Dormant origin firing promotes head-on transcription-replication conflicts at transcription termination sites in response to BRCA2 deficiency.
Journal
Nature communications
ISSN: 2041-1723
Titre abrégé: Nat Commun
Pays: England
ID NLM: 101528555
Informations de publication
Date de publication:
03 Jun 2024
03 Jun 2024
Historique:
received:
19
08
2023
accepted:
24
04
2024
medline:
4
6
2024
pubmed:
4
6
2024
entrez:
3
6
2024
Statut:
epublish
Résumé
BRCA2 is a tumor suppressor protein responsible for safeguarding the cellular genome from replication stress and genotoxicity, but the specific mechanism(s) by which this is achieved to prevent early oncogenesis remains unclear. Here, we provide evidence that BRCA2 acts as a critical suppressor of head-on transcription-replication conflicts (HO-TRCs). Using Okazaki-fragment sequencing (Ok-seq) and computational analysis, we identified origins (dormant origins) that are activated near the transcription termination sites (TTS) of highly expressed, long genes in response to replication stress. Dormant origins are a source for HO-TRCs, and drug treatments that inhibit dormant origin firing led to a reduction in HO-TRCs, R-loop formation, and DNA damage. Using super-resolution microscopy, we showed that HO-TRC events track with elongating RNA polymerase II, but not with transcription initiation. Importantly, RNase H2 is recruited to sites of HO-TRCs in a BRCA2-dependent manner to help alleviate toxic R-loops associated with HO-TRCs. Collectively, our results provide a mechanistic basis for how BRCA2 shields against genomic instability by preventing HO-TRCs through both direct and indirect means occurring at predetermined genomic sites based on the pre-cancer transcriptome.
Identifiants
pubmed: 38830843
doi: 10.1038/s41467-024-48286-1
pii: 10.1038/s41467-024-48286-1
doi:
Substances chimiques
BRCA2 Protein
0
BRCA2 protein, human
0
Ribonuclease H
EC 3.1.26.4
RNA Polymerase II
EC 2.7.7.-
Types de publication
Journal Article
Langues
eng
Sous-ensembles de citation
IM
Pagination
4716Informations de copyright
© 2024. The Author(s).
Références
Siegel, R. L., Miller, K. D., Fuchs, H. E. & Jemal, A. Cancer statistics, 2022. CA Cancer J. Clin. 72, 7–33 (2022).
doi: 10.3322/caac.21708
pubmed: 35020204
Labidi-Galy, S. I. et al. High grade serous ovarian carcinomas originate in the fallopian tube. Nat. Commun. 8, 1093 (2017).
pubmed: 29061967
pmcid: 5653668
doi: 10.1038/s41467-017-00962-1
Lee, Y. et al. A candidate precursor to serous carcinoma that originates in the distal fallopian tube. J. Pathol. 211, 26–35 (2007).
pubmed: 17117391
doi: 10.1002/path.2091
Zhang, S. et al. Both fallopian tube and ovarian surface epithelium are cells-of-origin for high-grade serous ovarian carcinoma. Nat. Commun. 10, 5367 (2019).
pubmed: 31772167
pmcid: 6879755
doi: 10.1038/s41467-019-13116-2
Kroeger, P. T. & Drapkin, R. Pathogenesis and heterogeneity of ovarian cancer. Curr. Opin. Obstet. Gynecol. 29, 26–34 (2017).
pubmed: 27898521
doi: 10.1097/GCO.0000000000000340
Konstantinopoulos, P. A., Ceccaldi, R., Shapiro, G. I. & D’Andrea, A. D. Homologous recombination deficiency: exploiting the fundamental vulnerability of ovarian cancer. Cancer Discov. 5, 1137–1154 (2015).
pubmed: 26463832
pmcid: 4631624
doi: 10.1158/2159-8290.CD-15-0714
Mijic, S. et al. Replication fork reversal triggers fork degradation in BRCA2-defective cells. Nat. Commun. 8, 859 (2017).
pubmed: 29038466
pmcid: 5643541
doi: 10.1038/s41467-017-01164-5
Schlacher, K. et al. Double-strand break repair-independent role for BRCA2 in blocking stalled replication fork degradation by MRE11. Cell 145, 529–542 (2011).
pubmed: 21565612
pmcid: 3261725
doi: 10.1016/j.cell.2011.03.041
Rickman, K. A. et al. Distinct roles of BRCA2 in replication fork protection in response to hydroxyurea and DNA interstrand cross-links. Genes Dev. 34, 832–846 (2020).
pubmed: 32354836
pmcid: 7263144
doi: 10.1101/gad.336446.120
Feng, W. & Jasin, M. BRCA2 suppresses replication stress-induced mitotic and G1 abnormalities through homologous recombination. Nat. Commun. 8, 525 (2017).
pubmed: 28904335
pmcid: 5597640
doi: 10.1038/s41467-017-00634-0
Lemaçon, D. et al. MRE11 and EXO1 nucleases degrade reversed forks and elicit MUS81-dependent fork rescue in BRCA2-deficient cells. Nat. Commun. 8, 860 (2017).
pubmed: 29038425
pmcid: 5643552
doi: 10.1038/s41467-017-01180-5
Panzarino, N. J. et al. Replication gaps underlie BRCA deficiency and therapy response. Cancer Res. 81, 1388–1397 (2021).
pubmed: 33184108
doi: 10.1158/0008-5472.CAN-20-1602
Crossley, M. P., Bocek, M. & Cimprich, K. A. R-loops as cellular regulators and genomic threats. Mol. Cell 73, 398–411 (2019).
pubmed: 30735654
pmcid: 6402819
doi: 10.1016/j.molcel.2019.01.024
Skourti-Stathaki, K. & Proudfoot, N. J. A double-edged sword: R loops as threats to genome integrity and powerful regulators of gene expression. Genes Dev. 28, 1384–1396 (2014).
pubmed: 24990962
pmcid: 4083084
doi: 10.1101/gad.242990.114
Sollier, J. & Cimprich, K. A. Breaking bad: R-loops and genome integrity. Trends Cell Biol. 25, 514–522 (2015).
pubmed: 26045257
pmcid: 4554970
doi: 10.1016/j.tcb.2015.05.003
Hamperl, S. & Cimprich, K. A. Conflict resolution in the genome: how transcription and replication make it work. Cell 167, 1455–1467 (2016).
pubmed: 27912056
pmcid: 5141617
doi: 10.1016/j.cell.2016.09.053
Santos-Pereira, J. M. & Aguilera, A. R loops: new modulators of genome dynamics and function. Nat. Rev. Genet. 16, 583–597 (2015).
pubmed: 26370899
doi: 10.1038/nrg3961
Marnef, A. & Legube, G. R-loops as Janus-faced modulators of DNA repair. Nat. Cell Biol. 23, 305–313 (2021).
pubmed: 33837288
doi: 10.1038/s41556-021-00663-4
Kemiha, S., Poli, J., Lin, Y.-L., Lengronne, A. & Pasero, P. Toxic R-loops: cause or consequence of replication stress? DNA Repair 107, 103199 (2021).
pubmed: 34399314
doi: 10.1016/j.dnarep.2021.103199
Petermann, E., Lan, L. & Zou, L. Sources, resolution and physiological relevance of R-loops and RNA–DNA hybrids. Nat. Rev. Mol. Cell Biol. https://doi.org/10.1038/s41580-022-00474-x (2022).
Cristini, A. et al. RNase H2, mutated in Aicardi‐Goutières syndrome, resolves co-transcriptional R-loops to prevent DNA breaks and inflammation. Nat. Commun. 13, 2961 (2022).
pubmed: 35618715
pmcid: 9135716
doi: 10.1038/s41467-022-30604-0
D’Alessandro, G. et al. BRCA2 controls DNA:RNA hybrid level at DSBs by mediating RNase H2 recruitment. Nat. Commun. 9, 5376 (2018).
pubmed: 30560944
pmcid: 6299093
doi: 10.1038/s41467-018-07799-2
Shivji, M. K. K., Renaudin, X., Williams, Ç. H. & Venkitaraman, A. R. BRCA2 regulates transcription elongation by RNA polymerase II to prevent R-loop accumulation. Cell Rep. 22, 1031–1039 (2018).
pubmed: 29386125
pmcid: 5846855
doi: 10.1016/j.celrep.2017.12.086
Bhatia, V. et al. BRCA2 prevents R-loop accumulation and associates with TREX-2 mRNA export factor PCID2. Nature 511, 362–365 (2014).
pubmed: 24896180
doi: 10.1038/nature13374
Prioleau, M.-N. & MacAlpine, D. M. DNA replication origins—where do we begin? Genes Dev. 30, 1683–1697 (2016).
pubmed: 27542827
pmcid: 5002974
doi: 10.1101/gad.285114.116
Fragkos, M., Ganier, O., Coulombe, P. & Méchali, M. DNA replication origin activation in space and time. Nat. Rev. Mol. Cell Biol. 16, 360–374 (2015).
pubmed: 25999062
doi: 10.1038/nrm4002
Chen, Y.-H. et al. Transcription shapes DNA replication initiation and termination in human cells. Nat. Struct. Mol. Biol. 26, 67–77 (2019).
pubmed: 30598550
doi: 10.1038/s41594-018-0171-0
Zeman, M. K. & Cimprich, K. A. Causes and consequences of replication stress. Nat. Cell Biol. 16, 2–9 (2014).
pubmed: 24366029
pmcid: 4354890
doi: 10.1038/ncb2897
Kawabata, T. et al. Stalled fork rescue via dormant replication origins in unchallenged S phase promotes proper chromosome segregation and tumor suppression. Mol. Cell 41, 543–553 (2011).
pubmed: 21362550
pmcid: 3062258
doi: 10.1016/j.molcel.2011.02.006
Bester, A. C. et al. Nucleotide deficiency promotes genomic instability in early stages of cancer development. Cell 145, 435–446 (2011).
pubmed: 21529715
pmcid: 3740329
doi: 10.1016/j.cell.2011.03.044
Courtot, L., Hoffmann, J.-S. & Bergoglio, V. The protective role of dormant origins in response to replicative stress. Int J. Mol. Sci. 19, 3569 (2018).
pubmed: 30424570
pmcid: 6274952
doi: 10.3390/ijms19113569
Técher, H., Koundrioukoff, S., Nicolas, A. & Debatisse, M. The impact of replication stress on replication dynamics and DNA damage in vertebrate cells. Nat. Rev. Genet. 18, 535–550 (2017).
pubmed: 28714480
doi: 10.1038/nrg.2017.46
Gómez-González, B. & Aguilera, A. Transcription-mediated replication hindrance: a major driver of genome instability. Genes Dev. 33, 1008–1026 (2019).
pubmed: 31123061
pmcid: 6672053
doi: 10.1101/gad.324517.119
Merrikh, H., Zhang, Y., Grossman, A. D. & Wang, J. D. Replication–transcription conflicts in bacteria. Nat. Rev. Microbiol. 10, 449–458 (2012).
pubmed: 22669220
pmcid: 3467967
doi: 10.1038/nrmicro2800
Lang, K. S. & Merrikh, H. The clash of macromolecular titans: replication-transcription conflicts in bacteria. Annu. Rev. Microbiol. 72, 1–18 (2018).
doi: 10.1146/annurev-micro-090817-062514
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.e19 (2017).
pubmed: 28802045
pmcid: 5570545
doi: 10.1016/j.cell.2017.07.043
Stoy, H. et al. Direct visualization of transcription-replication conflicts reveals post-replicative DNA:RNA hybrids. Nat. Struct. Mol. Biol. 30, 348–359 (2023).
pubmed: 36864174
pmcid: 10023573
doi: 10.1038/s41594-023-00928-6
Kumar, C., Batra, S., Griffith, J. D. & Remus, D. The interplay of RNA:DNA hybrid structure and G-quadruplexes determines the outcome of R-loop-replisome collisions. Elife 10, e72286 (2021).
pubmed: 34494544
pmcid: 8479836
doi: 10.7554/eLife.72286
Merrikh, H., Machón, C., Grainger, W. H., Grossman, A. D. & Soultanas, P. Co-directional replication–transcription conflicts lead to replication restart. Nature 470, 554–557 (2011).
pubmed: 21350489
pmcid: 3059490
doi: 10.1038/nature09758
Lang, K. S. et al. Replication-transcription conflicts generate R-loops that orchestrate bacterial stress survival and pathogenesis. Cell 170, 787–799.e18 (2017).
pubmed: 28802046
pmcid: 5630229
doi: 10.1016/j.cell.2017.07.044
Lang, K. S. & Merrikh, H. Topological stress is responsible for the detrimental outcomes of head-on replication-transcription conflicts. Cell Rep. 34, 108797 (2021).
pubmed: 33657379
pmcid: 7986047
doi: 10.1016/j.celrep.2021.108797
Germain, C. P. S. et al. Genomic patterns of transcription–replication interactions in mouse primary B cells. Nucleic Acids Res. 50, 2051–2073 (2022).
doi: 10.1093/nar/gkac035
García-Muse, T. & Aguilera, A. Transcription–replication conflicts: how they occur and how they are resolved. Nat. Rev. Mol. Cell Biol. 17, 553–563 (2016).
pubmed: 27435505
doi: 10.1038/nrm.2016.88
Aguilera, A. & Gaillard, H. Transcription and recombination: when RNA meets DNA. Cold Spring Harb. Perspect. Biol. 6, a016543 (2014).
pubmed: 25085910
pmcid: 4107990
doi: 10.1101/cshperspect.a016543
Barlow, J. H. & Nussenzweig, A. Replication initiation and genome instability: a crossroads for DNA and RNA synthesis. Cell Mol. Life Sci. 71, 4545–4559 (2014).
pubmed: 25238783
pmcid: 6289259
doi: 10.1007/s00018-014-1721-1
Petryk, N. et al. Replication landscape of the human genome. Nat. Commun. 7, 10208 (2016).
pubmed: 26751768
pmcid: 4729899
doi: 10.1038/ncomms10208
Guilbaud, G. et al. Determination of human DNA replication origin position and efficiency reveals principles of initiation zone organisation. Nucleic Acids Res. 50, 7436–7450 (2022).
pubmed: 35801867
pmcid: 9303276
doi: 10.1093/nar/gkac555
Akerman, I. et al. A predictable conserved DNA base composition signature defines human core DNA replication origins. Nat. Commun. 11, 4826 (2020).
pubmed: 32958757
pmcid: 7506530
doi: 10.1038/s41467-020-18527-0
Koyanagi, E. et al. Global landscape of replicative DNA polymerase usage in the human genome. Nat. Commun. 13, 7221 (2022).
pubmed: 36434012
pmcid: 9700718
doi: 10.1038/s41467-022-34929-8
Karst, A. M. & Drapkin, R. Primary culture and immortalization of human fallopian tube secretory epithelial cells. Nat. Protoc. 7, 1755–1764 (2012).
pubmed: 22936217
pmcid: 7433321
doi: 10.1038/nprot.2012.097
Lui, S. K. L. et al. Monitoring genome-wide replication fork directionality by Okazaki fragment sequencing in mammalian cells. Nat. Protoc. 16, 1193–1218 (2021).
pmcid: 8792808
doi: 10.1038/s41596-020-00454-5
Favorov, A. et al. Exploring massive, genome scale datasets with the GenometriCorr package. PLoS Comput. Biol. 8, e1002529 (2012).
pubmed: 22693437
pmcid: 3364938
doi: 10.1371/journal.pcbi.1002529
Ge, X. Q., Jackson, D. A. & Blow, J. J. Dormant origins licensed by excess Mcm2–7 are required for human cells to survive replicative stress. Genes Dev. 21, 3331–3341 (2007).
pubmed: 18079179
pmcid: 2113033
doi: 10.1101/gad.457807
Chen, Y.-H. et al. ATR-Mediated phosphorylation of FANCI regulates dormant origin firing in response to replication stress. Mol. Cell 58, 323–338 (2015).
pubmed: 25843623
pmcid: 4408929
doi: 10.1016/j.molcel.2015.02.031
Duan, H. et al. E3 ligase RFWD3 is a novel modulator of stalled fork stability in BRCA2-deficient cells. J. Cell Biol. 219 p.e201908192 (2020).
Kirstein, N. et al. Human ORC/MCM density is low in active genes and correlates with replication time but does not delimit initiation zones. eLife 10, e62161 (2021).
pubmed: 33683199
pmcid: 7993996
doi: 10.7554/eLife.62161
Scherr, M. J., Wahab, S. A., Remus, D. & Duderstadt, K. E. Mobile origin-licensing factors confer resistance to conflicts with RNA polymerase. Cell Rep. 38, 110531 (2022).
pubmed: 35320708
pmcid: 8961423
doi: 10.1016/j.celrep.2022.110531
Liu, Y. et al. Transcription shapes DNA replication initiation to preserve genome integrity. Genome Biol. 22, 176 (2021).
pubmed: 34108027
pmcid: 8188667
doi: 10.1186/s13059-021-02390-3
Macheret, M. & Halazonetis, T. D. Intragenic origins due to short G1 phases underlie oncogene-induced DNA replication stress. Nature 555, 112–116 (2018).
pubmed: 29466339
pmcid: 5837010
doi: 10.1038/nature25507
Hsin, J.-P. & Manley, J. L. The RNA polymerase II CTD coordinates transcription and RNA processing. Genes Dev. 26, 2119–2137 (2012).
pubmed: 23028141
pmcid: 3465734
doi: 10.1101/gad.200303.112
Egloff, S. & Murphy, S. Cracking the RNA polymerase II CTD code. Trends Genet. 24, 280–288 (2008).
pubmed: 18457900
doi: 10.1016/j.tig.2008.03.008
Yin, Y. et al. A basal-level activity of ATR links replication fork surveillance and stress response. Mol. Cell 81, 4243–4257.e6 (2021).
pubmed: 34473946
pmcid: 8541912
doi: 10.1016/j.molcel.2021.08.009
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).
pubmed: 31570834
pmcid: 6859070
doi: 10.1038/s41556-019-0392-4
Mas, A. M. et al. ORC1 binds to cis-transcribed RNAs for efficient activation of replication origins. Nat. Commun. 14, 4447 (2023).
pubmed: 37488096
pmcid: 10366126
doi: 10.1038/s41467-023-40105-3
Tonzi, P., Yin, Y., Lee, C. W. T., Rothenberg, E. & Huang, T. T. Translesion polymerase kappa-dependent DNA synthesis underlies replication fork recovery. eLife 7, e41426 (2018).
pubmed: 30422114
pmcid: 6251625
doi: 10.7554/eLife.41426
Whelan, D. R. et al. Spatiotemporal dynamics of homologous recombination repair at single collapsed replication forks. Nat. Commun. 9, 3882 (2018).
pubmed: 30250272
pmcid: 6155164
doi: 10.1038/s41467-018-06435-3
Coleman, K. E. et al. USP1-trapping lesions as a source of DNA replication stress and genomic instability. Nat. Commun. 13, 1740 (2022).
pubmed: 35365626
pmcid: 8975806
doi: 10.1038/s41467-022-29369-3
Lee, W. T. C. et al. Single-molecule imaging reveals replication fork coupled formation of G-quadruplex structures hinders local replication stress signaling. Nat. Commun. 12, 2525 (2021).
pubmed: 33953191
pmcid: 8099879
doi: 10.1038/s41467-021-22830-9
Yin, Y., Lee, W. T. C. & Rothenberg, E. Ultrafast data mining of molecular assemblies in multiplexed high-density super-resolution images. Nat. Commun. 10, 119 (2019).
pubmed: 30631072
pmcid: 6328550
doi: 10.1038/s41467-018-08048-2
Rodriguez-Acebes, S., Mourón, S. & Méndez, J. Uncoupling fork speed and origin activity to identify the primary cause of replicative stress phenotypes. J. Biol. Chem. 293, 12855–12861 (2018).
pubmed: 29959228
pmcid: 6102153
doi: 10.1074/jbc.RA118.003740
Jiang, W., McDonald, D., Hope, T. J. & Hunter, T. Mammalian Cdc7–Dbf4 protein kinase complex is essential for initiation of DNA replication. EMBO J. 18, 5703–5713 (1999).
pubmed: 10523313
pmcid: 1171637
doi: 10.1093/emboj/18.20.5703
Toledo, L. I. et al. ATR prohibits replication catastrophe by preventing global exhaustion of RPA. Cell 155, 1088–1103 (2013).
pubmed: 24267891
doi: 10.1016/j.cell.2013.10.043
Petermann, E., Woodcock, M. & Helleday, T. Chk1 promotes replication fork progression by controlling replication initiation. Proc. Natl Acad. Sci. USA 107, 16090–16095 (2010).
pubmed: 20805465
pmcid: 2941317
doi: 10.1073/pnas.1005031107
Rose, M., Burgess, J. T., O’Byrne, K., Richard, D. J. & Bolderson, E. PARP inhibitors: clinical relevance, mechanisms of action and tumor resistance. Front. Cell Dev. Biol. 8, 564601 (2020).
pubmed: 33015058
pmcid: 7509090
doi: 10.3389/fcell.2020.564601
Cong, K. et al. Replication gaps are a key determinant of PARP inhibitor synthetic lethality with BRCA deficiency. Mol. Cell https://doi.org/10.1016/j.molcel.2021.06.011 (2021).
Maya-Mendoza, A. et al. High speed of fork progression induces DNA replication stress and genomic instability. Nature 559, 279–284 (2018).
pubmed: 29950726
doi: 10.1038/s41586-018-0261-5
Cimprich, K. A. & Cortez, D. ATR: an essential regulator of genome integrity. Nat. Rev. Mol. Cell Biol. 9, 616–627 (2008).
pubmed: 18594563
pmcid: 2663384
doi: 10.1038/nrm2450
Moiseeva, T. et al. ATR kinase inhibition induces unscheduled origin firing through a Cdc7-dependent association between GINS and And-1. Nat. Commun. 8, 1392 (2017).
pubmed: 29123096
pmcid: 5680267
doi: 10.1038/s41467-017-01401-x
Montagnoli, A. et al. A Cdc7 kinase inhibitor restricts initiation of DNA replication and has antitumor activity. Nat. Chem. Biol. 4, 357–365 (2008).
pubmed: 18469809
doi: 10.1038/nchembio.90
Promonet, A. et al. Topoisomerase 1 prevents replication stress at R-loop-enriched transcription termination sites. Nat. Commun. 11, 3940 (2020).
pubmed: 32769985
pmcid: 7414224
doi: 10.1038/s41467-020-17858-2
Cerritelli, S. M. & Crouch, R. J. Ribonuclease H: the enzymes in eukaryotes. FEBS J. 276, 1494–1505 (2009).
pubmed: 19228196
doi: 10.1111/j.1742-4658.2009.06908.x
Cerritelli, S. M., Sakhuja, K. & Crouch, R. J. RNase H1, the gold standard for R-loop detection. Methods Mol. Biol. 2528, 91–114 (2022).
pubmed: 35704187
doi: 10.1007/978-1-0716-2477-7_7
Zhang, H. et al. Integrated proteogenomic characterization of human high-grade serous ovarian cancer. Cell 166, 755–765 (2016).
pubmed: 27372738
pmcid: 4967013
doi: 10.1016/j.cell.2016.05.069
Powell, S. K. et al. Dynamic loading and redistribution of the Mcm2‐7 helicase complex through the cell cycle. EMBO J. 34, 531–543 (2015).
pubmed: 25555795
pmcid: 4331006
doi: 10.15252/embj.201488307
Gros, J. et al. Post-licensing specification of eukaryotic replication origins by facilitated Mcm2-7 sliding along DNA. Mol. Cell 60, 797–807 (2015).
pubmed: 26656162
pmcid: 4680849
doi: 10.1016/j.molcel.2015.10.022
Stork, C. T. et al. Co-transcriptional R-loops are the main cause of estrogen-induced DNA damage. eLife 5, e17548 (2016).
pubmed: 27552054
pmcid: 5030092
doi: 10.7554/eLife.17548
Niehrs, C. & Luke, B. Regulatory R-loops as facilitators of gene expression and genome stability. Nat. Rev. Mol. Cell Biol. 21, 167–178 (2020).
pubmed: 32005969
pmcid: 7116639
doi: 10.1038/s41580-019-0206-3
Sohn, M.-H. et al. Classification of high-grade serous ovarian carcinoma by epithelial-to-mesenchymal transition signature and homologous recombination repair genes. Genes 12, 1103 (2021).
pubmed: 34356119
pmcid: 8303300
doi: 10.3390/genes12071103
Marcus, J. et al. Septin 9 isoforms promote tumorigenesis in mammary epithelial cells by increasing migration and ECM degradation through metalloproteinase secretion at focal adhesions. Oncogene 38, 5839–5859 (2019).
pubmed: 31285548
pmcid: 6859949
doi: 10.1038/s41388-019-0844-0
Karst, A. M., Levanon, K. & Drapkin, R. Modeling high-grade serous ovarian carcinogenesis from the fallopian tube. Proc. Natl Acad. Sci. USA 108, 7547–7552 (2011).
pubmed: 21502498
pmcid: 3088633
doi: 10.1073/pnas.1017300108
Dolgalev, I. Seq-N-Slide https://doi.org/10.5281/zenodo.5550459 (2022).
Sanz, L. A. & Chédin, F. High-resolution, strand-specific R-loop mapping via S9.6-based DNA–RNA immunoprecipitation and high-throughput sequencing. Nat. Protoc. 14, 1734–1755 (2019).
pubmed: 31053798
pmcid: 6615061
doi: 10.1038/s41596-019-0159-1
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).
pubmed: 25699710
pmcid: 4351672
doi: 10.1016/j.molcel.2015.01.011
McDermott, J. E. et al. Proteogenomic characterization of ovarian HGSC implicates mitotic kinases, replication stress in observed chromosomal instability. Cell Rep. Med. 1, 100004 (2020).
pubmed: 32529193
pmcid: 7289043
doi: 10.1016/j.xcrm.2020.100004