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  • Review Article
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Nuclear compartmentalization as a mechanism of quantitative control of gene expression

Abstract

Gene regulation requires the dynamic coordination of hundreds of regulatory factors at precise genomic and RNA targets. Although many regulatory factors have specific affinity for their nucleic acid targets, molecular diffusion and affinity models alone cannot explain many of the quantitative features of gene regulation in the nucleus. One emerging explanation for these quantitative properties is that DNA, RNA and proteins organize within precise, 3D compartments in the nucleus to concentrate groups of functionally related molecules. Recently, nucleic acids and proteins involved in many important nuclear processes have been shown to engage in cooperative interactions, which lead to the formation of condensates that partition the nucleus. In this Review, we discuss an emerging perspective of gene regulation, which moves away from classic models of stoichiometric interactions towards an understanding of how spatial compartmentalization can lead to non-stoichiometric molecular interactions and non-linear regulatory behaviours. We describe key mechanisms of nuclear compartment formation, including emerging roles for non-coding RNAs in facilitating their formation, and discuss the functional role of nuclear compartments in transcription regulation, co-transcriptional and post-transcriptional RNA processing, and higher-order chromatin regulation. More generally, we discuss how compartmentalization may explain important quantitative aspects of gene regulation.

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Fig. 1: Mechanisms of nuclear compartments formation.
Fig. 2: Spatially constrained non-coding RNAs can drive compartmentalization in the nucleus.
Fig. 3: Enhancers, promoters and transcription factors can form condensates that may facilitate rapid target search in the genome.
Fig. 4: Compartmentalization and chromatin regulation.
Fig. 5: Spatial and kinetic coupling of RNA polymerase II transcription and mRNA splicing.
Fig. 6: Involvement of nuclear bodies in RNA processing.

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References

  1. Mirny, L. et al. How a protein searches for its site on DNA: the mechanism of facilitated diffusion. J. Phys. A Math. Theor. 42, 434013 (2009).

    Article  CAS  Google Scholar 

  2. Jana, T., Brodsky, S. & Barkai, N. Speed–specificity trade-offs in the transcription factors search for their genomic binding sites. Trends Genet. 37, 421–432 (2021).

    Article  CAS  PubMed  Google Scholar 

  3. Strom, A. R. & Brangwynne, C. P. The liquid nucleome—phase transitions in the nucleus at a glance. J. Cell Sci. 132, jcs235093 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. Pombo, A. & Dillon, N. Three-dimensional genome architecture: players and mechanisms. Nat. Rev. Mol. Cell Biol. 16, 245–257 (2015).

    Article  CAS  PubMed  Google Scholar 

  5. Bonev, B. & Cavalli, G. Organization and function of the 3D genome. Nat. Rev. Genet. 17, 772–772 (2016).

    Article  CAS  PubMed  Google Scholar 

  6. Gibcus, J. H. & Dekker, J. The hierarchy of the 3D genome. Mol. Cell 49, 773–782 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  7. Schoenfelder, S. & Fraser, P. Long-range enhancer–promoter contacts in gene expression control. Nat. Rev. Genet. 20, 437–455 (2019).

    Article  CAS  PubMed  Google Scholar 

  8. Nora, E. P. et al. Spatial partitioning of the regulatory landscape of the X-inactivation centre. Nature 485, 381–385 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  9. Dundr, M. & Misteli, T. Biogenesis of nuclear bodies. Cold Spring Harb. Perspect. Biol. 2, a000711 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  10. Quinodoz, S. A. et al. Higher-order inter-chromosomal hubs shape 3D genome organization in the nucleus. Cell 174, 744–757.e24 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  11. Cho, W. K. et al. Mediator and RNA polymerase II clusters associate in transcription-dependent condensates. Science 361, 412–415 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. Boija, A. et al. Transcription factors activate genes through the phase-separation capacity of their activation domains. Cell 175, 1842–1855.e16 (2018).

    Article  CAS  PubMed  Google Scholar 

  13. Strom, A. R. et al. Phase separation drives heterochromatin domain formation. Nature 547, 241–245 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Larson, A. G. et al. Liquid droplet formation by HP1α suggests a role for phase separation in heterochromatin. Nature 547, 236–240 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  15. Huo, X. et al. The nuclear matrix protein SAFB cooperates with major satellite RNAs to stabilize heterochromatin architecture partially through phase separation. Mol. Cell 77, 368–383.e7 (2020).

    Article  CAS  PubMed  Google Scholar 

  16. Murray, D. T. et al. Structure of FUS protein fibrils and its relevance to self-assembly and phase separation of low-complexity domains. Cell 171, 615–627.e16 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  17. Sabari, B. R., Dall’Agnese, A. & Young, R. A. Biomolecular condensates in the nucleus. Trends Biochem. Sci. 45, 961–977 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  18. Hyman, A. A., Weber, C. A. & Jülicher, F. Liquid–liquid phase separation in biology. Annu. Rev. Cell Dev. Biol. 30, 39–58 (2014).

    Article  CAS  PubMed  Google Scholar 

  19. Banani, S. F., Lee, H. O., Hyman, A. A. & Rosen, M. K. Biomolecular condensates: organizers of cellular biochemistry. Nat. Rev. Mol. Cell Biol. 18, 285–298 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Lyon, A. S., Peeples, W. B. & Rosen, M. K. A framework for understanding the functions of biomolecular condensates across scales. Nat. Rev. Mol. Cell Biol. 22, 215–235 (2021).

    Article  CAS  PubMed  Google Scholar 

  21. Zhang, Z. et al. Rapid dynamics of general transcription factor TFIIB binding during preinitiation complex assembly revealed by single-molecule analysis. Genes Dev. 30, 2106–2118 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  22. Struhl, K. Helix-turn-helix, zinc-finger, and leucine-zipper motifs for eukaryotic transcriptional regulatory proteins. Trends Biochem. Sci. 14, 137–140 (1989).

    Article  CAS  PubMed  Google Scholar 

  23. Maris, C., Dominguez, C. & Allain, F. H. T. The RNA recognition motif, a plastic RNA-binding platform to regulate post-transcriptional gene expression. FEBS J. 272, 2118–2131 (2005).

    Article  CAS  PubMed  Google Scholar 

  24. García-Mayoral, M. F. et al. The structure of the C-terminal KH domains of KSRP reveals a noncanonical motif important for mRNA degradation. Structure 15, 485–498 (2007).

    Article  PubMed  CAS  Google Scholar 

  25. Stevens, S. W. et al. Biochemical and genetic analyses of the U5, U6, and U4/U6•U5 small nuclear ribonucleoproteins from Saccharomyces cerevisiae. RNA 7, 1543–1553 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  26. Derewenda, Z. S. Application of protein engineering to enhance crystallizability and improve crystal properties. Acta Crystallogr. D 66, 604–615 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  27. Derewenda, Z. S. Rational protein crystallization by mutational surface engineering. Structure 12, 529–535 (2004).

    Article  CAS  PubMed  Google Scholar 

  28. Deller, M. C., Kong, L. & Rupp, B. Protein stability: a crystallographer’s perspective. Acta Crystallogr. F 72, 72–95 (2016).

    Article  CAS  Google Scholar 

  29. Boeynaems, S. et al. Protein phase separation: a new phase in cell biology. Trends Cell Biol. 28, 420–435 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  30. Brangwynne, C. P., Tompa, P. & Pappu, R. V. Polymer physics of intracellular phase transitions. Nat. Phys. 11, 899–904 (2015).

    Article  CAS  Google Scholar 

  31. Li, P. et al. Phase transitions in the assembly of multivalent signalling proteins. Nature 483, 336–340 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Kitov, P. I. & Bundle, D. R. On the nature of the multivalency effect:  a thermodynamic model. J. Am. Chem. Soc. 125, 16271–16284 (2003).

    Article  CAS  PubMed  Google Scholar 

  33. McSwiggen, D. T., Mir, M., Darzacq, X. & Tjian, R. Evaluating phase separation in live cells: diagnosis, caveats, and functional consequences. Genes Dev. 33, 1619–1634 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Hnisz, D., Shrinivas, K., Young, R. A., Chakraborty, A. K. & Sharp, P. A. A phase separation model for transcriptional control. Cell 169, 13–23 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  35. Van Der Lee, R. et al. Classification of intrinsically disordered regions and proteins. Chem. Rev. 114, 6589–6631 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  36. Jain, A. & Vale, R. D. RNA phase transitions in repeat expansion disorders. Nature 546, 243–247 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Van Treeck, B. & Parker, R. Emerging roles for intermolecular RNA–RNA interactions in RNP assemblies. Cell 174, 791–802 (2018).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  38. Brangwynne, C. P. et al. Germline P granules are liquid droplets that localize by controlled dissolution/condensation. Science 324, 1729–1732 (2009).

    Article  CAS  PubMed  Google Scholar 

  39. Lafontaine, D. L. J., Riback, J. A., Bascetin, R. & Brangwynne, C. P. The nucleolus as a multiphase liquid condensate. Nat. Rev. Mol. Cell Biol. 22, 165–182 (2021).

    Article  CAS  PubMed  Google Scholar 

  40. Seif, E. et al. Phase separation by the polyhomeotic sterile α motif compartmentalizes Polycomb group proteins and enhances their activity. Nat. Commun. 11, 5609 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. 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).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  42. Tatavosian, R. et al. Nuclear condensates of the Polycomb protein chromobox 2 (CBX2) assemble through phase separation. J. Biol. Chem. 294, 1451–1463 (2019).

    Article  CAS  PubMed  Google Scholar 

  43. Guo, Y. E. et al. Pol II phosphorylation regulates a switch between transcriptional and splicing condensates. Nature 572, 543–548 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  44. Lu, Y. et al. Phase separation of TAZ compartmentalizes the transcription machinery to promote gene expression. Nat. Cell Biol. 22, 453–464 (2020).

    Article  CAS  PubMed  Google Scholar 

  45. Sabari, B. R. et al. Coactivator condensation at super-enhancers links phase separation and gene control. Science 361, eaar3958 (2018).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  46. Altmeyer, M. et al. Liquid demixing of intrinsically disordered proteins is seeded by poly(ADP-ribose). Nat. Commun. 6, 8088 (2015).

    Article  CAS  PubMed  Google Scholar 

  47. Pandya-Jones, A. et al. A protein assembly mediates Xist localization and gene silencing. Nature 587, 145–151 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  48. Padeken, J. & Heun, P. Nucleolus and nuclear periphery: Velcro for heterochromatin. Curr. Opin. Cell Biol. 28, 54–60 (2014).

    Article  CAS  PubMed  Google Scholar 

  49. Tzur, Y. B., Wilson, K. L. & Gruenbaum, Y. SUN-domain proteins: ‘Velcro’ that links the nucleoskeleton to the cytoskeleton. Nat. Rev. Mol. Cell Biol. 7, 782–788 (2006).

    Article  CAS  PubMed  Google Scholar 

  50. Falahati, H., Pelham-Webb, B., Blythe, S. & Wieschaus, E. Nucleation by rRNA dictates the precision of nucleolus assembly. Curr. Biol. 26, 277–285 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  51. Berry, J. et al. RNA transcription modulates phase transition-driven nuclear body assembly. Proc. Natl Acad. Sci. USA 112, E5237–E5245 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  52. Mao, Y. S., Sunwoo, H., Zhang, B. & Spector, D. L. Direct visualization of the co-transcriptional assembly of a nuclear body by noncoding RNAs. Nat. Cell Biol. 13, 95–101 (2011).

    Article  CAS  PubMed  Google Scholar 

  53. Nizami, Z., Deryusheva, S. & Gall, J. G. The Cajal body and histone locus body. Cold Spring Harb. Perspect. Biol. 2, a000653 (2010).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  54. Chaumeil, J., Le Baccon, P., Wutz, A. & Heard, E. A novel role for Xist RNA in the formation of a repressive nuclear compartment into which genes are recruited when silenced. Genes Dev. 20, 2223–2237 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  55. Maison, C. et al. Higher-order structure in pericentric heterochromatin involves a distinct pattern of histone modification and an RNA component. Nat. Genet. 30, 329–334 (2002).

    Article  PubMed  Google Scholar 

  56. Quinodoz, S. A. et al. RNA promotes the formation of spatial compartments in the nucleus. Preprint at bioRxiv https://doi.org/10.1101/2020.08.25.267435 (2020).

    Article  Google Scholar 

  57. Bernstein, E. & Allis, C. D. RNA meets chromatin. Genes Dev. 19, 1635–1655 (2005).

    Article  CAS  PubMed  Google Scholar 

  58. Bernstein, E. et al. Mouse Polycomb proteins bind differentially to methylated histone H3 and RNA and are enriched in facultative heterochromatin. Mol. Cell. Biol. 26, 2560–2569 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  59. McHugh, C. A. C. A. et al. The Xist lncRNA interacts directly with SHARP to silence transcription through HDAC3. Nature 521, 232–236 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  60. Markaki, Y. et al. Xist-seeded nucleation sites form local concentration gradients of silencing proteins to inactivate the X-chromosome. Preprint at bioRxiv https://doi.org/10.1101/2020.11.22.393546 (2020).

    Article  Google Scholar 

  61. Wutz, A., Rasmussen, T. P. & Jaenisch, R. Chromosomal silencing and localization are mediated by different domains of Xist RNA. Nat. Genet. 30, 167–174 (2002).

    Article  CAS  PubMed  Google Scholar 

  62. Hacisuleyman, E. et al. Topological organization of multichromosomal regions by the long intergenic noncoding RNA Firre. Nat. Struct. Mol. Biol. 21, 198–206 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  63. Lu, Z. et al. RNA duplex map in living cells reveals higher-order transcriptome structure. Cell 165, 1267–1279 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  64. Wang, L. et al. Histone modifications regulate chromatin compartmentalization by contributing to a phase separation mechanism. Mol. Cell 76, 646–659.e6 (2019).

    Article  CAS  PubMed  Google Scholar 

  65. Fan, C. et al. Rett mutations attenuate phase separation of MeCP2. Cell Discov. 6, 646 (2020).

    Article  CAS  Google Scholar 

  66. Li, C. H. et al. MeCP2 links heterochromatin condensates and neurodevelopmental disease. Nature 586, 440–444 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  67. Wang, L. et al. Rett syndrome-causing mutations compromise MeCP2-mediated liquid–liquid phase separation of chromatin. Cell Res. 30, 393–407 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  68. Engreitz, J. M. et al. The Xist lncRNA exploits three-dimensional genome architecture to spread across the X chromosome. Science 341, 1237973 (2013).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  69. Dossin, F. et al. SPEN integrates transcriptional and epigenetic control of X-inactivation. Nature 578, 455–460 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  70. McSwiggen, D. T. et al. Evidence for DNA-mediated nuclear compartmentalization distinct from phase separation. eLife 8, e47098 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  71. Gasch, A. P. et al. Genomic expression programs in the response of yeast cells to environmental changes. Mol. Biol. Cell 11, 4241–4257 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  72. Chapal, M., Mintzer, S., Brodsky, S., Carmi, M. & Barkai, N. Resolving noise-control conflict by gene duplication. PLoS Biol. 17, e3000289 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  73. Zobeck, K. L., Buckley, M. S., Zipfel, W. R. & Lis, J. T. Recruitment timing and dynamics of transcription factors at the Hsp70 loci in living cells. Mol. Cell 40, 965–975 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  74. Soutourina, J. Transcription regulation by the Mediator complex. Nat. Rev. Mol. Cell Biol. 19, 262–274 (2018).

    Article  CAS  PubMed  Google Scholar 

  75. Banerji, J., Olson, L. & Schaffner, W. A lymphocyte-specific cellular enhancer is located downstream of the joining region in immunoglobulin heavy chain genes. Cell 33, 729–740 (1983).

    Article  CAS  PubMed  Google Scholar 

  76. Amano, T. et al. Chromosomal dynamics at the Shh locus: limb bud-specific differential regulation of competence and active transcription. Dev. Cell 16, 47–57 (2009).

    Article  CAS  PubMed  Google Scholar 

  77. Benabdallah, N. S. et al. Decreased enhancer–promoter proximity accompanying enhancer activation. Mol. Cell 76, 473–484.e7 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  78. Furlong, E. E. M. & Levine, M. Developmental enhancers and chromosome topology. Science 361, 1341–1345 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  79. Chong, S. et al. Imaging dynamic and selective low-complexity domain interactions that control gene transcription. Science 361, eaar2555 (2018).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  80. Zamudio, A. V. et al. Mediator condensates localize signaling factors to key cell identity genes. Mol. Cell 76, 753–766.e6 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  81. Dixon, J. R. et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 485, 376–380 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  82. Szabo, Q., Bantignies, F. & Cavalli, G. Principles of genome folding into topologically associating domains. Sci. Adv. 5, eaaw1668 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  83. 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).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  84. Flavahan, W. A. et al. Altered chromosomal topology drives oncogenic programs in SDH-deficient GISTs. Nature 575, 229–233 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  85. Flavahan, W. A. et al. Insulator dysfunction and oncogene activation in IDH mutant gliomas. Nature 529, 110–114 (2016).

    Article  CAS  PubMed  Google Scholar 

  86. Williamson, I. et al. Developmentally regulated Shh expression is robust to TAD perturbations. Development 146, dev179523 (2019).

    Article  CAS  PubMed  Google Scholar 

  87. Hnisz, D. et al. Super-enhancers in the control of cell identity and disease. Cell 155, 934 (2013).

    Article  CAS  PubMed  Google Scholar 

  88. Whyte, W. A. et al. Master transcription factors and mediator establish super-enhancers at key cell identity genes. Cell 153, 307–319 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  89. Lovén, J. et al. Selective inhibition of tumor oncogenes by disruption of super-enhancers. Cell 153, 320–334 (2013).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  90. Van Steensel, B. et al. Localization of the glucocorticoid receptor in discrete clusters in the cell nucleus. J. Cell Sci. 108, 3003–3011 (1995).

    Article  PubMed  Google Scholar 

  91. Grande, M. A., Van Der Kraan, I., De Jong, L. & Van Driel, R. Nuclear distribution of transcription factors in relation to sites of transcription and RNA polymerase II. J. Cell Sci. 110, 1781–1791 (1997).

    Article  CAS  PubMed  Google Scholar 

  92. Bregman, D. B., Du, L., Van Der Zee, S. & Warren, S. L. Transcription-dependent redistribution of the large subunit of RNA polymerase II to discrete nuclear domains. J. Cell Biol. 129, 287–298 (1995).

    Article  CAS  PubMed  Google Scholar 

  93. Papantonis, A. & Cook, P. R. Transcription factories: genome organization and gene regulation. Chem. Rev. 113, 8683–8705 (2013).

    Article  CAS  PubMed  Google Scholar 

  94. Iborra, F. J., Pombo, A., Jackson, D. A. & Cook, P. R. Active RNA polymerases are localized within discrete transcription ‘factories’ in human nuclei. J. Cell Sci. 109, 1427–1436 (1996).

    Article  CAS  PubMed  Google Scholar 

  95. Cisse, I. I. et al. Real-time dynamics of RNA polymerase II clustering in live human cells. Science 341, 664–667 (2013).

    Article  CAS  PubMed  Google Scholar 

  96. Wang, X., Cairns, M. J. & Yan, J. Super-enhancers in transcriptional regulation and genome organization. Nucleic Acids Res. 47, 11481–11496 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  97. Cho, W. K. et al. RNA Polymerase II cluster dynamics predict mRNA output in living cells. eLife 5, e13617 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  98. Berg, O. G., Winter, R. B. & von Hippel, P. H. Diffusion-driven mechanisms of protein translocation on nucleic acids. 1. models and theory. Biochemistry 20, 6929–6948 (1981).

    Article  CAS  PubMed  Google Scholar 

  99. Elf, J., Li, G. W. & Xie, X. S. Probing transcription factor dynamics at the single-molecule level in a living cell. Science 316, 1191–1194 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  100. Li, G. W. & Xie, X. S. Central dogma at the single-molecule level in living cells. Nature 475, 308–315 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  101. Brodsky, S. et al. Intrinsically disordered regions direct transcription factor in vivo binding specificity. Mol. Cell 79, 459–471.e4 (2020).

    Article  CAS  PubMed  Google Scholar 

  102. Valsecchi, C. I. K. et al. RNA nucleation by MSL2 induces selective X chromosome compartmentalization. Nature 589, 137–142 (2021).

    Article  CAS  PubMed  Google Scholar 

  103. Branco, M. R. & Pombo, A. Intermingling of chromosome territories in interphase suggests role in translocations and transcription-dependent associations. PLoS Biol. 4, 780–788 (2006).

    Article  CAS  Google Scholar 

  104. Shopland, L. S., Johnson, C. V., Byron, M., McNeil, J. & Lawrence, J. B. Clustering of multiple specific genes and gene-rich R-bands around SC-35 domains: evidence for local euchromatic neighborhoods. J. Cell Biol. 162, 981–990 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  105. Lomvardas, S. et al. Interchromosomal interactions and olfactory receptor choice. Cell 126, 403–413 (2006).

    Article  CAS  PubMed  Google Scholar 

  106. Strehle, M. & Guttman, M. Xist drives spatial compartmentalization of DNA and protein to orchestrate initiation and maintenance of X inactivation. Curr. Opin. Cell Biol. 64, 139–147 (2020).

    Article  CAS  PubMed  Google Scholar 

  107. Loda, A. & Heard, E. Xist RNA in action: past, present, and future. PLoS Genet. 15, e1008333 (2019).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  108. Chu, C. et al. Systematic discovery of Xist RNA binding proteins. Cell 161, 404–416 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  109. Minajigi, A. et al. A comprehensive Xist interactome reveals cohesin repulsion and an RNA-directed chromosome conformation. Science 349, aab2276 (2015).

    Article  CAS  Google Scholar 

  110. Moindrot, B. et al. A pooled shRNA screen identifies Rbm15, Spen, and Wtap as factors required for Xist RNA-mediated silencing. Cell Rep. 12, 562–572 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  111. Monfort, A. et al. Identification of Spen as a crucial factor for Xist function through forward genetic screening in haploid embryonic stem cells. Cell Rep. 12, 554–561 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  112. Wutz, A. & Jaenisch, R. A shift from reversible to irreversible X inactivation is triggered during ES cell differentiation. Mol. Cell 5, 695–705 (2000).

    Article  CAS  PubMed  Google Scholar 

  113. Pandey, R. R. et al. Kcnq1ot1 antisense noncoding RNA mediates lineage-specific transcriptional silencing through chromatin-level regulation. Mol. Cell 32, 232–246 (2008).

    Article  CAS  PubMed  Google Scholar 

  114. Kanduri, C. Kcnq1ot1: a chromatin regulatory RNA. Semin. Cell Dev.Biol. 22, 343–350 (2011).

    Article  CAS  PubMed  Google Scholar 

  115. Nagano, T. & Fraser, P. Emerging similarities in epigenetic gene silencing by long noncoding RNAs. Mamm. Genome 20, 557–562 (2009).

    Article  CAS  PubMed  Google Scholar 

  116. Cheutin, T. et al. Maintenance of stable heterochromatin domains by dynamic HP1 binding. Science 299, 721–725 (2003).

    Article  CAS  PubMed  Google Scholar 

  117. Maison, C. & Almouzni, G. HP1 and the dynamics of heterochromatin maintenance. Nat. Rev. Mol. Cell Biol. 5, 296–304 (2004).

    Article  CAS  PubMed  Google Scholar 

  118. Zhao, J., Sun, B. K., Erwin, J. A., Song, J. J. & Lee, J. T. Polycomb proteins targeted by a short repeat RNA to the mouse X chromosome. Science 322, 750–756 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  119. Plath, K., Mlynarczyk-Evans, S., Nusinow, D. A. & Panning, B. Xist RNA and the mechanism of X chromosome inactivation. Annu. Rev. Genet. 36, 233–278 (2002).

    Article  CAS  PubMed  Google Scholar 

  120. Mira-Bontenbal, H. & Gribnau, J. New Xist-interacting proteins in X-chromosome inactivation. Curr. Biol. 26, R338–R342 (2016).

    Article  CAS  PubMed  Google Scholar 

  121. Cerase, A. et al. Phase separation drives X-chromosome inactivation: a hypothesis. Nat. Struct. Mol. Biol. 26, 331–334 (2019).

    Article  CAS  PubMed  Google Scholar 

  122. Wutz, A. Xist function: bridging chromatin and stem cells. Trends Genet. 23, 457–464 (2007).

    Article  CAS  PubMed  Google Scholar 

  123. Cao, R. et al. Role of histone H3 lysine 27 methylation in Polycomb-group silencing. Science 298, 1039–1043 (2002).

    Article  CAS  PubMed  Google Scholar 

  124. Min, J., Zhang, Y. & Xu, R. M. Structural basis for specific binding of polycomb chromodomain to histone H3 methylated at Lys 27. Genes Dev. 17, 1823–1828 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  125. Fischle, W. et al. Molecular basis for the discrimination of repressive methyl-lysine marks in histone H3 by Polycomb and HP1 chromodomains. Genes. Dev. 17, 1870–1881 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  126. Illingworth, R. S. Chromatin folding and nuclear architecture: PRC1 function in 3D. Curr. Opin. Genet. Dev. 55, 82–90 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  127. Pirrotta, V. & Li, H. B. A view of nuclear Polycomb bodies. Curr. Opin. Genet. Dev. 22, 101–109 (2012).

    Article  CAS  PubMed  Google Scholar 

  128. Eeftens, J. M., Kapoor, M. & Brangwynne, C. P. Epigenetic memory as a time integral over prior history of Polycomb phase separation. bioRxiv https://doi.org/10.1101/2020.08.19.254706 (2020).

    Article  Google Scholar 

  129. 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).

    Article  CAS  PubMed  Google Scholar 

  130. Girard, C., Will, C., Peng, J. et al. Post-transcriptional spliceosomes are retained in nuclear speckles until splicing completion. Nat. Commun. 3, 994 (2012).

    Article  PubMed  CAS  Google Scholar 

  131. Napolitano, G., Lania, L. & Majello, B. RNA polymerase II CTD modifications: how many tales from a single tail. J. Cell. Physiol. 229, 538–544 (2014).

    Article  CAS  PubMed  Google Scholar 

  132. Hsin, J. P. & Manley, J. L. The RNA polymerase II CTD coordinates transcription and RNA processing. Genes Dev. 26, 2119–2137 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  133. Ahn, S. H., Kim, M. & Buratowski, S. Phosphorylation of serine 2 within the RNA polymerase II C-terminal domain couples transcription and 3′ end processing. Mol. Cell 13, 67–76 (2004).

    Article  CAS  PubMed  Google Scholar 

  134. Merkhofer, E. C., Hu, P. & Johnson, T. L. Introduction to cotranscriptional RNA splicing. Methods Mol. Biol. 1126, 83–96 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  135. Han, J., Xiong, J., Wang, D. & Fu, X. D. Pre-mRNA splicing: where and when in the nucleus. Trends Cell Biol. 21, 336–343 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  136. Herzel, L., Ottoz, D. S. M., Alpert, T. & Neugebauer, K. M. Splicing and transcription touch base: co-transcriptional spliceosome assembly and function. Nat. Rev. Mol. Cell Biol. 18, 637–650 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  137. McCracken, S. et al. The C-terminal domain of RNA polymerase II couples mRNA processing to transcription. Nature 385, 357–360 (1997).

    Article  CAS  PubMed  Google Scholar 

  138. Zhang, S. et al. Structure of a transcribing RNA polymerase II-U1 snRNP complex. Science 371, 305–309 (2021).

    Article  CAS  PubMed  Google Scholar 

  139. Ding, F. & Elowitz, M. B. Constitutive splicing and economies of scale in gene expression. Nat. Struct. Mol. Biol. 26, 424–432 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  140. Chen, Y. et al. Mapping 3D genome organization relative to nuclear compartments using TSA-Seq as a cytological ruler. J. Cell Biol. 217, 4025–4048 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  141. Dopie, J., Sweredoski, M. J., Moradian, A. & Belmont, A. S. Tyramide signal amplification mass spectrometry (TSA-MS) ratio identifies nuclear speckle proteins. J. Cell Biol. 219, e201910207 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  142. Saitoh, N. et al. Proteomic analysis of interchromatin granule clusters. Mol. Biol. Cell 15, 3876–3890 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  143. Lamond, A. I. & Spector, D. L. Nuclear speckles: a model for nuclear organelles. Nat. Rev. Mol. Cell Biol. 4, 605–612 (2003).

    Article  CAS  PubMed  Google Scholar 

  144. Kim, J., Venkata, N. C., Gonzalez, G. A. H., Khanna, N. & Belmont, A. S. Gene expression amplification by nuclear speckle association. J. Cell Biol. 219, e201904046 (2020).

    PubMed  Google Scholar 

  145. Alexander, K. A. et al. p53 mediates target gene association with nuclear speckles for amplified RNA expression. Mol. Cell 81, 1666–1681 (2021).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  146. Misteli, T. & Spector, D. L. RNA polymerase II targets pre-mRNA splicing factors to transcription sites in vivo. Mol. Cell 3, 697–705 (1999).

    Article  CAS  PubMed  Google Scholar 

  147. Pederson, T. The nucleolus. Cold Spring Harb. Perspect. Biol. 3, 1–15 (2011).

    Google Scholar 

  148. Carmo-Fonseca, M. & Rino, J. RNA seeds nuclear bodies. Nat. Cell Biol. 13, 110–112 (2011).

    Article  CAS  PubMed  Google Scholar 

  149. Shevtsov, S. P. & Dundr, M. Nucleation of nuclear bodies by RNA. Nat. Cell Biol. 13, 167–173 (2011).

    Article  CAS  PubMed  Google Scholar 

  150. Marzluff, W. F. & Koreski, K. P. Birth and death of histone mRNAs. Trends Genet. 33, 745–759 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  151. Scheer, U. & Benavente, R. Functional and dynamic aspects of the mammalian nucleolus. BioEssays 12, 14–21 (1990).

    Article  CAS  PubMed  Google Scholar 

  152. Schlegel, R. A., Miller, L. S. & Rose, K. M. Reduction in RNA synthesis following red cell-mediated microinjection of antibodies to RNA polymerase I. Cell Biol. Int. Rep. 9, 341–350 (1985).

    Article  CAS  PubMed  Google Scholar 

  153. Watkins, N. J. & Bohnsack, M. T. The box C/D and H/ACA snoRNPs: key players in the modification, processing and the dynamic folding of ribosomal RNA. Wiley Interdiscip. Rev. RNA 3, 397–414 (2012).

    Article  CAS  PubMed  Google Scholar 

  154. Kiss-László, Z., Henry, Y., Bachellerie, J. P., Caizergues-Ferrer, M. & Kiss, T. Site-specific ribose methylation of preribosomal RNA: a novel function for small nucleolar RNAs. Cell 85, 1077–1088 (1996).

    Article  PubMed  Google Scholar 

  155. Ni, J., Tien, A. L. & Fournier, M. J. Small nucleolar RNAs direct site-specific synthesis of pseudouridine in ribosomal RNA. Cell 89, 565–573 (1997).

    Article  CAS  PubMed  Google Scholar 

  156. Spycher, C. et al. 3′ end processing of mouse histone pre-mRNA: evidence for additional base-pairing between U7 snRNA and pre-mRNA. Nucleic Acids Res. 22, 4023–4030 (1994).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  157. Kolev, N. G. & Steitz, J. A. Symplekin and multiple other polyadenylation factors participate in 3′-end maturation of histone mRNAs. Genes Dev. 19, 2583–2592 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  158. Mowry, K. L. & Steitz, J. A. Identification of the human U7 snRNP as one of several factors involved in the 3′ end maturation of histone premessenger RNA’s. Science 238, 1682–1687 (1987).

    Article  CAS  PubMed  Google Scholar 

  159. Machyna, M. et al. The coilin interactome identifies hundreds of small noncoding RNAs that traffic through cajal bodies. Mol. Cell 56, 389–399 (2014).

    Article  CAS  PubMed  Google Scholar 

  160. Darzacq, X. et al. Cajal body-specific small nuclear RNAs: a novel class of 2′-O-methylation and pseudouridylation guide RNAs. EMBO J. 21, 2746–2756 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  161. Kiss, A. M. et al. A Cajal body-specific pseudouridylation guide RNA is composed of two box H/ACA snoRNA-like domains. Nucleic Acids Res. 30, 4643–4649 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  162. 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.e7 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  163. van Steensel, B. & Belmont, A. S. Lamina-associated domains: links with chromosome architecture, heterochromatin, and gene repression. Cell 169, 780–791 (2017).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  164. Reddy, K. L., Zullo, J. M., Bertolino, E. & Singh, H. Transcriptional repression mediated by repositioning of genes to the nuclear lamina. Nature 452, 243–247 (2008).

    Article  CAS  PubMed  Google Scholar 

  165. Kumaran, R. I. & Spector, D. L. A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence. J. Cell Biol. 180, 51–65 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  166. Finlan, L. E. et al. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet. 4, e1000039 (2008).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  167. Kim, J. H. et al. LADL: light-activated dynamic looping for endogenous gene expression control. Nat. Methods 16, 633–639 (2019).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  168. Wang, H. et al. CRISPR-mediated programmable 3D genome positioning and nuclear organization. Cell 175, 1405–1417.e14 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  169. Shin, Y. et al. Spatiotemporal control of intracellular phase transitions using light-activated optodroplets. Cell 168, 159–171.e14 (2017).

    Article  CAS  PubMed  Google Scholar 

  170. Shin, Y. et al. Liquid nuclear condensates mechanically sense and restructure the genome. Cell 175, 1481–1491.e13 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  171. Takei, Y. et al. Integrated spatial genomics reveals global architecture of single nuclei. Nature 590, 344–350 (2021).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  172. Eng, C. H. L. et al. Transcriptome-scale super-resolved imaging in tissues by RNA seqFISH+. Nature 568, 235–239 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  173. Collombet, S. et al. RNA polymerase II depletion from the inactive X chromosome territory is not mediated by physical compartmentalization. Preprint at bioRxiv https://doi.org/10.1101/2021.03.26.437188 (2021).

    Article  Google Scholar 

  174. Jerkovic’, I. & Cavalli, G. Understanding 3D genome organization by multidisciplinary methods. Nat. Rev. Mol. Cell Biol. https://doi.org/10.1038/s41580-021-00362-w (2021).

    Article  Google Scholar 

  175. Ganser, L. R. & Myong, S. Methods to study phase-separated condensates and the underlying molecular interactions. Trends Biochem. Sci. 45, 1004–1005 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  176. Robinson, C. V., Sali, A. & Baumeister, W. The molecular sociology of the cell. Nature 450, 973–982 (2007).

    Article  CAS  PubMed  Google Scholar 

  177. Shah, S., Lubeck, E., Zhou, W. & Cai, L. In situ transcription profiling of single cells reveals spatial organization of cells in the mouse hippocampus. Neuron 92, 342–357 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  178. Shah, S. et al. Dynamics and spatial genomics of the nascent transcriptome by intron seqFISH. Cell 174, 363–376.e16 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  179. Lubeck, E., Coskun, A. F., Zhiyentayev, T., Ahmad, M. & Cai, L. Single-cell in situ RNA profiling by sequential hybridization. Nat. Methods 11, 360–361 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  180. Frieda, K. L. et al. Synthetic recording and in situ readout of lineage information in single cells. Nature 541, 107–111 (2017).

    Article  CAS  PubMed  Google Scholar 

  181. Xia, C., Fan, J., Emanuel, G., Hao, J. & Zhuang, X. Spatial transcriptome profiling by MERFISH reveals subcellular RNA compartmentalization and cell cycle-dependent gene expression. Proc. Natl Acad. Sci. USA 116, 19490–19499 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  182. Beliveau, B. J. et al. Versatile design and synthesis platform for visualizing genomes with Oligopaint FISH probes. Proc. Natl Acad. Sci. USA 109, 21301–21306 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  183. Beliveau, B. J. et al. Single-molecule super-resolution imaging of chromosomes and in situ haplotype visualization using Oligopaint FISH probes. Nat. Commun. 6, 7147 (2015).

    Article  CAS  PubMed  Google Scholar 

  184. Beliveau, B. J., Apostolopoulos, N. & Wu, C. T. Visualizing genomes with Oligopaint FISH probes. Curr. Protoc. Mol. Biol. 105, 14.23.1–14.23.20 (2014).

    Article  Google Scholar 

  185. Takei, Y. et al. Global architecture of the nucleus in single cells by DNA seqFISH+ and multiplexed immunofluorescence. Preprint at bioRxiv https://doi.org/10.1101/2020.11.29.403055 (2020).

    Article  Google Scholar 

  186. Dey, S. & Maiti, S. Single-molecule photobleaching: instrumentation and applications. J. Biosci. 43, 447–454 (2018).

    Article  CAS  PubMed  Google Scholar 

  187. Hsia, Y. et al. Design of a hyperstable 60-subunit protein icosahedron. Nature 535, 136–139 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  188. Elson, E. L. Introduction to fluorescence correlation spectroscopy — brief and simple. Methods 140–141, 3–9 (2018).

    Article  PubMed  CAS  Google Scholar 

  189. Huang, B., Babcock, H. & Zhuang, X. Breaking the diffraction barrier: super-resolution imaging of cells. Cell 143, 1047–1058 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  190. Godin, A. G., Lounis, B. & Cognet, L. Super-resolution microscopy approaches for live cell imaging. Biophys. J. 107, 1777–1784 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  191. McCall, P. M. et al. Quantitative phase microscopy enables precise and efficient determination of biomolecular condensate composition. Preprint at bioRxiv https://doi.org/10.1101/2020.10.25.352823 (2020).

    Article  Google Scholar 

  192. Lieberman-Aiden, E. et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326, 289–293 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  193. Beagrie, R. A. et al. Complex multi-enhancer contacts captured by genome architecture mapping. Nature 543, 519–524 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

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Acknowledgements

The authors thank members of the Guttman laboratory, especially M. Strehle, J. Jachowicz, S. Quinodoz and I. Goronzy for helpful comments and discussions, I.-M. Strazhnik for figures and S. Hiley for editing. P.B. is supported by the University of California, Los Angeles (UCLA)-Caltech Medical Scientist Training Program (MSTP), National Institutes of Health (NIH) F30CA247447 and a Chen Graduate Innovator Grant. M.G. is a New York Stem Cell Foundation Robertson Investigator and an investigator at the Heritage Medical Research Institute. Research in the Guttman laboratory is funded by the NIH 4DN programme, an NIH Director’s Transformative R01 Award, the Chan-Zuckerberg Initiative and funds from Caltech.

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Glossary

Affinity

The strength of a non-covalent biochemical interaction, defined as the ratio of the association and dissociation rates.

Diffusion and affinity models

Models describing how a molecule (such as a transcription factor) proceeding on a random walk through the nucleus samples many possible binding partners until it finds its high-affinity cognate target site.

Nuclear territories

A catch-all term for 3D regions contained within the nucleus.

Compartments

Nuclear territories enriched in specific DNA, RNA and/or protein molecules.

Mediator complex

A protein complex that facilitates enhancer–promoter interactions, RNA polymerase II (Pol II) loading and transcription initiation.

Stoichiometric interaction

Biochemical interaction that occurs with defined ratios of components, generally with high affinity and specificity, including in protein complexes or the binding of a transcription factor to its DNA motif.

Non-linear regulatory behaviours

Responses to alteration in the reaction rate or efficiency that exceed those expected under a linear model for the underlying changes in the amounts of reactants or catalysts.

Phase transitions

The biophysical process that leads to phase separation, referring to a discontinuous change in the thermodynamic equilibrium state of a system in response to a change in a parameter such as temperature, pressure or molecular concentration.

Multivalent interactions

Molecular associations between multiple binding sites on the interacting molecules; can result in variable stoichiometric ratios.

Intrinsically disordered regions

(IDRs). Protein regions that do not have a single preferred structural conformation.

Biomolecular condensates

Concentration-dependent assemblies of molecules of variable stoichiometry, usually driven by multivalent and cooperative interactions that can form with or without phase separation.

Avidity

(Also known as ‘functional affinity’). The collective strength of multiple non-covalent molecular interactions. Avidity represents the overall force conferred by multiple affinities in concert, which exceeds the sum of the strength of those interactions.

Phase separation

Thermodynamically driven partitioning of a homogeneous mixture into locally distinct chemical sub-mixtures (phases) with distinct properties.

Homotypic interactions

Interactions occurring between two or more copies of the same type of molecule.

Heterotypic interactions

Interactions occurring between at least two molecules of different types.

Valency

The number of non-covalent interactions with other molecules that a single molecule or domain can support.

Liquid–liquid phase separation

(LLPS). A specific form of phase separation defined by the formation of a liquid compartment within a larger liquid environment.

Granules

Small (<1 µm) condensates that generally have a simple composition compared with the larger nuclear bodies.

Diffusible

Refers to molecules that proceed on a random walk throughout the volume that contains them.

Constrained molecules

Molecules that proceed on a random walk preferentially within a sub-volume of their overall environment, often owing to having high affinity to other molecules in that sub-volume.

Bodies

In the nucleus, large (≥1 µm), functionally distinct territories, often involved in molecular biogenesis, such as the nucleolus (ribosome biogenesis) and Cajal bodies (biogenesis of small nuclear RNAs (snRNAs)).

Transcriptional condensates

Assemblies of general transcription factors and Mediator complexes around enhancers and promoters that facilitate transcription activation.

Facilitated diffusion

In the context of compartmentalization, the process by which compartments restrict the random walks of diffusible molecules to a smaller volume. For example, constraining the diffusion of a transcription factor to a small nuclear volume around its target.

Super-stoichiometric

Refers to interactions that occur without fixed ratios of components and can exceed the binding capacity of any individual molecule, often involving high-avidity, multivalent interactions.

X-inactivation centre

A region on the X chromosome containing X-inactive specific transcript (XIST) and its cis-regulators; necessary and sufficient for initiation of X-chromosome inactivation (XCI) and protected from it to allow continual expression of XIST.

Epigenetic memory

The set of DNA and histone modifications that are heritable either from parents to offspring or from a mother cell to daughter cells.

Splicing condensates

High concentrations of splicing factors localized around nascent RNA transcripts, often in proximity to (but distinct from) nuclear speckles.

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Bhat, P., Honson, D. & Guttman, M. Nuclear compartmentalization as a mechanism of quantitative control of gene expression. Nat Rev Mol Cell Biol 22, 653–670 (2021). https://doi.org/10.1038/s41580-021-00387-1

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