Topoisomerase 1 prevents replication stress at R-loop-enriched transcription termination sites.
Ataxia Telangiectasia Mutated Proteins
/ metabolism
DNA Breaks, Double-Stranded
DNA Replication
DNA Topoisomerases, Type I
/ genetics
Gene Knockdown Techniques
Genomic Instability
HEK293 Cells
HeLa Cells
Humans
Phosphorylation
Promoter Regions, Genetic
R-Loop Structures
/ genetics
RNA, Small Interfering
/ metabolism
Terminator Regions, Genetic
/ genetics
Transcription, Genetic
Journal
Nature communications
ISSN: 2041-1723
Titre abrégé: Nat Commun
Pays: England
ID NLM: 101528555
Informations de publication
Date de publication:
07 08 2020
07 08 2020
Historique:
received:
19
08
2019
accepted:
14
07
2020
entrez:
10
8
2020
pubmed:
10
8
2020
medline:
22
9
2020
Statut:
epublish
Résumé
R-loops have both positive and negative impacts on chromosome functions. To identify toxic R-loops in the human genome, here, we map RNA:DNA hybrids, replication stress markers and DNA double-strand breaks (DSBs) in cells depleted for Topoisomerase I (Top1), an enzyme that relaxes DNA supercoiling and prevents R-loop formation. RNA:DNA hybrids are found at both promoters (TSS) and terminators (TTS) of highly expressed genes. In contrast, the phosphorylation of RPA by ATR is only detected at TTS, which are preferentially replicated in a head-on orientation relative to the direction of transcription. In Top1-depleted cells, DSBs also accumulate at TTS, leading to persistent checkpoint activation, spreading of γ-H2AX on chromatin and global replication fork slowdown. These data indicate that fork pausing at the TTS of highly expressed genes containing R-loops prevents head-on conflicts between replication and transcription and maintains genome integrity in a Top1-dependent manner.
Identifiants
pubmed: 32769985
doi: 10.1038/s41467-020-17858-2
pii: 10.1038/s41467-020-17858-2
pmc: PMC7414224
doi:
Substances chimiques
RNA, Small Interfering
0
ATR protein, human
EC 2.7.11.1
Ataxia Telangiectasia Mutated Proteins
EC 2.7.11.1
DNA Topoisomerases, Type I
EC 5.99.1.2
TOP1 protein, human
EC 5.99.1.2
Types de publication
Journal Article
Research Support, N.I.H., Extramural
Research Support, Non-U.S. Gov't
Langues
eng
Sous-ensembles de citation
IM
Pagination
3940Subventions
Organisme : NIGMS NIH HHS
ID : R01 GM112131
Pays : United States
Organisme : NIGMS NIH HHS
ID : R01 GM120607
Pays : United States
Références
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
Macheret, M. & Halazonetis, T. D. DNA replication stress as a hallmark of cancer. Annu. Rev. Pathol. 10, 425–448 (2015).
pubmed: 25621662
doi: 10.1146/annurev-pathol-012414-040424
Fragkos, M., Ganier, O., Coulombe, P. & Mechali, M. DNA replication origin activation in space and time. Nat. Rev. Mol. Cell Biol. 16, 360–374 (2015).
pubmed: 25999062
doi: 10.1038/nrm4002
Zou, L. & Elledge, S. J. Sensing DNA damage through ATRIP recognition of RPA-ssDNA complexes. Science 300, 1542–1548 (2003).
pubmed: 12791985
doi: 10.1126/science.1083430
Pasero, P. & Vindigni, A. Nucleases acting at stalled forks: how to reboot the replication program with a few shortcuts. Annu. Rev. Genet. 51, 477–499 (2017).
pubmed: 29178820
doi: 10.1146/annurev-genet-120116-024745
Saldivar, J. C. et al. An intrinsic S/G2 checkpoint enforced by ATR. Science 361, 806–810 (2018).
pubmed: 30139873
pmcid: 6365305
doi: 10.1126/science.aap9346
Gaillard, H. & Aguilera, A. Transcription as a threat to genome integrity. Annu. Rev. Biochem. 85, 291–317 (2016).
pubmed: 27023844
doi: 10.1146/annurev-biochem-060815-014908
Merrikh, H. Spatial and temporal control of evolution through replication–transcription conflicts. Trends Microbiol. 25, 515–521 (2017).
pubmed: 28216294
pmcid: 5474121
doi: 10.1016/j.tim.2017.01.008
Prado, F. & Aguilera, A. Impairment of replication fork progression mediates RNA polII transcription-associated recombination. EMBO J. 24, 1267–1276 (2005).
pubmed: 15775982
pmcid: 556405
doi: 10.1038/sj.emboj.7600602
Garcia-Muse, T. & Aguilera, A. Transcription-replication conflicts: how they occur and how they are resolved. Nat. Rev. Mol. Cell Biol. 17, 553–563 (2016).
pubmed: 27435505
doi: 10.1038/nrm.2016.88
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.e719 (2017).
pubmed: 28802045
pmcid: 5570545
doi: 10.1016/j.cell.2017.07.043
Lang, K. S. et al. Replication-transcription conflicts generate R-loops that orchestrate bacterial stress survival and pathogenesis. Cell 170, 787–799.e718 (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. Preprint at https://www.biorxiv.org/content/10.1101/691188v1 (2019).
Petryk, N. et al. Replication landscape of the human genome. Nat. Commun. 7, 10208 (2016).
pubmed: 26751768
pmcid: 4729899
doi: 10.1038/ncomms10208
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
Pourkarimi, E., Bellush, J. M. & Whitehouse, I. Spatiotemporal coupling and decoupling of gene transcription with DNA replication origins during embryogenesis in C. elegans. Elife 5, e21728.stol (2016).
doi: 10.7554/eLife.21728
Drolet, M. et al. Overexpression of RNase H partially complements the growth defect of an Escherichia coli delta topA mutant: R-loop formation is a major problem in the absence of DNA topoisomerase I. Proc. Natl Acad. Sci. USA 92, 3526–3530 (1995).
pubmed: 7536935
doi: 10.1073/pnas.92.8.3526
pmcid: 42200
Chédin, F. Nascent connections: R-loops and chromatin patterning. Trends Genet. 32, 828–838 (2016).
pubmed: 27793359
pmcid: 5123964
doi: 10.1016/j.tig.2016.10.002
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
Stolz, R. et al. Interplay between DNA sequence and negative superhelicity drives R-loop structures. Proc. Natl Acad. Sci. USA 116, 6260–6269 (2019).
pubmed: 30850542
doi: 10.1073/pnas.1819476116
pmcid: 6442632
Chedin, F. & Benham, C. J. Emerging roles for R-loop structures in the management of topological stress. J. Biol. Chem. 295, 4684–4695 (2020).
pubmed: 32107311
pmcid: 7135976
doi: 10.1074/jbc.REV119.006364
Sanz, LionelA. et al. Prevalent, dynamic, and conserved R-loop structures associate with specific epigenomic signatures in mammals. Mol. Cell 63, 167–178 (2016).
pubmed: 27373332
pmcid: 4955522
doi: 10.1016/j.molcel.2016.05.032
Ginno, P. A., Lim, Y. W., Lott, P. L., Korf, I. & Chédin, 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).
pubmed: 23868195
pmcid: 3787257
doi: 10.1101/gr.158436.113
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).
pubmed: 21700224
pmcid: 3145960
doi: 10.1016/j.molcel.2011.04.026
Arab, K. et al. GADD45A binds R-loops and recruits TET1 to CpG island promoters. Nat. Genet. 51, 217–223 (2019).
Yu, K., Chedin, F., Hsieh, C.-L., Wilson, T. E. & Lieber, M. R. R-loops at immunoglobulin class switch regions in the chromosomes of stimulated B cells. Nat. Immunol. 4, 442–451 (2003).
pubmed: 12679812
doi: 10.1038/ni919
Graf, M. et al. Telomere length determines TERRA and R-loop regulation through the cell cycle. Cell 170, 72–85.e14 (2017).
pubmed: 28666126
doi: 10.1016/j.cell.2017.06.006
Costantino, L. et al. Break-induced replication repair of damaged forks induces genomic duplications in human cells. Science 343, 88–91 (2014).
pubmed: 24310611
doi: 10.1126/science.1243211
Costantino, L. & Koshland, D. The Yin and Yang of R-loop biology. Curr. Opin. Cell Biol. 34, 39–45 (2015).
pubmed: 25938907
pmcid: 4522345
doi: 10.1016/j.ceb.2015.04.008
Gomez-Gonzalez, B. et al. Genome-wide function of THO/TREX in active genes prevents R-loop-dependent replication obstacles. EMBO J. 30, 3106–3119 (2011).
pubmed: 21701562
pmcid: 3160181
doi: 10.1038/emboj.2011.206
El Hage, A., French, S. L., Beyer, A. L. & Tollervey, D. Loss of Topoisomerase I leads to R-loop-mediated transcriptional blocks during ribosomal RNA synthesis. Genes Dev. 24, 1546–1558 (2010).
pubmed: 20634320
pmcid: 2904944
doi: 10.1101/gad.573310
Chang, E. & Stirling, P. Replication fork protection factors controlling R-loop bypass and suppression. Genes 8, 33 (2017).
pmcid: 5295027
doi: 10.3390/genes8010033
Alzu, A. et al. Senataxin associates with replication forks to protect fork integrity across RNA-polymerase-II-transcribed genes. Cell 151, 835–846 (2012).
pubmed: 23141540
pmcid: 3494831
doi: 10.1016/j.cell.2012.09.041
Wellinger, R. E., Prado, F. & Aguilera, A. Replication fork progression is impaired by transcription in hyperrecombinant yeast cells lacking a functional THO complex. Mol. Cell. Biol. 26, 3327–3334 (2006).
pubmed: 16581804
pmcid: 1446968
doi: 10.1128/MCB.26.8.3327-3334.2006
Tuduri, S. et al. Topoisomerase I suppresses genomic instability by preventing interference between replication and transcription. Nat. Cell Biol. 11, 1315–1324 (2009).
pubmed: 19838172
pmcid: 2912930
doi: 10.1038/ncb1984
Gan, W. et al. R-loop-mediated genomic instability is caused by impairment of replication fork progression. Genes Dev. 25, 2041–2056 (2011).
pubmed: 21979917
pmcid: 3197203
doi: 10.1101/gad.17010011
Manzo, S. G. et al. DNA topoisomerase I differentially modulates R-loops across the human genome. Genome Biol. 19, 100 (2018).
pubmed: 30060749
pmcid: 6066927
doi: 10.1186/s13059-018-1478-1
Bianco, J. N. et al. Analysis of DNA replication profiles in budding yeast and mammalian cells using DNA combing. Methods 57, 149–157 (2012).
pubmed: 22579803
doi: 10.1016/j.ymeth.2012.04.007
Boguslawski, S. J. et al. Characterization of monoclonal antibody to DNA.RNA and its application to immunodetection of hybrids. J. Immunol. Methods 89, 123–130 (1986).
pubmed: 2422282
doi: 10.1016/0022-1759(86)90040-2
Sartori, A. A. et al. Human CtIP promotes DNA end resection. Nature 450, 509–514 (2007).
pubmed: 17965729
pmcid: 2409435
doi: 10.1038/nature06337
Iacovoni, J. S. et al. High-resolution profiling of [gamma]H2AX around DNA double strand breaks in the mammalian genome. EMBO J. 29, 1446–1457 (2010).
pubmed: 20360682
pmcid: 2868577
doi: 10.1038/emboj.2010.38
Hansen, R. S. et al. Sequencing newly replicated DNA reveals widespread plasticity in human replication timing. Proc. Natl Acad. Sci. USA 107, 139–144 (2010).
pubmed: 19966280
doi: 10.1073/pnas.0912402107
Barlow, J. H. et al. Identification of early replicating fragile sites that contribute to genome instability. Cell 152, 620–632 (2013).
pubmed: 23352430
pmcid: 3629730
doi: 10.1016/j.cell.2013.01.006
Li, X. & Manley, J. L. Inactivation of the SR protein splicing factor ASF/SF2 results in genomic instability. Cell 122, 365–378 (2005).
pubmed: 16096057
doi: 10.1016/j.cell.2005.06.008
Crosetto, N. et al. Nucleotide-resolution DNA double-strand break mapping by next-generation sequencing. Nat. Methods 10, 361–365 (2013).
pubmed: 23503052
pmcid: 3651036
doi: 10.1038/nmeth.2408
Biernacka, A. et al. i-BLESS is an ultra-sensitive method for detection of DNA double-strand breaks. Commun. Biol. 1, 181 (2018).
pubmed: 30393778
pmcid: 6208412
doi: 10.1038/s42003-018-0165-9
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).
pubmed: 28363678
doi: 10.1016/j.jmb.2017.03.024
Chappidi, N. et al. Fork cleavage-religation cycle and active transcription mediate replication restart after fork stalling at co-transcriptional R-loops. Mol. Cell 77, 528–541.e528 (2020).
pubmed: 31759821
doi: 10.1016/j.molcel.2019.10.026
Canela, A. et al. Topoisomerase II-induced chromosome breakage and translocation is determined by chromosome architecture and transcriptional activity. Mol. Cell 75, 252–266 e258 (2019).
pubmed: 31202577
doi: 10.1016/j.molcel.2019.04.030
pmcid: 8170508
Gothe, H. J. et al. Spatial chromosome folding and active transcription drive DNA fragility and formation of oncogenic MLL translocations. Mol. Cell 75, 267–283 e212 (2019).
pubmed: 31202576
doi: 10.1016/j.molcel.2019.05.015
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
Matos, D. A. et al. ATR protects the genome against R loops through a MUS81-triggered feedback loop. Mol. Cell 77, 514–527.e514 (2020).
pubmed: 31708417
doi: 10.1016/j.molcel.2019.10.010
Seiler, J. A., Conti, C., Syed, A., Aladjem, M. I. & Pommier, Y. The intra-S-phase checkpoint affects both DNA replication initiation and elongation: single-cell and -DNA fiber analyses. Mol. Cell. Biol. 27, 5806–5818 (2007).
pubmed: 17515603
pmcid: 1952133
doi: 10.1128/MCB.02278-06
Mutreja, K. et al. ATR-mediated global fork slowing and reversal assist fork traverse and prevent chromosomal breakage at DNA interstrand cross-links. Cell Rep. 24, 2629–2642.e2625 (2018).
pubmed: 30184498
pmcid: 6137818
doi: 10.1016/j.celrep.2018.08.019
Bacal, J. et al. Mrc1 and Rad9 cooperate to regulate initiation and elongation of DNA replication in response to DNA damage. EMBO J. 37, e99319 (2018).
pubmed: 30158111
pmcid: 6213276
doi: 10.15252/embj.201899319
Osmundson, J. S., Kumar, J., Yeung, R. & Smith, D. J. Pif1-family helicases cooperatively suppress widespread replication-fork arrest at tRNA genes. Nat. Struct. Mol. Biol. 24, 162–170 (2017).
pubmed: 27991904
doi: 10.1038/nsmb.3342
Cohen, S. et al. Senataxin resolves RNA:DNA hybrids forming at DNA double-strand breaks to prevent translocations. Nat. Commun. 9, 533 (2018).
pubmed: 29416069
pmcid: 5803260
doi: 10.1038/s41467-018-02894-w
Tran, P. L. T. et al. PIF1 family DNA helicases suppress R-loop mediated genome instability at tRNA genes. Nat. Commun. 8, 15025 (2017).
pubmed: 28429714
pmcid: 5413955
doi: 10.1038/ncomms15025
Nguyen, V. C. et al. Replication stress checkpoint signaling controls tRNA gene transcription. Nat. Struct. Mol. Biol. 17, 976–981 (2010).
pubmed: 20639887
doi: 10.1038/nsmb.1857
Poli, J. et al. Mec1, INO80, and the PAF1 complex cooperate to limit transcription replication conflicts through RNAPII removal during replication stress. Genes Dev. 30, 337–354 (2016).
pubmed: 26798134
pmcid: 4743062
doi: 10.1101/gad.273813.115
Lafon, A. et al. INO80 chromatin remodeler facilitates release of RNA polymerase II from chromatin for ubiquitin-mediated proteasomal degradation. Mol. Cell 60, 784–796 (2015).
pubmed: 26656161
pmcid: 4760348
doi: 10.1016/j.molcel.2015.10.028
Tantale, K. et al. A single-molecule view of transcription reveals convoys of RNA polymerases and multi-scale bursting. Nat. Commun. 7, 12248 (2016).
pubmed: 27461529
pmcid: 4974459
doi: 10.1038/ncomms12248
Rodriguez, J. et al. Intrinsic dynamics of a human gene reveal the basis of expression heterogeneity. Cell 176, 213–226.e218 (2019).
pubmed: 30554876
doi: 10.1016/j.cell.2018.11.026
Teloni, F. et al. Efficient pre-mRNA cleavage prevents replication-stress-associated genome instability. Mol. Cell 73, 670–683 (2018).
doi: 10.1016/j.molcel.2018.11.036
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.e483 (2018).
pubmed: 30078723
pmcid: 6264797
doi: 10.1016/j.molcel.2018.06.037
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
Lin, Y. L. et al. Feline immunodeficiency virus vectors for efficient transduction of primary human synoviocytes: application to an original model of rheumatoid arthritis. Hum. Gene Ther. 15, 588–596 (2004).
pubmed: 15212717
doi: 10.1089/104303404323142033
Jackson, D. A. & Pombo, A. Replicon clusters are stable units of chromosome structure: evidence that nuclear organization contributes to the efficient activation and propagation of S phase in human cells. J. Cell Biol. 140, 1285–1295 (1998).
pubmed: 9508763
pmcid: 2132671
doi: 10.1083/jcb.140.6.1285
Caron, P. et al. Cohesin protects genes against gammaH2AX Induced by DNA double-strand breaks. PLoS Genet. 8, e1002460 (2012).
pubmed: 22275873
pmcid: 3261922
doi: 10.1371/journal.pgen.1002460
Ginno, P. A., Lott, P. L., Christensen, H. C., Korf, I. & Chédin, F. R-loop formation is a distinctive characteristic of unmethylated human CpG island promoters. Mol. Cell 45, 814–825 (2012).
pubmed: 22387027
pmcid: 3319272
doi: 10.1016/j.molcel.2012.01.017
Mitra, A., Skrzypczak, M., Ginalski, K. & Rowicka, M. Strategies for achieving high sequencing accuracy for low diversity samples and avoiding sample bleeding using illumina platform. PLoS ONE 10, e0120520 (2015).
pubmed: 25860802
pmcid: 4393298
doi: 10.1371/journal.pone.0120520
Langmead, B. & Salzberg, S. L. Fast gapped-read alignment with Bowtie 2. Nat. Methods 9, 357–359 (2012).
pubmed: 22388286
pmcid: 3322381
doi: 10.1038/nmeth.1923
Dobin, A. et al. STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).
pubmed: 23104886
doi: 10.1093/bioinformatics/bts635
Li, H. et al. The sequence alignment/map format and SAMtools. Bioinformatics 25, 2078–2079 (2009).
pubmed: 19505943
pmcid: 2723002
doi: 10.1093/bioinformatics/btp352
Zhang, Y. et al. Model-based analysis of ChIP-Seq (MACS). Genome Biol. 9, R137 (2008).
pubmed: 18798982
pmcid: 2592715
doi: 10.1186/gb-2008-9-9-r137
Landt, S. G. et al. ChIP-seq guidelines and practices of the ENCODE and modENCODE consortia. Genome Res 22, 1813–1831 (2012).
pubmed: 22955991
pmcid: 3431496
doi: 10.1101/gr.136184.111
Quinlan, A. R. BEDTools: The Swiss-army tool for genome feature analysis. Curr. Protoc. Bioinformatics 47, 11.12.1–11.12.34 (2014).
doi: 10.1002/0471250953.bi1112s47
Ramirez, F., Dundar, F., Diehl, S., Gruning, B. A. & Manke, T. deepTools: a flexible platform for exploring deep-sequencing data. Nucleic Acids Res. 42, W187–W191 (2014).
pubmed: 24799436
pmcid: 4086134
doi: 10.1093/nar/gku365
Gentleman, R. C. et al. Bioconductor: open software development for computational biology and bioinformatics. Genome Biol. 5, R80 (2004).
pubmed: 15461798
pmcid: 545600
doi: 10.1186/gb-2004-5-10-r80