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.

RNA transcripts stimulate homologous recombination by forming DR-loops

Abstract

Homologous recombination (HR) repairs DNA double-strand breaks (DSBs) in the S and G2 phases of the cell cycle1,2,3. Several HR proteins are preferentially recruited to DSBs at transcriptionally active loci4,5,6,7,8,9,10, but how transcription promotes HR is poorly understood. Here we develop an assay to assess the effect of local transcription on HR. Using this assay, we find that transcription stimulates HR to a substantial extent. Tethering RNA transcripts to the vicinity of DSBs recapitulates the effects of local transcription, which suggests that transcription enhances HR through RNA transcripts. Tethered RNA transcripts stimulate HR in a sequence- and orientation-dependent manner, indicating that they function by forming DNA–RNA hybrids. In contrast to most HR proteins, RAD51-associated protein 1 (RAD51AP1) only promotes HR when local transcription is active. RAD51AP1 drives the formation of R-loops in vitro and is required for tethered RNAs to stimulate HR in cells. Notably, RAD51AP1 is necessary for the DSB-induced formation of DNA–RNA hybrids in donor DNA, linking R-loops to D-loops. In vitro, RAD51AP1-generated R-loops enhance the RAD51-mediated formation of D-loops locally and give rise to intermediates that we term ‘DR-loops’, which contain both DNA–DNA and DNA–RNA hybrids and favour RAD51 function. Thus, at DSBs in transcribed regions, RAD51AP1 promotes the invasion of RNA transcripts into donor DNA, and stimulates HR through the formation of DR-loops.

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

Relevant articles

Open Access articles citing this article.

Access options

Buy article

Get time limited or full article access on ReadCube.

$32.00

All prices are NET prices.

Fig. 1: HR is stimulated by local transcription.
Fig. 2: Tethering of RNA transcripts to DSB recapitulates the effects of local transcription on HR.
Fig. 3: RAD51AP1 promotes HR in a transcription- and RNA-dependent manner.
Fig. 4: RAD51AP1 drives R-loop formation in vitro and in cells.
Fig. 5: RAD51AP1 and RAD51 act together to form DR-loops.

Data availability

All relevant data are included in the Article and/or Supplementary Figs. 1, 2.

References

  1. Prakash, R., Zhang, Y., Feng, W. & Jasin, M. Homologous recombination and human health: the roles of BRCA1, BRCA2, and associated proteins. Cold Spring Harb. Perspect. Biol. 7, a016600 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  2. Heyer, W. D., Ehmsen, K. T. & Liu, J. Regulation of homologous recombination in eukaryotes. Annu. Rev. Genet. 44, 113–139 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Daley, J. M., Gaines, W. A., Kwon, Y. & Sung, P. Regulation of DNA pairing in homologous recombination. Cold Spring Harb. Perspect. Biol. 6, a017954 (2014).

    Article  PubMed  PubMed Central  Google Scholar 

  4. 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).

    Article  CAS  PubMed  Google Scholar 

  5. Ouyang, J., Lan, L. & Zou, L. Regulation of DNA break repair by transcription and RNA. Sci. China Life Sci. 60, 1081–1086 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  7. Aymard, F. et al. Transcriptionally active chromatin recruits homologous recombination at DNA double-strand breaks. Nat. Struct. Mol. Biol. 21, 366–374 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  8. 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).

    Article  CAS  PubMed  Google Scholar 

  9. Teng, Y. et al. ROS-induced R loops trigger a transcription-coupled but BRCA1/2-independent homologous recombination pathway through CSB. Nat. Commun. 9, 4115 (2018).

    Article  ADS  PubMed  PubMed Central  CAS  Google Scholar 

  10. Chen, H. et al. m5C modification of mRNA serves a DNA damage code to promote homologous recombination. Nat. Commun. 11, 2834 (2020).

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  11. Pierce, A. J., Johnson, R. D., Thompson, L. H. & Jasin, M. XRCC3 promotes homology-directed repair of DNA damage in mammalian cells. Genes Dev. 13, 2633–2638 (1999).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. Zetsche, B. et al. Cpf1 is a single RNA-guided endonuclease of a class 2 CRISPR–Cas system. Cell 163, 759–771 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  13. Pinder, J., Salsman, J. & Dellaire, G. Nuclear domain ‘knock-in’ screen for the evaluation and identification of small molecule enhancers of CRISPR-based genome editing. Nucleic Acids Res. 43, 9379–9392 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Chavez, A. et al. Highly efficient Cas9-mediated transcriptional programming. Nat. Methods 12, 326–328 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  15. Modesti, M. et al. RAD51AP1 is a structure-specific DNA binding protein that stimulates joint molecule formation during RAD51-mediated homologous recombination. Mol. Cell 28, 468–481 (2007).

    Article  CAS  PubMed  Google Scholar 

  16. Wiese, C. et al. Promotion of homologous recombination and genomic stability by RAD51AP1 via RAD51 recombinase enhancement. Mol. Cell 28, 482–490 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  17. Liang, F. et al. Promotion of RAD51-mediated homologous DNA pairing by the RAD51AP1–UAF1 complex. Cell Rep. 15, 2118–2126 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  18. Kovalenko, O. V., Golub, E. I., Bray-Ward, P., Ward, D. C. & Radding, C. M. A novel nucleic acid-binding protein that interacts with human rad51 recombinase. Nucleic Acids Res. 25, 4946–4953 (1997).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Dunlop, M. H. et al. Mechanistic insights into RAD51-associated protein 1 (RAD51AP1) action in homologous DNA repair. J. Biol. Chem. 287, 12343–12347 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Taylor, S. C., Laperriere, G. & Germain, H. Droplet digital PCR versus qPCR for gene expression analysis with low abundant targets: from variable nonsense to publication quality data. Sci. Rep. 7, 2409 (2017).

    Article  ADS  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

  23. 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).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  24. Sharma, S. et al. MRE11-RAD50-NBS1 complex is sufficient to promote transcription by RNA polymerase II at double-strand breaks by melting DNA ends. Cell Rep. 34, 108565 (2021).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Petukhova, G., Sung, P. & Klein, H. Promotion of Rad51-dependent D-loop formation by yeast recombination factor Rdh54/Tid1. Genes Dev. 14, 2206–2215 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  26. Van Komen, S., Petukhova, G., Sigurdsson, S., Stratton, S. & Sung, P. Superhelicity-driven homologous DNA pairing by yeast recombination factors Rad51 and Rad54. Mol. Cell 6, 563–572 (2000).

    Article  PubMed  Google Scholar 

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

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  29. 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.e3 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  30. Meers, C. et al. Genetic characterization of three distinct mechanisms supporting RNA-driven DNA repair and modification reveals major role of DNA polymerase ζ. Mol. Cell 79, 1037–1050 (2020).

    Article  PubMed  CAS  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

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

    Article  ADS  PubMed  PubMed Central  CAS  Google Scholar 

  33. Hatchi, E. et al. BRCA1 and RNAi factors promote repair mediated by small RNAs and PALB2–RAD52. Nature 591, 665–670 (2021).

    Article  ADS  CAS  PubMed  Google Scholar 

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

    Article  ADS  PubMed  PubMed Central  CAS  Google Scholar 

  35. Shanbhag, N. M., Rafalska-Metcalf, I. U., Balane-Bolivar, C., Janicki, S. M. & Greenberg, R. A. ATM-dependent chromatin changes silence transcription in cis to DNA double-strand breaks. Cell 141, 970–981 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  36. Greene, E. C. DNA sequence alignment during homologous recombination. J. Biol. Chem. 291, 11572–11580 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Baumann, P., Benson, F. E. & West, S. C. Human Rad51 protein promotes ATP-dependent homologous pairing and strand transfer reactions in vitro. Cell 87, 757–766 (1996).

    Article  CAS  PubMed  Google Scholar 

  38. Cohen, S. et al. Senataxin resolves RNA:DNA hybrids forming at DNA double-strand breaks to prevent translocations. Nat. Commun. 9, 533 (2018).

    Article  ADS  PubMed  PubMed Central  CAS  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

  40. O’Connor, M. J. Targeting the DNA damage response in cancer. Mol. Cell 60, 547–560 (2015).

    Article  PubMed  CAS  Google Scholar 

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

    Article  CAS  PubMed  Google Scholar 

  42. Meerbrey, K. L. et al. The pINDUCER lentiviral toolkit for inducible RNA interference in vitro and in vivo. Proc. Natl Acad. Sci. USA 108, 3665–3670 (2011).

    Article  ADS  CAS  PubMed  Google Scholar 

  43. Chailleux, C. et al. Quantifying DNA double-strand breaks induced by site-specific endonucleases in living cells by ligation-mediated purification. Nat. Protocols 9, 517–528 (2014).

    Article  CAS  PubMed  Google Scholar 

  44. Kleinstiver, B. P. et al. Engineered CRISPR–Cas12a variants with increased activities and improved targeting ranges for gene, epigenetic and base editing. Nat. Biotechnol. 37, 276–282 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  45. Tumini, E. & Aguilera, A. The sister-chromatid exchange assay in human cells. Methods Mol. Biol. 2153, 383–393 (2021).

    Article  CAS  PubMed  Google Scholar 

  46. Moquin, D. M. et al. Localized protein biotinylation at DNA damage sites identifies ZPET, a repressor of homologous recombination. Genes Dev. 33, 75–89 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  47. 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.e4 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

Download references

Acknowledgements

We thank P. Sung, G. Legube, J. K. Joung, J. Jin and G. Gill for reagents, and N. Dyson, A. Elia and members of the L.Z. and N. Dyson laboratories for discussions. L.Z. is the James & Patricia Poitras Endowed Chair in Cancer Research. This work is supported by grants from the NIH (CA197779 and CA218856) to L.Z.

Author information

Authors and Affiliations

Authors

Contributions

J.O., T.Y. and L.Z. designed the study. J.O. and T.Y. performed the experiments and data analyses. L.Z. supervised the experiments and data analyses. J.-M.Z., H.Y. and H.G. provided technical support. E.R. analysed gene expression data. D.A.H. and L.L. helped to supervise the study. J.O., T.Y. and L.Z. prepared the manuscript with contributions from all authors.

Corresponding authors

Correspondence to Jian Ouyang or Lee Zou.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Peer review information Nature thanks the 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.

Extended data figures and tables

Extended Data Fig. 1 Characterizations of Tet-DR-GFP and ASCL1-mClover HR reporters.

a, Flow cytometry analysis of a U2OS-derivative cell line in which the Tet-DR-GFP reporter is stably integrated. Cells were mock transfected, transfected with a control plasmid expressing mCherry or transfected with a plasmid expressing I-SceI-T2A-mCherry. GFP+ cells were only detected when I-SceI-T2A-mCherry was expressed and transcription of sceGFP was induced by Dox. b, Detection of repaired eGFP by qPCR. Cells carrying the Tet-DR-GFP reporter were mock transfected or transfected with a plasmid expressing I-Sce-T2A-mCherry (−/+ DSBs) in the absence and presence of Dox (−/+Dox). Genomic DNA was analysed for the repaired eGFP sequence by qPCR. c, FACS- and qPCR-based HR assays detect changes of HR efficiency similarly. Cells were transfected with siRNAs to knock down the indicated HR proteins and analysed by FACS- and qPCR-based HR assays. d, e, The transcription status of sceGFP does not affect I-SceI induced DSB formation. The I-SceI-induced DNA ends in sceGFP were ligated to a sequence-specific adaptor and quantified by qPCR. d, Amplification plot of qPCR. e, Linear quantification of DSBs by qPCR. The indicated amounts of genomic DNA containing I-SceI-generated DSBs were mixed with uncut genomic DNA, so that a total of 1 μg genomic DNA was analysed in each qPCR sample. f, Two additional stable U2OS clones carrying the Tet-DR-GFP reporter were used to analyse the ratio of HR levels in transcriptionally off and on states (−Dox/+Dox). Data are mean (n = 2 independent experiments). g, Total RNA isolated from HEK293T cells was reversely transcribed using random primers. cDNA was subjected to qPCR analysis to determine the relative expression levels of CRISPR-dCas9-VPR-activated ASCL1 and a panel of endogenous genes using the GAPDH transcript as a reference. Data of ASCL1 are mean (n = 4 independent experiments).

Extended Data Fig. 2 Effects of tethered RNA on HR.

a, The indicated fusion RNAs were tethered to a unique sequence 5′ to the I-SceI site in sceGFP. HR efficiency in cells expressing various fusion RNAs was analysed by qPCR in the transcriptionally on and off states. For each RNA, the HR level in the transcriptionally on state is defined as 1. The ratios of HR levels between the transcriptionally off and on states (−Dox/+Dox) were determined. Data are mean ± s.e.m. (n = 3 independent experiments). b, Two fusion RNAs containing 120-nt or 81-nt GFP sequence were tethered to a unique sequence 5′ to the I-SceI site in sceGFP. Relative HR efficiencies in cells expressing either of the two fusion RNAs or no fusion RNA were measured by qPCR in the transcriptionally off state (−Dox). The HR efficiency of cells expressing no fusion RNA in the transcriptionally on state (+Dox) serves as a reference. Data are mean ± s.e.m. (n = 3 independent experiments). c, Schematic to explain why only sense GFP RNA and not anti-sense GFP RNA can hybridize DNA when tethered 5′ to the I-SceI site. It also explains why anti-sense GFP RNA can hybridize DNA when tethered 3′ to the I-SceI site.

Extended Data Fig. 3 Effects of depletion of RAD51AP1, UAF1 and RAD52 on HR.

a, Cells were transfected with control or two independent RAD51AP1 siRNAs. Levels of endogenous RAD51AP1 and β-actin (loading control) were analysed by western blot. A representative western blot of three similar experiments is shown. b, HR efficiency was measured by FACS using U2OS-Tet-DR-GFP reporter cells after knockdown of RAD51AP1. Data are mean ± s.e.m (n = 3 independent experiments). c, Cells were transfected with control or UAF1 siRNA. Levels of UAF1 mRNA were analysed by RT–qPCR. Data are mean (n = 2 independent experiments). d, Cells were transfected with control, RAD51AP1 or UAF1 siRNA. Levels of endogenous RAD51AP1 and β-actin (loading control) were analysed by western blot. A representative western blot of three similar experiments is shown. e, HR efficiency was measured by FACS using U2OS-Tet-DR-GFP reporter cells after knockdown of RAD51AP1 or UAF1. Data are mean (n = 2 independent experiments). f, Cells were transfected with control or two independent RAD52 siRNAs. Levels of endogenous RAD52 and KU80 (loading control) were analysed by western blot. A representative western blot of three similar experiments is shown. g, HR efficiency was measured by FACS using U2OS-Tet-DR-GFP reporter cells after knockdown of RAD52. Data are mean ± s.e.m (n = 3 independent experiments). h, HR efficiency was measured by qPCR in transcriptionally on and off states (+/−Dox) after knockdown of RAD52. The HR efficiency of control siRNA transfected cells in the transcriptionally on state (+Dox) serves as a reference. Data are mean ± s.e.m (n = 3 independent experiments).

Extended Data Fig. 4 Functions and localization of RAD51AP1 in the DSB response.

a, Representative mitotic spreads showing SCEs. Samples were prepared from U2OS cells transfected with control or RAD51AP1 siRNA, and treated with 3 nM CPT or vehicle for 24 h. Scale bars, 10 μm. b, c, Survival analyses of cells treated with etoposide (b) or CPT (c). Data are mean (n = 2 independent experiments). d, e, ChIP–qPCR analysis of RAD51AP1 (d) or RAD51 (e) at sites of AsiSI-induced DSBs in transcriptionally active and inactive regions. AsiSI-ER was activated by 4-OHT. f, RAD51AP1 ChIP signals at several transcriptionally active and inactive AsiSI sites were normalized to XRCC4 ChIP signals. XRCC4, which binds to DSBs independently of transcription, serves as a reference for RAD51AP1. RAD51AP1 knockdown substantially reduced RAD51AP1 ChIP signals, confirming the specificity of RAD51AP1 ChIP. Data in ac are mean (n = 2 independent experiments).

Extended Data Fig. 5 Regulation of RAD51AP1 localization to DSBs.

a, U2OS cells carrying an array of tetracycline responsive elements (TREs) were transfected with a plasmid expressing the TetR-VP16-KillerRed (TA-KR) fusion protein. The TA-KR fusion protein binds the TRE array and induces DSBs upon light activation. The formation of RAD51AP1 or RAD51 foci at the TA-KR-marked locus was analysed by immunostaining. Scale bars, 10 μm. b, Immunostaining of RAD51AP1 in cells that were not irradiated or were irradiated with 2 Gy IR. Cells were analysed 2 h after IR. Scale bars, 10 μm. c, Correlation between the numbers of RAD51AP1 and RPA32 foci in IR-treated cells as determined by linear regression. The numbers of RAD51AP1 and RPA32 foci were quantified in individual cells (n = 260 cells analysed in one experiment). Individual cells were plotted according to the numbers of RAD51AP1 and RPA32 foci in them. dg, Asynchronously growing U2OS cells were treated with or without DRB for 4 h, exposed to 2 Gy IR or mock-treated, and analysed in 2 h. d, Immunostaining of RAD51AP1 and γH2AX. Scale bars, 10 μm. e, Numbers of RAD51AP1 foci in individual cells were plotted as mean ± s.d. (n = 401 cells for no IR, n = 408 cells for +IR, and n = 408 cells for +IR +DRB, analysed in one experiment). f, Numbers of γH2AX foci in individual cells were plotted as mean ± s.d. (n = 313 cells for −DRB and n = 294 cells for +DRB, analysed in one experiment). g, Numbers of RAD51AP1 foci in PCNA+ cells were plotted as mean ± s.d. (n = 360 cells for −DRB and n = 303 cells for +DRB, analysed in one experiment). h, i, U2OS cells transfected with control, UAF1 (h) or CtIP (i) siRNA were irradiated with 2 Gy IR. Immunostaining of RAD51AP1 was done 2 h after IR. Numbers of RAD51AP1 foci in individual cells were plotted as mean ±s.d. ***P < 0.001 (two-sided Student’s t test; P < 0.0001 in h, i. h, n = 483 cells for siCTRL, n = 527 cells for siUAF1 analysed in one experiment. i, n = 200 cells for siCTRL, n = 182 cells for siCtIP analysed in one experiment.

Extended Data Fig. 6 Characterizations of the ssDNA-, ssRNA- and dsDNA-binding activities of RAD51AP1.

a, Purified GST, GST-RAD51AP1WT and GST-RAD51AP1DBM were analysed by SDS–PAGE stained with Coomassie blue. Representative gel images from 3 similar experiments are shown. b, Increasing concentrations of RAD51AP1WT or RAD51AP1DBM (0, 12.5, 25, 50 and 100 nM) were incubated with labelled 63-nt ssDNA. Formation of the RAD51AP1–ssDNA complex was analysed by EMSA. The efficiency of complex formation was determined by quantifying the reduction in free ssDNA. Representative results from three similar experiments are shown. c, Increasing concentrations of RAD51AP1WT or RAD51AP1DBM (0, 50, 100, 200 and 400 nM) were incubated with labelled 55-bp dsDNA. Formation of the RAD51AP1–dsDNA complex was analysed by EMSA. The efficiency of complex formation was determined by quantifying the reduction in free dsDNA. Data are mean (n = 2 independent experiments). d, Increasing concentrations of RAD51AP1WT (0, 6.25, 12.5, 25 and 50 nM) were incubated with labelled 63-nt ssRNA and 80-nt ssDNA. Formation of the complex was analysed by EMSA. The efficiency of complex formation was determined by quantifying the reduction in free ssDNA or ssRNA. Representative results from three similar experiments are shown. e, In Fig. 4a, the efficiency of RAD51AP1–ssRNA complex formation was determined by quantifying the reduction in free ssRNA. Data are mean ± s.d. (n = 3 independent experiments).

Extended Data Fig. 7 Characterizations of the RNA-binding and R-loop-formation activities of RAD51AP1.

a, In Fig. 4b, the efficiency of RAD51AP1–hybrid complex formation was determined by quantifying the reduction in free DNA–RNA hybrid. Data are mean ± s.d. (n = 3 independent experiments). b, Increasing concentrations of RAD51AP1WT or RAD51AP1DBM (0, 50, 100, 200 and 400 nM) were incubated with labelled 25-bp dsRNA. Formation of the RAD51AP1–dsRNA complex was analysed by EMSA. The efficiency of complex formation was determined by quantifying the reduction in free dsRNA. Data are presented as mean (n = 2 independent experiments). c, Increasing concentrations of RAD51AP1WT (0, 50, 100 200 and 400 nM) were incubated with labelled 25-nt ssRNA. Formation of the RAD51AP1–ssRNA complex was analysed by EMSA. The efficiency of complex formation was determined by quantifying the reduction in free ssRNA. Data are mean (n = 2 independent experiments). d, In Fig. 4c, the efficiency of R-loop formation was determined by quantifying the shifted and unshifted bands in light exposures of the gel. Data are mean (n = 2 independent experiments). e, In Fig. 4d, the efficiency of R-loop formation was determined as in d. Data are mean (n = 2 independent experiments). f, In vitro R-loop formation with RAD51AP1 (0.4 μM) and labelled scrambled ssRNA or ssRNA (50 nM) with homology to dsDNA. Formation of R-loops was analysed by native gel electrophoresis. Representative results from three similar experiments are shown. g, R-loop formation with increasing concentration of RAD51AP1 (0.1, 0.2 μM), labelled ssRNA (50 nM) and a dsDNA plasmid containing a sequence homologous to the ssRNA in buffer D without ATP. Formation of R-loops was analysed by native gel electrophoresis. Representative results from three similar experiments are shown. h, In vitro strand exchange between ssRNA and dsDNA. Increasing concentrations of RAD51AP1 (0.25, 0.5, 1, 2 μM) were first incubated with ssRNA (63 nt, 200 nM) and then with fluorescent labelled linear dsDNA (30 nM). Strand-exchanged products were separated by 10% native polyacrylamide-TBE gel and imaged. Data are mean (n = 2 independent experiments).

Extended Data Fig. 8 Comparing RAD51AP1, RAD51 and RAD52 in R-loop formation.

a, In vitro D-loop formation with RAD51AP1 and RAD51. RAD51 was incubated with labelled 90-nt ssDNA for 5 min and then with RAD51AP1 for another 5 min. A dsDNA plasmid containing a sequence homologous to the ssDNA was then added to the reactions. Formation of D-loops was analysed by native gel electrophoresis. The efficiency of D-loop formation was determined by quantifying the shifted and unshifted bands in a light exposure of the gel. Representative results from three similar experiments are shown. b, In vitro R-loop formation with RAD51AP1 and RAD51. RAD51 was incubated with labelled 63-nt ssRNA and then with RAD51AP1 for another 5 min. A dsDNA plasmid containing a sequence homologous to the ssRNA was then added to the reactions. The concentrations of RAD51 and RAD51AP1 are indicated. Formation of R-loops was analysed by native gel electrophoresis. Representative results from three similar experiments are shown. c, In vitro R-loop formation with RAD51AP1 and RAD51 was analysed as in b. The concentrations of RAD51 and RAD51AP1 are indicated. Representative results from three similar experiments are shown. d, RAD51AP1 was incubated with labelled 63-nt ssRNA for 5 min and then with increasing concentrations of RAD51 for another 5 min. A dsDNA plasmid containing a sequence homologous to the ssRNA was then added to the reactions. Formation of R-loops were analysed by native gel electrophoresis. Representative results from two similar experiments are shown. e, In vitro R-loop formation activities of RAD51AP1 and RAD52. Increasing concentrations of RAD51AP1 (0.1, 0.2 and 0.4 μM) or RAD52 (0.1, 0.2 and 0.4 μM) were incubated with labelled 63-nt ssRNA and then with a dsDNA plasmid containing a sequence homologous to the ssRNA. Formation of R-loops were analysed by native gel electrophoresis. The efficiency of R-loop formation was determined by quantifying the shifted and unshifted bands in a light exposure of the gel. Data are mean (n = 2 independent experiments).

Extended Data Fig. 9 UAF1 stimulates RAD51AP1-mediated R-loop formation in vitro.

a, In vitro D-loop formation with RAD51 or the RAD51AP1–UAF1 complex. RAD51 was incubated with labelled 60-nt ssDNA and then with RAD51AP1 or the RAD51AP1–UAF1 complex. A dsDNA plasmid containing a sequence homologous to the ssDNA was then added to the reactions. Formation of D-loops was analysed by native gel electrophoresis. Representative results from 2 similar experiments are shown. b, In vitro R-loop formation with the RAD51AP1–UAF1 complex. Preformed RAD51AP1–UAF1 complexes were incubated with labelled 63-nt ssRNA and then with a dsDNA plasmid containing a sequence homologous to the ssRNA. Formation of R-loops was analysed by native gel electrophoresis. The efficiency of R-loop formation was determined by quantifying the shifted and unshifted bands in a light exposure of the gel. Data are mean ± s.d. (n = 3 independent experiments).

Extended Data Fig. 10 Characterizations of RNA transcripts and DNA–RNA hybrids at sceGFP and iGFP loci.

a, Total RNA isolated from U2OS-Tet-DR-GFP cells treated or untreated with Dox were digested with dsDNase and reverse-transcribed using random primers. cDNA was subjected to qPCR analysis to determine the relative levels of sceGFP and iGFP transcripts using the GAPDH transcript as a reference. The sceGFP transcript was increased by more than 60-fold by Dox. In the absence of Dox, the sceGFP transcript was more than 300-fold more abundant than the iGFP transcript. In the presence of Dox, the sceGFP transcript was more than 2,500-fold more abundant than the iGFP transcript. These results suggest that only sceGFP, and not iGFP, is transcribed in the presence of Dox. b, The accumulation of DNA–RNA hybrids in sceGFP and iGFP was analysed by DRIP–ddPCR as in Fig. 4f. In the +RNH (RNaseH) samples, extracted total nucleic acids were treated with RNaseH before being subjected to DRIP analysis. Data are mean (n = 2 independent experiments). c, The levels of DNA–RNA hybrids were tested by DRIP–ddPCR using primers that specifically detect an internal region between sceGFP and iGFP (orange arrowheads). These specific primers do not detect any sceGFP or iGFP containing DNA fragments after the restriction digestion during DRIP–ddPCR. The levels of DNA–RNA hybrids were measured in transcriptionally on and off states (+/− Dox) after DSB induction (+I-SceI). Data are mean (n = 2 independent experiments).

Extended Data Fig. 11 RAD51AP1 promotes the formation of DR-loops in donor DNA.

a, Experimental design for in vitro DR-loop formation using biotinylated ssRNA and fluorescently labelled ssDNA. b, In vitro DR-loop formation with biotinylated ssRNA (63 nt, 30 nM), RAD51AP1 (1 μM), RAD51 (0.25 μM), fluorescently labelled ssDNA (60 nt, 20 nM) and a dsDNA plasmid. The presence or absence of the indicated reaction components is shown above the dot blot. The levels of ssDNA captured by biotinylated ssRNA via DR-loops were measured by dot blot and quantified over blot background. Data are mean (n = 2–3 independent experiments). c, In vitro DR-loop formation with biotinylated ssRNA (50 nt, 30 nM), RAD51AP1WT (0.5 μM) or RAD51AP1DBM (0.5 μM), RAD51 (0.25 μM), fluorescently labelled ssDNA (50 nt, 20 nM) and a dsDNA plasmid. The presence or absence of the indicated reaction components is shown above the dot blot. The levels of ssDNA captured by biotinylated ssRNA via DR-loops were measured by dot blot and quantified over blot background. Representative results from two similar experiments are shown. d, In vitro DR-loop confirmation with RNaseH treatment. Data are mean (n = 2–3 independent experiments). e, In vitro DR-loop formation with different ssRNA and ssDNA oligos and dsDNA plasmids. Top, schematic of the ssRNA and ssDNA oligos tested. Red, biotinylated ssRNA; purple, fluorescently labelled ssDNA. The lengths and relative annealing positions of the oligos are indicated. pBSK+ and pMLM-mini are two different dsDNA plasmids that were used in the reactions. Both of them contain sequences homologous to the ssRNA and ssDNA oligos. f, Schematic of the three ssDNA oligos (red) and the ssRNA oligo (green) used in Fig. 5c, d. g, In vitro D-loop formation with RAD51 and ssDNA oligos A–C. RAD51 (0.25 μM) was incubated with labelled 60-nt ssDNA oligos A, B or C (30 nM) and then with a dsDNA plasmid containing sequences homologous to oligos A and B but not C. Formation of D-loops was analysed by native gel electrophoresis. The efficiency of D-loop formation was determined by quantifying the shifted and unshifted bands in a light exposure of the gel. Data are mean ± s.d. (n = 3 independent experiments).

Extended Data Fig. 12 Model of the role of DR-loops in HR.

The formation of DSBs in transcribed regions triggers the recruitment of both RAD51 and RAD51AP1. RAD51AP1 may associate with RNA transcripts through direct RNA binding. In S and G2 cells, RAD51AP1 promotes the invasion of RNA transcripts into donor DNA on sister chromatids. The presence of R-loops and RAD51AP1 in donor DNA may facilitate the invasion of RAD51 filaments, stabilize invaded ssDNA, help RAD51 find homologous sequences and promote the extension of ssDNA ends in DR-loops. It is also possible that RNA transcripts anneal with the displaced ssDNA in D-loops, helping to stabilize invaded ssDNA and extend ssDNA ends in DR-loops. The DNA–RNA hybrids in DR-loops are probably removed in a later step to allow the completion of HR.

Supplementary information

Supplementary Figures

This file contains Supplementary Figs 1-2 - the uncropped blots and FACS gating strategy.

Reporting Summary

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Ouyang, J., Yadav, T., Zhang, JM. et al. RNA transcripts stimulate homologous recombination by forming DR-loops. Nature 594, 283–288 (2021). https://doi.org/10.1038/s41586-021-03538-8

Download citation

  • Received:

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41586-021-03538-8

This article is cited by

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.

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