The core spliceosome as target and effector of non-canonical ATM signalling


In response to DNA damage, tissue homoeostasis is ensured by protein networks promoting DNA repair, cell cycle arrest or apoptosis. DNA damage response signalling pathways coordinate these processes, partly by propagating gene-expression-modulating signals. DNA damage influences not only the abundance of messenger RNAs, but also their coding information through alternative splicing. Here we show that transcription-blocking DNA lesions promote chromatin displacement of late-stage spliceosomes and initiate a positive feedback loop centred on the signalling kinase ATM. We propose that initial spliceosome displacement and subsequent R-loop formation is triggered by pausing of RNA polymerase at DNA lesions. In turn, R-loops activate ATM, which signals to impede spliceosome organization further and augment ultraviolet-irradiation-triggered alternative splicing at the genome-wide level. Our findings define R-loop-dependent ATM activation by transcription-blocking lesions as an important event in the DNA damage response of non-replicating cells, and highlight a key role for spliceosome displacement in this process.

Access options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Figure 1: DNA-damage-triggered chromatin displacement of activated spliceosomes.
Figure 2: Mobilization and displacement of mature spliceosomes from sites of UVC-induced DNA damage.
Figure 3: Chromatin displacement of mature spliceosomes is caused by RNAPII-blocking lesions and is NER-independent.
Figure 4: ATM modulates spliceosome mobilization and influences splicing decisions upon DNA damage.
Figure 5: Reciprocal regulation between spliceosome mobilization and R-loop-dependent ATM signalling.

Accession codes

Primary accessions

Sequence Read Archive

Data deposits

RNA-seq data have been deposited in the Sequence Read Archive with accession number SRP053034.


  1. 1

    Sirbu, B. M. & Cortez, D. DNA damage response: three levels of DNA repair regulation. Cold Spring Harb. Perspect. Biol. 5, a012724 (2013)

  2. 2

    Ciccia, A. & Elledge, S. J. The DNA damage response: making it safe to play with knives. Mol. Cell 40, 179–204 (2010)

  3. 3

    Hoeijmakers, J. H. DNA damage, aging, and cancer. N. Engl. J. Med. 361, 1475–1485 (2009)

  4. 4

    Shiloh, Y. & Ziv, Y. The ATM protein kinase: regulating the cellular response to genotoxic stress, and more. Nature Rev. Mol. Cell Biol. 14, 197–210 (2013)

  5. 5

    Kaidi, A. & Jackson, S. P. KAT5 tyrosine phosphorylation couples chromatin sensing to ATM signalling. Nature 498, 70–74 (2013)

  6. 6

    Lenzken, S. C., Loffreda, A. & Barabino, S. M. RNA splicing: a new player in the DNA damage response. Int. J. Cell Biol. 2013, 153634 (2013)

  7. 7

    Hoskins, A. A. & Moore, M. J. The spliceosome: a flexible, reversible macromolecular machine. Trends Biochem. Sci. 37, 179–188 (2012)

  8. 8

    Valadkhan, S. & Jaladat, Y. The spliceosomal proteome: at the heart of the largest cellular ribonucleoprotein machine. Proteomics 10, 4128–4141 (2010)

  9. 9

    Kornblihtt, A. R. et al. Alternative splicing: a pivotal step between eukaryotic transcription and translation. Nature Rev. Mol. Cell Biol. 14, 153–165 (2013)

  10. 10

    Schor, I. E., Gomez Acuna, L. I. & Kornblihtt, A. R. Coupling between transcription and alternative splicing. Cancer Treat. Res. 158, 1–24 (2013)

  11. 11

    Zhou, H. L., Luo, G., Wise, J. A. & Lou, H. Regulation of alternative splicing by local histone modifications: potential roles for RNA-guided mechanisms. Nucleic Acids Res. 42, 701–713 (2014)

  12. 12

    Alexander, R. & Beggs, J. D. Cross-talk in transcription, splicing and chromatin: who makes the first call? Biochem. Soc. Trans. 38, 1251–1256 (2010)

  13. 13

    Dutertre, M., Sanchez, G., Barbier, J., Corcos, L. & Auboeuf, D. The emerging role of pre-messenger RNA splicing in stress responses: sending alternative messages and silent messengers. RNA Biol. 8, 740–747 (2011)

  14. 14

    Paronetto, M. P., Minana, B. & Valcarcel, J. The Ewing sarcoma protein regulates DNA damage-induced alternative splicing. Mol. Cell 43, 353–368 (2011)

  15. 15

    Dutertre, M. et al. Cotranscriptional exon skipping in the genotoxic stress response. Nature Struct. Mol. Biol. 17, 1358–1366 (2010)

  16. 16

    Muñoz, M. J. et al. DNA damage regulates alternative splicing through inhibition of RNA polymerase II elongation. Cell 137, 708–720 (2009)

  17. 17

    Cox, J. & Mann, M. MaxQuant enables high peptide identification rates, individualized p.p.b.-range mass accuracies and proteome-wide protein quantification. Nature Biotechnol. 26, 1367–1372 (2008)

  18. 18

    Rino, J. et al. A stochastic view of spliceosome assembly and recycling in the nucleus. PLOS Comput. Biol. 3, 2019–2031 (2007)

  19. 19

    Dinant, C. et al. Activation of multiple DNA repair pathways by sub-nuclear damage induction methods. J. Cell Sci. 120, 2731–2740 (2007)

  20. 20

    Lagerwerf, S., Vrouwe, M. G., Overmeer, R. M., Fousteri, M. I. & Mullenders, L. H. DNA damage response and transcription. DNA Repair (Amst.) 10, 743–750 (2011)

  21. 21

    Vermeulen, W. & Fousteri, M. Mammalian transcription-coupled excision repair. Cold Spring Harb. Perspect. Biol. 5, a012625 (2013)

  22. 22

    Matsuoka, S. et al. ATM and ATR substrate analysis reveals extensive protein networks responsive to DNA damage. Science 316, 1160–1166 (2007)

  23. 23

    Blasius, M. et al. A phospho-proteomic screen identifies substrates of the checkpoint kinase Chk1. Genome Biol. 12, R78 (2011)

  24. 24

    Ahn, E. Y. et al. SON controls cell-cycle progression by coordinated regulation of RNA splicing. Mol. Cell 42, 185–198 (2011)

  25. 25

    Lee, J. H. & Paull, T. T. ATM activation by DNA double-strand breaks through the Mre11-Rad50-Nbs1 complex. Science 308, 551–554 (2005)

  26. 26

    Bakkenist, C. J. & Kastan, M. B. DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation. Nature 421, 499–506 (2003)

  27. 27

    Matsuoka, S., Huang, M. & Elledge, S. J. Linkage of ATM to cell cycle regulation by the Chk2 protein kinase. Science 282, 1893–1897 (1998)

  28. 28

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

  29. 29

    Hanasoge, S. & Ljungman, M. H2AX phosphorylation after UV irradiation is triggered by DNA repair intermediates and is mediated by the ATR kinase. Carcinogenesis 28, 2298–2304 (2007)

  30. 30

    Aguilera, A. & Garcia-Muse, T. R loops: from transcription byproducts to threats to genome stability. Mol. Cell 46, 115–124 (2012)

  31. 31

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

  32. 32

    Kotake, Y. et al. Splicing factor SF3b as a target of the antitumor natural product pladienolide. Nature Chem. Biol. 3, 570–575 (2007)

  33. 33

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

  34. 34

    Wu, H., Lima, W. F. & Crooke, S. T. Investigating the structure of human RNase H1 by site-directed mutagenesis. J. Biol. Chem. 276, 23547–23553 (2001)

  35. 35

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

  36. 36

    Reijns, M. A. M. et al. Enzymatic Removal of Ribonucleotides from DNA Is Essential for Mammalian Genome Integrity and Development. Cell 149, 1008–1022 (2012)

  37. 37

    McManus, C. J. & Graveley, B. R. RNA structure and the mechanisms of alternative splicing. Curr. Opin. Genet. Dev. 21, 373–379 (2011)

  38. 38

    Biton, S., Barzilai, A. & Shiloh, Y. The neurological phenotype of ataxia-telangiectasia: solving a persistent puzzle. DNA Repair (Amst.) 7, 1028–1038 (2008)

  39. 39

    Schwertman, P. et al. UV-sensitive syndrome protein UVSSA recruits USP7 to regulate transcription-coupled repair. Nature Genet. 44, 598–602 (2012)

  40. 40

    Nakazawa, Y., Yamashita, S., Lehmann, A. R. & Ogi, T. A semi-automated non-radioactive system for measuring recovery of RNA synthesis and unscheduled DNA synthesis using ethynyluracil derivatives. DNA Repair (Amst.) 9, 506–516 (2010)

  41. 41

    Houtsmuller, A. B. & Vermeulen, W. Macromolecular dynamics in living cell nuclei revealed by fluorescence redistribution after photobleaching. Histochem. Cell Biol. 115, 13–21 (2001)

  42. 42

    Wuarin, J. & Schibler, U. Physical isolation of nascent RNA chains transcribed by RNA polymerase II: evidence for cotranscriptional splicing. Mol. Cell. Biol. 14, 7219–7225 (1994)

  43. 43

    Galiveti, C. R., Rozhdestvensky, T. S., Brosius, J., Lehrach, H. & Konthur, Z. Application of housekeeping npcRNAs for quantitative expression analysis of human transcriptome by real-time PCR. RNA 16, 450–461 (2010)

  44. 44

    Trapnell, C., Pachter, L. & Salzberg, S. L. TopHat: discovering splice junctions with RNA-Seq. Bioinformatics 25, 1105–1111 (2009)

  45. 45

    Shen, S. et al. MATS: a Bayesian framework for flexible detection of differential alternative splicing from RNA-Seq data. Nucleic Acids Res. 40, e61 (2012)

Download references


We acknowledge the Optical Imaging Center of ErasmusMC for technical support; M. Reijns and A. Jackson for the S9.6 antibody; L. Marufu for technical assistance; and N. G. J. Jaspers for intellectual input. This work was funded by the Netherlands Organization for Scientific Research (NWO) ZonMW TOP Grants 912.08.031 and 912.12.132, Horizon Zenith 935.11.042, ALW 854.11.002 and 823.02.013, the Association for International Cancer Research 10-594, European Research Council Advanced Investigator Grants 233424 and 340988, and an ErasmusMC fellowship.

Author information

M.T. designed the study, performed the majority of experiments, analysed the data and authored the manuscript with contributions from W.V. and J.A.M. J.A.A.D. performed the liquid-chromatography tandem mass spectrometry analysis, L.S. assisted in fractionation/immunoblotting experiments, J.A.M. performed S9.6 antibody immunofluorescence and assisted in UVC micro-irradiation experiments, D.W. and R.H.M performed RT–PCR splicing assays, P.K., F.G.G. and W.v.IJ. performed RNA-seq experiments, L.H.M and M.G.V. generated RNaseH1 constructs and cell lines. L.H.M. and J.H.J.H. provided advice. All authors reviewed and commended on the manuscript.

Correspondence to Maria Tresini or Wim Vermeulen or Jurgen A. Marteijn.

Ethics declarations

Competing interests

The authors declare no competing financial interests.

Extended data figures and tables

Extended Data Figure 1 Chromatin association of splicing factors.

a, Schematic overview of the proteomic experiments for the identification of proteins that display UV-dependant chromatin association. b, Schematic outline of cell fractionation. c, Validation of the chromatin-isolation protocol for NER proteins that are recruited to chromatin in response to DNA damage. Mock-treated or UV-irradiated quiescent HDFs (20 J m−2, 1 h post-irradiation) were fractionated as outlined in b. Equal protein amounts from each fraction were analysed by immunoblotting using antibodies against the indicated NER proteins. Abundance of H2A is shown as a control for chromatin-isolation efficiency. d, UV-triggered changes in chromatin association of core splicing factors, identified by quantitative SILAC proteomics. Proteomic experiments were performed with HDFs as outlined in a. The table lists representative examples of splicing factors that participate in distinct snRNP complexes and their chromatin association in response to UV irradiation (20 J m−2, 1 h). U2 and U5 snRNP splicing factors show significantly reduced chromatin association (P ≤ 0.05, significance B17) and are indicated with a cross. Significance B was calculated by estimating the variance of the distribution of all protein ratios, taking into account the dependency of the distribution on the summed protein intensity17. ND, not detected. e, Abundance of splicing factors in total cell lysates. Total lysates were prepared from U2OS cells that were mock-treated or UV-irradiated (20 J m−2, 1 h post-irradiation) and splicing factor abundance was assayed by immunobloting. Abundance of H2A is shown as a loading control. Right, immunoblots; left, quantification of splicing factor signal intensities normalized to H2A (n = 3, mean ± s.d., one-way ANOVA/Bonferroni). f, UV-dependent interaction of splicing proteins with elongating RNAPII. Quiescent HDFs were prepared as outlined in b except that, instead of MNase digestion, chromatin was mechanically sheared. Elongating RNAPII was immunoprecipitated with an antibody that recognizes specifically the Ser2-phosphorylated RNAPII C-terminal domain (CTD) and its interaction with the U2 snRNP splicing factors SF3a1 and SF3b2 was assayed by immunoblotting.

Extended Data Figure 2 Validation of HDFs stably expressing GFP-tagged splicing factors.

a, Whole-cell lysates from HDFs stably expressing eGFP tagged PRP8, SF3a1, SNRNP40 or free eGFP, were analysed by immunoblotting using antibodies against GFP (left) or against PRP8, SF3a1 and SNRNP40 (right). Ectopically produced proteins were expressed at near or below endogenous levels. b, Fluorescent microscopy images of GFP-tagged splicing factors showing the expected punctuated nuclear distribution. Images were obtained at 40× magnification. c, Localization of SNRNP40–GFP in nuclear speckles which were visualized by immunofluorescence detection of the speckle marker SRSF2/SC35. Images were obtained at 63× magnification. d, Interaction of SNRNP40–GFP with endogenous splicing factors and elongating RNAPII. Quiescent SNRNP40–GFP-expressing HDFs were mock-treated or irradiated with 20 J m−2 UVC. After a 3-h recovery period, cells were lysed under native conditions and chromatin was sheared by mechanical force. SNRNP40–GFP was immunoprecipitated from whole-cell lysates using GFP-Trap agarose beads, and its association with endogenous splicing factors and the large subunit of RNAPII was assayed by immunoblotting. Non-transfected cells are shown as a negative control. SNRNP40–GFP interacts with U2 and U5 snRNP components, suggesting that the GFP tag does not interfere with complex formation. Interaction of SNRNP40 with its U5 snRNP partner PRP8 is partially maintained even after MNase digestion, consistent with its presence in U4/U6.U5 tri-snRNP complexes. Participation of SNRNP40–GFP in co-transcriptional splicing complexes is confirmed by co-immunoprecipitation of the active (hyperphosphorylated RNAPIIo) large subunit of RNAPII.

Extended Data Figure 3 Displacement of mature spliceosomes from subnuclear sites of UV-inflicted DNA damage.

a, U2OS cells stably expressing GFP-tagged splicing factors were UV-irradiated (60 J m−2) through isopore membranes resulting in DNA lesion formation in small subnuclear areas. DNA damage sites (circled) were visualized by immunofluorescence using an antibody against the NER recognition factor XPC. Scale bar, 5 μm. b, SF3a1–GFP and PRP8–GFP depletion from UVC laser microbeam irradiation sites. Quantification of 20 cells from two independent experiments. eGFP localization at sites of DNA damage is used to demonstrate that depletion of eGFP-tagged splicing factors is not caused by photobleaching. c, UVC laser microbeam irradiation results in preferential displacement of U2- and U5-associated splicing factors from DNA damage sites. Quiescent HDFs were irradiated in a 1-µm-diameter nuclear area via a UVC laser. GFP signal intensity, reflecting the abundance of GFP-tagged U1, U2, U4 and U5 snRNP components at UVC DNA damage sites, was quantified in the irradiated and in a non-irradiated nuclear area (undamaged control). Plotted is the fluorescence signal intensity expressed as a percentage of that before irradiation, at the 1-min time point. Cells expressing free eGFP were used as negative control. Representative from three independent experiment (n = 12, mean ± s.e.m., paired t-test). d, Depletion of splicing factors from UVC laser irradiation sites depends on active transcription. Transcription initiation was inhibited in quiescent HDFs by prolonged α-amanitin treatment (10 µM, ≥24 h) before subnuclear UVC laser irradiation. Plotted is the SNRNP40–GFP abundance in irradiated and non-irradiated nuclear areas at 1-min post-irradiation. Representative from three independent experiments (n = 12, mean ± s.e.m., one-way ANOVA/Bonferroni).

Extended Data Figure 4 SNRNP40 reorganization and speckle enlargement in response to UV irradiation.

Representative microscopic images showing SNRNP40–GFP distribution in quiescent HDFs before, and 1 h post UVC irradiation with 20 J m−2. a, Live cells. b, Fixed cells. Images were obtained at 63× magnification.

Extended Data Figure 5 Transcription stalling mobilizes spliceosomes independent from NER complex assembly and proteasome activity.

a, RNA synthesis is inhibited preferentially by genotoxins that inflict bulky DNA lesions. Influence of genotoxins on RNA synthesis of quiescent HDFs was measured by 5EU pulse labelling combined with click chemistry. Top, representative images; bottom, quantification of fluorescence intensity (n = 150, mean ± s.e.m., one-way ANOVA/Bonferroni). Images were obtained at 40× magnification. b, Mobilization of U2 and U5 snRNPs by genotoxins inflicting transcription-blocking DNA lesions. Mobilization of GFP-tagged SF3a1 (left) and PRP8 (right) assayed by FRAP in quiescent HDFs exposed to different types of genotoxins (n = 30, mean ± s.e.m., one-way ANOVA/Dunnett’s). IR, ionizing radiation. c, Chromatin displacement of mature spliceosomes is not TC–NER-dependent. Left, chromatin abundance of U2 and U5 snRNP splicing factors was assayed by immunoblotting in XPA-deficient (left), XPA–GFP-corrected (middle) and CSB-deficient (right) HDFs. Cells were UV-irradiated (20 J m−2) and chromatin was isolated at the indicated times. Top, immunoblots; bottom, quantification of splicing factor signal intensities normalized to H2A (n = 3, mean ± s.d., one-way ANOVA/Bonferroni). d, Proteasome activity is not required for UV-damage-induced spliceosome mobilization. Mobilization of SNRNP40–GFP assayed by FRAP in quiescent HDFs exposed to UV radiation in the presence or absence of the proteasome inhibitor MG132 (50 µM) (n = 30, mean ± s.e.m., t-test). e, SNRNP40–GFP mobilization by transcription inhibition. FRAP of SNRNP40–GFP in quiescent HDFS after inhibition of transcription initiation (10 µg ml−1 α-amanitin, 24 h) or elongation (1 µg ml−1 actinomycin D or 50 µM DRB, 1 h) (n = 30, mean ± s.e.m., one-way ANOVA/Dunnett’s).

Extended Data Figure 6 ATM-dependency of UV induced spliceosome mobilization, alternative splicing and gene expression changes.

a, UV irradiation and DRB-dependent mobilization of SNRNP40. Quiescent HDFs expressing SNRNP40–GFP were UV-irradiated or DRB treated with doses that inhibit transcription to similar levels. Splicing factor mobility was assayed by FRAP. b, Additive effect of combined UV and DRB treatments. FRAP of SNRNP40–GFP in quiescent HDFs treated with DRB, UV, or a combination of both, each at a dose that inhibits RNA synthesis by 50%. c, Impaired UV-dependent SF3a1 mobilization in cells lacking ATM activity. SF3a1–GFP mobilization was measured by FRAP in quiescent HDFs derived from an ataxia telangiectasia (AT) patient or a healthy donor. d, ATM-dependent spliceosome mobilization. Quiescent HDFs were treated with 10 µM ATM (KU55933), ATR (VE821) or DNA-PK (NU7441) inhibitors before irradiation. GFP-tagged SF3a1 or PRP8 mobility was assayed by FRAP. ATM, but not ATR or DNA-PK inhibition partially prevented the UV-induced splicing factor mobilization. a–d, n = 25, mean ± s.e.m., one-way ANOVA/Bonferroni. e, Reduced UV-induced intron retention in response to ATM silencing. Intron inclusion in retina pigment epithelium (RPE) cells transfected either with control or ATM-silencing siRNAs and subsequently mock-treated or UV-irradiated (20 J m−2, 6 h) was assayed by RT–PCR. f, ATM-dependent changes in intron retention. Intron inclusion was assayed by RT–PCR in untreated, UV-irradiated and DRB-treated quiescent cells in the presence or absence of 10 µM ATM inhibitor. g, Heat map of UV-triggered and ATM-dependent transcriptome changes. Quiescent cells were mock-treated or UV-irradiated in the presence or absence of the ATM inhibitor. Transcriptome profiles were generated by RNA-seq. Differentially expressed genes between untreated and UV-irradiated cells (P < 0.05) and UV-irradiated cells in the presence or absence of the ATM inhibitor (P < 0.05), were clustered in a heat map using Pearson correlation. n = 1,676 differentially expressed transcripts. The observed correlation indicates that UV-inducible transcriptome changes can be, in part, prevented by ATM inhibition. h, Lack of influence of ATM inhibition on DRB-dependent splicing factor mobility. Splicing factor mobility was measured by FRAP in untreated or DRB-treated HDFs in the presence or absence of 10 µM ATM inhibitor (n = 30, mean ± s.e.m., one-way ANOVA/Bonferroni).

Extended Data Figure 7 Canonical and non-canonical ATM activation.

a, ATM autophosphorylation (Ser1981) was assayed in quiescent HDFs 1 h after the indicated treatments. In non-replicating cells UV and trichostatin A (TSA) activate ATM via non-canonical pathways. Transcription inhibition by DRB has no influence on ATM activity. b, The quiescent status of serum-deprived HDFs was verified by immunodetection of the cell cycle marker Ki67, which is not expressed by quiescent (G0) cells. c, Immunofluorescence detection of active ATM in quiescent HDFs treated with DDR kinase inhibitors. d, Immunoblotting analysis of nuclear extracts derived from quiescent HDFs treated as in c using a phospho-specific ATM (Ser1981) antibody (top) and an antibody recognizing ATM (bottom). e, Differences in autophosphorylated-ATM distribution in quiescent HDFs treated with various ATM activators. Left, multiple cells; right, single-cell magnification illustrating pan-nuclear localization of phosphorylated ATM after UV irradiation and focal accumulation after CPT or ionizing radiation treatments. Magnified cells are indicated by arrows (left panel). f, Differences in amounts of DNA damage-foci formation indicative of DSBs, in response to CPT, UV and ionizing radiation. Quiescent HDFs were pre-treated with the ATR inhibitor (10 µM, 1 h) and subsequently exposed to the indicated genotoxins. DSB foci were visualized by immunofluorescence using antibodies against γH2AX and p53BP1. Left, multiple cells; right, single-cell magnification; images were obtained at 40× and 63× magnification, respectively. Magnified cells are indicated by arrows in the left panel.

Extended Data Figure 8 ATM activation by interference with spliceosome assembly or RNaseH1/H2A silencing.

a, ATM autophosphorylation was assayed by immunofluorescence in HDFs after silencing of SF3a1, PRP8 or combined silencing of RNaseH1 and RNaseH2A. b, Immunoblotting analysis of silenced proteins in total cell lysates. Tubulin is shown as a loading control. c, Splicing factor mobilization by the spliceosome inhibitor pladienolide B (PL) was assayed by FRAP in quiescent HDFs. Consistent with its function in interfering with spliceosome maturation following pre-spliceosome assembly, cell treatment with pladienolide B resulted in extensive mobilization of U5 snRNP factors (PRP8 and SNRNP40), partial mobilization of the U2 snRNP SF3a1, and had no influence on the U1 snRNP factor U1A (n = 30, mean ± s.e.m., one-way ANOVA/Bonferroni). d, e, ATM activation by Pladienolide B. Quiescent HDFs were either treated with 5 µM pladienolide B or exposed to 1 Gy ionizing radiation (IR) and autophosphorylated ATM was detected by immunofluorescence (d) or immunoblotting (e). f, Effect of pladienolide treatment on intron retention. RNA isolated from mock-treated, UV-irradiated or pladienolide B-treated RPE cells. Intron retention, as assayed by RT–PCR on transcripts of the indicated genes, shows that pladienolide B influences splicing to the same extent as UV irradiation. U/S, ratio of relative abundance of unspliced (U) to spliced (S) introns. g, Efficiency of RNaseH1 and H2A silencing at the single-cell level, assayed by immunofluorescence. Images were obtained at 40× magnification.

Extended Data Figure 9 UV-induced R-loop formation enhances spliceosome mobilization.

a, Recruitment of RNaseH1(D145N)–GFP at local DNA damage sites depends on endogenous levels of RNaseH activity. DNA damage was inflicted via a UVC laser in 1-µm-diameter subnuclear areas of cells after silencing of RNaseH2A or overexpression of RNaseH1–mCherry. Recruitment of RNaseH1(D145N)–GFP at the irradiated sites was monitored by live-cell imaging. Plotted is the fluorescence intensity of RNaseH1(D145N)–GFP at 1 min post-irradiation, at the irradiated and in a non-irradiated nuclear area. Representative from three independent experiments (n = 10, mean ± s.e.m., one-way ANOVA/Bonferroni). b, c, R-loop formation at sites of local UVC laser irradiation. Immunofluorescence detection of R-loops using the DNA–RNA hybrid-specific S9.6 antibody. Sites of irradiation are visualized by XPC immunodetection. b, Dashed boxes indicate the magnified areas shown in the right panels. The dashed lines indicate the line-scan track used to quantify fluorescence intensity of S9.6 and anti-XPC (shown in in the graph). c, Specificity of the antibody was confirmed by its increased sensitivity after RNaseH2A silencing and its ability to detect R-loops when suboptimal doses of UVC irradiation were applied. d, RNaseH1 accumulation at local DNA damage sites depends on active transcription but not ATM activity. Transcription initiation was inhibited in quiescent HDFs by α-amanitin (10 µg ml−1, 24 h) before local UVC laser irradiation. Plotted is the fluorescence intensity at 1 min post-irradiation of RNaseH1(D145N)–GFP at the irradiated area and in a non-irradiated nuclear area for untreated, ATM-inhibitor- and α-amanitin-treated cells. Representative from three experiments (n = 10, mean ± s.e.m., one-way ANOVA/Bonferroni). e, RNaseH1 overexpression inhibits the UV-dependent spliceosome mobilization. FRAP of U2OS cells stably expressing GFP-tagged SF3a1 and PRP8 and transiently transfected with RNaseH1–mCherry. f, RNaseH1 and H2A silencing potentiates the UV-dependent spliceosome mobilization. RNaseH1 and H2 were silenced in U2OS cells expressing SF3a1–GFP or PRP8–GFP and splicing factor mobility was assayed by FRAP. g, FRAP of SNRNP40–GFP in quiescent HDFs after RNaseH1/H2 silencing. eg, n = 30, mean ± s.e.m., one-way ANOVA/Bonferroni.

Extended Data Figure 10 Combined transcription inhibition and ATM activation results in extensive mobilization of mature spliceosomes.

a, Combinatorial effect of DRB and ionizing radiation on spliceosome mobilization. Quiescent HDFs were exposed to ionizing radiation in the presence or absence of DRB, and SF3a1–GFP and PRP8–GFP mobility was assayed by FRAP. b, The ionizing-radiation-mediated increase of DRB-dependent spliceosome mobilization depends on ATM activity. FRAP of GFP-tagged SNRNP40 in quiescent HDFs treated with DRB and/or ionizing radiation in the presence or absence of an ATM inibitor. c, Spliceosome mobilization by CPT. Quiescent HDFs were treated with 25 µg ml−1 CPT, 25 µM DRB and 20 J m−2 UV at doses that inhibit transcription to approximately 30% and their influence on SF3a1, PRP8 and SNRNP40 mobilization was measured by FRAP. Mobilization of GFP-tagged SF3a1, PRP8 and SNRNP40 in quiescent HDFs was measured by FRAP. ac, n = 30, mean ± s.e.m., one-way ANOVA/Bonferroni. d, Inhibition of RNA synthesis by the treatments shown in c was assayed in quiescent HDFs by 5EU incorporation and click chemistry (n = 150, mean ± s.e.m., one-way ANOVA/Dunnett’s).

Supplementary information

Supplementary information

This file contains Supplementary Tables 1-2. (PDF 1698 kb)

PowerPoint slides

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

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

Download citation

Further reading


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.