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:

RNA: a double-edged sword in genome maintenance

Nature Reviews Genetics volume 21, pages 651–670 (2020)Cite this article

Subjects

Abstract

All organisms must safeguard the integrity of their DNA to avoid deleterious consequences of genome instability, which have been linked to human diseases such as autoimmune disorders, neurodegenerative diseases and cancer. Traditionally, genome maintenance has been viewed largely in terms of DNA–protein interactions. However, emerging evidence points to RNA as a key modulator of genome stability, with seemingly opposing roles in promoting chromosomal instability and protecting genome integrity. Unravelling the mechanistic and contextual basis of this duality will not only improve our understanding of the interfaces between RNA and the genome but will also provide important insights into how disrupted RNA metabolism contributes to disease origin, laying the foundation for targeted intervention.

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: RNA can potentially affect genome stability at each stage of its life cycle.
Fig. 2: Co-transcriptional R-loops and genome stability.
Fig. 3: RNA-templated DSB repair.
Fig. 4: RNA-mediated DNA damage signalling.
Fig. 5: RNA processing and genome stability.
Fig. 6: RNA-mediated chromatin organization.
Fig. 7: RNA-mediated suppression of retrotransposons.
Fig. 8: Genomic ribonucleotides and mechanisms for their removal.

Similar content being viewed by others

References

  1. Jackson, S. P. & Bartek, J. The DNA-damage response in human biology and disease. Nature 461, 1071–1078 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  2. Bunting, S. F. & Nussenzweig, A. End-joining, translocations and cancer. Nat. Rev. Cancer 13, 443–454 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  3. McKinnon, P. J. Genome integrity and disease prevention in the nervous system. Genes. Dev. 31, 1180–1194 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  4. Tubbs, A. & Nussenzweig, A. Endogenous DNA damage as a source of genomic instability in cancer. Cell 168, 644–656 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  5. Bader, A. S., Hawley, B. R., Wilczynska, A. & Bushell, M. The roles of RNA in DNA double-strand break repair. Br. J. Cancer (2020).

  6. Belotserkovskii, B. P., Tornaletti, S., D’Souza, A. D. & Hanawalt, P. C. R-loop generation during transcription: Formation, processing and cellular outcomes. DNA Repair 71, 69–81 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  7. Huertas, P. & Aguilera, A. Cotranscriptionally formed DNA:RNA hybrids mediate transcription elongation impairment and transcription-associated recombination. Mol. Cell 12, 711–721 (2003).

    CAS  PubMed  Google Scholar 

  8. Chen, X., Yang, J. R., Zhang, J. & Nascent, R. N. A. folding mitigates transcription-associated mutagenesis. Genome Res. 26, 50–59 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  9. 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).

    CAS  PubMed  PubMed Central  Google Scholar 

  10. Stolz, R. et al. Interplay between DNA sequence and negative superhelicity drives R-loop structures. Proc. Natl Acad. Sci. USA 116, 6260–6269 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  11. Wahba, L., Costantino, L., Tan, F. J., Zimmer, A. & Koshland, D. S1-DRIP-seq identifies high expression and polyA tracts as major contributors to R-loop formation. Genes Dev. 30, 1327–1338 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  12. Salas-Armenteros, I. et al. Human THO-Sin3A interaction reveals new mechanisms to prevent R-loops that cause genome instability. EMBO J. 36, 3532–3547 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. Yang, X. et al. m6A promotes R-loop formation to facilitate transcription termination. Cell Res. 29, 1035–1038 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  14. Kotsantis, P. et al. Increased global transcription activity as a mechanism of replication stress in cancer. Nat. Commun. 7, 13087 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  15. Stork, C. T. et al. Co-transcriptional R-loops are the main cause of estrogen-induced DNA damage. Elife https://doi.org/10.7554/eLife.17548 (2016). This study reports that activation of transcription by oestrogen (oestradiol) can induce R-loop-dependent DNA damage at genomic loci that are enriched in rearrangements in breast cancer.

    Article  PubMed  PubMed Central  Google Scholar 

  16. Hamperl, S., Bocek, M. J., Saldivar, J. C., Swigut, T. & Cimprich, K. A. Transcription-replication conflict orientation modulates R-loop levels and activates distinct DNA damage responses. Cell 170, 774–786 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  17. Sordet, O. et al. Ataxia telangiectasia mutated activation by transcription- and topoisomerase I-induced DNA double-strand breaks. EMBO Rep. 10, 887–893 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  18. Garcia-Benitez, F., Gaillard, H. & Aguilera, A. Physical proximity of chromatin to nuclear pores prevents harmful R loop accumulation contributing to maintain genome stability. Proc. Natl Acad. Sci. USA 114, 10942–10947 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  19. Wahba, L., Amon, J. D., Koshland, D., Vuica-Ross, M. & RNase, H. and multiple RNA biogenesis factors cooperate to prevent RNA:DNA hybrids from generating genome instability. Mol. Cell 44, 978–988 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  20. Nadel, J. et al. RNA:DNA hybrids in the human genome have distinctive nucleotide characteristics, chromatin composition, and transcriptional relationships. Epigenetics Chromatin 8, 46 (2015).

    PubMed  PubMed Central  Google Scholar 

  21. Wahba, L., Gore, S. K. & Koshland, D. The homologous recombination machinery modulates the formation of RNA-DNA hybrids and associated chromosome instability. Elife 2, e00505 (2013).

    PubMed  PubMed Central  Google Scholar 

  22. Chen, L. et al. R-ChIP using inactive RNase H reveals dynamic coupling of R-loops with transcriptional pausing at gene promoters. Mol. Cell 68, 745–757 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  23. Dumelie, J. G. & Jaffrey, S. R. Defining the location of promoter-associated R-loops at near-nucleotide resolution using bisDRIP-seq. Elife https://doi.org/10.7554/eLife.28306 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  24. 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).

    CAS  PubMed  PubMed Central  Google Scholar 

  25. Malig, M., Hartono, S. R., Giafaglione, J. M., Sanz, L. A. & Chedin, F. Ultra-deep coverage single-molecule R-loop footprinting reveals principles of R-loop formation. J. Mol. Biol. https://doi.org/10.1016/j.jmb.2020.02.014 (2020). This study demonstrates the feasibility of mapping R-loops genome-wide at high resolution by capturing and sequencing the single-stranded DNA of the displaced strand, providing additional insights into the principles underlying R-loop formation. It is a valuable complementary approach to DNA–RNA immunoprecipitation coupled with high-throughput sequencing (described in Sanz et al., 2016).

    Article  PubMed  PubMed Central  Google Scholar 

  26. 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). This study is among the first to report genome-wide mapping of R-loops in human cells using the S9.6 antibody, which specifically recognizes RNA–DNA hybrids. This approach is known as DNA–RNA immunoprecipitation coupled with high-throughput sequencing.

    CAS  PubMed  PubMed Central  Google Scholar 

  27. Yan, Q., Shields, E. J., Bonasio, R. & Sarma, K. Mapping native R-loops genome-wide using a targeted nuclease approach. Cell Rep. 29, 1369–1380 (2019). This study, along with the studies by Chen et al. (2017) and Ginno et al. (2012), demonstrates the feasibility of mapping R-loops genome-wide by capturing them with a catalytically inactive RNase H that is conjugated either to an epitope tag or to the micrococcal nuclease.

    CAS  PubMed  PubMed Central  Google Scholar 

  28. Chedin, F. Nascent connections: R-loops and chromatin patterning. Trends Genet. 32, 828–838 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. Crossley, M. P., Bocek, M. & Cimprich, K. A. R-loops as cellular regulators and genomic threats. Mol. Cell 73, 398–411 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  30. Garcia-Muse, T. & Aguilera, A. R loops: from physiological to pathological roles. Cell 179, 604–618 (2019).

    CAS  PubMed  Google Scholar 

  31. Niehrs, C. & Luke, B. Regulatory R-loops as facilitators of gene expression and genome stability. Nat. Rev. Mol. Cell Biol. 21, 167–178 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  32. Grunseich, C. et al. Senataxin mutation reveals how R-loops promote transcription by blocking DNA methylation at gene promoters. Mol. Cell 69, 426–437 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  33. Skourti-Stathaki, K., Proudfoot, N. J. & Gromak, N. Human senataxin resolves RNA/DNA hybrids formed at transcriptional pause sites to promote Xrn2-dependent termination. Mol. Cell 42, 794–805 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  34. Sun, Q., Csorba, T., Skourti-Stathaki, K., Proudfoot, N. J. & Dean, C. R-loop stabilization represses antisense transcription at the Arabidopsis FLC locus. Science 340, 619–621 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  35. Tan-Wong, S. M., Dhir, S. & Proudfoot, N. J. R-loops promote antisense transcription across the mammalian genome. Mol. Cell 76, 600–616 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  36. Holt, I. J. The mitochondrial R-loop. Nucleic Acids Res. 47, 5480–5489 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  37. Posse, V. et al. RNase H1 directs origin-specific initiation of DNA replication in human mitochondria. PLoS Genet. 15, e1007781 (2019).

    PubMed  PubMed Central  Google Scholar 

  38. Wongsurawat, T., Jenjaroenpun, P., Kwoh, C. K. & Kuznetsov, V. Quantitative model of R-loop forming structures reveals a novel level of RNA-DNA interactome complexity. Nucleic Acids Res. 40, e16 (2012).

    CAS  PubMed  Google Scholar 

  39. Belotserkovskii, B. P. & Hanawalt, P. C. Anchoring nascent RNA to the DNA template could interfere with transcription. Biophys. J. 100, 675–684 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  40. Gomez-Gonzalez, B. & Aguilera, A. Transcription-mediated replication hindrance: a major driver of genome instability. Genes Dev. 33, 1008–1026 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. Gan, W. et al. R-loop-mediated genomic instability is caused by impairment of replication fork progression. Genes Dev. 25, 2041–2056 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  42. 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).

    CAS  PubMed  Google Scholar 

  43. Freudenreich, C. H. R-loops: targets for nuclease cleavage and repeat instability. Curr. Genet. 64, 789–794 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  44. Sollier, J. et al. Transcription-coupled nucleotide excision repair factors promote R-loop-induced genome instability. Mol. Cell 56, 777–785 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  45. Saini, N. & Gordenin, D. A. Hypermutation in single-stranded DNA. DNA Repair 91–92, 102868 (2020).

    PubMed  Google Scholar 

  46. Yu, K. & Lieber, M. R. Current insights into the mechanism of mammalian immunoglobulin class switch recombination. Crit. Rev. Biochem. Mol. Biol. 54, 333–351 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  47. Costantino, L. & Koshland, D. Genome-wide map of r-loop-induced damage reveals how a subset of R-loops contributes to genomic instability. Mol. Cell 71, 487–497 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  48. Castellano-Pozo, M. et al. R loops are linked to histone H3 S10 phosphorylation and chromatin condensation. Mol. Cell 52, 583–590 (2013).

    CAS  PubMed  Google Scholar 

  49. Garcia-Pichardo, D. et al. Histone mutants separate R loop formation from genome instability induction. Mol. Cell 66, 597–609 (2017). This study, along with that by Salas-Armenteros et al. (2017), suggests that specific chromatin contexts (such as histone modifications) may alter the properties of R-loops, making them more harmful.

    CAS  PubMed  Google Scholar 

  50. Garcia-Rubio, M. et al. Yra1-bound RNA-DNA hybrids cause orientation-independent transcription-replication collisions and telomere instability. Genes Dev. 32, 965–977 (2018). This study, along with that by Hamperl et al. (2017), reports that co-directional transcription–replication conflicts are associated with enhanced R-loop clearance, providing a plausible explanation as to why transcription and replication tend to be co-oriented in mammalian cells.

    CAS  PubMed  PubMed Central  Google Scholar 

  51. Stuckey, R., Garcia-Rodriguez, N., Aguilera, A. & Wellinger, R. E. Role for RNA:DNA hybrids in origin-independent replication priming in a eukaryotic system. Proc. Natl Acad. Sci. USA 112, 5779–5784 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  52. Gorthi, A. et al. EWS-FLI1 increases transcription to cause R-loops and block BRCA1 repair in Ewing sarcoma. Nature 555, 387–391 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  53. Nergadze, S. G. et al. CpG-island promoters drive transcription of human telomeres. RNA 15, 2186–2194 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  54. Azzalin, C. M., Reichenbach, P., Khoriauli, L., Giulotto, E. & Lingner, J. Telomeric repeat containing RNA and RNA surveillance factors at mammalian chromosome ends. Science 318, 798–801 (2007).

    CAS  PubMed  Google Scholar 

  55. Luke, B. et al. The Rat1p 5’ to 3’ exonuclease degrades telomeric repeat-containing RNA and promotes telomere elongation in Saccharomyces cerevisiae. Mol. Cell 32, 465–477 (2008).

    CAS  PubMed  Google Scholar 

  56. Schoeftner, S. & Blasco, M. A. Developmentally regulated transcription of mammalian telomeres by DNA-dependent RNA polymerase II. Nat. Cell Biol. 10, 228–236 (2008).

    CAS  PubMed  Google Scholar 

  57. Pfeiffer, V. & Lingner, J. TERRA promotes telomere shortening through exonuclease 1-mediated resection of chromosome ends. PLoS Genet. 8, e1002747 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  58. Herold, S. et al. Recruitment of BRCA1 limits MYCN-driven accumulation of stalled RNA polymerase. Nature 567, 545–549 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  59. Shivji, M. K. K., Renaudin, X., Williams, C. H. & Venkitaraman, A. R. BRCA2 regulates transcription elongation by RNA polymerase II to prevent R-loop accumulation. Cell Rep. 22, 1031–1039 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  60. Zhang, X. et al. Attenuation of RNA polymerase II pausing mitigates BRCA1-associated R-loop accumulation and tumorigenesis. Nat. Commun. 8, 15908 (2017).

    PubMed  PubMed Central  Google Scholar 

  61. Kim, N. & Jinks-Robertson, S. The Top1 paradox: friend and foe of the eukaryotic genome. DNA Repair 56, 33–41 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  62. Nakazawa, Y. et al. Ubiquitination of DNA damage-stalled RNAPII promotes transcription-coupled repair. Cell 180, 1228–1244 (2020).

    CAS  PubMed  Google Scholar 

  63. Tufegdzic Vidakovic, A. et al. Regulation of the RNAPII Pool Is Integral to the DNA damage response. Cell 180, e1221 (2020).

    Google Scholar 

  64. Cerritelli, S. M., Crouch, R. J. & Ribonuclease, H. the enzymes in eukaryotes. FEBS J. 276, 1494–1505 (2009).

    CAS  PubMed  Google Scholar 

  65. Nguyen, H. D. et al. Functions of replication protein a as a sensor of R loops and a regulator of RNaseH1. Mol. Cell 65, 832–847 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  66. Hatchi, E. et al. BRCA1 recruitment to transcriptional pause sites is required for R-loop-driven DNA damage repair. Mol. Cell 57, 636–647 (2015). This study, along with the studies by Herold et al. (2019) and Zhang et al. (2017), shows that BRCA1 promotes genome stability and tumour suppression in part by resolving R-loops at transcription pause sites.

    CAS  PubMed  PubMed Central  Google Scholar 

  67. Allison, D. F. & Wang, G. G. R-loops: formation, function, and relevance to cell stress. Cell Stress. 3, 38–46 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  68. Petryk, N. et al. Replication landscape of the human genome. Nat. Commun. 7, 10208 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  69. Barroso, S. et al. The DNA damage response acts as a safeguard against harmful DNA-RNA hybrids of different origins. EMBO Rep. 20, e47250 (2019).

    PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  Google Scholar 

  71. Zhao, H., Zhu, M., Limbo, O. & Russell, P. RNase H eliminates R-loops that disrupt DNA replication but is nonessential for efficient DSB repair. EMBO Rep. https://doi.org/10.15252/embr.201745335 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  72. Perego, M. G. L., Taiana, M., Bresolin, N., Comi, G. P. & Corti, S. R-loops in motor neuron diseases. Mol. Neurobiol. 56, 2579–2589 (2019).

    CAS  PubMed  Google Scholar 

  73. Richard, P. & Manley, J. L. R loops and links to human disease. J. Mol. Biol. 429, 3168–3180 (2017).

    CAS  PubMed  Google Scholar 

  74. Colak, D. et al. Promoter-bound trinucleotide repeat mRNA drives epigenetic silencing in fragile X syndrome. Science 343, 1002–1005 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  75. Groh, M., Lufino, M. M., Wade-Martins, R. & Gromak, N. R-loops associated with triplet repeat expansions promote gene silencing in Friedreich ataxia and fragile X syndrome. PLoS Genet. 10, e1004318 (2014).

    PubMed  PubMed Central  Google Scholar 

  76. Sagie, S. et al. Telomeres in ICF syndrome cells are vulnerable to DNA damage due to elevated DNA:RNA hybrids. Nat. Commun. 8, 14015 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  77. Zhu, Q. et al. Heterochromatin-encoded satellite RNAs induce breast cancer. Mol. Cell 70, 842–853 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  78. Zhu, Q. et al. BRCA1 tumour suppression occurs via heterochromatin-mediated silencing. Nature 477, 179–184 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  79. Bhatia, V. et al. BRCA2 prevents R-loop accumulation and associates with TREX-2 mRNA export factor PCID2. Nature 511, 362–365 (2014).

    CAS  PubMed  Google Scholar 

  80. Garcia-Rubio, M. L. et al. The Fanconi anemia pathway protects genome integrity from R-loops. PLoS Genet. 11, e1005674 (2015).

    PubMed  PubMed Central  Google Scholar 

  81. Schwab, R. A. et al. The Fanconi anemia pathway maintains genome stability by coordinating replication and transcription. Mol. Cell 60, 351–361 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  82. Bhatia, V., Herrera-Moyano, E., Aguilera, A. & Gomez-Gonzalez, B. The role of replication-associated repair factors on R-loops. Genes https://doi.org/10.3390/genes8070171 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  83. Chang, E. Y. & Stirling, P. C. Replication fork protection factors controlling R-loop bypass and suppression. Genes https://doi.org/10.3390/genes8010033 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  84. Okamoto, Y., Hejna, J. & Takata, M. Regulation of R-loops and genome instability in Fanconi anemia. J. Biochem. 165, 465–470 (2019).

    CAS  PubMed  Google Scholar 

  85. Paull, T. T. RNA-DNA hybrids and the convergence with DNA repair. Crit. Rev. Biochem. Mol. Biol. 54, 371–384 (2019).

    CAS  PubMed  Google Scholar 

  86. Wells, J. P., White, J. & Stirling, P. C. R loops and their composite cancer connections. Trends Cancer 5, 619–631 (2019).

    CAS  PubMed  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  88. Akman, G. et al. Pathological ribonuclease H1 causes R-loop depletion and aberrant DNA segregation in mitochondria. Proc. Natl Acad. Sci. USA 113, E4276–E4285 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  89. Holt, I. J. The Jekyll and Hyde character of RNase H1 and its multiple roles in mitochondrial DNA metabolism. DNA Repair 84, 102630 (2019).

    CAS  PubMed  Google Scholar 

  90. Kannan, A., Jiang, X., He, L., Ahmad, S. & Gangwani, L. ZPR1 prevents R-loop accumulation, upregulates SMN2 expression and rescues spinal muscular atrophy. Brain 143, 69–93 (2020).

    PubMed  Google Scholar 

  91. Wang, I. X. et al. Human proteins that interact with RNA/DNA hybrids. Genome Res. 28, 1405–1414 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  92. Cristini, A., Groh, M., Kristiansen, M. S. & Gromak, N. RNA/DNA hybrid interactome identifies DXH9 as a molecular player in transcriptional termination and R-loop-associated DNA damage. Cell Rep. 23, 1891–1905 (2018). This study, and that by Wang et al. (2018), reports the identification of human proteins that exhibit preferential interaction with RNA–DNA hybrids, providing a useful resource for future studies aimed at elucidating how RNA–DNA hybrids are regulated and their possible physiological functions.

    CAS  PubMed  PubMed Central  Google Scholar 

  93. Marnef, A., Cohen, S. & Legube, G. Transcription-coupled DNA double-strand break repair: active genes need special care. J. Mol. Biol. 429, 1277–1288 (2017).

    CAS  PubMed  Google Scholar 

  94. Adelman, C. A. & Petrini, J. H. Division of labor: DNA repair and the cell cycle specific functions of the Mre11 complex. Cell Cycle 8, 1510–1514 (2009).

    CAS  PubMed  Google Scholar 

  95. Keskin, H., Meers, C. & Storici, F. Transcript RNA supports precise repair of its own DNA gene. RNA Biol. 13, 157–165 (2016).

    PubMed  Google Scholar 

  96. Aguilera, A. & Gomez-Gonzalez, B. DNA-RNA hybrids: the risks of DNA breakage during transcription. Nat. Struct. Mol. Biol. 24, 439–443 (2017).

    CAS  PubMed  Google Scholar 

  97. Bader, A. S. & Bushell, M. DNA:RNA hybrids form at DNA double-strand breaks in transcriptionally active loci. Cell Death Dis. 11, 280 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  98. Puget, N., Miller, K. M. & Legube, G. Non-canonical DNA/RNA structures during transcription-coupled double-strand break repair: roadblocks or bona fide repair intermediates? DNA Repair 81, 102661 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  99. Derr, L. K., Strathern, J. N. & Garfinkel, D. J. RNA-mediated recombination in S. cerevisiae. Cell 67, 355–364 (1991).

    CAS  PubMed  Google Scholar 

  100. Keskin, H. et al. Transcript-RNA-templated DNA recombination and repair. Nature 515, 436–439 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  101. Lee, M. H. et al. Somatic APP gene recombination in Alzheimer’s disease and normal neurons. Nature 563, 639–645 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  102. Li, S. et al. Precise gene replacement in rice by RNA transcript-templated homologous recombination. Nat. Biotechnol. 37, 445–450 (2019).

    CAS  PubMed  Google Scholar 

  103. Nowacki, M. et al. RNA-mediated epigenetic programming of a genome-rearrangement pathway. Nature 451, 153–158 (2008).

    CAS  PubMed  Google Scholar 

  104. Ono, R. et al. Double strand break repair by capture of retrotransposon sequences and reverse-transcribed spliced mRNA sequences in mouse zygotes. Sci. Rep. 5, 12281 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  105. Onozawa, M. et al. Repair of DNA double-strand breaks by templated nucleotide sequence insertions derived from distant regions of the genome. Proc. Natl Acad. Sci. USA 111, 7729–7734 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  106. Shen, Y. et al. RNA-driven genetic changes in bacteria and in human cells. Mutat. Res. 717, 91–98 (2011).

    CAS  PubMed  Google Scholar 

  107. Storici, F. et al. Repair. Nature 447, 338–341 (2007). This study and the study by Keskin et al. (2014) demonstrate that yeast can use RNA transcript as a template to repair a DNA DSB.

    CAS  PubMed  PubMed Central  Google Scholar 

  108. Mazina, O. M., Keskin, H., Hanamshet, K., Storici, F. & Mazin, A. V. Rad52 inverse strand exchange drives RNA-templated DNA double-strand break repair. Mol. Cell 67, 19–29 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  109. McDevitt, S., Rusanov, T., Kent, T., Chandramouly, G. & Pomerantz, R. T. How RNA transcripts coordinate DNA recombination and repair. Nat. Commun. 9, 1091 (2018).

    PubMed  PubMed Central  Google Scholar 

  110. Keskin, H. & Storici, F. Defects in RNase H2 stimulate DNA break repair by RNA reverse transcribed into cDNA. Microrna 4, 109–116 (2015).

    CAS  PubMed  Google Scholar 

  111. Chakraborty, A. et al. Classical non-homologous end-joining pathway utilizes nascent RNA for error-free double-strand break repair of transcribed genes. Nat. Commun. 7, 13049 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  112. Jang, Y. et al. Intrinsically disordered protein RBM14 plays a role in generation of RNA:DNA hybrids at double-strand break sites. Proc. Natl Acad. Sci. USA 117, 5329–5338 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  113. Gout, J. F. et al. The landscape of transcription errors in eukaryotic cells. Sci. Adv. 3, e1701484 (2017).

    PubMed  PubMed Central  Google Scholar 

  114. Amon, J. D. & Koshland, D. RNase H enables efficient repair of R-loop induced DNA damage. Elife https://doi.org/10.7554/eLife.20533 (2016).

    Article  PubMed  PubMed Central  Google Scholar 

  115. D’Alessandro, G. et al. BRCA2 controls DNA:RNA hybrid level at DSBs by mediating RNase H2 recruitment. Nat. Commun. 9, 5376 (2018).

    PubMed  PubMed Central  Google Scholar 

  116. Domingo-Prim, J. et al. EXOSC10 is required for RPA assembly and controlled DNA end resection at DNA double-strand breaks. Nat. Commun. 10, 2135 (2019).

    PubMed  PubMed Central  Google Scholar 

  117. Li, L. et al. DEAD box 1 facilitates removal of RNA and homologous recombination at DNA double-strand breaks. Mol. Cell Biol. 36, 2794–2810 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  118. Ohle, C. et al. Transient RNA-DNA hybrids are required for efficient double-strand break repair. Cell 167, 1001–1013 (2016).

    CAS  PubMed  Google Scholar 

  119. Yasuhara, T. et al. Human Rad52 promotes XPG-mediated R-loop processing to initiate transcription-associated homologous recombination repair. Cell 175, 558–570 (2018).

    CAS  PubMed  Google Scholar 

  120. Lu, W. T. et al. Drosha drives the formation of DNA:RNA hybrids around DNA break sites to facilitate DNA repair. Nat. Commun. 9, 532 (2018).

    PubMed  PubMed Central  Google Scholar 

  121. Wei, W. et al. A role for small RNAs in DNA double-strand break repair. Cell 149, 101–112 (2012).

    CAS  PubMed  Google Scholar 

  122. Wei, L. et al. DNA damage during the G0/G1 phase triggers RNA-templated, Cockayne syndrome B-dependent homologous recombination. Proc. Natl Acad. Sci. USA 112, E3495–E3504 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  123. Welty, S. et al. RAD52 is required for RNA-templated recombination repair in post-mitotic neurons. J. Biol. Chem. 293, 1353–1362 (2018).

    CAS  PubMed  Google Scholar 

  124. Hanamshet, K., Mazina, O. M. & Mazin, A. V. Reappearance from obscurity: mammalian Rad52 in homologous recombination. Genes https://doi.org/10.3390/genes7090063 (2016).

    Article  PubMed  PubMed Central  Google Scholar 

  125. Jalan, M., Olsen, K. S. & Powell, S. N. Emerging roles of RAD52 in genome maintenance. Cancers https://doi.org/10.3390/cancers11071038 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  126. Malacaria, E., Honda, M., Franchitto, A., Spies, M. & Pichierri, P. Physiological and pathological roles of RAD52 at DNA replication Forks. Cancers https://doi.org/10.3390/cancers12020402 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  127. Nogueira, A., Fernandes, M., Catarino, R. & Medeiros, R. RAD52 functions in homologous recombination and its importance on genomic integrity maintenance and cancer therapy. Cancers https://doi.org/10.3390/cancers11111622 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  128. Caron, P., van der Linden, J. & van Attikum, H. Bon voyage: a transcriptional journey around DNA breaks. DNA Repair 82, 102686 (2019).

    CAS  PubMed  Google Scholar 

  129. Machour, F. E. & Ayoub, N. Transcriptional regulation at DSBs: mechanisms and consequences. Trends Genet. https://doi.org/10.1016/j.tig.2020.01.001 (2020).

  130. Kim, J., Sturgill, D., Tran, A. D., Sinclair, D. A. & Oberdoerffer, P. Controlled DNA double-strand break induction in mice reveals post-damage transcriptome stability. Nucleic Acids Res. 44, e64 (2016).

    PubMed  Google Scholar 

  131. Kruhlak, M. et al. The ATM repair pathway inhibits RNA polymerase I transcription in response to chromosome breaks. Nature 447, 730–734 (2007).

    CAS  PubMed  Google Scholar 

  132. Purman, C. E. et al. Regional gene repression by DNA double-strand breaks in G1 phase cells. Mol. Cell Biol. https://doi.org/10.1128/MCB.00181-19 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  133. Gutbrod, M. J. & Martienssen, R. A. Conserved chromosomal functions of RNA interference. Nat. Rev. Genet. 21, 311–331 (2020).

    CAS  PubMed  Google Scholar 

  134. Michelini, F. et al. From “cellular” RNA to “smart” RNA: multiple roles of RNA in genome stability and beyond. Chem. Rev. 118, 4365–4403 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  135. Bonath, F., Domingo-Prim, J., Tarbier, M., Friedlander, M. R. & Visa, N. Next-generation sequencing reveals two populations of damage-induced small RNAs at endogenous DNA double-strand breaks. Nucleic Acids Res. 46, 11869–11882 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  136. Francia, S. et al. Site-specific DICER and DROSHA RNA products control the DNA-damage response. Nature 488, 231–235 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  137. Michalik, K. M., Bottcher, R. & Forstemann, K. A small RNA response at DNA ends in Drosophila. Nucleic Acids Res. 40, 9596–9603 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  138. Miki, D. et al. Efficient generation of diRNAs requires components in the posttranscriptional gene silencing pathway. Sci. Rep. 7, 301 (2017).

    PubMed  PubMed Central  Google Scholar 

  139. Pessina, F. et al. Functional transcription promoters at DNA double-strand breaks mediate RNA-driven phase separation of damage-response factors. Nat. Cell Biol. 21, 1286–1299 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  140. Vitor, A. C. et al. Single-molecule imaging of transcription at damaged chromatin. Sci. Adv. 5, eaau1249 (2019).

    PubMed  PubMed Central  Google Scholar 

  141. Michelini, F. et al. Damage-induced lncRNAs control the DNA damage response through interaction with DDRNAs at individual double-strand breaks. Nat. Cell Biol. 19, 1400–1411 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  142. Rossiello, F. et al. DNA damage response inhibition at dysfunctional telomeres by modulation of telomeric DNA damage response RNAs. Nat. Commun. 8, 13980 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  143. Francia, S., Cabrini, M., Matti, V., Oldani, A. & d’Adda di Fagagna, F. DICER, DROSHA and DNA damage response RNAs are necessary for the secondary recruitment of DNA damage response factors. J. Cell Sci. 129, 1468–1476 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  144. Wang, Q. & Goldstein, M. Small RNAs Recruit chromatin-modifying enzymes MMSET and Tip60 to reconfigure damaged DNA upon double-strand break and facilitate repair. Cancer Res. 76, 1904–1915 (2016).

    CAS  PubMed  Google Scholar 

  145. Gao, M. et al. Ago2 facilitates Rad51 recruitment and DNA double-strand break repair by homologous recombination. Cell Res. 24, 532–541 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  146. Mirman, Z. & de Lange, T. 53BP1: a DSB escort. Genes Dev. 34, 7–23 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  147. Tarsounas, M. & Sung, P. The antitumorigenic roles of BRCA1-BARD1 in DNA repair and replication. Nat. Rev. Mol. Cell Biol. 21, 284–299 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  148. Kilic, S. et al. Phase separation of 53BP1 determines liquid-like behavior of DNA repair compartments. EMBO J. 38, e101379 (2019). This study and the study by Pessina et al. (2019) demonstrate that focal assembly of 53BP1 at DNA damage sites is driven by phase separation, with Pessina et al. suggesting a role for diRNA in this process.

    PubMed  PubMed Central  Google Scholar 

  149. Botuyan, M. V. et al. Mechanism of 53BP1 activity regulation by RNA-binding TIRR and a designer protein. Nat. Struct. Mol. Biol. 25, 591–600 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  150. Dai, Y., Zhang, A., Shan, S., Gong, Z. & Zhou, Z. Structural basis for recognition of 53BP1 tandem Tudor domain by TIRR. Nat. Commun. 9, 2123 (2018).

    PubMed  PubMed Central  Google Scholar 

  151. Drane, P. et al. TIRR regulates 53BP1 by masking its histone methyl-lysine binding function. Nature 543, 211–216 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  152. Wang, J. et al. Molecular basis for the inhibition of the methyl-lysine binding function of 53BP1 by TIRR. Nat. Commun. 9, 2689 (2018).

    PubMed  PubMed Central  Google Scholar 

  153. Jacquet, K. et al. The TIP60 complex regulates bivalent chromatin recognition by 53BP1 through direct H4K20me binding and H2AK15 acetylation. Mol. Cell 62, 409–421 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  154. Tang, J. et al. Acetylation limits 53BP1 association with damaged chromatin to promote homologous recombination. Nat. Struct. Mol. Biol. 20, 317–325 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  155. Lenzken, S. C. et al. FUS-dependent phase separation initiates double-strand break repair. Preprint at bioRxiv https://doi.org/10.1101/798884 (2019).

  156. Wang, W. Y. et al. Interaction of FUS and HDAC1 regulates DNA damage response and repair in neurons. Nat. Neurosci. 16, 1383–1391 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  157. Altmeyer, M. et al. Liquid demixing of intrinsically disordered proteins is seeded by poly(ADP-ribose). Nat. Commun. 6, 8088 (2015).

    CAS  PubMed  Google Scholar 

  158. Patel, A. et al. A liquid-to-solid phase transition of the ALS protein FUS accelerated by disease mutation. Cell 162, 1066–1077 (2015).

    CAS  PubMed  Google Scholar 

  159. Naumann, M. et al. Impaired DNA damage response signaling by FUS-NLS mutations leads to neurodegeneration and FUS aggregate formation. Nat. Commun. 9, 335 (2018).

    PubMed  PubMed Central  Google Scholar 

  160. Chen, L. et al. The augmented R-loop is a unifying mechanism for myelodysplastic syndromes induced by high-risk splicing factor mutations. Mol. Cell 69, 412–425 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  161. Li, X. & Manley, J. L. Inactivation of the SR protein splicing factor ASF/SF2 results in genomic instability. Cell 122, 365–378 (2005).

    CAS  PubMed  Google Scholar 

  162. Bonnet, A. et al. Introns protect eukaryotic genomes from transcription-associated genetic instability. Mol. Cell 67, 608–621 (2017). This study and the study by Li and Manley (2005) provide evidence that splicing factors, rather than splicing per se, inhibit co-transcriptional R-loop formation by binding to pre-mRNA, which prevents stable RNA–DNA interaction.

    CAS  PubMed  Google Scholar 

  163. Gupta, S. K., Luo, L. & Yen, L. RNA-mediated gene fusion in mammalian cells. Proc. Natl Acad. Sci. USA 115, E12295–E12304 (2018). This study provides evidence that an RNA transcript containing sufficient sequence homologies to two disparate genomic loci can drive gene fusion.

    CAS  PubMed  PubMed Central  Google Scholar 

  164. Wu, H., Li, X. & Li, H. Gene fusions and chimeric RNAs, and their implications in cancer. Genes. Dis. 6, 385–390 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  165. Kandel, E. S. & Nudler, E. Template switching by RNA polymerase II in vivo. Evidence and implications from a retroviral system. Mol. Cell 10, 1495–1502 (2002).

    CAS  PubMed  Google Scholar 

  166. Yan, Z. et al. Genome-wide colocalization of RNA-DNA interactions and fusion RNA pairs. Proc. Natl Acad. Sci. USA 116, 3328–3337 (2019). This study maps RNA–DNA interactions genome-wide in normal cells and reports that spatial proximity between RNA and genomic DNA is associated with the creation of chimeric fusion transcripts.

    CAS  PubMed  PubMed Central  Google Scholar 

  167. Huang, H., Weng, H. & Chen, J. The biogenesis and precise control of RNA m6A methylation. Trends Genet. 36, 44–52 (2020).

    CAS  PubMed  Google Scholar 

  168. Xiang, Y. et al. RNA m6A methylation regulates the ultraviolet-induced DNA damage response. Nature 543, 573–576 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  169. Stern, H. R., Sefcikova, J., Chaparro, V. E. & Beuning, P. J. Mammalian DNA polymerase kappa activity and specificity. Molecules https://doi.org/10.3390/molecules24152805 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  170. Rastogi, R. P., Richa, Kumar, A., Tyagi, M. B. & Sinha, R. P. Molecular mechanisms of ultraviolet radiation-induced DNA damage and repair. J. Nucleic Acids 2010, 592980 (2010).

    PubMed  PubMed Central  Google Scholar 

  171. Fu, Y. & Zhuang, X. m6A-binding YTHDF proteins promote stress granule formation. Nat. Chem. Biol. https://doi.org/10.1038/s41589-020-0524-y (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  172. Gao, Y. et al. Multivalent m6A motifs promote phase separation of YTHDF proteins. Cell Res. 29, 767–769 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  173. Ries, R. J. et al. m6A enhances the phase separation potential of mRNA. Nature 571, 424–428 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  174. Abakir, A. et al. N6-methyladenosine regulates the stability of RNA:DNA hybrids in human cells. Nat. Genet. https://doi.org/10.1038/s41588-019-0549-x (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  175. Tresini, M. et al. The core spliceosome as target and effector of non-canonical ATM signalling. Nature 523, 53–58 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  176. Bell, J. C. et al. Chromatin-associated RNA sequencing (ChAR-seq) maps genome-wide RNA-to-DNA contacts. Elife https://doi.org/10.7554/eLife.27024 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  177. Li, X. et al. GRID-seq reveals the global RNA-chromatin interactome. Nat. Biotechnol. 35, 940–950 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  178. Mondal, T., Rasmussen, M., Pandey, G. K., Isaksson, A. & Kanduri, C. Characterization of the RNA content of chromatin. Genome Res. 20, 899–907 (2010). This study and the studies by Bell et al. (2018) and Li et al. (2017) describe different approaches for mapping the global RNA–chromatin interactome in D. melanogaster, mouse and human cells.

    CAS  PubMed  PubMed Central  Google Scholar 

  179. Rodriguez-Campos, A. & Azorin, F. RNA is an integral component of chromatin that contributes to its structural organization. PLoS ONE 2, e1182 (2007).

    PubMed  PubMed Central  Google Scholar 

  180. Caudron-Herger, M. & Rippe, K. Nuclear architecture by RNA. Curr. Opin. Genet. Dev. 22, 179–187 (2012).

    CAS  PubMed  Google Scholar 

  181. Johnson, W. L. & Straight, A. F. RNA-mediated regulation of heterochromatin. Curr. Opin. Cell Biol. 46, 102–109 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  182. Li, X. & Fu, X. D. Chromatin-associated RNAs as facilitators of functional genomic interactions. Nat. Rev. Genet. 20, 503–519 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  183. Mishra, K. & Kanduri, C. Understanding long noncoding RNA and chromatin interactions: what we know so far. Noncoding RNA https://doi.org/10.3390/ncrna5040054 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  184. Wang, Y. & Feigon, J. Structural biology of telomerase and its interaction at telomeres. Curr. Opin. Struct. Biol. 47, 77–87 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  185. Mangaonkar, A. A. & Patnaik, M. M. Short telomere syndromes in clinical practice: bridging bench and bedside. Mayo Clin. Proc. 93, 904–916 (2018).

    PubMed  Google Scholar 

  186. Toubiana, S. & Selig, S. DNA:RNA hybrids at telomeres - when it is better to be out of the (R) loop. FEBS J. 285, 2552–2566 (2018).

    CAS  PubMed  Google Scholar 

  187. Kabeche, L., Nguyen, H. D., Buisson, R. & Zou, L. A mitosis-specific and R loop-driven ATR pathway promotes faithful chromosome segregation. Science 359, 108–114 (2018).

    CAS  PubMed  Google Scholar 

  188. Montero, J. J., Lopez de Silanes, I., Grana, O. & Blasco, M. A. Telomeric RNAs are essential to maintain telomeres. Nat. Commun. 7, 12534 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  189. Caceres-Gutierrez, R. & Herrera, L. A. Centromeric non-coding transcription: opening the black box of chromosomal instability? Curr. Genomics 18, 227–235 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  190. Maicher, A., Kastner, L. & Luke, B. Telomeres and disease: enter TERRA. RNA Biol. 9, 843–849 (2012).

    CAS  PubMed  Google Scholar 

  191. Arora, R. et al. RNaseH1 regulates TERRA-telomeric DNA hybrids and telomere maintenance in ALT tumour cells. Nat. Commun. 5, 5220 (2014).

    CAS  PubMed  Google Scholar 

  192. Hansen, A. S. et al. Distinct classes of chromatin loops revealed by deletion of an RNA-binding region in CTCF. Mol. Cell 76, 395–411 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  193. Saldana-Meyer, R. et al. RNA interactions are essential for CTCF-mediated genome organization. Mol. Cell 76, 412–422 (2019). This study and the study by Hansen et al. (2019) show that CTCF-dependent genome organization is at least in part mediated by the interactions between CTCF and RNA.

    CAS  PubMed  PubMed Central  Google Scholar 

  194. Misteli, T. & Soutoglou, E. The emerging role of nuclear architecture in DNA repair and genome maintenance. Nat. Rev. Mol. Cell Biol. 10, 243–254 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  195. Li, Y., Syed, J. & Sugiyama, H. RNA-DNA triplex formation by long noncoding RNAs. Cell Chem. Biol. 23, 1325–1333 (2016).

    CAS  PubMed  Google Scholar 

  196. Klein, S. J. & O’Neill, R. J. Transposable elements: genome innovation, chromosome diversity, and centromere conflict. Chromosome Res. 26, 5–23 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  197. Platt, R. N. II, Vandewege, M. W. & Ray, D. A. Mammalian transposable elements and their impacts on genome evolution. Chromosome Res. 26, 25–43 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  198. Wimmer, K., Callens, T., Wernstedt, A. & Messiaen, L. The NF1 gene contains hotspots for L1 endonuclease-dependent de novo insertion. PLoS Genet. 7, e1002371 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  199. Burns, K. H. Our conflict with transposable elements and its implications for human disease. Annu. Rev. Pathol. 15, 51–70 (2020).

    CAS  PubMed  Google Scholar 

  200. Kazazian, H. H. Jr. & Moran, J. V. Mobile DNA in health and disease. N. Engl. J. Med. 377, 361–370 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  201. Hall, A. C., Ostrowski, L. A., Pietrobon, V. & Mekhail, K. Repetitive DNA loci and their modulation by the non-canonical nucleic acid structures R-loops and G-quadruplexes. Nucleus 8, 162–181 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  202. Ardeljan, D. et al. Cell fitness screens reveal a conflict between LINE-1 retrotransposition and DNA replication. Nat. Struct. Mol. Biol. 27, 168–178 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  203. Jeon, J. et al. Retroelement insertion in a CRISPR/Cas9 editing site in the early embryo intensifies genetic mosaicism. Front. Cell Dev. Biol. 7, 273 (2019).

    PubMed  PubMed Central  Google Scholar 

  204. Morrish, T. A. et al. DNA repair mediated by endonuclease-independent LINE-1 retrotransposition. Nat. Genet. 31, 159–165 (2002).

    CAS  PubMed  Google Scholar 

  205. Maizels, N. & Davis, L. Initiation of homologous recombination at DNA nicks. Nucleic Acids Res. 46, 6962–6973 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  206. Wang, Y. et al. BRCA1 intronic Alu elements drive gene rearrangements and PARP inhibitor resistance. Nat. Commun. 10, 5661 (2019). This study demonstrates that Alu-mediated genomic rearrangements upstream of a nonsense mutation in the BRCA1 gene can restore the correct reading frame, leading to the production of a stable BRCA1 protein and the development of chemoresistance.

    PubMed  PubMed Central  Google Scholar 

  207. Anwar, S. L., Wulaningsih, W. & Lehmann, U. Transposable elements in human cancer: causes and consequences of deregulation. Int. J. Mol. Sci. https://doi.org/10.3390/ijms18050974 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  208. Pavlicek, A. et al. Evolution of the tumor suppressor BRCA1 locus in primates: implications for cancer predisposition. Hum. Mol. Genet. 13, 2737–2751 (2004).

    CAS  PubMed  Google Scholar 

  209. Mita, P. et al. BRCA1 and S phase DNA repair pathways restrict LINE-1 retrotransposition in human cells. Nat. Struct. Mol. Biol. 27, 179–191 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  210. Goodier, J. L. Restricting retrotransposons: a review. Mob. DNA 7, 16 (2016).

    PubMed  PubMed Central  Google Scholar 

  211. Czech, B. et al. piRNA-guided genome defense: from biogenesis to silencing. Annu. Rev. Genet. 52, 131–157 (2018).

    CAS  PubMed  Google Scholar 

  212. Ernst, C., Odom, D. T. & Kutter, C. The emergence of piRNAs against transposon invasion to preserve mammalian genome integrity. Nat. Commun. 8, 1411 (2017).

    PubMed  PubMed Central  Google Scholar 

  213. Iwasaki, Y. W., Siomi, M. C. & Siomi, H. PIWI-interacting RNA: its biogenesis and functions. Annu. Rev. Biochem. 84, 405–433 (2015).

    CAS  PubMed  Google Scholar 

  214. Ozata, D. M., Gainetdinov, I., Zoch, A., O’Carroll, D. & Zamore, P. D. PIWI-interacting RNAs: small RNAs with big functions. Nat. Rev. Genet. 20, 89–108 (2019).

    CAS  PubMed  Google Scholar 

  215. Schmid, M. & Jensen, T. H. Controlling nuclear RNA levels. Nat. Rev. Genet. 19, 518–529 (2018).

    CAS  PubMed  Google Scholar 

  216. Dos Santos, R. F. et al. Major 3’-5’ exoribonucleases in the metabolism of coding and non-coding RNA. Prog. Mol. Biol. Transl. Sci. 159, 101–155 (2018).

    PubMed  Google Scholar 

  217. Houseley, J. & Tollervey, D. The many pathways of RNA degradation. Cell 136, 763–776 (2009).

    CAS  PubMed  Google Scholar 

  218. Kilchert, C., Wittmann, S. & Vasiljeva, L. The regulation and functions of the nuclear RNA exosome complex. Nat. Rev. Mol. Cell Biol. 17, 227–239 (2016).

    CAS  PubMed  Google Scholar 

  219. Levy, S. & Schuster, G. Polyadenylation and degradation of RNA in the mitochondria. Biochem. Soc. Trans. 44, 1475–1482 (2016).

    CAS  PubMed  Google Scholar 

  220. Nair, L., Chung, H. & Basu, U. Regulation of long non-coding RNAs and genome dynamics by the RNA surveillance machinery. Nat. Rev. Mol. Cell Biol. 21, 123–136 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  221. Pefanis, E. et al. RNA exosome-regulated long non-coding RNA transcription controls super-enhancer activity. Cell 161, 774–789 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  222. Silva, S., Camino, L. P. & Aguilera, A. Human mitochondrial degradosome prevents harmful mitochondrial R loops and mitochondrial genome instability. Proc. Natl Acad. Sci. USA 115, 11024–11029 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  223. Morton, D. J. et al. The RNA exosome and RNA exosome-linked disease. RNA 24, 127–142 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  224. Pashler, A. L., Towler, B. P., Jones, C. I. & Newbury, S. F. The roles of the exoribonucleases DIS3L2 and XRN1 in human disease. Biochem. Soc. Trans. 44, 1377–1384 (2016).

    CAS  PubMed  Google Scholar 

  225. Dhir, A. et al. Mitochondrial double-stranded RNA triggers antiviral signalling in humans. Nature 560, 238–242 (2018). This study and the study by Silva et al. (2018) report that the loss of mitochondrial RNA degradosome activity leads to R-loop-associated genome instability as well as the formation of immunogenic double-stranded RNA, highlighting the importance of RNA turnover in mitochondria.

    CAS  PubMed  PubMed Central  Google Scholar 

  226. Bresson, S. & Tollervey, D. Surveillance-ready transcription: nuclear RNA decay as a default fate. Open Biol. https://doi.org/10.1098/rsob.170270 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  227. Traut, T. W. Physiological concentrations of purines and pyrimidines. Mol. Cell Biochem. 140, 1–22 (1994).

    CAS  PubMed  Google Scholar 

  228. Nava, G. M. et al. One, no one, and one hundred thousand: the many forms of ribonucleotides in DNA. Int. J. Mol. Sci. https://doi.org/10.3390/ijms21051706 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  229. Cerritelli, S. M. & Crouch, R. J. The balancing act of ribonucleotides in DNA. Trends Biochem. Sci. 41, 434–445 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  230. Nick McElhinny, S. A. et al. Abundant ribonucleotide incorporation into DNA by yeast replicative polymerases. Proc. Natl Acad. Sci. USA 107, 4949–4954 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  231. Clausen, A. R., Zhang, S., Burgers, P. M., Lee, M. Y. & Kunkel, T. A. Ribonucleotide incorporation, proofreading and bypass by human DNA polymerase delta. DNA Repair 12, 121–127 (2013).

    CAS  PubMed  Google Scholar 

  232. Goksenin, A. Y. et al. Human DNA polymerase epsilon is able to efficiently extend from multiple consecutive ribonucleotides. J. Biol. Chem. 287, 42675–42684 (2012).

    PubMed  PubMed Central  Google Scholar 

  233. Kasiviswanathan, R. & Copeland, W. C. Ribonucleotide discrimination and reverse transcription by the human mitochondrial DNA polymerase. J. Biol. Chem. 286, 31490–31500 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  234. Koh, K. D., Balachander, S., Hesselberth, J. R. & Storici, F. Ribose-seq: global mapping of ribonucleotides embedded in genomic DNA. Nat. Methods 12, 251–257 (2015). This study describes an elegant protocol for mapping embedded ribonucleotides genome-wide in yeast, which takes advantage of alkali-dependent cleavage of the DNA backbone at the site of embedded ribonucleotides.

    CAS  PubMed  PubMed Central  Google Scholar 

  235. Crespan, E. et al. Impact of ribonucleotide incorporation by DNA polymerases beta and lambda on oxidative base excision repair. Nat. Commun. 7, 10805 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  236. Nick McElhinny, S. A. & Ramsden, D. A. Polymerase mu is a DNA-directed DNA/RNA polymerase. Mol. Cell Biol. 23, 2309–2315 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  237. Pryor, J. M. et al. Ribonucleotide incorporation enables repair of chromosome breaks by nonhomologous end joining. Science 361, 1126–1129 (2018). This study reports the striking finding that the insertion of ribonucleotides by error-prone DNA polymerases stimulates DSB ligation.

    CAS  PubMed  PubMed Central  Google Scholar 

  238. Su, Y. et al. Human DNA polymerase eta has reverse transcriptase activity in cellular environments. J. Biol. Chem. 294, 6073–6081 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  239. Kennedy, E. M., Amie, S. M., Bambara, R. A. & Kim, B. Frequent incorporation of ribonucleotides during HIV-1 reverse transcription and their attenuated repair in macrophages. J. Biol. Chem. 287, 14280–14288 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  240. Williams, J. S., Lujan, S. A. & Kunkel, T. A. Processing ribonucleotides incorporated during eukaryotic DNA replication. Nat. Rev. Mol. Cell Biol. 17, 350–363 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  241. Hyjek, M., Figiel, M., Nowotny, M. & RNases, H. Structure and mechanism. DNA Repair 84, 102672 (2019).

    CAS  PubMed  Google Scholar 

  242. Cerritelli, S. M. et al. Failure to produce mitochondrial DNA results in embryonic lethality in Rnaseh1 null mice. Mol. Cell 11, 807–815 (2003).

    CAS  PubMed  Google Scholar 

  243. Holmes, J. B. et al. Primer retention owing to the absence of RNase H1 is catastrophic for mitochondrial DNA replication. Proc. Natl Acad. Sci. USA 112, 9334–9339 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  244. Lima, W. F. et al. Viable RNaseH1 knockout mice show RNaseH1 is essential for R loop processing, mitochondrial and liver function. Nucleic Acids Res. 44, 5299–5312 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  245. Reijns, M. A. et al. Enzymatic removal of ribonucleotides from DNA is essential for mammalian genome integrity and development. Cell 149, 1008–1022 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  246. Grossman, L. I., Watson, R. & Vinograd, J. The presence of ribonucleotides in mature closed-circular mitochondrial DNA. Proc. Natl Acad. Sci. USA 70, 3339–3343 (1973).

    CAS  PubMed  PubMed Central  Google Scholar 

  247. Wong-Staal, F., Mendelsohn, J. & Goulian, M. Ribonucleotides in closed circular mitochondrial DNA from HeLa cells. Biochem. Biophys. Res. Commun. 53, 140–148 (1973).

    CAS  PubMed  Google Scholar 

  248. Kellner, V. & Luke, B. Molecular and physiological consequences of faulty eukaryotic ribonucleotide excision repair. EMBO J https://doi.org/10.15252/embj.2019102309 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  249. Klein, H. L. Genome instabilities arising from ribonucleotides in DNA. DNA Repair 56, 26–32 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  250. Chiu, H. C. et al. RNA intrusions change DNA elastic properties and structure. Nanoscale 6, 10009–10017 (2014).

    CAS  PubMed  Google Scholar 

  251. Lujan, S. A., Williams, J. S., Clausen, A. R., Clark, A. B. & Kunkel, T. A. Ribonucleotides are signals for mismatch repair of leading-strand replication errors. Mol. Cell 50, 437–443 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  252. Hiller, B. et al. Mammalian RNase H2 removes ribonucleotides from DNA to maintain genome integrity. J. Exp. Med. 209, 1419–1426 (2012). This study and the studies by Cerritelli et al. (2003) and Reijns et al. (2012) are among the first to describe the deleterious phenotypes associated with a complete loss of RNase H1 or RNase H2 in mice.

    CAS  PubMed  PubMed Central  Google Scholar 

  253. Uehara, R. et al. Two RNase H2 mutants with differential rNMP processing activity reveal a threshold of ribonucleotide tolerance for embryonic development. Cell Rep. 25, 1135–1145 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  254. Cilli, P., Minoprio, A., Bossa, C., Bignami, M. & Mazzei, F. Formation and repair of mismatches containing ribonucleotides and oxidized bases at repeated DNA sequences. J. Biol. Chem. 290, 26259–26269 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  255. Mentegari, E. et al. Ribonucleotide incorporation by human DNA polymerase eta impacts translesion synthesis and RNase H2 activity. Nucleic Acids Res. 45, 2600–2614 (2017).

    CAS  PubMed  Google Scholar 

  256. Malfatti, M. C. et al. Abasic and oxidized ribonucleotides embedded in DNA are processed by human APE1 and not by RNase H2. Nucleic Acids Res. 45, 11193–11212 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  257. Cai, Y., Geacintov, N. E. & Broyde, S. Ribonucleotides as nucleotide excision repair substrates. DNA Repair 13, 55–60 (2014).

    CAS  PubMed  Google Scholar 

  258. Vaisman, A. & Woodgate, R. Redundancy in ribonucleotide excision repair: competition, compensation, and cooperation. DNA Repair 29, 74–82 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  259. Lindsey-Boltz, L. A., Kemp, M. G., Hu, J. & Sancar, A. Analysis of ribonucleotide removal from DNA by human nucleotide excision repair. J. Biol. Chem. 290, 29801–29807 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  260. Sassa, A. et al. Processing of a single ribonucleotide embedded into DNA by human nucleotide excision repair and DNA polymerase eta. Sci. Rep. 9, 13910 (2019).

    PubMed  PubMed Central  Google Scholar 

  261. Lazzaro, F. et al. RNase H and postreplication repair protect cells from ribonucleotides incorporated in DNA. Mol. Cell 45, 99–110 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  262. Nick McElhinny, S. A. et al. Genome instability due to ribonucleotide incorporation into DNA. Nat. Chem. Biol. 6, 774–781 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  263. Sparks, J. L. & Burgers, P. M. Error-free and mutagenic processing of topoisomerase 1-provoked damage at genomic ribonucleotides. EMBO J. 34, 1259–1269 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  264. Epshtein, A., Potenski, C. J. & Klein, H. L. 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 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  265. Zimmermann, M. et al. CRISPR screens identify genomic ribonucleotides as a source of PARP-trapping lesions. Nature 559, 285–289 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  266. Huang, S. N., Williams, J. S., Arana, M. E., Kunkel, T. A. & Pommier, Y. Topoisomerase I-mediated cleavage at unrepaired ribonucleotides generates DNA double-strand breaks. EMBO J. 36, 361–373 (2017). This study and the studies by Nick McElhinny et al. (Nat. Chem. Biol., 2010) and Sparks and Burgers (2015) demonstrate how unexcised genomic ribonucleotides can lead to genome instability through a TOP1-mediated repair mechanism.

    CAS  PubMed  Google Scholar 

  267. Sciascia, N. et al. Suppressing proteasome mediated processing of topoisomerase II DNA-protein complexes preserves genome integrity. Elife https://doi.org/10.7554/eLife.53447 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  268. Berglund, A. K. et al. Nucleotide pools dictate the identity and frequency of ribonucleotide incorporation in mitochondrial DNA. PLoS Genet. 13, e1006628 (2017).

    PubMed  PubMed Central  Google Scholar 

  269. Moss, C. F. et al. Aberrant ribonucleotide incorporation and multiple deletions in mitochondrial DNA of the murine MPV17 disease model. Nucleic Acids Res. 45, 12808–12815 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  270. Reyes, A. et al. RNASEH1 mutations impair mtDNA Replication and cause adult-onset mitochondrial encephalomyopathy. Am. J. Hum. Genet. 97, 186–193 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  271. Gunther, C. et al. Defective removal of ribonucleotides from DNA promotes systemic autoimmunity. J. Clin. Invest. 125, 413–424 (2015).

    PubMed  Google Scholar 

  272. Pizzi, S. et al. Reduction of hRNase H2 activity in Aicardi-Goutieres syndrome cells leads to replication stress and genome instability. Hum. Mol. Genet. 24, 649–658 (2015).

    CAS  PubMed  Google Scholar 

  273. Crow, Y. J. et al. Mutations in genes encoding ribonuclease H2 subunits cause Aicardi-Goutieres syndrome and mimic congenital viral brain infection. Nat. Genet. 38, 910–916 (2006).

    CAS  PubMed  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  275. Pokatayev, V. et al. RNase H2 catalytic core Aicardi-Goutieres syndrome-related mutant invokes cGAS-STING innate immune-sensing pathway in mice. J. Exp. Med. 213, 329–336 (2016). This study and the study by Mackenzie et al. (2016) model disease-associated Rnaseh2 mutations in mice and show that deficient ribonucleotide excision leads to the activation of inflammatory immune responses.

    CAS  PubMed  PubMed Central  Google Scholar 

  276. Lim, Y. W. et al. hypomethylation and RNA:DNA hybrid accumulation in Aicardi-Goutieres syndrome. Elife https://doi.org/10.7554/eLife.08007 (2015).

    Article  PubMed  PubMed Central  Google Scholar 

  277. Chon, H. et al. RNase H2 roles in genome integrity revealed by unlinking its activities. Nucleic Acids Res. 41, 3130–3143 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  278. Cornelio, D. A., Sedam, H. N., Ferrarezi, J. A., Sampaio, N. M. & Argueso, J. L. Both R-loop removal and ribonucleotide excision repair activities of RNase H2 contribute substantially to chromosome stability. DNA Repair 52, 110–114 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  279. Storci, G. et al. Genomic stability, anti-inflammatory phenotype, and up-regulation of the RNAseH2 in cells from centenarians. Cell Death Differ. 26, 1845–1858 (2019). This study reports that cells of centenarians overexpress RNase H2, while various diseased tissues show reduced RNase H2 expression, potentially linking RNase H2 function to human longevity.

    CAS  PubMed  PubMed Central  Google Scholar 

  280. Alexandrov, L. B. et al. The repertoire of mutational signatures in human cancer. Nature 578, 94–101 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

Download references

Acknowledgements

The authors thank S. John and N. Johnson for critical reading of the manuscript. The authors acknowledge and apologize to peers whose work could not be cited owing to space limitation.

Author information

Authors and Affiliations

  1. Laboratory of Genome Integrity, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA

    Dali Zong & André Nussenzweig

  2. Division of Cancer Biology, National Cancer Institute, National Institutes of Health, Rockville, MD, USA

    Philipp Oberdoerffer

  3. Laboratory of Cell Biology, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA

    Pedro J. Batista

Authors
  1. Dali Zong
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Philipp Oberdoerffer
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Pedro J. Batista
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. André Nussenzweig
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

D.Z. researched data for article. D.Z., and A.N. wrote the article. All authors contributed substantially to discussion of the content and reviewed/edited the manuscript before submission.

Corresponding author

Correspondence to André Nussenzweig.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Peer review information

Nature Reviews Genetics thanks A. Aguilera, who co-reviewed with B. Gómez-González, F. Storici, who co-reviewed with C. Meers, and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

Publisher’s note

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

Glossary

Replisome

A large ensemble of proteins that together create a replication fork and initiate DNA synthesis from a replication origin.

Nucleolytic cleavage

A chemical or enzymatic process that breaks phosphodiester bonds in the DNA backbone.

Structure-specific nucleases

Enzymes that recognize and cleave specific types of secondary structure within DNA.

Nucleotide excision repair

A mode of DNA repair wherein a string of nucleotides containing the damaged bases are excised and the resultant gap is filled in by new DNA synthesis.

Transition mutations

A type of point mutation that changes a purine nucleotide to another purine or a pyrimidine nucleotide to another pyrimidine.

Immunoglobulin class switch recombination

A physiological gene rearrangement event in B lymphocytes that replaces the heavy-chain constant region of the expressed immunoglobulin isotype with a different heavy-chain constant region, thereby altering the effector function of the antibody.

CpG islands

Distinct regions of the genome that contain a large number of predominantly unmethylated CG dinucleotide repeats, which are frequently found near mammalian gene promoters.

Promoter-proximal pausing sites

Regions near the promoter of genes where RNA polymerase pauses until it is stimulated to resume transcriptional elongation.

Gene gating

The process of tethering genes to nuclear pore complexes, which enables rapid RNA maturation and export.

Gene conversion

A homology-directed DNA recombination event wherein a resected double-strand break invades the intact sister chromatid and uses it as a template to restore sequences around the break site.

Single-strand annealing

A mutagenic form of DNA repair wherein repetitive regions on either side of a double-strand break are converted into single-stranded DNA by end resection and annealed to one another, resulting in the deletion of the intervening sequences.

Break-induced replication

A non-reciprocal DNA recombination event wherein a resected one-ended double-strand break invades an intact genomic region and initiates long-range unidirectional templated DNA synthesis that typically proceeds to the end of the invaded chromosome.

Synthetic lethality

A condition where combined deficiencies in two or more genes result in cell death, while perturbation of any one of these genes is tolerated.

Phase separation

The separation of a single homogeneous mixture into two distinct phases.

Ribonucleoprotein

A macromolecular complex consisting of both RNA and protein.

Spliceosome

A large multisubunit ribonucleoprotein complex that removes introns from a precursor mRNA transcript.

trans splicing

An RNA splicing event in which exons of two disparate precursor mRNAs are joined together into a single chimeric transcript.

cis splicing of adjacent genes

An event in which two neighbouring genes are read as a single message due to readthrough transcription followed by intergenic splicing.

Epitranscriptomic

Pertaining to RNA modifications that potentially regulate gene expression but that do not involve alterations in the primary ribonucleotide sequence.

Translesion synthesis

A DNA damage tolerance mechanism that enables DNA synthesis past replication-blocking DNA lesions.

Base excision repair

A mode of repair that removes a small, non-helix-distorting base lesion from the genome, leaving an abasic site that is subsequently cleaved and filled in by either non-displacement (short-patch) or strand-displacement (long-patch) DNA synthesis.

Topologically associated domain

(TAD). A genomic domain created by chromosome folding that favours chromatin interactions within its borders.

CTCF-mediated chromatin looping

A process by which CTCF binding to DNA facilitates long-range chromatin interactions between regions that are far apart on the linear chromosome.

RNA–DNA triplex

A form of RNA–DNA interaction in which the RNA strand inserts itself into the major groove of the duplex DNA in a sequence-specific manner.

RNA exosome

A multiprotein ribonuclease complex capable of degrading various types of RNA molecule.

Degradosome

A multicomponent ribonuclease complex that degrades mitochondrial RNA.

Slipped-strand mispairing

Erroneous base pairing in regions of repetitive DNA due to sequence misalignment, resulting in a short stretch of unpaired nucleotides that form a bulge in the DNA.

Non-allelic recombination

A DNA recombination event between any two regions in the genome that have a high degree of sequence homology but are not alleles.

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Zong, D., Oberdoerffer, P., Batista, P.J. et al. RNA: a double-edged sword in genome maintenance. Nat Rev Genet 21, 651–670 (2020). https://doi.org/10.1038/s41576-020-0263-7

Download citation

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41576-020-0263-7

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