Many cellular processes require large-scale rearrangements of chromatin structure. Structural maintenance of chromosomes (SMC) protein complexes are molecular machines that can provide structure to chromatin. These complexes can connect DNA elements in cis, walk along DNA, build and processively enlarge DNA loops and connect DNA molecules in trans to hold together the sister chromatids. These DNA-shaping abilities place SMC complexes at the heart of many DNA-based processes, including chromosome segregation in mitosis, transcription control and DNA replication, repair and recombination. In this Review, we discuss the latest insights into how SMC complexes such as cohesin, condensin and the SMC5–SMC6 complex shape DNA to direct these fundamental chromosomal processes. We also consider how SMC complexes, by building chromatin loops, can counteract the natural tendency of alike chromatin regions to cluster. SMC complexes thus control nuclear organization by participating in a molecular tug of war that determines the architecture of our genome.
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Chuang, P. T., Albertson, D. G. & Meyer, B. J. DPY-27: a chromosome condensation protein homolog that regulates C. elegans dosage compensation through association with the X chromosome. Cell 79, 459–474 (1994).
Niki, H., Jaffe, A., Imamura, R., Ogura, T. & Hiraga, S. The new gene mukB codes for a 177 kD protein with coiled-coil domains involved in chromosome partitioning of E. coli. EMBO J. 10, 183–193 (1991).
Guacci, V., Koshland, D. & Strunnikov, A. A direct link between sister chromatid cohesion and chromosome condensation revealed through the analysis of MCD1 in S. cerevisiae. Cell 91, 47–57 (1997).
Hirano, T. & Mitchison, T. J. A heterodimeric coiled-coil protein required for mitotic chromosome condensation in vitro. Cell 79, 449–458 (1994).
Michaelis, C., Ciosk, R. & Nasmyth, K. Cohesins: chromosomal proteins that prevent premature separation of sister chromatids. Cell 91, 35–45 (1997).
Saka, Y. et al. Fission yeast cut3 and cut14, members of a ubiquitous protein family, are required for chromosome condensation and segregation in mitosis. EMBO J. 13, 4938–4952 (1994).
Strunnikov, A. V., Larionov, V. L. & Koshland, D. SMC1: an essential yeast gene encoding a putative head-rod-tail protein is required for nuclear division and defines a new ubiquitous protein family. J. Cell Biol. 123, 1635–1648 (1993).
Saitoh, N., Goldberg, I. G., Wood, E. R. & Earnshaw, W. C. ScII: an abundant chromosome scaffold protein is a member of a family of putative ATPases with an unusual predicted tertiary structure. J. Cell Biol. 127, 303–318 (1994).
Hirano, T., Kobayashi, R. & Hirano, M. Condensins, chromosome condensation protein complexes containing XCAP-C, XCAP-E and a xenopus homolog of the Drosophila barren protein. Cell 89, 511–521 (1997).
Losada, A., Hirano, M. & Hirano, T. Identification of Xenopus SMC protein complexes required for sister chromatid cohesion. Genes Dev. 12, 1986–1997 (1998).
Wells, J. N., Gligoris, T. G., Nasmyth, K. A. & Marsh, J. A. Evolution of condensin and cohesin complexes driven by replacement of Kite by Hawk proteins. Curr. Biol. 27, R17–R18 (2017).
Palecek, J. J. & Gruber, S. Kite proteins: a superfamily of SMC/kleisin partners conserved across bacteria, archaea, and eukaryotes. Structure 23, 2183–2190 (2015).
Petela, N. J. et al. Scc2 is a potent activator of cohesin’s ATPase that promotes loading by binding Scc1 without Pds5. Mol. Cell 70, 1134–1148 (2018).
van Ruiten, M. S. et al. The cohesin acetylation cycle controls chromatin loop length through a PDS5A brake mechanism. Nat. Struct. Mol. Biol. 29, 586–591 (2022).
Bastié, N. et al. Smc3 acetylation, Pds5 and Scc2 control the translocase activity that establishes cohesin-dependent chromatin loops. Nat. Struct. Mol. Biol. 29, 575–585 (2022).
Kolesar, P., Stejskal, K., Potesil, D., Murray, J. M. & Palecek, J. J. Role of Nse1 subunit of SMC5/6 complex as a ubiquitin ligase. Cells 11, 165 (2022).
Andrews, E. A. et al. Nse2, a component of the Smc5–6 complex, is a SUMO ligase required for the response to DNA damage. Mol. Cell Biol. 25, 185–196 (2005).
Zhao, X. & Blobel, G. A SUMO ligase is part of a nuclear multiprotein complex that affects DNA repair and chromosomal organization. Proc. Natl Acad. Sci. USA 102, 4777–4782 (2005).
Taschner, M. et al. Nse5/6 inhibits the Smc5/6 ATPase and modulates DNA substrate binding. EMBO J. 40, e107807 (2021).
Hallett, S. T. et al. Nse5/6 is a negative regulator of the ATPase activity of the Smc5/6 complex. Nucleic Acids Res. 49, 4534–4549 (2021).
Räschle, M. et al. Proteomics reveals dynamic assembly of repair complexes during bypass of DNA cross-links. Science 348, 1253671 (2015).
Nasmyth, K. Disseminating the genome: joining, resolving, and separating sister chromatids during mitosis and meiosis. Annu. Rev. Genet. 35, 673–745 (2001). This Review puts forward the hypothesis that SMC complexes shape DNA through a mechanism of processive loop enlargement.
Ryu, J.-K. et al. Condensin extrudes DNA loops in steps up to hundreds of base pairs that are generated by ATP binding events. Nucleic Acids Res. 50, 820–832 (2022).
Pradhan, B. et al. SMC complexes can traverse physical roadblocks bigger than their ring size. Cell Rep. 41, 111491 (2022).
Oldenkamp, R. & Rowland, B. D. A walk through the SMC cycle: from catching DNAs to shaping the genome. Mol. Cell 82, 1616–1630 (2022).
Davidson, I. F. & Peters, J. M. Genome folding through loop extrusion by SMC complexes. Nat. Rev. Mol. Cell Biol. 22, 445–464 (2021).
Birkenbihl, R. P. & Subramani, S. Cloning and characterization of rad21 an essential gene of Schizosaccharomyces pombe involved in DNA double-strand-break repair. Nucleic Acids Res. 20, 6605–6611 (1992).
Abramo, K. et al. A chromosome folding intermediate at the condensin-to-cohesin transition during telophase. Nat. Cell Biol. 21, 1393–1402 (2019).
Zhang, H. et al. Chromatin structure dynamics during the mitosis-to-G1 phase transition. Nature 576, 158–162 (2019).
Ciosk, R. et al. Cohesin’s binding to chromosomes depends on a separate complex consisting of Scc2 and Scc4 proteins. Mol. Cell 5, 243–254 (2000).
Davidson, I. F. et al. DNA loop extrusion by human cohesin. Science 366, 1338–1345 (2019).
Kim, Y., Shi, Z., Zhang, H., Finkelstein, I. J. & Yu, H. Human cohesin compacts DNA by loop extrusion. Science 366, 1345–1349 (2019).
Beckouët, F. et al. Releasing activity disengages cohesin’s Smc3/Scc1 interface in a process blocked by acetylation. Mol. Cell 61, 563–574 (2016).
Murayama, Y. & Uhlmann, F. DNA entry into and exit out of the cohesin ring by an interlocking gate mechanism. Cell 163, 1628–1640 (2015).
Eichinger, C. S., Kurze, A., Oliveira, R. A. & Nasmyth, K. Disengaging the Smc3/kleisin interface releases cohesin from Drosophila chromosomes during interphase and mitosis. EMBO J. 32, 656–665 (2013).
Chan, K. L. et al. Cohesin’s DNA exit gate is distinct from its entrance gate and is regulated by acetylation. Cell 150, 961–974 (2012).
Buheitel, J. & Stemmann, O. Prophase pathway-dependent removal of cohesin from human chromosomes requires opening of the Smc3–Scc1 gate. EMBO J. 32, 666–676 (2013).
Kueng, S. et al. Wapl controls the dynamic association of cohesin with chromatin. Cell 127, 955–967 (2006).
Gandhi, R., Gillespie, P. J. & Hirano, T. Human Wapl is a cohesin-binding protein that promotes sister-chromatid resolution in mitotic prophase. Curr. Biol. 16, 2406–2417 (2006).
Haarhuis, J. H. I. et al. The cohesin release factor WAPL restricts chromatin loop extension. Cell 169, 693–707 (2017). Together with Gassler et al. (2017) and Wutz et al. (2017), these papers reveal that increasing the residence time of cohesin on the DNA causes a genome-wide extension of chromatin loops.
Gassler, J. et al. A mechanism of cohesin dependent loop extrusion organizes zygotic genome architecture. EMBO J. 36, 3600–3618 (2017).
Wutz, G. et al. Topologically associating domains and chromatin loops depend on cohesin and are regulated by CTCF, WAPL, and PDS5 proteins. EMBO J. 36, 3573–3599 (2017).
Schwarzer, W. et al. Two independent modes of chromatin organization revealed by cohesin removal. Nature 551, 51–56 (2017).
Rao, S. S. P. et al. Cohesin loss eliminates all loop domains. Cell 171, 305–320 (2017). Together with Haarhuis et al. (2017), Gassler et al. (2017), Wutz et al. (2017) and Schwarzer et al. ( 2017), these papers reveal that cohesin counteracts compartmentalization. In the study by Rao et al., cohesin was revealed to be required for the formation of all TADs and able to rebuild such structures in under an hour.
Golfier, S., Quail, T., Kimura, H. & Brugués, J. Cohesin and condensin extrude DNA loops in a cell-cycle dependent manner. eLife 9, e53885 (2020).
Zhang, H. et al. CTCF and R-loops are boundaries of cohesin-mediated DNA looping. Preprint at bioRxiv https://doi.org/10.1101/2022.09.15.508177 (2022).
Davidson, I. F. et al. CTCF is a DNA-tension-dependent barrier to cohesin-mediated loop extrusion. Nature 616, 822–827 (2023).
Parelho, V. et al. Cohesins functionally associate with CTCF on mammalian chromosome arms. Cell 132, 422–433 (2008).
Wendt, K. S. et al. Cohesin mediates transcriptional insulation by CCCTC-binding factor. Nature 451, 796–801 (2008).
Davidson, I. F. et al. Rapid movement and transcriptional relocalization of human cohesin on DNA. EMBO J. 35, 2671–2685 (2016).
Rao, S. S. P. et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell 159, 1665–1680 (2014).
Vietri Rudan, M. et al. Comparative Hi-C reveals that CTCF underlies evolution of chromosomal domain architecture. Cell Rep. 10, 1297–1309 (2015).
Nakahashi, H. et al. A genome-wide map of CTCF multivalency redefines the CTCF code. Cell Rep. 3, 1678–1689 (2013).
Li, Y. et al. The structural basis for cohesin–CTCF-anchored loops. Nature 578, 472–476 (2020).
Pugacheva, E. M. et al. CTCF mediates chromatin looping via N-terminal domain-dependent cohesin retention. Proc. Natl Acad. Sci. USA 117, 2020–2031 (2020).
Nishana, M. et al. Defining the relative and combined contribution of CTCF and CTCFL to genomic regulation. Genome Biol. 21, 108 (2020).
Nora, E. P. et al. Molecular basis of CTCF binding polarity in genome folding. Nat. Commun. 11, 5612 (2020).
Liu, Y. & Dekker, J. CTCF–CTCF loops and intra-TAD interactions show differential dependence on cohesin ring integrity. Nat. Cell Biol. 24, 1516–1527 (2022).
Saldaña-Meyer, R. et al. RNA interactions are essential for CTCF-mediated genome organization. Mol. Cell 76, 412–422 (2019).
Hansen, A. S. et al. Distinct classes of chromatin loops revealed by deletion of an RNA-binding region in CTCF. Mol. Cell 76, 395–411 (2019).
Wutz, G. et al. ESCO1 and CTCF enable formation of long chromatin loops by protecting cohesin/stag1 from WAPL. eLife 9, e52091 (2020).
Psakhye, I. & Branzei, D. SMC complexes are guarded by the SUMO protease Ulp2 against SUMO-chain-mediated turnover. Cell Rep. 36, 109485 (2021).
Wagner, K. et al. The SUMO isopeptidase SENP6 functions as a rheostat of chromatin residency in genome maintenance and chromosome dynamics. Cell Rep. 29, 480–494 (2019).
Hansen, A. S., Pustova, I., Cattoglio, C., Tjian, R. & Darzacq, X. CTCF and cohesin regulate chromatin loop stability with distinct dynamics. eLife 6, e25776 (2017).
Cattoglio, C. et al. Determining cellular CTCF and cohesin abundances to constrain 3D genome models. eLife 8, e40164 (2019).
Holzmann, J. et al. Absolute quantification of cohesin, CTCF and their regulators in human cells. eLife 8, e46269 (2019).
Stevens, T. J. et al. 3D structures of individual mammalian genomes studied by single-cell Hi-C. Nature 544, 59–64 (2017).
Bintu, B. et al. Super-resolution chromatin tracing reveals domains and cooperative interactions in single cells. Science 362, eaau1783 (2018).
Su, J. H., Zheng, P., Kinrot, S. S., Bintu, B. & Zhuang, X. Genome-scale imaging of the 3D organization and transcriptional activity of chromatin. Cell 182, 1641–1659 (2020).
Flyamer, I. M. et al. Single-nucleus Hi-C reveals unique chromatin reorganization at oocyte-to-zygote transition. Nature 544, 110–114 (2017).
Finn, E. H. et al. Extensive heterogeneity and intrinsic variation in spatial genome organization. Cell 176, 1502–1515 (2019).
Luppino, J. M. et al. Cohesin promotes stochastic domain intermingling to ensure proper regulation of boundary-proximal genes. Nat. Genet. 52, 840–848 (2020).
Gabriele, M. et al. Dynamics of CTCF- and cohesin-mediated chromatin looping revealed by live-cell imaging. Science 376, 476–501 (2022). This study shows by imaging single cells that CTCF-anchored chromatin loops are highly dynamic and that loop domains are rarely fully looped.
Mach, P. et al. Cohesin and CTCF control the dynamics of chromosome folding. Nat. Genet. 54, 1907–1918 (2022).
Symmons, O. et al. The Shh topological domain facilitates the action of remote enhancers by reducing the effects of genomic distances. Dev. Cell 39, 529–543 (2016).
Flavahan, W. A. et al. Insulator dysfunction and oncogene activation in IDH mutant gliomas. Nature 529, 110–114 (2016).
Hnisz, D. et al. Activation of proto-oncogenes by disruption of chromosome neighborhoods. Science 351, 1454–1458 (2016).
Lupiáñez, D. G. et al. Disruptions of topological chromatin domains cause pathogenic rewiring of gene-enhancer interactions. Cell 161, 1012–1025 (2015).
Rinzema, N. J. et al. Building regulatory landscapes reveals that an enhancer can recruit cohesin to create contact domains, engage CTCF sites and activate distant genes. Nat. Struct. Mol. Biol. 29, 563–574 (2022).
Ing-Simmons, E. et al. Spatial enhancer clustering and regulation of enhancer-proximal genes by cohesin. Genome Res. 25, 504–513 (2015).
Kagey, M. H. et al. Mediator and cohesin connect gene expression and chromatin architecture. Nature 467, 430–435 (2010).
Thiecke, M. J. et al. Cohesin-dependent and -independent mechanisms mediate chromosomal contacts between promoters and enhancers. Cell Rep. 32, 107929 (2020).
Vian, L. et al. The energetics and physiological impact of Cohesin extrusion. Cell 173, 1165–1178 (2018).
Barrington, C. et al. Enhancer accessibility and CTCF occupancy underlie asymmetric TAD architecture and cell type specific genome topology. Nat. Commun. 10, 2908 (2019).
Kane, L. et al. Cohesin is required for long-range enhancer action at the Shh locus. Nat. Struct. Mol. Biol. 29, 891–897 (2022).
Robles-Rebollo, I. et al. Cohesin couples transcriptional bursting probabilities of inducible enhancers and promoters. Nat. Commun. 13, 4342 (2022).
Liu, N. Q. et al. WAPL maintains a cohesin loading cycle to preserve cell-type-specific distal gene regulation. Nat. Genet. 53, 100–109 (2021).
Cuartero, S. et al. Control of inducible gene expression links cohesin to hematopoietic progenitor self-renewal and differentiation. Nat. Immunol. 19, 932–941 (2018).
Calderon, L. et al. Cohesin-dependence of neuronal gene expression relates to chromatin loop length. eLife 11, e76539 (2022).
Rinaldi, L. et al. The glucocorticoid receptor associates with the cohesin loader NIPBL to promote long-range gene regulation. Sci. Adv. 8, 8360 (2022).
Yamada, T. et al. Sensory experience remodels genome architecture in neural circuit to drive motor learning. Nature 569, 708–713 (2019).
Sams, D. S. et al. Neuronal CTCF is necessary for basal and experience-dependent gene regulation, memory formation, and genomic structure of BDNF and arc. Cell Rep. 17, 2418–2430 (2016).
Wang, J., Bando, M., Shirahige, K. & Nakato, R. Large-scale multi-omics analysis suggests specific roles for intragenic cohesin in transcriptional regulation. Nat. Commun. 13, 3218 (2022).
Lupo, R., Breiling, A., Bianchi, M. E. & Orlando, V. Drosophila chromosome condensation proteins Topoisomerase II and Barren colocalize with Polycomb and maintain Fab-7 PRE silencing. Mol. Cell 7, 127–136 (2001).
Zhang, T. et al. Condensin I and II behaviour in interphase nuclei and cells undergoing premature chromosome condensation. Chromosome Res. 24, 243–269 (2016).
Lancaster, L., Patel, H., Kelly, G. & Uhlmann, F. A role for condensin in mediating transcriptional adaptation to environmental stimuli. Life Sci. Alliance 4, 202000961 (2021).
Yuen, K. C., Slaughter, B. D. & Gerton, J. L. Condensin II is anchored by TFIIIC and H3K4me3 in the mammalian genome and supports the expression of active dense gene clusters. Sci. Adv. 3, e1700191 (2017).
Li, W. et al. Condensin I and II complexes license full estrogen receptor α-dependent enhancer activation. Mol. Cell 59, 188–202 (2015).
Hoencamp, C. et al. 3D genomics across the tree of life reveals condensin II as a determinant of architecture type. Science 372, 984–989 (2021). This study shows that condensin II activity in mitosis establishes interphase genome organization, thereby promoting chromosome territories over Rabl-like organization.
Macdonald, L. et al. Rapid and specific degradation of endogenous proteins in mouse models using auxin-inducible degrons. eLife 11, e77987 (2022).
Abdennur, N. et al. Condensin II inactivation in interphase does not affect chromatin folding or gene expression. Preprint at bioRxiv https://doi.org/10.1101/437459 (2018).
Hocquet, C. et al. Condensin controls cellular RNA levels through the accurate segregation of chromosomes instead of directly regulating transcription. eLife 7, e38517 (2018).
Woodward, J. et al. Condensin II mutation causes T-cell lymphoma through tissue-specific genome instability. Genes Dev. 30, 2173–2186 (2016).
Zhang, Y., Zhang, X., Dai, H. Q., Hu, H. & Alt, F. W. The role of chromatin loop extrusion in antibody diversification. Nat. Rev. Immunol. 22, 550–566 (2022).
Guo, C. et al. CTCF-binding elements mediate control of V(D)J recombination. Nature 477, 424–431 (2011).
Dai, H. Q. et al. Loop extrusion mediates physiological Igh locus contraction for RAG scanning. Nature 590, 338–343 (2021).
Ba, Z. et al. CTCF orchestrates long-range cohesin-driven V(D)J recombinational scanning. Nature 586, 305–310 (2020).
Hill, L. et al. Wapl repression by Pax5 promotes V gene recombination by Igh loop extrusion. Nature 584, 142–147 (2020).
Jain, S., Ba, Z., Zhang, Y., Dai, H. Q. & Alt, F. W. CTCF-binding elements mediate accessibility of RAG substrates during chromatin scanning. Cell 174, 102–116 (2018).
Lin, S. G., Guo, C., Sua, A., Zhang, Y. & Alt, F. W. CTCF-binding elements 1 and 2 in the Igh intergenic control region cooperatively regulate V(D)J recombination. Proc. Natl Acad. Sci. USA 112, 1815–1820 (2015).
Medvedovic, J. et al. Flexible long-range loops in the VH gene region of the Igh locus facilitate the generation of a diverse antibody repertoire. Immunity 39, 229–244 (2013).
Zhang, X. et al. Fundamental roles of chromatin loop extrusion in antibody class switching. Nature 575, 385–389 (2019). This study reports that cohesin-mediated loop extrusion is vital for deletional class-switch recombination in B lymphocytes.
Seitan, V. C. et al. A role for cohesin in T-cell-receptor rearrangement and thymocyte differentiation. Nature 476, 467–473 (2011). This study shows that cohesin is important for the rearrangement of the T cell receptor locus and efficient thymocyte differentiation.
Haering, C. H., Farcas, A. M., Arumugam, P., Metson, J. & Nasmyth, K. The cohesin ring concatenates sister DNA molecules. Nature 454, 297–301 (2008). This study shows that cohesin can topologically entrap two sister DNA molecules within its ring-shaped structure.
Murayama, Y., Samora, C. P., Kurokawa, Y., Iwasaki, H. & Uhlmann, F. Establishment of DNA–DNA interactions by the cohesin ring. Cell 172, 465–469 (2018).
Xu, H., Boone, C. & Brown, G. W. Genetic dissection of parallel sister-chromatid cohesion pathways. Genetics 176, 1417–1429 (2007).
Srinivasan, M., Fumasoni, M., Petela, N. J., Murray, A. & Nasmyth, K. A. Cohesion is established during DNA replication utilising chromosome associated cohesin rings as well as those loaded de novo onto nascent DNAs. eLife 9, e56611 (2020). This study describes two pathways for the establishment of sister chromatid cohesion in budding yeast: the conversion pathway and the de novo pathway.
Cameron, G. et al. Sister chromatid cohesion establishment during DNA replication termination. Preprint at biorXiv https://doi.org/10.1101/2022.09.15.508094 (2022).
Srinivasan, M. et al. Scc2 counteracts a Wapl-independent mechanism that releases cohesin from chromosomes during G1. eLife 8, e44736 (2019).
Rhodes, J. D. P. et al. Cohesin can remain associated with chromosomes during DNA replication. Cell Rep. 20, 2749–2755 (2017).
van Schie, J. J. et al. MMS22L-TONSL functions in sister chromatid cohesion in a pathway parallel to DSCC1-RFC. Life Sci. Alliance 6, e2022201598 (2023).
Liu, H. W. et al. Division of labor between PCNA loaders in DNA replication and sister chromatid cohesion establishment. Mol. Cell 78, 725–738 (2020).
Kawasumi, R. et al. Vertebrate CTF18 and DDX11 essential function in cohesion is bypassed by preventing WAPL-mediated cohesin release. Genes Dev. 35, 1368–1382 (2021).
Faramarz, A. et al. Non-redundant roles in sister chromatid cohesion of the DNA helicase DDX11 and the SMC3 acetyl transferases ESCO1 and ESCO2. PLoS ONE 15, e0220348 (2020).
Psakhye, I., Kawasumi, R., Abe, T., Hirota, K. & Branzei, D. PCNA recruits cohesin loader Scc2/NIPBL to ensure sister chromatid cohesion. Preprint at biorXiv https://doi.org/10.1101/2022.09.16.508217 (2022).
Mitter, M. et al. Conformation of sister chromatids in the replicated human genome. Nature 586, 139–144 (2020). This study uses a novel technique, sister chromatid-sensitive Hi-C (scsHi-C) to reveal that sister chromatid cohesion is enriched at CTCF sites.
Collier, J. E. & Nasmyth, K. A. DNA passes through cohesin’s hinge as well as its Smc3–kleisin interface. eLife 11, e80310 (2022).
Nagasaka, K. et al. Cohesin mediates DNA loop extrusion and sister chromatid cohesion by distinct mechanisms. Preprint at bioRxiv https://doi.org/10.1101/2022.09.23.509019 (2022).
Ivanov, D. et al. Eco1 is a novel acetyltransferase that can acetylate proteins involved in cohesion. Curr. Biol. 12, 323–328 (2002).
Ben-Shahar, T. R. et al. Eco1-dependent cohesin acetylation during establishment of sister chromatid cohesion. Science 321, 563–566 (2008).
Unal, E. et al. Eco1-dependent cohesin acetylation during establishment of sister chromatid cohesion. Science 321, 566–569 (2008).
Zhang, J. et al. Acetylation of Smc3 by Eco1 is required for S phase sister chromatid cohesion in both human and yeast. Mol. Cell 31, 143–151 (2008).
Rowland, B. D. et al. Building sister chromatid cohesion: Smc3 acetylation counteracts an antiestablishment activity. Mol. Cell 33, 763–774 (2009).
Minamino, M., Bouchoux, C., Canal, B., Diffley, J. F. X. & Uhlmann, F. A replication fork determinant for the establishment of sister chromatid cohesion. Cell 186, 837–849 (2023).
Gerlich, D., Koch, B., Dupeux, F., Peters, J. M. & Ellenberg, J. Live-cell imaging reveals a stable cohesin–chromatin interaction after but not before DNA replication. Curr. Biol. 16, 1571–1578 (2006).
Sutani, T., Kawaguchi, T., Kanno, R., Itoh, T. & Shirahige, K. Budding yeast Wpl1(Rad61)–Pds5 complex counteracts sister chromatid cohesion-establishing reaction. Curr. Biol. 19, 492–497 (2009).
Feytout, A., Vaur, S., Genier, S., Vazquez, S. & Javerzat, J.-P. Psm3 acetylation on conserved lysine residues is dispensable for viability in fission yeast but contributes to eso1-mediated sister chromatid cohesion by antagonizing Wpl1. Mol. Cell Biol. 31, 1771–1786 (2011).
Rankin, S., Ayad, N. G. & Kirschner, M. W. Sororin, a substrate of the anaphase-promoting complex, is required for sister chromatid cohesion in vertebrates. Mol. Cell 18, 185–200 (2005).
Schmitz, J., Watrin, E., Lénárt, P., Mechtler, K. & Peters, J. M. Sororin is required for stable binding of cohesin to chromatin and for sister chromatid cohesion in interphase. Curr. Biol. 17, 630–636 (2007).
Nishiyama, T. et al. Sororin mediates sister chromatid cohesion by antagonizing Wapl. Cell 143, 737–749 (2010).
Ladurner, R. et al. Sororin actively maintains sister chromatid cohesion. EMBO J. 35, 635–653 (2016).
Emerson, D. J. et al. Cohesin-mediated loop anchors confine the locations of human replication origins. Nature 606, 812–819 (2022).
Dequeker, B. J. H. et al. MCM complexes are barriers that restrict cohesin-mediated loop extrusion. Nature 606, 197–203 (2022).
Jeppsson, K. et al. Cohesin-dependent chromosome loop extrusion is limited by transcription and stalled replication forks. Sci. Adv. 8, eabn7063 (2022).
Sonoda, E. et al. Scc1/Rad21/Mcd1 is required for sister chromatid cohesion and kinetochore function in vertebrate cells. Dev. Cell 1, 759–770 (2001).
Bauerschmidt, C. et al. Cohesin promotes the repair of ionizing radiation-induced DNA double-strand breaks in replicated chromatin. Nucleic Acids Res. 38, 477–487 (2009).
Sjögren, C. & Nasmyth, K. Sister chromatid cohesion is required for postreplicative double-strand break repair in Saccharomyces cerevisiae. Curr. Biol. 11, 991–995 (2001).
Kim, J. S. et al. Specific recruitment of human cohesin to laser-induced DNA damage. J. Biol. Chem. 277, 45149–45153 (2002).
Ström, L., Lindroos, H. B., Shirahige, K. & Sjögren, C. Postreplicative recruitment of cohesin to double-strand breaks is required for DNA repair. Mol. Cell 16, 1003–1015 (2004).
Ünal, E. et al. DNA damage response pathway uses histone modification to assemble a double-strand break-specific cohesin domain. Mol. Cell 16, 991–1002 (2004).
Ström, L. et al. Postreplicative formation of cohesion is required for repair and induced by a single DNA break. Science 317, 242–245 (2007).
Unal, E., Heidinger-Pauli, J. M. & Koshland, D. DNA double-strand breaks trigger genome-wide sister-chromatid cohesion through Eco1 (Ctf7). Science 317, 245–248 (2007). Together with Ström et al. (2007), these papers show that in response to DNA damage, sister chromatin cohesion is established genome-wide in a replication-independent manner.
Covo, S., Westmoreland, J. W., Gordenin, D. A. & Resnick, M. A. Cohesin is limiting for the suppression of DNA damage-induced recombination between homologous chromosomes. PLoS Genet. 6, e1001006 (2010).
Piazza, A. et al. Cohesin regulates homology search during recombinational DNA repair. Nat. Cell Biol. 23, 1176–1186 (2021).
Uhlmann, F. & Nasmyth, K. Cohesion between sister chromatids must be established during DNA replication. Curr. Biol. 8, 1095–1102 (1998).
Caron, P. et al. Cohesin protects genes against γH2AX induced by DNA double-strand breaks. PLoS Genet. 8, e1002460 (2012).
Potts, P. R., Porteus, M. H. & Yu, H. Human SMC5/6 complex promotes sister chromatid homologous recombination by recruiting the SMC1/3 cohesin complex to double-strand breaks. EMBO J. 25, 3377–3388 (2006).
Bot, C. et al. Independent mechanisms recruit the cohesin loader protein NIPBL to sites of DNA damage. J. Cell Sci. 130, 1134–1146 (2017).
Gelot, C. et al. The cohesin complex prevents the end joining of distant DNA double-strand ends. Mol. Cell 61, 15–26 (2016).
Arnould, C. et al. Loop extrusion as a mechanism for formation of DNA damage repair foci. Nature 590, 660–665 (2021).
Kong, X. et al. Distinct functions of human cohesin-SA1 and cohesin-SA2 in double-strand break repair. Mol. Cell Biol. 34, 685–698 (2014).
Verkade, H. M., Teli, T., Laursen, L. V., Murray, J. M. & O’Connell, M. J. A homologue of the Rad18 postreplication repair gene is required for DNA damage responses throughout the fission yeast cell cycle. Mol. Genet. Genomics 265, 993–1003 (2001).
Lehmann, A. R. et al. The rad18 gene of Schizosaccharomyces pombe defines a new subgroup of the SMC superfamily involved in DNA repair. Mol. Cell Biol. 15, 7067–7080 (1995).
Fujioka, Y., Kimata, Y., Nomaguchi, K., Watanabe, K. & Kohno, K. Identification of a novel non-structural maintenance of chromosomes (SMC) component of the SMC5–SMC6 complex involved in DNA repair. J. Biol. Chem. 277, 21585–21591 (2002).
Betts Lindroos, H. et al. Chromosomal association of the Smc5/6 complex reveals that it functions in differently regulated pathways. Mol. Cell 22, 755–767 (2006).
De Piccoli, G. et al. Smc5–Smc6 mediate DNA double-strand-break repair by promoting sister-chromatid recombination. Nat. Cell Biol. 8, 1032–1034 (2006).
Wu, N. et al. Scc1 sumoylation by Mms21 promotes sister chromatid recombination through counteracting Wapl. Genes Dev. 26, 1473–1485 (2012).
Bermúdez-López, M. et al. Sgs1’s roles in DNA end resection, HJ dissolution, and crossover suppression require a two-step SUMO regulation dependent on Smc5/6. Genes Dev. 30, 1339–1356 (2016).
Agashe, S. et al. Smc5/6 functions with Sgs1–Top3–Rmi1 to complete chromosome replication at natural pause sites. Nat. Commun. 12, 2111 (2021).
Bonner, J. N. et al. Smc5/6 mediated sumoylation of the Sgs1–Top3–Rmi1 complex promotes removal of recombination intermediates. Cell Rep. 16, 368–378 (2016).
Chavez, A., George, V., Agrawal, V. & Johnson, F. B. Sumoylation and the structural maintenance of chromosomes (Smc) 5/6 complex slow senescence through recombination intermediate resolution. J. Biol. Chem. 285, 11922–11930 (2010).
Bermúdez-López, M. et al. The Smc5/6 complex is required for dissolution of DNA-mediated sister chromatid linkages. Nucleic Acids Res. 38, 6502–6512 (2010).
Menolfi, D., Delamarre, A., Lengronne, A., Pasero, P. & Branzei, D. Essential roles of the Smc5/6 complex in replication through natural pausing sites and endogenous DNA damage tolerance. Mol. Cell 60, 835–846 (2015).
Chang, J. T. H. et al. Smc5/6’s multifaceted DNA binding capacities stabilize branched DNA structures. Nat. Commun. 13, 7179 (2022).
Tanasie, N. L., Gutiérrez-Escribano, P., Jaklin, S., Aragon, L. & Stigler, J. Stabilization of DNA fork junctions by Smc5/6 complexes revealed by single-molecule imaging. Cell Rep. 41, 111778 (2022).
Torres-Rosell, J. et al. SMC5 and SMC6 genes are required for the segregation of repetitive chromosome regions. Nat. Cell Biol. 7, 412–419 (2005).
Yong-Gonzales, V., Hang, L. E., Castellucci, F., Branzei, D. & Zhao, X. The Smc5–Smc6 complex regulates recombination at centromeric regions and affects kinetochore protein sumoylation during normal growth. PLoS ONE 7, e51540 (2012).
Venegas, A. B., Natsume, T., Kanemaki, M. & Hickson, I. D. Inducible degradation of the human SMC5/6 complex reveals an essential role only during interphase. Cell Rep. 31, 107533 (2020).
Rossi, F. et al. SMC5/6 acts jointly with Fanconi anemia factors to support DNA repair and genome stability. EMBO Rep. 21, e48222 (2020).
Grange, L. J. et al. Pathogenic variants in SLF2 and SMC5 cause segmented chromosomes and mosaic variegated hyperploidy. Nat. Commun. 13, 6664 (2022).
Wood, J. L., Singh, N., Mer, G. & Chen, J. MCPH1 functions in an H2AX-dependent but MDC1-independent pathway in response to DNA damage. J. Biol. Chem. 282, 35416–35423 (2007).
Kong, X. et al. Condensin I recruitment to base damage-enriched DNA lesions is modulated by PARP1. PLoS ONE 6, e23548 (2011).
Heale, J. T. et al. Condensin I interacts with the PARP-1–XRCC1 complex and functions in DNA single-strand break repair. Mol. Cell 21, 837–848 (2006).
Boteva, L. et al. Common fragile sites are characterized by faulty condensin loading after replication stress. Cell Rep. 32, 108177 (2020).
Ono, T., Yamashita, D. & Hirano, T. Condensin II initiates sister chromatid resolution during S phase. J. Cell Biol. 200, 429–441 (2013).
Nagasaka, K., Hossain, M. J., Roberti, M. J., Ellenberg, J. & Hirota, T. Sister chromatid resolution is an intrinsic part of chromosome organization in prophase. Nat. Cell Biol. 18, 692–699 (2016).
Batty, P. et al. Cohesin-mediated DNA loop extrusion resolves sister chromatids in G2 phase. Preprint at bioRxiv https://doi.org/10.1101/2023.01.12.523718 (2023).
D’Ambrosio, C. et al. Identification of cis-acting sites for condensin loading onto budding yeast chromosomes. Genes Dev. 22, 2215–2227 (2008).
Charbin, A., Bouchoux, C. & Uhlmann, F. Condensin aids sister chromatid decatenation by topoisomerase II. Nucleic Acids Res. 42, 340–348 (2014).
Dyson, S., Segura, J., Martínez‐García, B., Valdés, A. & Roca, J. Condensin minimizes topoisomerase II mediated entanglements of DNA in vivo. EMBO J. 40, e105393 (2021).
Baxter, J. et al. Positive supercoiling of mitotic DNA drives decatenation by topoisomerase II in eukaryotes. Sci. Rep. 331, 1328–1332 (2011).
Sen, N. et al. Physical proximity of sister chromatids promotes Top2-dependent intertwining. Mol. Cell 64, 134–147 (2016).
Piskadlo, E., Tavares, A. & Oliveira, R. A. Metaphase chromosome structure is dynamically maintained by condensin I-directed DNA (de)catenation. eLife 6, e26120 (2017).
Coelho, P. A., Queiroz-Machado, J. & Sunkel, C. E. Condensin-dependent localisation of topoisomerase II to an axial chromosomal structure is required for sister chromatid resolution during mitosis. J. Cell Sci. 116, 4763–4776 (2003).
Ono, T., Sakamoto, C., Nakao, M., Saitoh, N. & Hirano, T. Condensin II plays an essential role in reversible assembly of mitotic chromosomes in situ. Mol. Biol. Cell 28, 2875–2886 (2017).
Hudson, D. F., Vagnarelli, P., Gassmann, R. & Earnshaw, W. C. Condensin is required for nonhistone protein assembly and structural integrity of vertebrate mitotic chromosomes. Dev. Cell 5, 323–336 (2003).
Deiss, K. et al. A genome-wide RNAi screen identifies the SMC5/6 complex as a non-redundant regulator of a Topo2a-dependent G2 arrest. Nucleic Acids Res. 47, 2906–2921 (2019).
Verver, D. E. et al. Non-SMC element 2 (NSMCE2) of the SMC5/6 complex helps to resolve topological stress. Int. J. Mol. Sci. 17, 1782 (2016).
Shintomi, K. & Hirano, T. Guiding functions of the C-terminal domain of topoisomerase IIα advance mitotic chromosome assembly. Nat. Commun. 12, 2917 (2021).
Hildebrand, E. M. et al. Chromosome decompaction and cohesin direct topoisomerase II activity to establish and maintain an unentangled interphase genome. Preprint at biorXiv https://doi.org/10.1101/2022.10.15.511838 (2022).
Ono, T. et al. Differential contributions of condensin I and condensin II to mitotic chromosome architecture in vertebrate cells. Cell 115, 109–121 (2003).
Green, L. C. et al. Contrasting roles of condensin I and condensin II in mitotic chromosome formation. J. Cell Sci. 125, 1591–1604 (2012).
Shintomi, K. & Hirano, T. The relative ratio of condensing I to II determines chromosome shapes. Genes Dev. 25, 1464–1469 (2011).
Shintomi, K. et al. Mitotic chromosome assembly despite nucleosome depletion in Xenopus egg extracts. Science 356, 1284–1287 (2017).
Samejima, K. et al. Functional analysis after rapid degradation of condensins and 3D-EM reveals chromatin volume is uncoupled from chromosome architecture in mitosis. J. Cell Sci. 131, jcs210187 (2018).
Houlard, M. et al. Condensin confers the longitudinal rigidity of chromosomes. Nat. Cell Biol. 17, 771–781 (2015).
Gerlich, D., Hirota, T., Koch, B., Peters, J. M. & Ellenberg, J. Condensin I stabilizes chromosomes mechanically through a dynamic interaction in live cells. Curr. Biol. 16, 333–344 (2006).
Gibcus, J. H. et al. A pathway for mitotic chromosome formation. Science 359, eaao6135 (2018). By combining Hi-C, imaging and polymer modelling, together with Walther et al. (2018), these studies show that the mitotic chromosome is formed by condensin II-dependent formation of large chromatin loops that are subdivided by condensin I into smaller, nested loops.
Houlard, M. et al. MCPH1 inhibits condensin II during interphase by regulating its SMC2–kleisin interface. eLife 10, e73348 (2021). This study establishes microcephalin (MCPH1) as a condensin II-removal factor, which functions by opening up the condensin II SMC2–kleisin interface.
Hirota, T., Gerlich, D., Koch, B., Ellenberg, J. & Peters, J. M. Distinct functions of condensin I and II in mitotic chromosome assembly. J. Cell Sci. 117, 6435–6445 (2004). Together with Ono et al. (2003), these papers discovered condensin II and described distinct functions for the two condensin complexes.
Ono, T., Fang, Y., Spector, D. L. & Hirano, T. Spatial and temporal regulation of condensins I and II in mitotic chromosome assembly in human cells. Mol. Biol. Cell 15, 3296–3308 (2004).
Walther, N. et al. A quantitative map of human condensins provides new insights into mitotic chromosome architecture. J. Cell Biol. 217, 2309–2328 (2018). This study performs quantitative imaging of condensin complexes on the DNA of mitotic chromosomes, which supports the model that condensin II forms an axis of large chromatin loops that are subdivided by condensin I into smaller loops.
Oliveira, R. A., Coelho, P. A. & Sunkel, C. E. The condensin I subunit barren/CAP-H is essential for the structural integrity of centromeric heterochromatin during mitosis. Mol. Cell Biol. 25, 8971–8984 (2005).
Sacristan, C. et al. Condensin reorganizes centromeric chromatin during mitotic entry into a bipartite structure stabilized by cohesin. Preprint at bioRxiv https://doi.org/10.1101/2022.08.01.502248 (2022).
Waizenegger, I. C., Hauf, S., Meinke, A. & Peters, J. M. Two distinct pathways remove mammalian cohesin from chromosome arms in prophase and from centromeres in anaphase. Cell 103, 399–410 (2000). This study shows that in mammalian cells, cohesin is removed from chromatin in two distinct waves, with only the second being dependent on separase-mediated cleavage of cohesin.
Zhang, N., Panigrahi, A. K., Mao, Q. & Pati, D. Interaction of sororin protein with polo-like kinase 1 mediates resolution of chromosomal arm cohesion. J. Biol. Chem. 286, 41826–41837 (2011).
Sumara, I. et al. The dissociation of cohesin from chromosomes in prophase is regulated by polo-like kinase. Mol. Cell 9, 515–525 (2002).
Nishiyama, T., Sykora, M. M., Huis, P. J., Mechtler, K. & Peters, J. M. Aurora B and Cdk1 mediate Wapl activation and release of acetylated cohesin from chromosomes by phosphorylating sororin. Proc. Natl Acad. Sci. USA 110, 13404–13409 (2013).
Losada, A., Hirano, M. & Hirano, T. Cohesin release is required for sister chromatid resolution, but not for condensin-mediated compaction, at the onset of mitosis. Genes Dev. 16, 3004–3016 (2002).
Liu, H., Rankin, S. & Yu, H. Phosphorylation-enabled binding of SGO1–PP2A to cohesin protects sororin and centromeric cohesion during mitosis. Nat. Cell Biol. 15, 40–49 (2012).
Hauf, S. et al. Dissociation of cohesin from chromosome arms and loss of arm cohesion during early mitosis depends on phosphorylation of SA2. PLoS Biol. 3, e69 (2005).
Giménez-Abián, J. F. et al. Regulation of sister chromatid cohesion between chromosome arms. Curr. Biol. 14, 1187–1193 (2004).
Dreier, M. R., Bekier, M. E. & Taylor, W. R. Regulation of sororin by Cdk1-mediated phosphorylation. J. Cell Sci. 124, 2976–2987 (2011).
Haarhuis, J. H. I. et al. WAPL-mediated removal of cohesin protects against segregation errors and aneuploidy. Curr. Biol. 23, 2071–2077 (2013).
Chu, L., Zhang, Z., Mukhina, M., Zickler, D. & Kleckner, N. Sister chromatids separate during anaphase in a three-stage program as directed by interaxis bridges. Proc. Natl Acad. Sci. USA 119, e2123363119 (2022).
Tedeschi, A. et al. Wapl is an essential regulator of chromatin structure and chromosome segregation. Nature 501, 564–568 (2013). This study reveals that stabilization of cohesin on DNA in interphase leads to cohesin accumulation in an axial configuration and condensation of interphase chromosomes.
Perea-Resa, C., Bury, L., Cheeseman, I. M. & Blower, M. D. Cohesin removal reprograms gene expression upon mitotic entry. Mol. Cell 78, 127–140 (2020).
Salic, A., Waters, J. C. & Mitchison, T. J. Vertebrate shugoshin links sister centromere cohesion and kinetochore microtubule stability in mitosis. Cell 118, 567–578 (2004).
Tang, Z., Sun, Y., Harley, S. E., Zou, H. & Yu, H. Human Bub1 protects centromeric sister-chromatid cohesion through Shugoshin during mitosis. Proc. Natl Acad. Sci. USA 101, 18012–18017 (2004).
McGuinness, B. E., Hirota, T., Kudo, N. R., Peters, J. M. & Nasmyth, K. Shugoshin prevents dissociation of cohesin from centromeres during mitosis in vertebrate cells. PLoS Biol. 3, 0433–0449 (2005).
Riedel, C. G. et al. Protein phosphatase 2A protects centromeric sister chromatid cohesion during meiosis I. Nature 441, 53–61 (2006).
Kitajima, T. S. et al. Shugoshin collaborates with protein phosphatase 2A to protect cohesin. Nature 441, 46–52 (2006).
Hara, K. et al. Structure of cohesin subcomplex pinpoints direct shugoshin-Wapl antagonism in centromeric cohesion. Nat. Struct. Mol. Biol. 21, 864–870 (2014).
García-Nieto, A. et al. Structural basis of centromeric cohesion protection. Nat. Struct. Mol. Biol. https://doi.org/10.1038/s41594-023-00968-y (2023).
Peters, J. M. The anaphase promoting complex/cyclosome: a machine designed to destroy. Nat. Rev. Mol. Cell Biol. 7, 644–656 (2006).
Vázquez-Novelle, M. D. et al. Cdk1 inactivation terminates mitotic checkpoint surveillance and stabilizes kinetochore attachments in anaphase. Curr. Biol. 24, 638–645 (2014).
Oliveira, R. A., Hamilton, R. S., Pauli, A., Davis, I. & Nasmyth, K. Cohesin cleavage and Cdk inhibition trigger formation of daughter nuclei. Nat. Cell Biol. 12, 185–192 (2010).
Yu, J. et al. Structural basis of human separase regulation by securin and CDK1–cyclin B1. Nature 596, 138–142 (2021).
Hellmuth, S., Gómez-H, L., Pendás, A. M. & Stemmann, O. Securin-independent regulation of separase by checkpoint-induced shugoshin–MAD2. Nature 580, 536–541 (2020).
Uhlmann, F., Lottspelch, F. & Nasmyth, K. Sister-chromatid separation at anaphase onset is promoted by cleavage of the cohesin subunit Scc1. Nature 400, 37–42 (1999). This study reveals that separase-mediated cleavage of the Scc1 subunit of cohesin is required for anaphase onset.
Uhlmann, F., Wernic, D., Poupart, M. A., Koonin, E. V. & Nasmyth, K. Cleavage of cohesin by the CD clan protease separin triggers anaphase in yeast. Cell 103, 375–386 (2000).
Boveri, T. Die Blastomerenkerne von Ascaris megalocephala und die Theorie der Chromosomenindividualitat. Arch. Zellforsch. 3, 181–268 (1909).
Joyce, E. F., Williams, B. R., Xie, T. & Wu, C. T. Identification of genes that promote or antagonize somatic homolog pairing using a high-throughput FISH-based screen. PLoS Genet. 8, e1002667 (2012).
Bauer, C. R., Hartl, T. A. & Bosco, G. Condensin II promotes the formation of chromosome territories by inducing axial compaction of polyploid interphase chromosomes. PLoS Genet. 8, e1002873 (2012). This study shows that, in Drosophila melanogaster, condensin II promotes the formation of chromosome territories and regulates the spatial organization of heterochromatin domains.
Hartl, T. A., Smith, H. F. & Bosco, G. Chromosome alignment and transvection are antagonized by condensin II. Science 322, 1384–1387 (2008). This study shows that in D. melanogaster, condensin II negatively regulates transvection, possibly by restricting chromosomal interactions in trans.
Buster, D. W. et al. SCF(Slimb) ubiquitin ligase suppresses condensin II-mediated nuclear reorganization by degrading Cap-H2. J. Cell Biol. 201, 49–63 (2013).
Vernizzi, L. & Lehner, C. F. Bivalent individualization during chromosome territory formation in Drosophila spermatocytes by controlled condensin II protein activity and additional force generators. PLoS Genet. 17, e1009870 (2021).
Nguyen, H. Q. et al. Drosophila casein kinase I alpha regulates homolog pairing and genome organization by modulating condensin II subunit Cap-H2 levels. PLoS Genet. 11, e1005014 (2015).
Rosin, L. F., Nguyen, S. C., Joyce, E. F. & Bosco, G. Condensin II drives large-scale folding and spatial partitioning of interphase chromosomes in Drosophila nuclei. PLoS Genet. 14, 1007393 (2018).
Iwasaki, O., Corcoran, C. J. & Noma, K. I. Involvement of condensin-directed gene associations in the organization and regulation of chromosome territories during the cell cycle. Nucleic Acids Res. 44, 3618–3628 (2016).
Howard-Till, R. & Loidl, J. Condensins promote chromosome individualization and segregation during mitosis, meiosis, and amitosis in Tetrahymena thermophila. Mol. Biol. Cell 29, 466–478 (2018).
Municio, C. et al. The Arabidopsis condensin CAP-D subunits arrange interphase chromatin. N. Phytol. 230, 972–987 (2021).
Schubert, V., Lermontova, I. & Schubert, I. The Arabidopsis CAP-D proteins are required for correct chromatin organisation, growth and fertility. Chromosoma 122, 517–533 (2013).
Sakamoto, T., Sugiyama, T., Yamashita, T. & Matsunaga, S. Plant condensin II is required for the correct spatial relationship between centromeres and rDNA arrays. Nucleus 10, 116–125 (2019).
Sakamoto, T. et al. Two-step regulation of centromere distribution by condensin II and the nuclear envelope proteins. Res. Sq. https://doi.org/10.21203/rs.3.rs-793150/v1 (2021).
Nishide, K. & Hirano, T. Overlapping and non-overlapping functions of condensins I and II in neural stem cell divisions. PLoS Genet. 10, e1004847 (2014).
King, T. D. et al. Recurrent losses and rapid evolution of the condensin II complex in insects. Mol. Biol. Evol. 36, 2195–2204 (2019).
Hirano, T. Chromosome territories meet a condensin. PLoS Genet. 8, e1002939 (2012).
Lieberman-Aiden, E. et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326, 289–293 (2009).
Falk, M. et al. Heterochromatin drives compartmentalization of inverted and conventional nuclei. Nature 570, 395–399 (2019).
Biswas, S. et al. HP1 oligomerization compensates for low-affinity H3K9me recognition and provides a tunable mechanism for heterochromatin-specific localization. Sci. Adv. 8, eabk0793 (2022).
Canzio, D. et al. Chromodomain-mediated oligomerization of HP1 suggests a nucleosome-bridging mechanism for heterochromatin assembly. Mol. Cell 41, 67–81 (2011).
Erdel, F. et al. Mouse heterochromatin adopts digital compaction states without showing hallmarks of HP1-driven liquid–liquid phase separation. Mol. Cell 78, 236–249 (2020).
Gitler, A. D., Shorter, J., Ha, T. & Myong, S. Just took a DNA test, turns out 100% not that phase. Mol. Cell 78, 193–194 (2020).
Sanulli, S. et al. HP1 reshapes nucleosome core to promote phase separation of heterochromatin. Nature 575, 390–394 (2019).
Zenk, F. et al. HP1 drives de novo 3D genome reorganization in early Drosophila embryos. Nature 593, 289–293 (2021).
Plys, A. J. et al. Phase separation of polycomb-repressive complex 1 is governed by a charged disordered region of CBX2. Genes Dev. 33, 799–813 (2019).
Jagannathan, M., Cummings, R. & Yamashita, Y. M. The modular mechanism of chromocenter formation in Drosophila. eLife 8, e243938 (2019).
Chardon, F. et al. CENP-B-mediated DNA loops regulate activity and stability of human centromeres. Mol. Cell 82, 1751–1767.e8 (2022).
Gibson, B. A. et al. Organization of chromatin by intrinsic and regulated phase separation. Cell 179, 470–484 (2019).
Guo, Y. E. et al. Pol II phosphorylation regulates a switch between transcriptional and splicing condensates. Nature 572, 543–548 (2019).
Nuebler, J., Fudenberg, G., Imakaev, M., Abdennur, N. & Mirny, L. A. Chromatin organization by an interplay of loop extrusion and compartmental segregation. Proc. Natl Acad. Sci. USA 115, 6697–6706 (2018).
Haarhuis, J. H. I. et al. A mediator-cohesin axis controls heterochromatin domain formation. Nat. Commun. 13, 754 (2022).
Alipour, E. & Marko, J. F. Self-organization of domain structures by DNA-loop-extruding enzymes. Nucleic Acids Res. 40, 11202–11212 (2012).
Fudenberg, G. et al. Formation of chromosomal domains by loop extrusion. Cell Rep. 15, 2038–2049 (2016).
Sanborn, A. L. et al. Chromatin extrusion explains key features of loop and domain formation in wild-type and engineered genomes. Proc. Natl Acad. Sci. USA 112, E6456–E6465 (2015).
Goloborodko, A., Imakaev, M. V., Marko, J. F. & Mirny, L. Compaction and segregation of sister chromatids via active loop extrusion. eLife 5, e14864 (2016).
Wang, X., Brandão, H. B., Le, T. B. K., Laub, M. T. & Rudner, D. Z. Bacillus subtilis SMC complexes juxtapose chromosome arms as they travel from origin to terminus. Science 355, 524–527 (2017).
Ganji, M. et al. Real-time imaging of DNA loop extrusion by condensin. Science 360, 102–105 (2018). The first proof that SMC complexes, in this case budding yeast condensin, can perform loop extrusion in vitro.
Pradhan, B. et al. The Smc5/6 complex is a DNA loop extruding motor. Nature https://doi.org/10.1038/s41586-023-05963-3 (2023).
Kong, M. et al. Human condensin I and II drive extensive ATP-dependent compaction of nucleosome-bound DNA. Mol. Cell 79, 99–114.e9 (2020).
Guo, Y. et al. Chromatin jets define the properties of cohesin-driven in vivo loop extrusion. Mol. Cell 82, 3769–3780 (2022).
Naumova, N. et al. Organization of the mitotic chromosome. Science 342, 948–953 (2013).
Irwan, I. D., Bogerd, H. P. & Cullen, B. R. Epigenetic silencing by the SMC5/6 complex mediates HIV-1 latency. Nat. Microbiol. 7, 2101–2113 (2022).
Dupont, L. et al. The SMC5/6 complex compacts and silences unintegrated HIV-1 DNA and is antagonized by Vpr. Cell Host Microbe 29, 792–805 (2021).
Decorsière, A. et al. Hepatitis B virus X protein identifies the Smc5/6 complex as a host restriction factor. Nature 531, 386–389 (2016). This study identifies the SMC5–SMC6 complex as a restriction factor in hepatitis B virus infection that can be degraded with the aid of the viral protein HBx.
Gibson, R. T. & Androphy, E. J. The SMC5/6 complex represses the replicative program of high-risk human papillomavirus type 31. Pathogens 9, 786 (2020).
Bentley, P., Tan, M. J. A., McBride, A. A., White, E. A. & Howley, P. M. The SMC5/6 complex interacts with the papillomavirus E2 protein and influences maintenance of viral episomal DNA. J. Virol. 92, e00356-18 (2018).
Xu, W. et al. PJA1 coordinates with the SMC5/6 complex to restrict DNA viruses and episomal genes in an interferon-independent manner. J. Virol. 92, 825–843 (2018).
Yiu, S. P. T., Guo, R., Zerbe, C., Weekes, M. P. & Gewurz, B. E. Epstein–Barr virus BNRF1 destabilizes SMC5/6 cohesin complexes to evade its restriction of replication compartments. Cell Rep. 10, 110411 (2022).
Han, C. et al. KSHV RTA antagonizes SMC5/6 complex-induced viral chromatin compaction by hijacking the ubiquitin–proteasome system. PLoS Pathog. 18, e1010744 (2022).
Abdul, F. et al. Smc5/6 silences episomal transcription by a three-step function. Nat. Struct. Mol. Biol. 29, 922–931 (2022).
Murphy, C. M. et al. Hepatitis B virus X protein promotes degradation of SMC5/6 to enhance HBV replication. Cell Rep. 16, 2846–2854 (2016).
Abdul, F. et al. Smc5/6 antagonism by HBx is an evolutionarily conserved function of hepatitis B virus infection in mammals. J. Virol. 92, e00769-18 (2018).
Deep, A. et al. The condensin-like Wadjet complex protects bacteria from plasmid transformation by recognition and cleavage of closed-circular DNA. Mol. Cell 82, 4145–4159 (2022). This study shows that the bacterial SMC complex Wadjet has a role in antiplasmid defence by cleaving circular plasmid DNA, for which it requires its topoisomerase-like subunit.
Doron, S. et al. Systematic discovery of antiphage defense systems in the microbial pangenome. Science 359, eaar4120 (2018).
Liu, H. W. et al. DNA-measuring Wadjet SMC ATPases restrict smaller circular plasmids by DNA cleavage. Mol. Cell 82, 4727–4740 (2022).
Rabl, C. Über Zelltheilung. Morphol. Jahrb. 10, 214–330 (1885).
Longo, G. M. C. & Roukos, V. Territories or spaghetti? Chromosome organization exposed. Nat. Rev. Mol. Cell Biol. 22, 508–508 (2021).
Owing to space limitations, the authors apologize for not being able to cite all relevant articles or discuss all aspects of SMC biology. The authors acknowledge financial support from the European Research Council (772471-CohesinLooping) and from the Dutch Research Council (VI.C.202.098) and thank A. Friskes and members of the Rowland Laboratory for critical reading of the manuscript.
The authors declare no competing interests.
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A molecular linkage that naturally arises in processes such as DNA replication, in which two DNA molecules are knotted in such a way that can only be resolved by the breaking of the DNA chain.
An accumulation of different heterochromatin regions into a dense structure within the nuclei of some types of animal cells.
A chromatin-organizing protein that binds to a specific DNA motif and acts as a boundary for extruding cohesin complexes.
A chromatin conformation capture technique, in which regions that are close together in the nucleus are crosslinked and identified by sequencing, which provides genome-wide information about the 3D nuclear architecture of a population of cells.
- Malformation syndromes
A disorder characterized by congenital anomalies that are causally related.
- Promyelocytic leukaemia bodies
Nuclear membrane-less organelles that accumulate certain proteins and that have been implicated in processes ranging from DNA damage repair to antiviral defence.
- Topologically associating domains
(TADs). Self-interacting genomic regions as observed by chromatin capture techniques that are flanked by CTCF binding sites and consist of chromatin loops formed by cohesin.
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Hoencamp, C., Rowland, B.D. Genome control by SMC complexes. Nat Rev Mol Cell Biol 24, 633–650 (2023). https://doi.org/10.1038/s41580-023-00609-8
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