Three-dimensional chromatin packing and positioning of plant genomes

Abstract

Information and function of a genome are not only decorated with epigenetic marks in the linear DNA sequence but also in their non-random spatial organization in the nucleus. Recent research has revealed that three-dimensional (3D) chromatin organization is highly correlated with the functionality of the genome, contributing to many cellular processes. Driven by the improvements in chromatin conformation capture methods and visualization techniques, the past decade has been an exciting period for the study of plants’ 3D genome structures, and our knowledge in this area has been substantially advanced. This Review describes our current understanding of plant chromatin organization and positioning beyond the nucleosomal level, and discusses future directions.

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Fig. 1: Schematic representation of hierarchical chromatin organisation in plants.
Fig. 2: Comparison of Arabidopsis and rice chromatin organization at chromosomal level.

References

  1. 1.

    Dixon, J. R., Gorkin, D. U. & Ren, B. Chromatin Domains: the unit of chromosome organization. Mol. Cell 62, 668–680 (2016).

  2. 2.

    Probst, A. V. & Mittelsten Scheid, O. Stress-induced structural changes in plant chromatin. Curr. Opin. Plant Biol. 27, 8–16 (2015).

  3. 3.

    Rosa, S. et al. Physical clustering of FLC alleles during Polycomb-mediated epigenetic silencing in vernalization. Genes Dev. 27, 1845–1850 (2013).

  4. 4.

    Paweletz, N. Walther Flemming: pioneer of mitosis research. Nat. Rev. Mol. Cell Biol. 2, 72–75 (2001).

  5. 5.

    Flemming, W. Zellsubstanz, kern und zelltheilung. (Vogel, Leipzig, 1882).

  6. 6.

    Sexton, T. & Cavalli, G. The role of chromosome domains in shaping the functional genome. Cell 160, 1049–1059 (2015).

  7. 7.

    Grob, S. & Grossniklaus, U. Chromosome conformation capture-based studies reveal novel features of plant nuclear architecture. Curr. Opin. Plant Biol. 36, 149–157 (2017).

  8. 8.

    Liu, C. & Weigel, D. Chromatin in 3D: progress and prospects for plants. Genome Biol. 16, 170 (2015).

  9. 9.

    Dekker, J., Rippe, K., Dekker, M. & Kleckner, N. Capturing chromosome conformation. Science 295, 1306–1311 (2002).

  10. 10.

    Zhao, Z. et al. Circular chromosome conformation capture (4C) uncovers extensive networks of epigenetically regulated intra- and interchromosomal interactions. Nat. Genet. 38, 1341–1347 (2006).

  11. 11.

    Dostie, J. et al. Chromosome Conformation Capture Carbon Copy (5C): a massively parallel solution for mapping interactions between genomic elements. Genome Res. 16, 1299–1309 (2006).

  12. 12.

    Simonis, M., Kooren, J. & de Laat, W. An evaluation of 3C-based methods to capture DNA interactions. Nat. Methods 4, 895–901 (2007).

  13. 13.

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

  14. 14.

    Grob, S. & Cavalli, G. Technical review: a hitchhiker’s guide to chromosome conformation capture. Methods Mol. Biol. 1675, 233–246 (2018).

  15. 15.

    Sotelo-Silveira, M., Chávez Montes, R. A., Sotelo-Silveira, J. R., Marsch-Martínez, N. & de Folter, S. Entering the next dimension: plant penomes in 3D. Trends Plant Sci. http://doi.org/cqwt (2018).

  16. 16.

    Mumbach, M. R. et al. HiChIP: efficient and sensitive analysis of protein-directed genome architecture. Nat. Methods 13, 919–922 (2016).

  17. 17.

    Li, G. et al. ChIA-PET tool for comprehensive chromatin interaction analysis with paired-end tag sequencing. Genome Biol. 11, R22 (2010).

  18. 18.

    Mifsud, B. et al. Mapping long-range promoter contacts in human cells with high-resolution capture Hi-C. Nat. Genet. 47, 598–606 (2015).

  19. 19.

    Jäger, R. et al. Capture Hi-C identifies the chromatin interactome of colorectal cancer risk loci. Nat. Commun. 6, 6178 (2015).

  20. 20.

    Li, G. et al. Chromatin interaction analysis with paired-end tag (ChIA-PET) sequencing technology and application. BMC Genom. 15, S11 (2014).

  21. 21.

    Nagano, T. et al. Single-cell Hi-C reveals cell-to-cell variability in chromosome structure. Nature 502, 59–64 (2013).

  22. 22.

    Stevens, T. J. et al. 3D structures of individual mammalian genomes studied by single-cell Hi-C. Nature 544, 59–64 (2017).

  23. 23.

    Du, Z. et al. Allelic reprogramming of 3D chromatin architecture during early mammalian development. Nature 547, 232–235 (2017).

  24. 24.

    Flyamer, I. M. et al. Single-nucleus Hi-C reveals unique chromatin reorganization at oocyte-to-zygote transition. Nature 544, 110–114 (2017).

  25. 25.

    Nagano, T. et al. Cell-cycle dynamics of chromosomal organization at single-cell resolution. Nature 547, 61–67 (2017).

  26. 26.

    Dekker, J. et al. The 4D nucleome project. Nature 549, 219–226 (2017).

  27. 27.

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

  28. 28.

    Giorgetti, L. & Heard, E. Closing the loop: 3C versus DNA FISH. Genome Biol. 17, 215 (2016).

  29. 29.

    Fudenberg, G. & Imakaev, M. FISH-ing for captured contacts: towards reconciling FISH and 3C. Nat. Methods 14, 673–678 (2017).

  30. 30.

    Williamson, I. et al. Spatial genome organization: contrasting views from chromosome conformation capture and fluorescence in situ hybridization. Genes Dev. 28, 2778–2791 (2014).

  31. 31.

    Cui, C., Shu, W. & Li, P. Fluorescence in situ hybridization: cell-based genetic diagnostic and research applications. Front. Cell Dev. Biol. 4, 89 (2016).

  32. 32.

    Solovei, I. et al. Spatial preservation of nuclear chromatin architecture during three-dimensional fluorescence in situ hybridization (3D-FISH). Exp. Cell Res. 276, 10–23 (2002).

  33. 33.

    Cremer, M. et al. Multicolor 3D fluorescence in situ hybridization for imaging interphase chromosomes. Methods Mol. Biol. 463, 205–239 (2008).

  34. 34.

    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).

  35. 35.

    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).

  36. 36.

    Ni, Y. et al. Super-resolution imaging of a 2.5 kb non-repetitive DNA in situ in the nuclear genome using molecular beacon probes. eLife 6, e21660 (2017).

  37. 37.

    Boettiger, A. N. et al. Super-resolution imaging reveals distinct chromatin folding for different epigenetic states. Nature 529, 418–422 (2016).

  38. 38.

    Wang, S. et al. Spatial organization of chromatin domains and compartments in single chromosomes. Science 353, 598–602 (2016).

  39. 39.

    Qin, P. et al. Live cell imaging of low- and non-repetitive chromosome loci using CRISPR–Cas9. Nat. Commun. 8, 14725 (2017).

  40. 40.

    Ye, H., Rong, Z. & Lin, Y. Live cell imaging of genomic loci using dCas9-SunTag system and a bright fluorescent protein. Protein Cell 8, 853–855 (2017).

  41. 41.

    Ma, Y. et al. Live cell imaging of single genomic loci with quantum dot-labeled TALEs. Nat. Commun. 8, 15318 (2017).

  42. 42.

    Hong, Y., Lu, G., Duan, J., Liu, W. & Zhang, Y. Comparison and optimization of CRISPR/dCas9/gRNA genome-labeling systems for live cell imaging. Genome Biol. 19, 39 (2018).

  43. 43.

    Kind, J. et al. Single-cell dynamics of genome-nuclear lamina interactions. Cell 153, 178–192 (2013).

  44. 44.

    Yu, M. & Ren, B. The three-dimensional organization of mammalian genomes. Annu. Rev. Cell Dev. Biol. 33, 265–289 (2017).

  45. 45.

    Pecinka, A. et al. Chromosome territory arrangement and homologous pairing in nuclei of Arabidopsis thaliana are predominantly random except for NOR-bearing chromosomes. Chromosoma 113, 258–269 (2004).

  46. 46.

    Fransz, P., De Jong, J. H., Lysak, M., Castiglione, M. R. & Schubert, I. Interphase chromosomes in Arabidopsis are organized as well defined chromocenters from which euchromatin loops emanate. Proc. Natl Acad. Sci. USA 99, 14584–14589 (2002).

  47. 47.

    Tiang, C.-L., He, Y. & Pawlowski, W. P. Chromosome organization and dynamics during interphase, mitosis, and meiosis in plants. Plant Physiol. 158, 26–34 (2012).

  48. 48.

    Rodriguez-Granados, N. Y. et al. Put your 3D glasses on: plant chromatin is on show. J. Exp. Bot. 67, 3205–3221 (2016).

  49. 49.

    Dong, F. & Jiang, J. Non-Rabl patterns of centromere and telomere distribution in the interphase nuclei of plant cells. Chromosome Res. 6, 551–558 (1998).

  50. 50.

    Prieto, P., Santos, A. P., Moore, G. & Shaw, P. Chromosomes associate premeiotically and in xylem vessel cells via their telomeres and centromeres in diploid rice (Oryza sativa). Chromosoma 112, 300–307 (2004).

  51. 51.

    Liu, C., Cheng, Y.-J., Wang, J.-W. & Weigel, D. Prominent topologically associated domains differentiate global chromatin packing in rice from Arabidopsis. Nat. Plants 3, 742–748 (2017).

  52. 52.

    Bass, H. W. et al. Evidence for the coincident initiation of homolog pairing and synapsis during the telomere-clustering (bouquet) stage of meiotic prophase. J. Cell Sci. 113, 1033–1042 (2000).

  53. 53.

    Schwarzacher, T. Three stages of meiotic homologous chromosome pairing in wheat: cognition, alignment and synapsis. Sex. Plant Reprod. 10, 324–331 (1997).

  54. 54.

    Zhang, F. et al. The F-box protein ZYGO1 mediates Bouquet formation to promote homologous pairing, synapsis, and recombination in rice meiosis. Plant Cell 29, 2597–2609 (2017).

  55. 55.

    Bourbousse, C. et al. Light signaling controls nuclear architecture reorganization during seedling establishment. Proc. Natl Acad. Sci. USA 112, 2836–2844 (2015).

  56. 56.

    Pecinka, A. et al. Epigenetic regulation of repetitive elements is attenuated by prolonged heat stress in Arabidopsis. Plant Cell 22, 3118–3129 (2010).

  57. 57.

    Wang, L.-C., Wu, J.-R., Hsu, Y.-J. & Wu, S.-J. Arabidopsis HIT4, a regulator involved in heat-triggered reorganization of chromatin and release of transcriptional gene silencing, relocates from chromocenters to the nucleolus in response to heat stress. New Phytol. 205, 544–554 (2015).

  58. 58.

    Dong, P. et al. 3D chromatin architecture of large plant genomes determined by local A/B compartments. Mol. Plant 10, 1497–1509 (2017).

  59. 59.

    Grob, S., Schmid, M. W. & Grossniklaus, U. Hi-C analysis in Arabidopsis identifies the KNOT, a structure with similarities to the flamenco locus of Drosophila. Mol. Cell 55, 678–693 (2014).

  60. 60.

    Wang, J. et al. Genome-wide analysis of the distinct types of chromatin interactions in Arabidopsis thaliana. Plant Cell Physiol. 58, 57–70 (2017).

  61. 61.

    Wang, M. et al. Evolutionary dynamics of 3D genome architecture following polyploidization in cotton. Nat. Plants 4, 90–97 (2018).

  62. 62.

    Grob, S., Schmid, M. W., Luedtke, N. W., Wicker, T. & Grossniklaus, U. Characterization of chromosomal architecture in Arabidopsis by chromosome conformation capture. Genome Biol. 14, R129 (2013).

  63. 63.

    Concia, L. et al. Genome-wide analysis of the Arabidopsis thaliana replication timing program. Plant Physiol. 176, 2166–2185 (2018).

  64. 64.

    Feng, S. et al. Genome-wide Hi-C analyses in wild-type and mutants reveal high-resolution chromatin interactions in Arabidopsis. Mol. Cell 55, 694–707 (2014).

  65. 65.

    Zhu, W. et al. Altered chromatin compaction and histone methylation drive non-additive gene expression in an interspecific Arabidopsis hybrid. Genome Biol. 18, 157 (2017).

  66. 66.

    Szabo, Q. et al. TADs are 3D structural units of higher-order chromosome organization in Drosophila. Sci. Adv. 4, eaar8082 (2018).

  67. 67.

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

  68. 68.

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

  69. 69.

    Hou, C., Li, L., Qin, Z. S. & Corces, V. G. Gene density, transcription, and insulators contribute to the partition of the Drosophila genome into physical domains. Mol. Cell 48, 471–484 (2012).

  70. 70.

    Sexton, T. et al. Three-dimensional folding and functional organization principles of the Drosophila genome. Cell 148, 458–472 (2012).

  71. 71.

    Crane, E. et al. Condensin-driven remodelling of X chromosome topology during dosage compensation. Nature 523, 240–244 (2015).

  72. 72.

    Mizuguchi, T. et al. Cohesin-dependent globules and heterochromatin shape 3D genome architecture in S. pombe. Nature 516, 432–435 (2014).

  73. 73.

    Le, T. B. K., Imakaev, M. V., Mirny, L. A. & Laub, M. T. High-resolution mapping of the spatial organization of a bacterial chromosome. Science 342, 731–734 (2013).

  74. 74.

    Phillips-Cremins, J. E. et al. Architectural protein subclasses shape 3D organization of genomes during lineage commitment. Cell 153, 1281–1295 (2013).

  75. 75.

    Bonev, B. et al. Multiscale 3D genome rewiring during mouse neural development. Cell 171, 557–572 (2017).

  76. 76.

    Hug, C. B., Grimaldi, A. G., Kruse, K. & Vaquerizas, J. M. Chromatin architecture emerges during zygotic genome activation independent of transcription. Cell 169, 216–228 (2017).

  77. 77.

    Naumova, N. et al. Organization of the mitotic chromosome. Science 342, 948–953 (2013).

  78. 78.

    Phillips, J. E. & Corces, V. G. CTCF: master weaver of the genome. Cell 137, 1194–1211 (2009).

  79. 79.

    Fudenberg, G. et al. Formation of chromosomal domains by loop extrusion. Cell Rep. 15, 2038–2049 (2016).

  80. 80.

    Zuin, J. et al. Cohesin and CTCF differentially affect chromatin architecture and gene expression in human cells. Proc. Natl Acad. Sci. USA 111, 996–1001 (2014).

  81. 81.

    Sofueva, S. et al. Cohesin-mediated interactions organize chromosomal domain architecture. EMBO J. 32, 3119–3129 (2013).

  82. 82.

    de Wit, E. et al. CTCF binding polarity determines chromatin looping. Mol. Cell 60, 676–684 (2015).

  83. 83.

    Bonev, B. & Cavalli, G. Organization and function of the 3D genome. Nat. Rev. Genet. 17, 661–678 (2016).

  84. 84.

    Dekker, J. & Mirny, L. The 3D genome as moderator of chromosomal communication. Cell 164, 1110–1121 (2016).

  85. 85.

    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, 6456–6465 (2015).

  86. 86.

    Terakawa, T. et al. The condensin complex is a mechanochemical motor that translocates along DNA. Science 358, 672–676 (2017).

  87. 87.

    Ganji, M. et al. Real-time imaging of DNA loop extrusion by condensin. Science 360, 102–105 (2018).

  88. 88.

    Gibcus, J. H. et al. A pathway for mitotic chromosome formation. Science 359, eaao6135 (2018).

  89. 89.

    Haarhuis, J. H. I. et al. The cohesin release factor WAPL restricts chromatin loop extension. Cell 169, 693–707 (2017).

  90. 90.

    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).

  91. 91.

    Busslinger, G. A. et al. Cohesin is positioned in mammalian genomes by transcription, CTCF and Wapl. Nature 544, 503–507 (2017).

  92. 92.

    Schwarzer, W. et al. Two independent modes of chromatin organization revealed by cohesin removal. Nature 551, 51–56 (2017).

  93. 93.

    Rao, S. S. P. et al. Cohesin loss eliminates all loop domains. Cell 171, 305–320 (2017).

  94. 94.

    Nora, E. P. et al. Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169, 930–944 (2017).

  95. 95.

    Rowley, M. J. et al. Evolutionarily conserved principles predict 3D chromatin organization. Mol. Cell 67, 837–852 (2017).

  96. 96.

    Isoda, T. et al. Non-coding transcription instructs chromatin folding and compartmentalization to dictate enhancer-promoter communication and T cell fate. Cell 171, 103–119 (2017).

  97. 97.

    Benedetti, F., Racko, D., Dorier, J., Burnier, Y. & Stasiak, A. Transcription-induced supercoiling explains formation of self-interacting chromatin domains in S. pombe. Nucleic Acids Res. 45, 9850–9859 (2017).

  98. 98.

    Moissiard, G. et al. MORC family ATPases required for heterochromatin condensation and gene silencing. Science 336, 1448–1451 (2012).

  99. 99.

    Wang, C. et al. Genome-wide analysis of local chromatin packing in Arabidopsis thaliana. Genome Res. 25, 246–256 (2015).

  100. 100.

    Heger, P., Marin, B., Bartkuhn, M., Schierenberg, E. & Wiehe, T. The chromatin insulator CTCF and the emergence of metazoan diversity. Proc. Natl Acad. Sci. USA 109, 17507–17512 (2012).

  101. 101.

    Ulianov, S. V. et al. Active chromatin and transcription play a key role in chromosome partitioning into topologically associating domains. Genome Res. 26, 70–84 (2015).

  102. 102.

    Wang, M. et al. Asymmetric subgenome selection and cis-regulatory divergence during cotton domestication. Nat. Genet. 49, 579–587 (2017).

  103. 103.

    Dong, Q. et al. Genome-wide Hi-C analysis reveals extensive hierarchical chromatin interactions in rice. Plant J. 94, 1141–1156 (2018).

  104. 104.

    Vietri Rudan, M. et al. Comparative Hi-C reveals that CTCF underlies evolution of chromosomal domain architecture. Cell Rep. 10, 1297–1309 (2015).

  105. 105.

    Liu, C. et al. Genome-wide analysis of chromatin packing in Arabidopsis thaliana at single-gene resolution. Genome Res. 26, 1057–1068 (2016).

  106. 106.

    Hsieh, T.-H. S. et al. Mapping nucleosome resolution chromosome folding in yeast by micro-C. Cell 162, 108–119 (2015).

  107. 107.

    Xu, W. et al. The R-loop is a common chromatin feature of the Arabidopsis genome. Nat. Plants 3, 704–714 (2017).

  108. 108.

    Conn, V. M. et al. A circRNA from SEPALLATA3 regulates splicing of its cognate mRNA through R-loop formation. Nat. Plants 3, 17053 (2017).

  109. 109.

    Fransz, P. & de Jong, H. From nucleosome to chromosome: a dynamic organization of genetic information: Dynamic organization of genetic information. Plant J. 66, 4–17 (2011).

  110. 110.

    Weber, C. M. & Henikoff, S. Histone variants: dynamic punctuation in transcription. Genes Dev. 28, 672–682 (2014).

  111. 111.

    Vergara, Z. & Gutierrez, C. Emerging roles of chromatin in the maintenance of genome organization and function in plants. Genome Biol. 18, 96 (2017).

  112. 112.

    Bannister, A. J. & Kouzarides, T. Regulation of chromatin by histone modifications. Cell Res. 21, 381–395 (2011).

  113. 113.

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

  114. 114.

    Nguyen, H. Q. & Bosco, G. Gene positioning effects on expression in eukaryotes. Annu. Rev. Genet. 49, 627–646 (2015).

  115. 115.

    Groves, N. R., Biel, A. M., Newman-Griffis, A. H. & Meier, I. Dynamic changes in plant nuclear organization in response to environmental and developmental signals. Plant Physiol. 176, 230–241 (2018).

  116. 116.

    Németh, A. & Längst, G. Genome organization in and around the nucleolus. Trends Genet. 27, 149–156 (2011).

  117. 117.

    Durut, N. et al. A duplicated NUCLEOLIN gene with antagonistic activity is required for chromatin organization of silent 45S rDNA in Arabidopsis. Plant Cell 26, 1330–1344 (2014).

  118. 118.

    Pontvianne, F. et al. Subnuclear partitioning of rRNA genes between the nucleolus and nucleoplasm reflects alternative epiallelic states. Genes Dev. 27, 1545–1550 (2013).

  119. 119.

    Pontvianne, F. et al. Identification of nucleolus-associated chromatin domains reveals a role for the nucleolus in 3D organization of the A. thaliana genome. Cell Rep. 16, 1574–1587 (2016).

  120. 120.

    Armstrong, S. J., Franklin, F. C. & Jones, G. H. Nucleolus-associated telomere clustering and pairing precede meiotic chromosome synapsis in Arabidopsis thaliana. J. Cell Sci. 114, 4207–4217 (2001).

  121. 121.

    Németh, A. et al. Initial genomics of the human nucleolus. PLoS Genet. 6, e1000889 (2010).

  122. 122.

    van Koningsbruggen, S. et al. High-resolution whole-genome sequencing reveals that specific chromatin domains from most human chromosomes associate with nucleoli. Mol. Biol. Cell 21, 3735–3748 (2010).

  123. 123.

    Montacié, C. et al. Nucleolar proteome analysis and proteasomal activity assays reveal a link between nucleolus and 26S proteasome in A. thaliana. Front. Plant Sci. 8, 1815 (2017).

  124. 124.

    Pendle, A. F. et al. Proteomic analysis of the Arabidopsis nucleolus suggests novel nucleolar functions. Mol. Biol. Cell 16, 260–269 (2005).

  125. 125.

    Harr, J. C., Gonzalez-Sandoval, A. & Gasser, S. M. Histones and histone modifications in perinuclear chromatin anchoring: from yeast to man. EMBO Rep. 17, 139–155 (2016).

  126. 126.

    Ciska, M. & Moreno Díaz de la Espina, S. The intriguing plant nuclear lamina. Front. Plant Sci. 5, 166 (2014).

  127. 127.

    Zhou, X., Graumann, K. & Meier, I. The plant nuclear envelope as a multifunctional platform LINCed by SUN and KASH. J. Exp. Bot. 66, 1649–1659 (2015).

  128. 128.

    Meier, I., Richards, E. J. & Evans, D. E. Cell biology of the plant nucleus. Annu. Rev. Plant Biol. 68, 139–172 (2017).

  129. 129.

    Goto, C., Tamura, K., Fukao, Y., Shimada, T. & Hara-Nishimura, I. The novel nuclear envelope protein KAKU4 modulates nuclear morphology in Arabidopsis. Plant Cell 26, 2143–2155 (2014).

  130. 130.

    Pawar, V. et al. A novel family of plant nuclear envelope-associated proteins. J. Exp. Bot. 67, 5699–5710 (2016).

  131. 131.

    Ibarra, A. & Hetzer, M. W. Nuclear pore proteins and the control of genome functions. Genes Dev. 29, 337–349 (2015).

  132. 132.

    Strambio-De-Castillia, C., Niepel, M. & Rout, M. P. The nuclear pore complex: bridging nuclear transport and gene regulation. Nat. Rev. Mol. Cell Biol. 11, 490–501 (2010).

  133. 133.

    Parry, G. The plant nuclear envelope and regulation of gene expression. J. Exp. Bot. 66, 1673–1685 (2015).

  134. 134.

    Yang, Y., Wang, W., Chu, Z., Zhu, J.-K. & Zhang, H. Roles of nuclear pores and nucleo-cytoplasmic trafficking in plant stress responses. Front. Plant Sci. 8, 574 (2017).

  135. 135.

    Smith, S. et al. Marker gene tethering by nucleoporins affects gene expression in plants. Nucleus 6, 471–478 (2015).

  136. 136.

    Zhu, Y. et al. An Arabidopsis nucleoporin NUP85 modulates plant responses to ABA and salt stress. PLoS Genet. 13, e1007124 (2017).

  137. 137.

    Fang, Y. & Spector, D. L. Centromere positioning and dynamics in living Arabidopsis plants. Mol. Biol. Cell 16, 5710–5718 (2005).

  138. 138.

    Bi, X. et al. Nonrandom domain organization of the Arabidopsis genome at the nuclear periphery. Genome Res. 27, 1162–1173 (2017).

  139. 139.

    Tessadori, F. et al. Large-scale dissociation and sequential reassembly of pericentric heterochromatin in dedifferentiated Arabidopsis cells. J. Cell Sci. 120, 1200–1208 (2007).

  140. 140.

    van Zanten, M. et al. Photoreceptors CRYTOCHROME2 and phytochrome B control chromatin compaction in Arabidopsis. Plant Physiol. 154, 1686–1696 (2010).

  141. 141.

    Soppe, W. J. J. et al. DNA methylation controls histone H3 lysine 9 methylation and heterochromatin assembly in Arabidopsis. EMBO J. 21, 6549–6559 (2002).

  142. 142.

    Towbin, B. D. et al. Step-wise methylation of histone H3K9 positions heterochromatin at the nuclear periphery. Cell 150, 934–947 (2012).

  143. 143.

    Bian, Q., Khanna, N., Alvikas, J. & Belmont, A. S. β-Globin cis-elements determine differential nuclear targeting through epigenetic modifications. J. Cell Biol. 203, 767–783 (2013).

  144. 144.

    Gonzalez-Sandoval, A. et al. Perinuclear anchoring of H3K9-methylated chromatin stabilizes induced cell fate in C. elegans embryos. Cell 163, 1333–1347 (2015).

  145. 145.

    Harr, J. C. et al. Directed targeting of chromatin to the nuclear lamina is mediated by chromatin state and A-type lamins. J. Cell Biol. 208, 33–52 (2015).

  146. 146.

    Solovei, I. et al. LBR and lamin A/C sequentially tether peripheral heterochromatin and inversely regulate differentiation. Cell 152, 584–598 (2013).

  147. 147.

    Wang, H., Dittmer, T. A. & Richards, E. J. Arabidopsis CROWDED NUCLEI (CRWN) proteins are required for nuclear size control and heterochromatin organization. BMC Plant Biol. 13, 200 (2013).

  148. 148.

    Poulet, A. et al. The LINC complex contributes to heterochromatin organisation and transcriptional gene silencing in plants. J. Cell Sci. 130, 590–601 (2017).

  149. 149.

    Dittmer, T. A., Stacey, N. J., Sugimoto-Shirasu, K. & Richards, E. J. LITTLE NUCLEI genes affecting nuclear morphology in Arabidopsis thaliana. Plant Cell 19, 2793–2803 (2007).

  150. 150.

    Sakamoto, Y. & Takagi, S. LITTLE NUCLEI 1 and 4 regulate nuclear morphology in Arabidopsis thaliana. Plant Cell Physiol. 54, 622–633 (2013).

  151. 151.

    Feng, C.-M., Qiu, Y., Van Buskirk, E. K., Yang, E. J. & Chen, M. Light-regulated gene repositioning in Arabidopsis. Nat. Commun. 5, 3027 (2014).

  152. 152.

    Oka, R. et al. Genome-wide mapping of transcriptional enhancer candidates using DNA and chromatin features in maize. Genome Biol. 18, 137 (2017).

  153. 153.

    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).

  154. 154.

    Veluchamy, A. et al. LHP1 Regulates H3K27me3 spreading and shapes the three-dimensional conformation of the Arabidopsis genome. PLoS ONE 11, e0158936 (2016).

  155. 155.

    Chica, C., Louis, A., Roest Crollius, H., Colot, V. & Roudier, F. Comparative epigenomics in the Brassicaceae reveals two evolutionarily conserved modes of PRC2-mediated gene regulation. Genome Biol. 18, 207 (2017).

  156. 156.

    Xiong, J., Zhang, Z. & Zhu, B. Polycomb `polypacks' the chromatin. Proc. Natl Acad. Sci. USA 113, 14878–14880 (2016).

  157. 157.

    Hall, L. L. & Lawrence, J. B. RNA as a fundamental component of interphase chromosomes: could repeats prove key? Curr. Opin. Genet. Dev. 37, 137–147 (2016).

  158. 158.

    Marchese, F. P., Raimondi, I. & Huarte, M. The multidimensional mechanisms of long noncoding RNA function. Genome Biol. 18, 206 (2017).

  159. 159.

    Benoit, M., Layat, E., Tourmente, S. & Probst, A. V. Heterochromatin dynamics during developmental transitions in Arabidopsis - a focus on ribosomal DNA loci. Gene 526, 39–45 (2013).

  160. 160.

    Melé, M. & Rinn, J. L. ‘Cat’s Cradling’ the 3D genome by the act of lncRNA transcription. Mol. Cell 62, 657–664 (2016).

  161. 161.

    Arabidopsis Genome Initiative. Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408, 796–815 (2000).

  162. 162.

    Wu, J. et al. Physical maps and recombination frequency of six rice chromosomes. Plant J. 36, 720–730 (2003).

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Acknowledgements

This work has received funding from the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation programme (grant agreement No. 757600). We apologize to our colleagues whose work was not included or discussed sufficiently in this manuscript due to space constraints.

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E.D. and C.L. wrote the manuscript.

Correspondence to Chang Liu.

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Doğan, E.S., Liu, C. Three-dimensional chromatin packing and positioning of plant genomes. Nature Plants 4, 521–529 (2018). https://doi.org/10.1038/s41477-018-0199-5

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