Abstract
Recent development of innovative tools for live imaging of actin filaments (F-actin) enabled the detection of surprising nuclear structures responding to various stimuli, challenging previous models that actin is substantially monomeric in the nucleus. We review these discoveries, focusing on double-strand break (DSB) repair responses. These studies revealed a remarkable network of nuclear filaments and regulatory mechanisms coordinating chromatin dynamics with repair progression and led to a paradigm shift by uncovering the directed movement of repair sites.
This is a preview of subscription content, access via your institution
Access options
Access Nature and 54 other Nature Portfolio journals
Get Nature+, our best-value online-access subscription
$29.99 / 30 days
cancel any time
Subscribe to this journal
Receive 12 print issues and online access
$209.00 per year
only $17.42 per issue
Buy this article
- Purchase on SpringerLink
- Instant access to full article PDF
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
References
Pollard, T. D. Actin and actin-binding proteins. Cold Spring Harb. Perspect. Biol. 8, a018226 (2016).
Rottner, K., Faix, J., Bogdan, S., Linder, S. & Kerkhoff, E. Actin assembly mechanisms at a glance. J. Cell Sci. 130, 3427–3435 (2017).
Titus, M. A. Myosin-driven intracellular transport. Cold Spring Harb. Perspect. Biol. 10, a021972 (2018).
Pollard, T. D. Regulation of actin filament assembly by Arp2/3 complex and formins. Annu. Rev. Biophys. Biomol. Struct. 36, 451–477 (2007).
Verboon, J. M., Sugumar, B. & Parkhurst, S. M. Wiskott-Aldrich syndrome proteins in the nucleus: aWASH with possibilities. Nucleus 6, 349–359 (2015).
Belin, B. J., Cimini, B. A., Blackburn, E. H. & Mullins, R. D. Visualization of actin filaments and monomers in somatic cell nuclei. Mol. Biol. Cell 24, 982–994 (2013).
Melak, M., Plessner, M. & Grosse, R. Actin visualization at a glance. J. Cell Sci. 130, 525–530 (2017).
Baarlink, C., Wang, H. & Grosse, R. Nuclear actin network assembly by formins regulates the SRF coactivator MAL. Science 340, 864–867 (2013).
Belin, B. J., Lee, T. & Mullins, R. D. DNA damage induces nuclear actin filament assembly by Formin -2 and Spire-½ that promotes efficient DNA repair. eLife 4, e07735 (2015).
Plessner, M., Melak, M., Chinchilla, P., Baarlink, C. & Grosse, R. Nuclear F-actin formation and reorganization upon cell spreading. J. Biol. Chem. 290, 11209–11216 (2015).
Caridi, C. P. et al. Nuclear F-actin and myosins drive relocalization of heterochromatic breaks. Nature 559, 54–60 (2018).
Baarlink, C. et al. A transient pool of nuclear F-actin at mitotic exit controls chromatin organization. Nat. Cell Biol. 19, 1389–1399 (2017).
Spracklen, A. J., Fagan, T. N., Lovander, K. E. & Tootle, T. L. The pros and cons of common actin labeling tools for visualizing actin dynamics during Drosophila oogenesis. Dev. Biol. 393, 209–226 (2014).
Plessner, M. & Grosse, R. Dynamizing nuclear actin filaments. Curr. Opin. Cell Biol. 56, 1–6 (2019).
Clark, T. G. & Rosenbaum, J. L. An actin filament matrix in hand-isolated nuclei of X. laevis oocytes. Cell 18, 1101–1108 (1979).
Bohnsack, M. T., Stüven, T., Kuhn, C., Cordes, V. C. & Görlich, D. A selective block of nuclear actin export stabilizes the giant nuclei of Xenopus oocytes. Nat. Cell Biol. 8, 257–263 (2006).
Dopie, J., Skarp, K. P., Rajakylä, E. K., Tanhuanpää, K. & Vartiainen, M. K. Active maintenance of nuclear actin by importin 9 supports transcription. Proc. Natl Acad. Sci. USA 109, E544–E552 (2012).
Feric, M. & Brangwynne, C. P. A nuclear F-actin scaffold stabilizes ribonucleoprotein droplets against gravity in large cells. Nat. Cell Biol. 15, 1253–1259 (2013).
Oda, H., Shirai, N., Ura, N., Ohsumi, K. & Iwabuchi, M. Chromatin tethering to the nuclear envelope by nuclear actin filaments: a novel role of the actin cytoskeleton in the Xenopus blastula. Genes Cells 22, 376–391 (2017).
Miyamoto, K., Pasque, V., Jullien, J. & Gurdon, J. B. Nuclear actin polymerization is required for transcriptional reprogramming of Oct4 by oocytes. Genes Dev. 25, 946–958 (2011).
Miyamoto, K. et al. Nuclear Wave1 is required for reprogramming transcription in oocytes and for normal development. Science 341, 1002–1005 (2013).
Miralles, F., Posern, G., Zaromytidou, A. I. & Treisman, R. Actin dynamics control SRF activity by regulation of its coactivator MAL. Cell 113, 329–342 (2003).
Vartiainen, M. K., Guettler, S., Larijani, B. & Treisman, R. Nuclear actin regulates dynamic subcellular localization and activity of the SRF cofactor MAL. Science 316, 1749–1752 (2007).
Kircher, P. et al. Filamin A interacts with the coactivator MKL1 to promote the activity of the transcription factor SRF and cell migration. Sci. Signal. 8, ra112 (2015).
Tsopoulidis, N. et al. T cell receptor-triggered nuclear actin network formation drives CD4+ T cell effector functions. Sci. Immunol. 4, eaav1987 (2019).
Chuang, C. H. et al. Long-range directional movement of an interphase chromosome site. Curr. Biol. 16, 825–831 (2006).
Dundr, M. et al. Actin-dependent intranuclear repositioning of an active gene locus in vivo. J. Cell Biol. 179, 1095–1103 (2007).
Philimonenko, V. V. et al. Nuclear actin and myosin I are required for RNA polymerase I transcription. Nat. Cell Biol. 6, 1165–1172 (2004).
Yoo, Y., Wu, X. & Guan, J. L. A novel role of the actin-nucleating Arp2/3 complex in the regulation of RNA polymerase II-dependent transcription. J. Biol. Chem. 282, 7616–7623 (2007).
Serebryannyy, L. A. et al. Persistent nuclear actin filaments inhibit transcription by RNA polymerase II. J. Cell Sci. 129, 3412–3425 (2016).
Söderberg, E., Hessle, V., von Euler, A. & Visa, N. Profilin is associated with transcriptionally active genes. Nucleus 3, 290–299 (2012).
Sokolova, M. et al. Nuclear actin is required for transcription during Drosophila oogenesis. iScience 9, 63–70 (2018).
Tondeleir, D. et al. Cells lacking β-actin are genetically reprogrammed and maintain conditional migratory capacity. Mol. Cell. Proteomics 11, 255–271 (2012).
Xie, X. et al. β-actin-dependent global chromatin organization and gene expression programs control cellular identity. FASEB J. 32, 1296–1314 (2018).
Xie, X., Jankauskas, R., Mazari, A. M. A., Drou, N. & Percipalle, P. β-actin regulates a heterochromatin landscape essential for optimal induction of neuronal programs during direct reprograming. PLoS Genet. 14, e1007846 (2018).
Klages-Mundt, N. L., Kumar, A., Zhang, Y., Kapoor, P. & Shen, X. The nature of actin-family proteins in chromatin-modifying complexes. Front. Genet. 9, 398 (2018).
Fenn, S. et al. Structural biochemistry of nuclear actin-related proteins 4 and 8 reveals their interaction with actin. EMBO J. 30, 2153–2166 (2011).
Cao, T. et al. Crystal structure of a nuclear actin ternary complex. Proc. Natl Acad. Sci. USA 113, 8985–8990 (2016).
Liu, C., Zhu, R. & Mao, Y. Nuclear actin polymerized by mDia2 confines centromere movement during CENP-A loading. iScience 9, 314–327 (2018).
Parisis, N. et al. Initiation of DNA replication requires actin dynamics and formin activity. EMBO J. 36, 3212–3231 (2017).
Raghuraman, M. K., Brewer, B. J. & Fangman, W. L. Cell cycle-dependent establishment of a late replication program. Science 276, 806–809 (1997).
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).
Dimitrova, D. S. & Gilbert, D. M. The spatial position and replication timing of chromosomal domains are both established in early G1 phase. Mol. Cell 4, 983–993 (1999).
Heun, P., Laroche, T., Raghuraman, M. K. & Gasser, S. M. The positioning and dynamics of origins of replication in the budding yeast nucleus. J. Cell Biol. 152, 385–400 (2001).
Shermoen, A. W., McCleland, M. L. & O’Farrell, P. H. Developmental control of late replication and S phase length. Curr. Biol. 20, 2067–2077 (2010).
Dileep, V. et al. Topologically associating domains and their long-range contacts are established during early G1 coincident with the establishment of the replication-timing program. Genome Res. 25, 1104–1113 (2015).
Knott, S. R. et al. Forkhead transcription factors establish origin timing and long-range clustering in S. cerevisiae. Cell 148, 99–111 (2012).
Fang, D. et al. Dbf4 recruitment by forkhead transcription factors defines an upstream rate-limiting step in determining origin firing timing. Genes Dev. 31, 2405–2415 (2017).
Zhang, H. et al. Dynamic relocalization of replication origins by Fkh1 requires execution of DDK function and Cdc45 loading at origins. eLife 8, e45512 (2019).
Peace, J. M., Ter-Zakarian, A. & Aparicio, O. M. Rif1 regulates initiation timing of late replication origins throughout the S. cerevisiae genome. PLoS One 9, e98501 (2014).
Hafner, L. et al. Rif1 binding and control of chromosome-internal DNA replication origins is limited by telomere sequestration. Cell Rep. 23, 983–992 (2018).
Welch, M. D. & Way, M. Arp2/3-mediated actin-based motility: a tail of pathogen abuse. Cell Host Microbe 14, 242–255 (2013).
Wilkie, A. R., Lawler, J. L. & Coen, D. M. A role for nuclear F-actin induction in human cytomegalovirus nuclear egress. MBio 7, e01254–16 (2016).
Ohkawa, T. & Welch, M. D. Baculovirus actin-based motility drives nuclear envelope disruption and nuclear egress. Curr. Biol. 28, 2153–2159.e4 (2018).
Chang, H. H. Y., Pannunzio, N. R., Adachi, N. & Lieber, M. R. Non-homologous DNA end joining and alternative pathways to double-strand break repair. Nat. Rev. Mol. Cell Biol. 18, 495–506 (2017).
Kowalczykowski, S. C. An overview of the molecular mechanisms of recombinational DNA repair. Cold Spring Harb. Perspect. Biol. 7, a016410 (2015).
Zuchero, J. B., Coutts, A. S., Quinlan, M. E., Thangue, N. B. & Mullins, R. D. p53-cofactor JMY is a multifunctional actin nucleation factor. Nat. Cell Biol. 11, 451–459 (2009).
Yuan, Y. & Shen, Z. Interaction with BRCA2 suggests a role for filamin-1 (hsFLNa) in DNA damage response. J. Biol. Chem. 276, 48318–48324 (2001).
Yue, J. et al. The cytoskeleton protein filamin-A is required for an efficient recombinational DNA double strand break repair. Cancer Res. 69, 7978–7985 (2009).
Velkova, A., Carvalho, M. A., Johnson, J. O., Tavtigian, S. V. & Monteiro, A. N. Identification of Filamin A as a BRCA1-interacting protein required for efficient DNA repair. Cell Cycle 9, 1421–1433 (2010).
Hansen, R. K. et al. SCAI promotes DNA double-strand break repair in distinct chromosomal contexts. Nat. Cell Biol. 18, 1357–1366 (2016).
Isobe, S. Y., Nagao, K., Nozaki, N., Kimura, H. & Obuse, C. Inhibition of RIF1 by SCAI allows BRCA1-mediated repair. Cell Reports 20, 297–307 (2017).
Brandt, D. T. et al. SCAI acts as a suppressor of cancer cell invasion through the transcriptional control of β1-integrin. Nat. Cell Biol. 11, 557–568 (2009).
Sridharan, D., Brown, M., Lambert, W. C., McMahon, L. W. & Lambert, M. W. Nonerythroid alphaII spectrin is required for recruitment of FANCA and XPF to nuclear foci induced by DNA interstrand cross-links. J. Cell Sci. 116, 823–835 (2003).
Andrin, C. et al. A requirement for polymerized actin in DNA double-strand break repair. Nucleus 3, 384–395 (2012).
Kulashreshtha, M., Mehta, I. S., Kumar, P. & Rao, B. J. Chromosome territory relocation during DNA repair requires nuclear myosin 1 recruitment to chromatin mediated by ϒ-H2AX signaling. Nucleic Acids Res. 44, 8272–8291 (2016).
Spichal, M. et al. Evidence for a dual role of actin in regulating chromosome organization and dynamics in yeast. J. Cell Sci. 129, 681–692 (2016).
Evdokimova, V. N., Gandhi, M., Nikitski, A. V., Bakkenist, C. J. & Nikiforov, Y. E. Nuclear myosin/actin-motored contact between homologous chromosomes is initiated by ATM kinase and homology-directed repair proteins at double-strand DNA breaks to suppress chromosome rearrangements. Oncotarget 9, 13612–13622 (2018).
Marnef, A. et al. A cohesin/HUSH- and LINC-dependent pathway controls ribosomal DNA double-strand break repair. Genes Dev. 33, 1–16 (2019).
Kapoor, P. & Shen, X. Mechanisms of nuclear actin in chromatin-remodeling complexes. Trends Cell Biol. 24, 238–246 (2014).
Neumann, F. R. et al. Targeted INO80 enhances subnuclear chromatin movement and ectopic homologous recombination. Genes Dev. 26, 369–383 (2012).
Horigome, C. et al. SWR1 and INO80 chromatin remodelers contribute to DNA double-strand break perinuclear anchorage site choice. Mol. Cell 55, 626–639 (2014).
Park, E. J., Hur, S. K. & Kwon, J. Human INO80 chromatin-remodelling complex contributes to DNA double-strand break repair via the expression of Rad54B and XRCC3 genes. Biochem. J. 431, 179–187 (2010).
Wang, Y. H. et al. DNA damage causes rapid accumulation of phosphoinositides for ATR signaling. Nat. Commun. 8, 2118 (2017).
Sun, M. H. et al. DNA double-strand breaks induce the nuclear actin filaments formation in cumulus-enclosed oocytes but not in denuded oocytes. PLoS One 12, e0170308 (2017).
Hoskins, R. A. et al. Sequence finishing and mapping of Drosophila melanogaster heterochromatin. Science 316, 1625–1628 (2007).
Ho, J. W. et al. Comparative analysis of metazoan chromatin organization. Nature 512, 449–452 (2014).
Hoskins, R. A. et al. The Release 6 reference sequence of the Drosophila melanogaster genome. Genome Res. 25, 445–458 (2015).
Lachner, M., O’Carroll, D., Rea, S., Mechtler, K. & Jenuwein, T. Methylation of histone H3 lysine 9 creates a binding site for HP1 proteins. Nature 410, 116–120 (2001).
Riddle, N. C. et al. Plasticity in patterns of histone modifications and chromosomal proteins in Drosophila heterochromatin. Genome Res. 21, 147–163 (2011).
Guelen, L. et al. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453, 948–951 (2008).
Peric-Hupkes, D. et al. Molecular maps of the reorganization of genome-nuclear lamina interactions during differentiation. Mol. Cell 38, 603–613 (2010).
Sexton, T. et al. Three-dimensional folding and functional organization principles of the Drosophila genome. Cell 148, 458–472 (2012).
Goodarzi, A. A. et al. ATM signaling facilitates repair of DNA double-strand breaks associated with heterochromatin. Mol. Cell 31, 167–177 (2008).
Chiolo, I. et al. Double-strand breaks in heterochromatin move outside of a dynamic HP1a domain to complete recombinational repair. Cell 144, 732–744 (2011).
Chiolo, I., Tang, J., Georgescu, W. & Costes, S. V. Nuclear dynamics of radiation-induced foci in euchromatin and heterochromatin. Mutat. Res. 750, 56–66 (2013).
Ryu, T. et al. Heterochromatic breaks move to the nuclear periphery to continue recombinational repair. Nat. Cell Biol. 17, 1401–1411 (2015).
Tsouroula, K. et al. Temporal and spatial uncoupling of DNA double strand break repair pathways within mammalian heterochromatin. Mol. Cell 63, 293–305 (2016).
Li, Q. et al. The three-dimensional genome organization of Drosophila melanogaster through data integration. Genome Biol. 18, 145 (2017).
Caridi, P. C., Delabaere, L., Zapotoczny, G. & Chiolo, I. And yet, it moves: nuclear and chromatin dynamics of a heterochromatic double-strand break. Phil. Trans. R. Soc. Lond. B 372, 20160291 (2017).
Peng, J. C. & Karpen, G. H. H3K9 methylation and RNA interference regulate nucleolar organization and repeated DNA stability. Nat. Cell Biol. 9, 25–35 (2007).
Peng, J. C. & Karpen, G. H. Epigenetic regulation of heterochromatic DNA stability. Curr. Opin. Genet. Dev. 18, 204–211 (2008).
Peng, J. C. & Karpen, G. H. Heterochromatic genome stability requires regulators of histone H3 K9 methylation. PLoS Genet. 5, e1000435 (2009).
Ryu, T., Bonner, M. R. & Chiolo, I. Cervantes and Quijote protect heterochromatin from aberrant recombination and lead the way to the nuclear periphery. Nucleus 7, 485–497 (2016).
Dialynas, G., Delabaere, L. & Chiolo, I. Arp2/3 and Unc45 maintain heterochromatin stability in Drosophila polytene chromosomes. Exp. Biol. Med. https://doi.org/10.1177/1535370219862282 (2019).
Beucher, A. et al. ATM and Artemis promote homologous recombination of radiation-induced DNA double-strand breaks in G2. EMBO J. 28, 3413–3427 (2009).
Janssen, A. et al. A single double-strand break system reveals repair dynamics and mechanisms in heterochromatin and euchromatin. Genes Dev. 30, 1645–1657 (2016).
Amaral, N., Ryu, T., Li, X. & Chiolo, I. Nuclear dynamics of heterochromatin repair. Trends Genet. 33, 86–100 (2017).
Rawal, C. P.C. C. & Chiolo, I. Actin’ between phase separated domains for heterochromatin repair. DNA Repair https://doi.org/10.1016/j.dnarep.2019.102646 (2019).
Guenatri, M., Bailly, D., Maison, C. & Almouzni, G. Mouse centric and pericentric satellite repeats form distinct functional heterochromatin. J. Cell Biol. 166, 493–505 (2004).
Jakob, B. et al. DNA double-strand breaks in heterochromatin elicit fast repair protein recruitment, histone H2AX phosphorylation and relocation to euchromatin. Nucleic Acids Res. 39, 6489–6499 (2011).
Ayoub, N., Jeyasekharan, A. D., Bernal, J. A. & Venkitaraman, A. R. HP1-β mobilization promotes chromatin changes that initiate the DNA damage response. Nature 453, 682–686 (2008).
Dronamraju, R. & Mason, J. M. MU2 and HP1a regulate the recognition of double strand breaks in Drosophila melanogaster. PLoS One 6, e25439 (2011).
Colmenares, S. U. et al. Drosophila histone demethylase KDM4A has enzymatic and non-enzymatic roles in controlling heterochromatin integrity. Dev. Cell 42, 156–169.e5 (2017).
Goodarzi, A. A., Kurka, T. & Jeggo, P. A. KAP-1 phosphorylation regulates CHD3 nucleosome remodeling during the DNA double-strand break response. Nat. Struct. Mol. Biol. 18, 831–839 (2011).
Delabaere, L. & Chiolo, I. ReiNF4rcing repair pathway choice during cell cycle. Cell Cycle 15, 1182–1183 (2016).
Senaratne, T. N., Joyce, E. F., Nguyen, S. C. & Wu, C. T. Investigating the interplay between sister chromatid cohesion and homolog pairing in Drosophila nuclei. PLoS Genet. 12, e1006169 (2016).
See, C., Arya, D., Lin, E. & Chiolo, I. Live cell imaging of nuclear actin filaments and heterochromatic repair foci in Drosophila and mouse cells. Preprint at PeerJ Preprints https://doi.org/10.7287/peerj.preprints.27900v1 (2019).
Caridi, C. P. et al. Quantitative methods to investigate the 4D dynamics of heterochromatic repair sites in Drosophila cells. Methods Enzymol. 601, 359–389 (2018).
Schrank, B. R. et al. Nuclear ARP2/3 drives DNA break clustering for homology-directed repair. Nature 559, 61–66 (2018).
Lisby, M., Mortensen, U. H. & Rothstein, R. Colocalization of multiple DNA double-strand breaks at a single Rad52 repair centre. Nat. Cell Biol. 5, 572–577 (2003).
Aten, J. A. et al. Dynamics of DNA double-strand breaks revealed by clustering of damaged chromosome domains. Science 303, 92–95 (2004).
Kruhlak, M. J. et al. Changes in chromatin structure and mobility in living cells at sites of DNA double-strand breaks. J. Cell Biol. 172, 823–834 (2006).
Neumaier, T. et al. Evidence for formation of DNA repair centers and dose-response nonlinearity in human cells. Proc. Natl Acad. Sci. USA 109, 443–448 (2012).
Aymard, F. et al. Genome-wide mapping of long-range contacts unveils clustering of DNA double-strand breaks at damaged active genes. Nat. Struct. Mol. Biol. 24, 353–361 (2017).
Costes, S. V., Chiolo, I., Pluth, J. M., Barcellos-Hoff, M. H. & Jakob, B. Spatiotemporal characterization of ionizing radiation induced DNA damage foci and their relation to chromatin organization. Mutat. Res. 704, 78–87 (2010).
Clouaire, T. et al. Comprehensive mapping of histone modifications at DNA double-strand breaks deciphers repair pathway chromatin signatures. Mol. Cell 72, 250–262.e6 (2018).
Lottersberger, F., Karssemeijer, R. A., Dimitrova, N. & de Lange, T. 53BP1 and the LINC complex promote microtubule-dependent DSB mobility and DNA repair. Cell 163, 880–893 (2015).
Lawrimore, J. et al. Microtubule dynamics drive enhanced chromatin motion and mobilize telomeres in response to DNA damage. Mol. Biol. Cell 28, 1701–1711 (2017).
Oshidari, R. et al. Nuclear microtubule filaments mediate non-linear directional motion of chromatin and promote DNA repair. Nat. Commun. 9, 2567 (2018).
Swartz, R. K., Rodriguez, E. C. & King, M. C. A role for nuclear envelope-bridging complexes in homology-directed repair. Mol. Biol. Cell 25, 2461–2471 (2014).
Chung, D. K. et al. Perinuclear tethers license telomeric DSBs for a broad kinesin- and NPC-dependent DNA repair process. Nat. Commun. 6, 7742 (2015).
Nagai, S. et al. Functional targeting of DNA damage to a nuclear pore-associated SUMO-dependent ubiquitin ligase. Science 322, 597–602 (2008).
Su, X. A., Dion, V., Gasser, S. M. & Freudenreich, C. H. Regulation of recombination at yeast nuclear pores controls repair and triplet repeat stability. Genes Dev. 29, 1006–1017 (2015).
Quivy, J.-P., Gérard, A., Cook, A. J., Roche, D. & Almouzni, G. The HP1-p150/CAF-1 interaction is required for pericentric heterochromatin replication and S-phase progression in mouse cells. Nat. Struct. Mol. Biol. 15, 972–979 (2008).
Johnson, M. A., Sharma, M., Mok, M. T. & Henderson, B. R. Stimulation of in vivo nuclear transport dynamics of actin and its co-factors IQGAP1 and Rac1 in response to DNA replication stress. Biochim. Biophys. Acta 1833, 2334–2347 (2013).
Lamm, N., Masamsetti, V.P., Read, M.N., Biro, M. & Cesare, A.J. ATR and mTOR regulate F-actin to alter nuclear architecture and repair replication stress. bioRxiv preprint at https://doi.org/10.1101/451708 (2018).
Bermejo, R. et al. The replication checkpoint protects fork stability by releasing transcribed genes from nuclear pores. Cell 146, 233–246 (2011).
Kumar, A. et al. ATR mediates a checkpoint at the nuclear envelope in response to mechanical stress. Cell 158, 633–646 (2014).
Torres-Rosell, J. et al. The Smc5-Smc6 complex and SUMO modification of Rad52 regulates recombinational repair at the ribosomal gene locus. Nat. Cell Biol. 9, 923–931 (2007).
Harding, S. M., Boiarsky, J. A. & Greenberg, R. A. ATM dependent silencing links nucleolar chromatin reorganization to DNA damage recognition. Cell Rep. 13, 251–259 (2015).
van Sluis, M. & McStay, B. A localized nucleolar DNA damage response facilitates recruitment of the homology-directed repair machinery independent of cell cycle stage. Genes Dev. 29, 1151–1163 (2015).
Horigome, C., Unozawa, E., Ooki, T. & Kobayashi, T. Ribosomal RNA gene repeats associate with the nuclear pore complex for maintenance after DNA damage. PLoS Genet. 15, e1008103 (2019).
Therizols, P. et al. Telomere tethering at the nuclear periphery is essential for efficient DNA double strand break repair in subtelomeric region. J. Cell Biol. 172, 189–199 (2006).
Khadaroo, B. et al. The DNA damage response at eroded telomeres and tethering to the nuclear pore complex. Nat. Cell Biol. 11, 980–987 (2009).
Cho, N. W., Dilley, R. L., Lampson, M. A. & Greenberg, R. A. Interchromosomal homology searches drive directional ALT telomere movement and synapsis. Cell 159, 108–121 (2014).
Churikov, D. et al. SUMO-dependent relocalization of eroded telomeres to nuclear pore complexes controls telomere recombination. Cell Rep. 15, 1242–1253 (2016).
Kalocsay, M., Hiller, N. J. & Jentsch, S. Chromosome-wide Rad51 spreading and SUMO-H2A.Z-dependent chromosome fixation in response to a persistent DNA double-strand break. Mol. Cell 33, 335–343 (2009).
Oza, P., Jaspersen, S. L., Miele, A., Dekker, J. & Peterson, C. L. Mechanisms that regulate localization of a DNA double-strand break to the nuclear periphery. Genes Dev. 23, 912–927 (2009).
Roukos, V. et al. Spatial dynamics of chromosome translocations in living cells. Science 341, 660–664 (2013).
Miné-Hattab, J., Recamier, V., Izeddin, I., Rothstein, R. & Darzacq, X. in Molecular Biology of the Cell vol. 28 (eds Lidke, D., Lippincott-Schwartz, J. and Mogilner, A.) 3323–3332 (American Society for Cell Biology, 2017).
Dion, V., Kalck, V., Horigome, C., Towbin, B. D. & Gasser, S. M. Increased mobility of double-strand breaks requires Mec1, Rad9 and the homologous recombination machinery. Nat. Cell Biol. 14, 502–509 (2012).
Miné-Hattab, J. & Rothstein, R. Increased chromosome mobility facilitates homology search during recombination. Nat. Cell Biol. 14, 510–517 (2012).
Amitai, A., Seeber, A., Gasser, S. M. & Holcman, D. Visualization of chromatin decompaction and break site extrusion as predicted by statistical polymer modeling of single-locus trajectories. Cell Rep. 18, 1200–1214 (2017).
Spichal, M. & Fabre, E. The emerging role of the cytoskeleton in chromosome dynamics. Front. Genet. 8, 60 (2017).
Dion, V. & Gasser, S. M. Chromatin movement in the maintenance of genome stability. Cell 152, 1355–1364 (2013).
Hatakeyama, H., Nakahata, Y., Yarimizu, H. & Kanzaki, M. Live-cell single-molecule labeling and analysis of myosin motors with quantum dots. Mol. Biol. Cell 28, 173–181 (2017).
Li, H., Guo, F., Rubinstein, B. & Li, R. Actin-driven chromosomal motility leads to symmetry breaking in mammalian meiotic oocytes. Nat. Cell Biol. 10, 1301–1308 (2008).
Larson, A. G. et al. Liquid droplet formation by HP1α suggests a role for phase separation in heterochromatin. Nature 547, 236–240 (2017).
Strom, A. R. et al. Phase separation drives heterochromatin domain formation. Nature 547, 241–245 (2017).
Azzaz, A. M. et al. Human heterochromatin protein 1α promotes nucleosome associations that drive chromatin condensation. J. Biol. Chem. 289, 6850–6861 (2014).
Cahill, D. P., Kinzler, K. W., Vogelstein, B. & Lengauer, C. Genetic instability and darwinian selection in tumours. Trends Cell Biol. 9, M57–M60 (1999).
Zhang, C. Z. et al. Chromothripsis from DNA damage in micronuclei. Nature 522, 179–184 (2015).
Yang, X. & Lin, Y. Functions of nuclear actin-binding proteins in human cancer. Oncol. Lett. 15, 2743–2748 (2018).
Buchbinder, D., Nugent, D. J. & Fillipovich, A. H. Wiskott-Aldrich syndrome: diagnosis, current management, and emerging treatments. Appl. Clin. Genet. 7, 55–66 (2014).
The Huntington’s Disease Collaborative Research Group. A novel gene containing a trinucleotide repeat that is expanded and unstable on Huntington’s disease chromosomes. Cell 72, 971–983 (1993).
Mangiarini, L. et al. Exon 1 of the HD gene with an expanded CAG repeat is sufficient to cause a progressive neurological phenotype in transgenic mice. Cell 87, 493–506 (1996).
Munsie, L. et al. Mutant huntingtin causes defective actin remodeling during stress: defining a new role for transglutaminase 2 in neurodegenerative disease. Hum. Mol. Genet. 20, 1937–1951 (2011).
Maiuri, T. et al. Huntingtin is a scaffolding protein in the ATM oxidative DNA damage response complex. Hum. Mol. Genet. 26, 395–406 (2017).
Prochniewicz, E., Thompson, L. V. & Thomas, D. D. Age-related decline in actomyosin structure and function. Exp. Gerontol. 42, 931–938 (2007).
White, R. R. et al. Double-strand break repair by interchromosomal recombination: an in vivo repair mechanism utilized by multiple somatic tissues in mammals. PLoS One 8, e84379 (2013).
Sukup-Jackson, M. R. et al. Rosa26-GFP direct repeat (RaDR-GFP) mice reveal tissue- and age-dependence of homologous recombination in mammals in vivo. PLoS Genet. 10, e1004299 (2014).
Delabaere, L. et al. Aging impairs double-strand break repair by homologous recombination in Drosophila germ cells. Aging Cell 16, 320–328 (2017).
Simon, D. N., Zastrow, M. S. & Wilson, K. L. Direct actin binding to A- and B-type lamin tails and actin filament bundling by the lamin A tail. Nucleus 1, 264–272 (2010).
Scaffidi, P. & Misteli, T. Reversal of the cellular phenotype in the premature aging disease Hutchinson-Gilford progeria syndrome. Nat. Med. 11, 440–445 (2005).
Scaffidi, P. & Misteli, T. Lamin A-dependent nuclear defects in human aging. Science 312, 1059–1063 (2006).
Zada, D., Bronshtein, I., Lerer-Goldshtein, T., Garini, Y. & Appelbaum, L. Sleep increases chromosome dynamics to enable reduction of accumulating DNA damage in single neurons. Nat. Commun. 10, 895 (2019).
Acknowledgements
We apologize to our colleagues whose work could not be cited owing to space limitations. We thank S. Keagy for useful comments on the manuscript. This work is supported by NIH R01GM117376 and NSF Career 1751197 to I.C. and the DFG GR 2111/7-1 to R.G.
Author information
Authors and Affiliations
Contributions
C.P.C., R.G., and I.C. contributed to manuscript and figure preparation. M.P. contributed to Fig. 1.
Corresponding author
Ethics declarations
Competing interests
The authors declare no competing interests.
Additional information
Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Rights and permissions
About this article
Cite this article
Caridi, C.P., Plessner, M., Grosse, R. et al. Nuclear actin filaments in DNA repair dynamics. Nat Cell Biol 21, 1068–1077 (2019). https://doi.org/10.1038/s41556-019-0379-1
Received:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1038/s41556-019-0379-1
This article is cited by
-
Bioinformatics Analysis of Actin Interactome: Characterization of the Nuclear and Cytoplasmic Actin-Binding Proteins
The Protein Journal (2024)
-
DNA replication and replication stress response in the context of nuclear architecture
Chromosoma (2024)
-
Loss of p53 function promotes DNA damage-induced formation of nuclear actin filaments
Cell Death & Disease (2023)
-
Nuclear myosin VI maintains replication fork stability
Nature Communications (2023)
-
Cancer cells remodel nuclear actin filaments to resist chemotherapy
Nature (2023)