Role of Protein Linked DNA Breaks in Cancer

  • Walaa R. AllamEmail author
  • Mohamed E. Ashour
  • Amr A. Waly
  • Sherif El-KhamisyEmail author
Part of the Advances in Experimental Medicine and Biology book series (AEMB, volume 1007)


Topoisomerases are a group of specialized enzymes that function to maintain DNA topology by introducing transient DNA breaks during transcription and replication. As a result of abortive topoisomerases activity, topoisomerases catalytic intermediates may be trapped on the DNA forming topoisomerase cleavage complexes (Topcc). Topoisomerases trapping on the DNA is the mode of action of several anticancer drugs, it lead to formation of protein linked DAN breaks (PDBs). PDBs are now considered as one of the most dangerous forms of endogenous DNA damage and a major threat to genomic stability. The repair of PDBs involves both the sensing and repair pathways. Unsuccessful repair of PDBs leads to different signs of genomic instabilities such as chromosomal rearrangements and cancer predisposition. In this chapter we will summarize the role of topoisomerases induced PDBs, identification and signaling, repair, role in transcription. We will also discuss the role of PDBs in cancer with a special focus on prostate cancer.


Protein linked DNA breaks Topoisomerases DAN repair Topoisomerases poisons Genome integrity 


  1. 1.
    Schilsky RL (1996) Methotrexate: an effective agent for treating cancer and building careers. The polyglutamate era. Stem Cells 14:29–32. doi: 10.1002/stem.140029 PubMedCrossRefGoogle Scholar
  2. 2.
    Miura S, Yoshimura Y, Satoh H, Izuta S (2001) The antitumor mechanism of 1-(2-deoxy-2-fluoro-4-thio-beta-D-arabinofuranosyl)-cytosine: effects of its triphosphate on mammalian DNA polymerases. Jpn J Cancer Res 92:562–567PubMedCrossRefGoogle Scholar
  3. 3.
    Povirk LF (1996) DNA damage and mutagenesis by radiomimetic DNA-cleaving agents: bleomycin, neocarzinostatin and other enediynes. Mutat Res 355:71–89PubMedCrossRefGoogle Scholar
  4. 4.
    Zorbas H, Keppler BK (2005) Cisplatin damage: are DNA repair proteins saviors or traitors to the cell? Chembiochem 6:1157–1166. doi: 10.1002/cbic.200400427 PubMedCrossRefGoogle Scholar
  5. 5.
    Chen T, Stephens PA, Middleton FK, Curtin NJ (2012) Targeting the S and G2 checkpoint to treat cancer. Drug Discov Today 17:194–202. doi: 10.1016/j.drudis.2011.12.009 PubMedCrossRefGoogle Scholar
  6. 6.
    Biss M, Xiao W (2012) Selective tumor killing based on specific DNA-damage response deficiencies. Cancer Biol Ther 13:239–246. doi: 10.4161/cbt.18921 PubMedPubMedCentralCrossRefGoogle Scholar
  7. 7.
    Yang X, Wood PA, Hrushesky WJ (2010) Mammalian TIMELESS is required for ATM-dependent CHK2 activation and G2/M checkpoint control. J Biol Chem 285:3030–3034. doi: 10.1074/jbc.M109.050237 PubMedCrossRefGoogle Scholar
  8. 8.
    Bakkenist CJ, Kastan MB (2003) DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation. Nature 421:499–506. doi: 10.1038/nature01368 PubMedCrossRefGoogle Scholar
  9. 9.
    Lee JH, Paull TT (2007) Activation and regulation of ATM kinase activity in response to DNA double-strand breaks. Oncogene 26:7741–7748. doi: 10.1038/sj.onc.1210872 PubMedCrossRefGoogle Scholar
  10. 10.
    Buscemi G et al (2001) Chk2 activation dependence on Nbs1 after DNA damage. Mol Cell Biol 21:5214–5222. doi: 10.1128/MCB.21.15.5214-5222.2001 PubMedPubMedCentralCrossRefGoogle Scholar
  11. 11.
    Nakanishi K et al (2002) Interaction of FANCD2 and NBS1 in the DNA damage response. Nat Cell Biol 4:913–920. doi: 10.1038/ncb879 PubMedCrossRefGoogle Scholar
  12. 12.
    Uziel T et al (2003) Requirement of the MRN complex for ATM activation by DNA damage. EMBO J 22:5612–5621. doi: 10.1093/emboj/cdg541 PubMedPubMedCentralCrossRefGoogle Scholar
  13. 13.
    Lee JH, Paull TT (2005) ATM activation by DNA double-strand breaks through the Mre11-Rad50-Nbs1 complex. Science 308:551–554. doi: 10.1126/science.1108297 PubMedCrossRefGoogle Scholar
  14. 14.
    Woods D, Turchi JJ (2013) Chemotherapy induced DNA damage response: convergence of drugs and pathways. Cancer Biol Ther 14:379–389. doi: 10.4161/cbt.23761 PubMedPubMedCentralCrossRefGoogle Scholar
  15. 15.
    Liu X et al (2012) Overlapping functions between XLF repair protein and 53BP1 DNA damage response factor in end joining and lymphocyte development. Proc Natl Acad Sci U S A 109:3903–3908. doi: 10.1073/pnas.1120160109 PubMedPubMedCentralCrossRefGoogle Scholar
  16. 16.
    McGowan CH, Russell P (2004) The DNA damage response: sensing and signaling. Curr Opin Cell Biol 16:629–633. doi: 10.1016/ PubMedCrossRefGoogle Scholar
  17. 17.
    Harrison JC, Haber JE (2006) Surviving the breakup: the DNA damage checkpoint. Annu Rev Genet 40:209–235. doi: 10.1146/annurev.genet.40.051206.105231 PubMedCrossRefGoogle Scholar
  18. 18.
    Stewart L, Redinbo MR, Qiu X, Hol WG, Champoux JJ (1998) A model for the mechanism of human topoisomerase I. Science 279:1534–1541PubMedCrossRefGoogle Scholar
  19. 19.
    Staker BL et al (2002) The mechanism of topoisomerase I poisoning by a camptothecin analog. Proc Natl Acad Sci U S A 99:15387–15392. doi: 10.1073/pnas.242259599 PubMedPubMedCentralCrossRefGoogle Scholar
  20. 20.
    Wang JC (2002) Cellular roles of DNA topoisomerases: a molecular perspective. Nat Rev Mol Cell Biol 3:430–440. doi: 10.1038/nrm831 PubMedCrossRefGoogle Scholar
  21. 21.
    Flatten K et al (2005) The role of checkpoint kinase 1 in sensitivity to topoisomerase I poisons. J Biol Chem 280:14349–14355. doi: 10.1074/jbc.M411890200 PubMedCrossRefGoogle Scholar
  22. 22.
    Wu CS et al (2014) SUMOylation of ATRIP potentiates DNA damage signaling by boosting multiple protein interactions in the ATR pathway. Genes Dev 28:1472–1484. doi: 10.1101/gad.238535.114 PubMedPubMedCentralCrossRefGoogle Scholar
  23. 23.
    Wu CS, Zou L (2016) The SUMO (Small Ubiquitin-like Modifier) ligase PIAS3 primes ATR for checkpoint activation. J Biol Chem 291:279–290. doi: 10.1074/jbc.M115.691170 PubMedCrossRefGoogle Scholar
  24. 24.
    Liu S et al (2013) PIAS3 promotes homology-directed repair and distal non-homologous end joining. Oncol Lett 6:1045–1048. doi: 10.3892/ol.2013.1472 PubMedPubMedCentralGoogle Scholar
  25. 25.
    Sakasai R, Teraoka H, Tibbetts RS (2010) Proteasome inhibition suppresses DNA-dependent protein kinase activation caused by camptothecin. DNA Repair (Amst) 9:76–82. doi: 10.1016/j.dnarep.2009.10.008 CrossRefGoogle Scholar
  26. 26.
    Kocher S, Spies-Naumann A, Kriegs M, Dahm-Daphi J, Dornreiter I (2013) ATM is required for the repair of Topotecan-induced replication-associated double-strand breaks. Radiother Oncol 108:409–414. doi: 10.1016/j.radonc.2013.06.024 PubMedCrossRefGoogle Scholar
  27. 27.
    Serrano MA et al (2013) DNA-PK, ATM and ATR collaboratively regulate p53-RPA interaction to facilitate homologous recombination DNA repair. Oncogene 32:2452–2462. doi: 10.1038/onc.2012.257 PubMedCrossRefGoogle Scholar
  28. 28.
    Das BB et al (2009) Optimal function of the DNA repair enzyme TDP1 requires its phosphorylation by ATM and/or DNA-PK. EMBO J 28:3667–3680. doi: 10.1038/emboj.2009.302 PubMedPubMedCentralCrossRefGoogle Scholar
  29. 29.
    Chiang SC, Carroll J, El-Khamisy SF (2010) TDP1 serine 81 promotes interaction with DNA ligase IIIalpha and facilitates cell survival following DNA damage. Cell Cycle 9:588–595. doi: 10.4161/cc.9.3.10598 PubMedCrossRefGoogle Scholar
  30. 30.
    Zhou Y et al (2017) Regulation of the DNA damage response by DNA-PKcs inhibitory phosphorylation of ATM. Mol Cell 65:91–104. doi: 10.1016/j.molcel.2016.11.004 PubMedCrossRefGoogle Scholar
  31. 31.
    Sordet O et al (2009) Ataxia telangiectasia mutated activation by transcription- and topoisomerase I-induced DNA double-strand breaks. EMBO Rep 10:887–893. doi: 10.1038/embor.2009.97 PubMedPubMedCentralCrossRefGoogle Scholar
  32. 32.
    Cristini A et al (2016) DNA-PK triggers histone ubiquitination and signaling in response to DNA double-strand breaks produced during the repair of transcription-blocking topoisomerase I lesions. Nucleic Acids Res 44:1161–1178. doi: 10.1093/nar/gkv1196 PubMedCrossRefGoogle Scholar
  33. 33.
    Alagoz M, Chiang SC, Sharma A, El-Khamisy SF (2013) ATM deficiency results in accumulation of DNA-topoisomerase I covalent intermediates in neural cells. PLoS One 8:e58239. doi: 10.1371/journal.pone.0058239 PubMedPubMedCentralCrossRefGoogle Scholar
  34. 34.
    Huelsenbeck SC et al (2012) Rac1 protein signaling is required for DNA damage response stimulated by topoisomerase II poisons. J Biol Chem 287:38590–38599. doi: 10.1074/jbc.M112.377903 PubMedPubMedCentralCrossRefGoogle Scholar
  35. 35.
    Forrest RA et al (2012) Activation of DNA damage response pathways as a consequence of anthracycline-DNA adduct formation. Biochem Pharmacol 83:1602–1612. doi: 10.1016/j.bcp.2012.02.026 PubMedCrossRefGoogle Scholar
  36. 36.
    Maede Y et al (2014) Differential and common DNA repair pathways for topoisomerase I- and II-targeted drugs in a genetic DT40 repair cell screen panel. Mol Cancer Ther 13:214–220. doi: 10.1158/1535-7163.MCT-13-0551 PubMedCrossRefGoogle Scholar
  37. 37.
    Rossi R, Lidonnici MR, Soza S, Biamonti G, Montecucco A (2006) The dispersal of replication proteins after Etoposide treatment requires the cooperation of Nbs1 with the ataxia telangiectasia Rad3-related/Chk1 pathway. Cancer Res 66:1675–1683. doi: 10.1158/0008-5472.CAN-05-2741 PubMedCrossRefGoogle Scholar
  38. 38.
    Quennet V, Beucher A, Barton O, Takeda S, Lobrich M (2011) CtIP and MRN promote non-homologous end-joining of etoposide-induced DNA double-strand breaks in G1. Nucleic Acids Res 39:2144–2152. doi: 10.1093/nar/gkq1175 PubMedCrossRefGoogle Scholar
  39. 39.
    Alvarez-Quilon A et al (2014) ATM specifically mediates repair of double-strand breaks with blocked DNA ends. Nat Commun 5:3347. doi: 10.1038/ncomms4347 PubMedPubMedCentralCrossRefGoogle Scholar
  40. 40.
    Korwek Z et al (2012) Inhibition of ATM blocks the etoposide-induced DNA damage response and apoptosis of resting human T cells. DNA Repair (Amst) 11:864–873. doi: 10.1016/j.dnarep.2012.08.006 CrossRefGoogle Scholar
  41. 41.
    Cliby WA, Lewis KA, Lilly KK, Kaufmann SH (2002) S phase and G2 arrests induced by topoisomerase I poisons are dependent on ATR kinase function. J Biol Chem 277:1599–1606. doi: 10.1074/jbc.M106287200 PubMedCrossRefGoogle Scholar
  42. 42.
    Costanzo V et al (2003) An ATR- and Cdc7-dependent DNA damage checkpoint that inhibits initiation of DNA replication. Mol Cell 11:203–213PubMedCrossRefGoogle Scholar
  43. 43.
    Siu WY et al (2004) Topoisomerase poisons differentially activate DNA damage checkpoints through ataxia-telangiectasia mutated-dependent and -independent mechanisms. Mol Cancer Ther 3:621–632PubMedGoogle Scholar
  44. 44.
    Ho CC et al (2005) The relative contribution of CHK1 and CHK2 to Adriamycin-induced checkpoint. Exp Cell Res 304:1–15. doi: 10.1016/j.yexcr.2004.10.016 PubMedCrossRefGoogle Scholar
  45. 45.
    Martensson S, Nygren J, Osheroff N, Hammarsten O (2003) Activation of the DNA-dependent protein kinase by drug-induced and radiation-induced DNA strand breaks. Radiat Res 160:291–301PubMedCrossRefGoogle Scholar
  46. 46.
    Das BB et al (2014) PARP1-TDP1 coupling for the repair of topoisomerase I-induced DNA damage. Nucleic Acids Res 42:4435–4449. doi: 10.1093/nar/gku088 PubMedPubMedCentralCrossRefGoogle Scholar
  47. 47.
    Das SK et al (2016) Poly(ADP-ribose) polymers regulate DNA topoisomerase I (Top1) nuclear dynamics and camptothecin sensitivity in living cells. Nucleic Acids Res 44:8363–8375. doi: 10.1093/nar/gkw665 PubMedPubMedCentralCrossRefGoogle Scholar
  48. 48.
    Cuya SM, Comeaux EQ, Wanzeck K, Yoon KJ, van Waardenburg RC (2016) Dysregulated human Tyrosyl-DNA phosphodiesterase I acts as cellular toxin. Oncotarget 7:86660–86674. doi: 10.18632/oncotarget.13528 PubMedPubMedCentralGoogle Scholar
  49. 49.
    Takashima H et al (2002) Mutation of TDP1, encoding a topoisomerase I-dependent DNA damage repair enzyme, in spinocerebellar ataxia with axonal neuropathy. Nat Genet 32:267–272. doi: 10.1038/ng987 PubMedCrossRefGoogle Scholar
  50. 50.
    He X et al (2007) Mutation of a conserved active site residue converts tyrosyl-DNA phosphodiesterase I into a DNA topoisomerase I-dependent poison. J Mol Biol 372:1070–1081. doi: 10.1016/j.jmb.2007.07.055 PubMedCrossRefGoogle Scholar
  51. 51.
    Huang SY et al (2013) TDP1 repairs nuclear and mitochondrial DNA damage induced by chain-terminating anticancer and antiviral nucleoside analogs. Nucleic Acids Res 41:7793–7803. doi: 10.1093/nar/gkt483 PubMedPubMedCentralCrossRefGoogle Scholar
  52. 52.
    Hudson JJ, Chiang SC, Wells OS, Rookyard C, El-Khamisy SF (2012) SUMO modification of the neuroprotective protein TDP1 facilitates chromosomal single-strand break repair. Nat Commun 3:733. doi: 10.1038/ncomms1739 PubMedPubMedCentralCrossRefGoogle Scholar
  53. 53.
    El-Khamisy SF, Hartsuiker E, Caldecott KW (2007) TDP1 facilitates repair of ionizing radiation-induced DNA single-strand breaks. DNA Repair 6:1485–1495. doi: 10.1016/j.dnarep.2007.04.015 PubMedCrossRefGoogle Scholar
  54. 54.
    Alagoz M, Wells OS, El-Khamisy SF (2014) TDP1 deficiency sensitizes human cells to base damage via distinct topoisomerase I and PARP mechanisms with potential applications for cancer therapy. Nucleic Acids Res 42:3089–3103. doi: 10.1093/nar/gkt1260 PubMedCrossRefGoogle Scholar
  55. 55.
    Davies DR, Interthal H, Champoux JJ, Hol WG (2002) The crystal structure of human tyrosyl-DNA phosphodiesterase, Tdp1. Structure 10:237–248PubMedCrossRefGoogle Scholar
  56. 56.
    Comeaux EQ, van Waardenburg RC (2014) Tyrosyl-DNA phosphodiesterase I resolves both naturally and chemically induced DNA adducts and its potential as a therapeutic target. Drug Metab Rev 46:494–507. doi: 10.3109/03602532.2014.971957 PubMedCrossRefGoogle Scholar
  57. 57.
    Malik M, Nitiss KC, Enriquez-Rios V, Nitiss JL (2006) Roles of nonhomologous end-joining pathways in surviving topoisomerase II-mediated DNA damage. Mol Cancer Ther 5:1405–1414. doi: 10.1158/1535-7163.MCT-05-0263 PubMedCrossRefGoogle Scholar
  58. 58.
    Gomez-Herreros F et al (2013) TDP2-dependent non-homologous end-joining protects against topoisomerase II-induced DNA breaks and genome instability in cells and in vivo. PLoS Genet 9:e1003226. doi: 10.1371/journal.pgen.1003226 PubMedPubMedCentralCrossRefGoogle Scholar
  59. 59.
    Zeng Z et al (2012) TDP2 promotes repair of topoisomerase I-mediated DNA damage in the absence of TDP1. Nucleic Acids Res 40:8371–8380. doi: 10.1093/nar/gks622 PubMedPubMedCentralCrossRefGoogle Scholar
  60. 60.
    Bian K et al (2016) ERK3 regulates TDP2-mediated DNA damage response and chemoresistance in lung cancer cells. Oncotarget 7:6665–6675. doi: 10.18632/oncotarget.6682
  61. 61.
    Pommier Y et al (2014) Tyrosyl-DNA-phosphodiesterases (TDP1 and TDP2). DNA Repair 19:114–129. doi: 10.1016/j.dnarep.2014.03.020 PubMedPubMedCentralCrossRefGoogle Scholar
  62. 62.
    Gao R, Huang SY, Marchand C, Pommier Y (2012) Biochemical characterization of human tyrosyl-DNA phosphodiesterase 2 (TDP2/TTRAP): a Mg(2+)/Mn(2+)-dependent phosphodiesterase specific for the repair of topoisomerase cleavage complexes. J Biol Chem 287:30842–30852. doi: 10.1074/jbc.M112.393983 PubMedPubMedCentralCrossRefGoogle Scholar
  63. 63.
    Gao R et al (2014) Proteolytic degradation of topoisomerase II (Top2) enables the processing of Top2.DNA and Top2.RNA covalent complexes by tyrosyl-DNA-phosphodiesterase 2 (TDP2). J Biol Chem 289:17960–17969. doi: 10.1074/jbc.M114.565374 PubMedPubMedCentralCrossRefGoogle Scholar
  64. 64.
    Heo J et al (2015) TDP1 promotes assembly of non-homologous end joining protein complexes on DNA. DNA Repair 30:28–37. doi: 10.1016/j.dnarep.2015.03.003 PubMedPubMedCentralCrossRefGoogle Scholar
  65. 65.
    Elsayed W, El-Shafie L, Hassan MK, Farag MA, El-Khamisy SF (2016) Isoeugenol is a selective potentiator of camptothecin cytotoxicity in vertebrate cells lacking TDP1. Sci Rep 6:26626. doi: 10.1038/srep26626 PubMedPubMedCentralCrossRefGoogle Scholar
  66. 66.
    Beck DE et al (2016) Synthesis and biological evaluation of new fluorinated and chlorinated indenoisoquinoline topoisomerase I poisons. Bioorg Med Chem 24:1469–1479. doi: 10.1016/j.bmc.2016.02.015 PubMedPubMedCentralCrossRefGoogle Scholar
  67. 67.
    Liao S, Tammaro M, Yan H (2016) The structure of ends determines the pathway choice and Mre11 nuclease dependency of DNA double-strand break repair. Nucleic Acids Res 44:5689–5701. doi: 10.1093/nar/gkw274 PubMedPubMedCentralCrossRefGoogle Scholar
  68. 68.
    Aparicio T, Baer R, Gottesman M, Gautier J (2016) MRN, CtIP, and BRCA1 mediate repair of topoisomerase II-DNA adducts. J Cell Biol 212:399–408. doi: 10.1083/jcb.201504005 PubMedPubMedCentralCrossRefGoogle Scholar
  69. 69.
    Wang J et al (2016) Loss of CtIP disturbs homologous recombination repair and sensitizes breast cancer cells to PARP inhibitors. Oncotarget 7:7701–7714. doi: 10.18632/oncotarget.6715
  70. 70.
    Chanut P, Britton S, Coates J, Jackson SP, Calsou P (2016) Coordinated nuclease activities counteract Ku at single-ended DNA double-strand breaks. Nat Commun 7:12889. doi: 10.1038/ncomms12889 PubMedPubMedCentralCrossRefGoogle Scholar
  71. 71.
    Guirouilh-Barbat J et al (2016) 53BP1 protects against CtIP-dependent capture of ectopic chromosomal sequences at the junction of distant double-strand breaks. PLoS Genet 12:e1006230. doi: 10.1371/journal.pgen.1006230 PubMedPubMedCentralCrossRefGoogle Scholar
  72. 72.
    Ahrabi S et al (2016) A role for human homologous recombination factors in suppressing microhomology-mediated end joining. Nucleic Acids Res 44:5743–5757. doi: 10.1093/nar/gkw326 PubMedPubMedCentralCrossRefGoogle Scholar
  73. 73.
    Anand R, Ranjha L, Cannavo E, Cejka P (2016) Phosphorylated CtIP functions as a co-factor of the MRE11-RAD50-NBS1 endonuclease in DNA end resection. Mol Cell 64:940–950. doi: 10.1016/j.molcel.2016.10.017 PubMedCrossRefGoogle Scholar
  74. 74.
    Chen YJ et al (2016) S. cerevisiae Mre11 recruits conjugated SUMO moieties to facilitate the assembly and function of the Mre11-Rad50-Xrs2 complex. Nucleic Acids Res 44:2199–2213. doi: 10.1093/nar/gkv1523 PubMedPubMedCentralCrossRefGoogle Scholar
  75. 75.
    Deshpande RA, Lee JH, Arora S, Paull TT (2016) Nbs1 converts the human Mre11/Rad50 nuclease complex into an endo/exonuclease machine specific for protein-DNA adducts. Mol Cell 64:593–606. doi: 10.1016/j.molcel.2016.10.010 PubMedCrossRefGoogle Scholar
  76. 76.
    Broderick R et al (2016) EXD2 promotes homologous recombination by facilitating DNA end resection. Nat Cell Biol 18:271–280. doi: 10.1038/ncb3303 PubMedPubMedCentralCrossRefGoogle Scholar
  77. 77.
    Keyamura K, Arai K, Hishida T (2016) Srs2 and Mus81-Mms4 prevent accumulation of toxic inter-homolog recombination intermediates. PLoS Genet 12:e1006136. doi: 10.1371/journal.pgen.1006136 PubMedPubMedCentralCrossRefGoogle Scholar
  78. 78.
    Higashide M, Shinohara M (2016) Budding yeast SLX4 contributes to the appropriate distribution of crossovers and meiotic double-strand break formation on bivalents during meiosis. G3 (Bethesda) 6:2033–2042. doi: 10.1534/g3.116.029488
  79. 79.
    Dibitetto D et al (2016) Slx4 and Rtt107 control checkpoint signalling and DNA resection at double-strand breaks. Nucleic Acids Res 44:669–682. doi: 10.1093/nar/gkv1080 PubMedCrossRefGoogle Scholar
  80. 80.
    Guervilly JH, Gaillard PH (2016) SLX4 gains weight with SUMO in genome maintenance. Mol Cell Oncol 3:e1008297. doi: 10.1080/23723556.2015.1008297 PubMedCrossRefGoogle Scholar
  81. 81.
    Regairaz M et al (2011) Mus81-mediated DNA cleavage resolves replication forks stalled by topoisomerase I-DNA complexes. J Cell Biol 195:739–749. doi: 10.1083/jcb.201104003 PubMedPubMedCentralCrossRefGoogle Scholar
  82. 82.
    Sebesta M et al (2017) Esc2 promotes Mus81 complex-activity via its SUMO-like and DNA binding domains. Nucleic Acids Res 45:215–230. doi: 10.1093/nar/gkw882 PubMedCrossRefGoogle Scholar
  83. 83.
    Zhang YW et al (2011) Poly(ADP-ribose) polymerase and XPF-ERCC1 participate in distinct pathways for the repair of topoisomerase I-induced DNA damage in mammalian cells. Nucleic Acids Res 39:3607–3620. doi: 10.1093/nar/gkq1304 PubMedPubMedCentralCrossRefGoogle Scholar
  84. 84.
    LoRusso PM et al (2016) Phase I safety, pharmacokinetic, and pharmacodynamic study of the poly(ADP-ribose) polymerase (PARP) inhibitor veliparib (ABT-888) in combination with irinotecan in patients with advanced solid tumors. Clin Cancer Res 22:3227–3237. doi: 10.1158/1078-0432.CCR-15-0652 PubMedPubMedCentralCrossRefGoogle Scholar
  85. 85.
    Mullen JR, Chen CF, Brill SJ (2010) Wss1 is a SUMO-dependent isopeptidase that interacts genetically with the Slx5-Slx8 SUMO-targeted ubiquitin ligase. Mol Cell Biol 30:3737–3748. doi: 10.1128/MCB.01649-09 PubMedPubMedCentralCrossRefGoogle Scholar
  86. 86.
    O’Neill BM, Hanway D, Winzeler EA, Romesberg FE (2004) Coordinated functions of WSS1, PSY2 and TOF1 in the DNA damage response. Nucleic Acids Res 32:6519–6530. doi: 10.1093/nar/gkh994 PubMedPubMedCentralCrossRefGoogle Scholar
  87. 87.
    Stingele J, Schwarz MS, Bloemeke N, Wolf PG, Jentsch S (2014) A DNA-dependent protease involved in DNA-protein crosslink repair. Cell 158:327–338. doi: 10.1016/j.cell.2014.04.053 PubMedCrossRefGoogle Scholar
  88. 88.
    Balakirev MY et al (2015) Wss1 metalloprotease partners with Cdc48/Doa1 in processing genotoxic SUMO conjugates. Elife 4. doi: 10.7554/eLife.06763
  89. 89.
    Duxin JP, Dewar JM, Yardimci H, Walter JC (2014) Repair of a DNA-protein crosslink by replication-coupled proteolysis. Cell 159:346–357. doi: 10.1016/j.cell.2014.09.024 PubMedPubMedCentralCrossRefGoogle Scholar
  90. 90.
    Stingele J et al (2016) Mechanism and regulation of DNA-protein crosslink repair by the DNA-dependent metalloprotease SPRTN. Mol Cell 64:688–703. doi: 10.1016/j.molcel.2016.09.031 PubMedPubMedCentralCrossRefGoogle Scholar
  91. 91.
    Lopez-Mosqueda J et al (2016) SPRTN is a mammalian DNA-binding metalloprotease that resolves DNA-protein crosslinks. Elife 5. doi: 10.7554/eLife.21491
  92. 92.
    Lessel D et al (2014) Mutations in SPRTN cause early onset hepatocellular carcinoma, genomic instability and progeroid features. Nat Genet 46:1239–1244. doi: 10.1038/ng.3103 PubMedPubMedCentralCrossRefGoogle Scholar
  93. 93.
    Vaz B et al (2016) Metalloprotease SPRTN/DVC1 orchestrates replication-coupled DNA-protein crosslink repair. Mol Cell 64:704–719. doi: 10.1016/j.molcel.2016.09.032 PubMedPubMedCentralCrossRefGoogle Scholar
  94. 94.
    Mosbech A et al (2012) DVC1 (C1orf124) is a DNA damage-targeting p97 adaptor that promotes ubiquitin-dependent responses to replication blocks. Nat Struct Mol Biol 19:1084–1092. doi: 10.1038/nsmb.2395 PubMedCrossRefGoogle Scholar
  95. 95.
    Morocz M et al (2017) DNA-dependent protease activity of human Spartan facilitates replication of DNA-protein crosslink-containing DNA. Nucleic Acids Res. doi: 10.1093/nar/gkw1315
  96. 96.
    Ray Chaudhuri A et al (2012) Topoisomerase I poisoning results in PARP-mediated replication fork reversal. Nat Struct Mol Biol 19:417–423. doi: 10.1038/nsmb.2258 PubMedCrossRefGoogle Scholar
  97. 97.
    Solier S et al (2013) Transcription poisoning by Topoisomerase I is controlled by gene length, splice sites, and miR-142-3p. Cancer Res 73:4830–4839. doi: 10.1158/0008-5472.CAN-12-3504 PubMedCrossRefGoogle Scholar
  98. 98.
    Berti M et al (2013) Human RECQ1 promotes restart of replication forks reversed by DNA topoisomerase I inhibition. Nat Struct Mol Biol 20:347–354. doi: 10.1038/nsmb.2501 PubMedPubMedCentralCrossRefGoogle Scholar
  99. 99.
    Zellweger R et al (2015) Rad51-mediated replication fork reversal is a global response to genotoxic treatments in human cells. J Cell Biol 208:563–579. doi: 10.1083/jcb.201406099 PubMedPubMedCentralCrossRefGoogle Scholar
  100. 100.
    Hashimoto Y, Puddu F, Costanzo V (2011) RAD51- and MRE11-dependent reassembly of uncoupled CMG helicase complex at collapsed replication forks. Nat Struct Mol Biol 19:17–24. doi: 10.1038/nsmb.2177 PubMedPubMedCentralCrossRefGoogle Scholar
  101. 101.
    Petermann E, Orta ML, Issaeva N, Schultz N, Helleday T (2010) Hydroxyurea-stalled replication forks become progressively inactivated and require two different RAD51-mediated pathways for restart and repair. Mol Cell 37:492–502. doi: 10.1016/j.molcel.2010.01.021 PubMedPubMedCentralCrossRefGoogle Scholar
  102. 102.
    Hu J et al (2012) The intra-S phase checkpoint targets Dna2 to prevent stalled replication forks from reversing. Cell 149:1221–1232. doi: 10.1016/j.cell.2012.04.030 PubMedCrossRefGoogle Scholar
  103. 103.
    Ying S, Hamdy FC, Helleday T (2012) Mre11-dependent degradation of stalled DNA replication forks is prevented by BRCA2 and PARP1. Cancer Res 72:2814–2821. doi: 10.1158/0008-5472.CAN-11-3417 PubMedCrossRefGoogle Scholar
  104. 104.
    Schlacher K et al (2011) Double-strand break repair-independent role for BRCA2 in blocking stalled replication fork degradation by MRE11. Cell 145:529–542. doi: 10.1016/j.cell.2011.03.041 PubMedPubMedCentralCrossRefGoogle Scholar
  105. 105.
    Shiotani B et al (2013) Two distinct modes of ATR activation orchestrated by Rad17 and Nbs1. Cell Rep 3:1651–1662. doi: 10.1016/j.celrep.2013.04.018 PubMedPubMedCentralCrossRefGoogle Scholar
  106. 106.
    Murina O et al (2014) FANCD2 and CtIP cooperate to repair DNA interstrand crosslinks. Cell Rep 7:1030–1038. doi: 10.1016/j.celrep.2014.03.069 PubMedCrossRefGoogle Scholar
  107. 107.
    Yeo JE, Lee EH, Hendrickson EA, Sobeck A (2014) CtIP mediates replication fork recovery in a FANCD2-regulated manner. Hum Mol Genet 23:3695–3705. doi: 10.1093/hmg/ddu078 PubMedPubMedCentralCrossRefGoogle Scholar
  108. 108.
    Kuo C, Nuang H, Campbell JL (1983) Isolation of yeast DNA replication mutants in permeabilized cells. Proc Natl Acad Sci U S A 80:6465–6469PubMedPubMedCentralCrossRefGoogle Scholar
  109. 109.
    Budd ME, Campbell JL (1995) A yeast gene required for DNA replication encodes a protein with homology to DNA helicases. Proc Natl Acad Sci U S A 92:7642–7646PubMedPubMedCentralCrossRefGoogle Scholar
  110. 110.
    Gravel S, Chapman JR, Magill C, Jackson SP (2008) DNA helicases Sgs1 and BLM promote DNA double-strand break resection. Genes Dev 22:2767–2772. doi: 10.1101/gad.503108 PubMedPubMedCentralCrossRefGoogle Scholar
  111. 111.
    Zhu Z, Chung WH, Shim EY, Lee SE, Ira G (2008) Sgs1 helicase and two nucleases Dna2 and Exo1 resect DNA double-strand break ends. Cell 134:981–994. doi: 10.1016/j.cell.2008.08.037 PubMedPubMedCentralCrossRefGoogle Scholar
  112. 112.
    Nicolette ML et al (2010) Mre11-Rad50-Xrs2 and Sae2 promote 5′ strand resection of DNA double-strand breaks. Nat Struct Mol Biol 17:1478–1485. doi: 10.1038/nsmb.1957 PubMedPubMedCentralCrossRefGoogle Scholar
  113. 113.
    Nimonkar AV et al (2011) BLM-DNA2-RPA-MRN and EXO1-BLM-RPA-MRN constitute two DNA end resection machineries for human DNA break repair. Genes Dev 25:350–362. doi: 10.1101/gad.2003811 PubMedPubMedCentralCrossRefGoogle Scholar
  114. 114.
    Peng G et al (2012) Human nuclease/helicase DNA2 alleviates replication stress by promoting DNA end resection. Cancer Res 72:2802–2813. doi: 10.1158/0008-5472.CAN-11-3152 PubMedPubMedCentralCrossRefGoogle Scholar
  115. 115.
    Thangavel S et al (2015) DNA2 drives processing and restart of reversed replication forks in human cells. J Cell Biol 208:545–562. doi: 10.1083/jcb.201406100 PubMedPubMedCentralCrossRefGoogle Scholar
  116. 116.
    Iannascoli C, Palermo V, Murfuni I, Franchitto A, Pichierri P (2015) The WRN exonuclease domain protects nascent strands from pathological MRE11/EXO1-dependent degradation. Nucleic Acids Res 43:9788–9803. doi: 10.1093/nar/gkv836 PubMedPubMedCentralGoogle Scholar
  117. 117.
    Shamanna RA et al (2016) WRN regulates pathway choice between classical and alternative non-homologous end joining. Nat Commun 7:13785. doi: 10.1038/ncomms13785 PubMedPubMedCentralCrossRefGoogle Scholar
  118. 118.
    Shamanna RA et al. (2016) Camptothecin targets WRN protein: mechanism and relevance in clinical breast cancer. Oncotarget 7:13269–13284. doi: 10.18632/oncotarget.7906
  119. 119.
    Ribeyre C et al (2016) Nascent DNA proteomics reveals a chromatin remodeler required for topoisomerase I loading at replication forks. Cell Rep 15:300–309. doi: 10.1016/j.celrep.2016.03.027 PubMedCrossRefGoogle Scholar
  120. 120.
    Pommier Y, Kohlhagen G, Wu C, Simmons DT (1998) Mammalian DNA topoisomerase I activity and poisoning by camptothecin are inhibited by simian virus 40 large T antigen. Biochemistry 37:3818–3823. doi: 10.1021/bi972067d PubMedCrossRefGoogle Scholar
  121. 121.
    Seinsoth S, Uhlmann-Schiffler H, Stahl H (2003) Bidirectional DNA unwinding by a ternary complex of T antigen, nucleolin and topoisomerase I. EMBO Rep 4:263–268. doi: 10.1038/sj.embor.embor770 PubMedPubMedCentralCrossRefGoogle Scholar
  122. 122.
    Gavin I, Horn PJ, Peterson CL (2001) SWI/SNF chromatin remodeling requires changes in DNA topology. Mol Cell 7:97–104PubMedCrossRefGoogle Scholar
  123. 123.
    Walfridsson J, Khorosjutina O, Matikainen P, Gustafsson CM, Ekwall K (2007) A genome-wide role for CHD remodelling factors and Nap1 in nucleosome disassembly. EMBO J 26:2868–2879. doi: 10.1038/sj.emboj.7601728 PubMedPubMedCentralCrossRefGoogle Scholar
  124. 124.
    Durand-Dubief M, Svensson JP, Persson J, Ekwall K (2011) Topoisomerases, chromatin and transcription termination. Transcription 2:66–70. doi: 10.4161/trns.2.2.14411 PubMedPubMedCentralCrossRefGoogle Scholar
  125. 125.
    Krawczyk C, Dion V, Schar P, Fritsch O (2014) Reversible Top1 cleavage complexes are stabilized strand-specifically at the ribosomal replication fork barrier and contribute to ribosomal DNA stability. Nucleic Acids Res 42:4985–4995. doi: 10.1093/nar/gku148 PubMedPubMedCentralCrossRefGoogle Scholar
  126. 126.
    Dykhuizen EC et al (2013) BAF complexes facilitate decatenation of DNA by topoisomerase IIalpha. Nature 497:624–627. doi: 10.1038/nature12146 PubMedPubMedCentralCrossRefGoogle Scholar
  127. 127.
    Husain A et al (2016) Chromatin remodeller SMARCA4 recruits topoisomerase 1 and suppresses transcription-associated genomic instability. Nat Commun 7:10549. doi: 10.1038/ncomms10549 PubMedPubMedCentralCrossRefGoogle Scholar
  128. 128.
    Stoll G et al (2013) Deletion of TOP3beta, a component of FMRP-containing mRNPs, contributes to neurodevelopmental disorders. Nat Neurosci 16:1228–1237. doi: 10.1038/nn.3484 PubMedPubMedCentralCrossRefGoogle Scholar
  129. 129.
    Xu D et al (2013) Top3beta is an RNA topoisomerase that works with fragile X syndrome protein to promote synapse formation. Nat Neurosci 16:1238–1247. doi: 10.1038/nn.3479 PubMedPubMedCentralCrossRefGoogle Scholar
  130. 130.
    Yang Y et al (2014) Arginine methylation facilitates the recruitment of TOP3B to chromatin to prevent R loop accumulation. Mol Cell 53:484–497. doi: 10.1016/j.molcel.2014.01.011 PubMedPubMedCentralCrossRefGoogle Scholar
  131. 131.
    Ahmad M et al (2016) RNA topoisomerase is prevalent in all domains of life and associates with polyribosomes in animals. Nucleic Acids Res 44:6335–6349. doi: 10.1093/nar/gkw508 PubMedPubMedCentralCrossRefGoogle Scholar
  132. 132.
    Siaw GE, Liu IF, Lin PY, Been MD, Hsieh TS (2016) DNA and RNA topoisomerase activities of Top3beta are promoted by mediator protein Tudor domain-containing protein 3. Proc Natl Acad Sci U S A 113:E5544–E5551. doi: 10.1073/pnas.1605517113 PubMedPubMedCentralCrossRefGoogle Scholar
  133. 133.
    Meisenberg C et al. (2016) Epigenetic changes in histone acetylation underpin resistance to the topoisomerase I inhibitor irinotecan. Nucleic Acids Res. doi: 10.1093/nar/gkw1026
  134. 134.
    Maxwell A, Costenaro L, Mitelheiser S, Bates AD (2005) Coupling ATP hydrolysis to DNA strand passage in type IIA DNA topoisomerases. Biochem Soc Trans 33:1460–1464. doi: 10.1042/BST20051460 PubMedCrossRefGoogle Scholar
  135. 135.
    Joshi RS, Pina B, Roca J (2012) Topoisomerase II is required for the production of long Pol II gene transcripts in yeast. Nucleic Acids Res 40:7907–7915. doi: 10.1093/nar/gks626 PubMedPubMedCentralCrossRefGoogle Scholar
  136. 136.
    King IF et al (2013) Topoisomerases facilitate transcription of long genes linked to autism. Nature 501:58–62. doi: 10.1038/nature12504 PubMedPubMedCentralCrossRefGoogle Scholar
  137. 137.
    Roedgaard M, Fredsoe J, Pedersen JM, Bjergbaek L, Andersen AH (2015) DNA topoisomerases are required for preinitiation complex assembly during GAL gene activation. PLoS One 10:e0132739. doi: 10.1371/journal.pone.0132739 PubMedPubMedCentralCrossRefGoogle Scholar
  138. 138.
    Ju BG et al (2006) A topoisomerase IIbeta-mediated dsDNA break required for regulated transcription. Science 312:1798–1802. doi: 10.1126/science.1127196 PubMedCrossRefGoogle Scholar
  139. 139.
    Puc J et al (2015) Ligand-dependent enhancer activation regulated by topoisomerase-I activity. Cell 160:367–380. doi: 10.1016/j.cell.2014.12.023 PubMedPubMedCentralCrossRefGoogle Scholar
  140. 140.
    Bunch H et al (2015) Transcriptional elongation requires DNA break-induced signalling. Nat Commun 6:10191. doi: 10.1038/ncomms10191 PubMedPubMedCentralCrossRefGoogle Scholar
  141. 141.
    Madabhushi R et al (2015) Activity-induced DNA breaks govern the expression of neuronal early-response genes. Cell 161:1592–1605. doi: 10.1016/j.cell.2015.05.032 PubMedPubMedCentralCrossRefGoogle Scholar
  142. 142.
    Williams JS, Lujan SA, Kunkel TA (2016) Processing ribonucleotides incorporated during eukaryotic DNA replication. Nat Rev Mol Cell Biol 17:350–363. doi: 10.1038/nrm.2016.37 PubMedPubMedCentralCrossRefGoogle Scholar
  143. 143.
    Williams JS et al (2013) Topoisomerase 1-mediated removal of ribonucleotides from nascent leading-strand DNA. Mol Cell 49:1010–1015. doi: 10.1016/j.molcel.2012.12.021 PubMedPubMedCentralCrossRefGoogle Scholar
  144. 144.
    Sparks JL, Burgers PM (2015) Error-free and mutagenic processing of topoisomerase 1-provoked damage at genomic ribonucleotides. EMBO J 34:1259–1269. doi: 10.15252/embj.201490868
  145. 145.
    Kim N et al (2011) Mutagenic processing of ribonucleotides in DNA by yeast topoisomerase I. Science 332:1561–1564. doi: 10.1126/science.1205016 PubMedPubMedCentralCrossRefGoogle Scholar
  146. 146.
    Williams JS, Kunkel TA (2014) Ribonucleotides in DNA: origins, repair and consequences. DNA Repair (Amst) 19:27–37. doi: 10.1016/j.dnarep.2014.03.029 PubMedCentralCrossRefGoogle Scholar
  147. 147.
    Potenski CJ, Niu H, Sung P, Klein HL (2014) Avoidance of ribonucleotide-induced mutations by RNase H2 and Srs2-Exo1 mechanisms. Nature 511:251–254. doi: 10.1038/nature13292 PubMedPubMedCentralCrossRefGoogle Scholar
  148. 148.
    Lazzaro F et al (2012) RNase H and postreplication repair protect cells from ribonucleotides incorporated in DNA. Mol Cell 45:99–110. doi: 10.1016/j.molcel.2011.12.019 PubMedPubMedCentralCrossRefGoogle Scholar
  149. 149.
    Sekiguchi J, Shuman S (1997) Site-specific ribonuclease activity of eukaryotic DNA topoisomerase I. Mol Cell 1:89–97PubMedCrossRefGoogle Scholar
  150. 150.
    Huang SY, Ghosh S, Pommier Y (2015) Topoisomerase I alone is sufficient to produce short DNA deletions and can also reverse nicks at ribonucleotide sites. J Biol Chem 290:14068–14076. doi: 10.1074/jbc.M115.653345 PubMedPubMedCentralCrossRefGoogle Scholar
  151. 151.
    Arana ME et al (2012) Transcriptional responses to loss of RNase H2 in Saccharomyces cerevisiae. DNA Repair (Amst) 11:933–941. doi: 10.1016/j.dnarep.2012.09.006 PubMedCentralCrossRefGoogle Scholar
  152. 152.
    Conover HN et al (2015) Stimulation of chromosomal rearrangements by ribonucleotides. Genetics 201:951–961. doi: 10.1534/genetics.115.181149 PubMedPubMedCentralCrossRefGoogle Scholar
  153. 153.
    Epshtein A, Potenski CJ, Klein HL (2016) Increased spontaneous recombination in RNase H2-deficient cells arises from multiple contiguous rNMPs and not from single rNMP residues incorporated by DNA polymerase epsilon. Microb Cell 3:248–254. doi: 10.15698/mic2016.06.506
  154. 154.
    Niu H, Potenski CJ, Epshtein A, Sung P, Klein HL (2016) Roles of DNA helicases and Exo1 in the avoidance of mutations induced by Top1-mediated cleavage at ribonucleotides in DNA. Cell Cycle 15:331–336. doi: 10.1080/15384101.2015.1128594 PubMedCrossRefGoogle Scholar
  155. 155.
    Pendleton M, Lindsey RH Jr, Felix CA, Grimwade D, Osheroff N (2014) Topoisomerase II and leukemia. Ann N Y Acad Sci 1310:98–110. doi: 10.1111/nyas.12358 PubMedPubMedCentralCrossRefGoogle Scholar
  156. 156.
    Cowell IG et al (2012) Model for MLL translocations in therapy-related leukemia involving topoisomerase IIbeta-mediated DNA strand breaks and gene proximity. Proc Natl Acad Sci U S A 109:8989–8994. doi: 10.1073/pnas.1204406109 PubMedPubMedCentralCrossRefGoogle Scholar
  157. 157.
    Azarova AM et al (2007) Roles of DNA topoisomerase II isozymes in chemotherapy and secondary malignancies. Proc Natl Acad Sci U S A 104:11014–11019. doi: 10.1073/pnas.0704002104 PubMedPubMedCentralCrossRefGoogle Scholar
  158. 158.
    Lovett BD et al (2001) Near-precise interchromosomal recombination and functional DNA topoisomerase II cleavage sites at MLL and AF-4 genomic breakpoints in treatment-related acute lymphoblastic leukemia with t(4;11) translocation. Proc Natl Acad Sci U S A 98:9802–9807. doi: 10.1073/pnas.171309898 PubMedPubMedCentralCrossRefGoogle Scholar
  159. 159.
    Mahler M, Silverman ED, Schulte-Pelkum J, Fritzler MJ (2010) Anti-Scl-70 (topo-I) antibodies in SLE: myth or reality? Autoimmun Rev 9:756–760. doi: 10.1016/j.autrev.2010.06.005 PubMedCrossRefGoogle Scholar
  160. 160.
    Haffner MC et al (2010) Androgen-induced TOP2B-mediated double-strand breaks and prostate cancer gene rearrangements. Nat Genet 42:668–675. doi: 10.1038/ng.613 PubMedPubMedCentralCrossRefGoogle Scholar
  161. 161.
    Tomlins SA et al (2005) Recurrent fusion of TMPRSS2 and ETS transcription factor genes in prostate cancer. Science 310:644–648. doi: 10.1126/science.1117679 PubMedCrossRefGoogle Scholar
  162. 162.
    Helgeson BE et al (2008) Characterization of TMPRSS2:ETV5 and SLC45A3:ETV5 gene fusions in prostate cancer. Cancer Res 68:73–80. doi: 10.1158/0008-5472.CAN-07-5352 PubMedCrossRefGoogle Scholar
  163. 163.
    Hermans KG et al (2008) Truncated ETV1, fused to novel tissue-specific genes, and full-length ETV1 in prostate cancer. Cancer Res 68:7541–7549. doi: 10.1158/0008-5472.CAN-07-5930 PubMedCrossRefGoogle Scholar
  164. 164.
    Lin C et al (2009) Nuclear receptor-induced chromosomal proximity and DNA breaks underlie specific translocations in cancer. Cell 139:1069–1083. doi: 10.1016/j.cell.2009.11.030 PubMedPubMedCentralCrossRefGoogle Scholar
  165. 165.
    Kumar-Sinha C, Tomlins SA, Chinnaiyan AM (2008) Recurrent gene fusions in prostate cancer. Nat Rev Cancer 8:497–511. doi: 10.1038/nrc2402 PubMedPubMedCentralCrossRefGoogle Scholar
  166. 166.
    Bastus NC et al (2010) Androgen-induced TMPRSS2:ERG fusion in nonmalignant prostate epithelial cells. Cancer Res 70:9544–9548. doi: 10.1158/0008-5472.CAN-10-1638 PubMedPubMedCentralCrossRefGoogle Scholar
  167. 167.
    Bowen C, Gelmann E (2010) P. NKX3.1 activates cellular response to DNA damage. Cancer Res 70:3089–3097. doi: 10.1158/0008-5472.CAN-09-3138 PubMedCrossRefGoogle Scholar
  168. 168.
    Bowen C, Zheng T, Gelmann EP (2015) NKX3.1 suppresses TMPRSS2-ERG gene rearrangement and mediates repair of androgen receptor-induced DNA damage. Cancer Res 75:2686–2698. doi: 10.1158/0008-5472.CAN-14-3387 PubMedPubMedCentralCrossRefGoogle Scholar
  169. 169.
    Zhang H, Zheng T, Chua CW, Shen M, Gelmann EP (2016) Nkx3.1 controls the DNA repair response in the mouse prostate. Prostate 76:402–408. doi: 10.1002/pros.23131 PubMedCrossRefGoogle Scholar
  170. 170.
    Mani RS et al (2016) Inflammation-induced oxidative stress mediates gene fusion formation in prostate cancer. Cell Rep 17:2620–2631. doi: 10.1016/j.celrep.2016.11.019 PubMedPubMedCentralCrossRefGoogle Scholar
  171. 171.
    Laxman B et al (2006) Noninvasive detection of TMPRSS2:ERG fusion transcripts in the urine of men with prostate cancer. Neoplasia 8:885–888. doi: 10.1593/neo.06625 PubMedPubMedCentralCrossRefGoogle Scholar
  172. 172.
    Merdan S et al (2015) Assessment of long-term outcomes associated with urinary prostate cancer antigen 3 and TMPRSS2:ERG gene fusion at repeat biopsy. Cancer 121:4071–4079. doi: 10.1002/cncr.29611 PubMedCrossRefGoogle Scholar
  173. 173.
    Geybels MS et al (2015) Epigenomic profiling of prostate cancer identifies differentially methylated genes in TMPRSS2:ERG fusion-positive versus fusion-negative tumors. Clin Epigenetics 7:128. doi: 10.1186/s13148-015-0161-6 PubMedPubMedCentralCrossRefGoogle Scholar
  174. 174.
    Walker C, Herranz-Martin S, Karyka E, Liao C, Lewis K, Elsayed W, Lukashchuk V, Chiang SC, Ray S, Mulcahy PJ, Jurga M, Tsagakis I, Iannitti T, Chandran J, Coldicott I, De Vos KJ, Hassan MK, Higginbottom A, Shaw PJ, Hautbergue GM, Azzouz M, El-Khamisy SF (2017, July) C9orf72 expansion disrupts ATM-mediated chromosomal break repair. Nat Neurosci. doi: 10.1038/nn.4604. [Epub ahead of print]

Copyright information

© American Association of Pharmaceutical Scientists 2017

Authors and Affiliations

  1. 1.Center for Genomics, Helmy Institute for Medical SciencesZewail City of Science and TechnologyGizaEgypt
  2. 2.Krebs Institute and Sheffield Institute for Nucleic Acids, Department of Molecular Biology and Biotechnology, Firth CourtUniversity of SheffieldSheffieldUK

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