB1 and CEB25 minisatellites in Saccharomyces cerevisiae depends on G-quadruplexes or Cdc13. PLoS Genet. 8, e1003033 (2012).
208. Lopes, J. et al. G-Quadruplex-induced instability during leading-strand replication. EMBO J. 30, 4033-4046 (2011).
209. Badie, S. et al. BRCA2 acts as a RAD51 loader to facilitate telomere replication and capping. Nat. Struct. Mol. Biol. 17, 1461-1469 (2010).
210. Zimmer, J. et al. Targeting BRCA1 and BRCA2 deficiencies with G-quadruplex-interacting compounds. Mol. Cell 61, 449-460 (2016).
211. 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).
212. Schiavone, D. et al. Determinants of G quadruplex-induced epigenetic instability in REV1-deficient cells. EMBO J. 33, 2507-2520 (2014).
213. Papadopoulou, C., Guilbaud, G., Schiavone, D. & Sale, J. E. Nucleotide pool depletion induces G-quadruplex-dependent perturbation of gene expression. Cell Rep. 13, 2491-2503 (2015).
214. Clynes, D. & Gibbons, R. J. ATRX and the replication of structured DNA. Curr. Opin. Genet. Dev. 23, 289-294 (2013).
215. Schiavone, D. et al. PrimPol is required for replicative tolerance of G quadruplexes in vertebrate cells. Mol. Cell 61, 161-169 (2016).
216. Pontarin, G. et al. Ribonucleotide reduction is a cytosolic process in mammalian cells independently of DNA damage. Proc. Natl Acad. Sci. USA 105, 17801-17806 (2008).
217. Prem veer Reddy, G. & Pardee, A. B. Multienzyme complex for metabolic channeling in mammalian DNA replication. Proc. Natl Acad. Sci. USA 77, 3312-3316 (1980). This pioneer work shows that enzymes involved in dNTP and DNA synthesis physically interact, suggesting that precursors can be channelled to replication foci.
218. Baril, E., Baril, B. & Laszlo, J. DNA polymerases in normal and regenerating rat liver and hepatomas: Targets for chemotherapy. Adv. Enzyme Regul. 12, 355-372 (1974).
219. Wickremasinghe, R. G., Yaxley, J. C. & Hoffbrand, A. V. Gel filtration of a complex of DNA polymerase and DNA precursor-synthesizing enzymes from a human lymphoblastoid cell line. Biochim. Biophys. Acta 740, 243-248 (1983).
220. Alabert, C. et al. Nascent chromatin capture proteomics determines chromatin dynamics during DNA replication and identifies unknown fork components. Nat. Cell Biol. 16, 281-293 (2014).
221. Taricani, L., Shanahan, F., Malinao, M. C., Beaumont, M. & Parry, D. A. Functional approach reveals a genetic and physical interaction between ribonucleotide reductase and CHK1 in mammalian cells. PLoS ONE 9, e111714 (2014).
222. veer Reddy, G. P. & Pardee, A. B. Coupled ribonucleoside diphosphate reduction, channeling, and incorporation into DNA of mammalian cells. J. Biol. Chem. 257, 12526-12531 (1982).
223. Zhang, Y. W., Jones, T. L., Martin, S. E., Caplen, N. J. & Pommier, Y. Implication of checkpoint kinase-dependent up-regulation of ribonucleotide reductase R2 in DNA damage response. J. Biol. Chem. 284, 18085-18095 (2009).
224. Sun, Y., Jiang, X., Chen, S., Fernandes, N. & Price, B. D. A role for the Tip60 histone acetyltransferase in the acetylation and activation of ATM. Proc. Natl Acad. Sci. USA 102, 13182-13187 (2005).
225. Altmeyer, M. et al. Liquid demixing of intrinsically disordered proteins is seeded by poly(ADP-ribose). Nat. Commun. 6, 8088 (2015). This study shows that DNA breaks are the sites of the transient and reversible assembly of proteins with large intrinsically disordered domains, which creates dynamic nuclear pseudo-compartments.
226. Ragland, R. L. et al. RNF4 and PLK1 are required for replication fork collapse in ATR-deficient cells. Genes Dev. 27, 2259-2273 (2013).
227. Ammazzalorso, F., Pirzio, L. M., Bignami, M., Franchitto, A. & Pichierri, P. ATR and ATM differently regulate WRN to prevent DSBs at stalled replication forks and promote replication fork recovery. EMBO J. 29, 3156-3169 (2010).
228. 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).
229. 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).
230. Kais, Z. et al. FANCD2 maintains fork stability in BRCA1/2-deficient tumors and promotes alternative end-joining DNA repair. Cell Rep. 15, 2488-2499 (2016).
231. Michl, J., Zimmer, J., Buffa, F. M., McDermott, U. & Tarsounas, M. FANCD2 limits replication stress and genome instability in cells lacking BRCA2. Nat. Struct. Mol. Biol. 23, 755-757 (2016).
232. Ceccaldi, R., Sarangi, P. & D'Andrea, A. D. The Fanconi anaemia pathway: New players and new functions. Nat. Rev. Mol. Cell Biol. 17, 337-349 (2016).
233. Guervilly, J. H., Mace-Aime, G. & Rosselli, F. Loss of CHK1 function impedes DNA damage-induced FANCD2 monoubiquitination but normalizes the abnormal G2 arrest in Fanconi anemia. Hum. Mol. Genet. 17, 679-689 (2008).