A guide to studying 3D genome structure and dynamics in the kidney.

Yang, J. H. et al. Loss of epigenetic information as a cause of mammalian aging. Cell 186, 305–326.e27 (2023).

pubmed: 36638792 pmcid: 10166133 doi: 10.1016/j.cell.2022.12.027

Krijger, P. H. & de Laat, W. Regulation of disease-associated gene expression in the 3D genome. Nat. Rev. Mol. Cell Biol. 17, 771–782 (2016).

pubmed: 27826147 doi: 10.1038/nrm.2016.138

Maurano, M. T. et al. Systematic localization of common disease-associated variation in regulatory DNA. Science 337, 1190–1195 (2012).

pubmed: 22955828 pmcid: 3771521 doi: 10.1126/science.1222794

Lambert, S. A. et al. The human transcription factors. Cell 175, 598–599 (2018).

pubmed: 30290144 doi: 10.1016/j.cell.2018.09.045

Kim, S. & Shendure, J. Mechanisms of interplay between transcription factors and the 3D genome. Mol. Cell 76, 306–319 (2019).

pubmed: 31521504 doi: 10.1016/j.molcel.2019.08.010

Dekker, J. et al. Spatial and temporal organization of the genome: current state and future aims of the 4D nucleome project. Mol. Cell 83, 2624–2640 (2023).

pubmed: 37419111 pmcid: 10528254 doi: 10.1016/j.molcel.2023.06.018

Mescher, A. L. in: Junqueira’s Basic Histology: Text and Atlas, 17th Edition. (McGraw Hill, 2024).

Markaki, Y. et al. Functional nuclear organization of transcription and DNA replication: a topographical marriage between chromatin domains and the interchromatin compartment. Cold Spring Harb. Symp. Quant. Biol. 75, 475–492 (2010).

pubmed: 21467142 doi: 10.1101/sqb.2010.75.042

Gall, J. G. & Pardue, M. L. Formation and detection of RNA-DNA hybrid molecules in cytological preparations. Proc. Natl Acad. Sci. USA 63, 378–383 (1969).

pubmed: 4895535 pmcid: 223575 doi: 10.1073/pnas.63.2.378

Rudkin, G. T. & Stollar, B. D. High resolution detection of DNA-RNA hybrids in situ by indirect immunofluorescence. Nature 265, 472–473 (1977).

pubmed: 401954 doi: 10.1038/265472a0

Singer, R. H. & Ward, D. C. Actin gene expression visualized in chicken muscle tissue culture by using in situ hybridization with a biotinated nucleotide analog. Proc. Natl Acad. Sci. USA 79, 7331–7335 (1982).

pubmed: 6961411 pmcid: 347333 doi: 10.1073/pnas.79.23.7331

Tanner, M. et al. Chromogenic in situ hybridization: a practical alternative for fluorescence in situ hybridization to detect HER-2/neu oncogene amplification in archival breast cancer samples. Am. J. Pathol. 157, 1467–1472 (2000).

pubmed: 11073807 pmcid: 1885742 doi: 10.1016/S0002-9440(10)64785-2

Moyzis, R. K. et al. A highly conserved repetitive DNA sequence, (TTAGGG)n, present at the telomeres of human chromosomes. Proc. Natl Acad. Sci. USA 85, 6622–6626 (1988).

pubmed: 3413114 pmcid: 282029 doi: 10.1073/pnas.85.18.6622

Rigby, P. W., Dieckmann, M., Rhodes, C. & Berg, P. Labeling deoxyribonucleic acid to high specific activity in vitro by nick translation with DNA polymerase I. J. Mol. Biol. 113, 237–251 (1977).

pubmed: 881736 doi: 10.1016/0022-2836(77)90052-3

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).

pubmed: 23236188 pmcid: 3535588 doi: 10.1073/pnas.1213818110

Boyle, S., Rodesch, M. J., Halvensleben, H. A., Jeddeloh, J. A. & Bickmore, W. A. Fluorescence in situ hybridization with high-complexity repeat-free oligonucleotide probes generated by massively parallel synthesis. Chromosome Res. 19, 901–909 (2011).

pubmed: 22006037 pmcid: 3210351 doi: 10.1007/s10577-011-9245-0

Yamada, N. A. et al. Visualization of fine-scale genomic structure by oligonucleotide-based high-resolution FISH. Cytogenet. Genome Res. 132, 248–254 (2011).

pubmed: 21178330 doi: 10.1159/000322717

Raj, A., van den Bogaard, P., Rifkin, S. A., van Oudenaarden, A. & Tyagi, S. Imaging individual mRNA molecules using multiple singly labeled probes. Nat. Methods 5, 877–879 (2008).

pubmed: 18806792 pmcid: 3126653 doi: 10.1038/nmeth.1253

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).

pubmed: 11978004 doi: 10.1006/excr.2002.5513

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

pubmed: 33505024 pmcid: 7878433 doi: 10.1038/s41586-020-03126-2

Nguyen, H. Q. et al. 3D mapping and accelerated super-resolution imaging of the human genome using in situ sequencing. Nat. Methods 17, 822–832 (2020).

pubmed: 32719531 pmcid: 7537785 doi: 10.1038/s41592-020-0890-0

Chen, K. H., Boettiger, A. N., Moffitt, J. R., Wang, S. & Zhuang, X. R. N. A imaging. Spatially resolved, highly multiplexed RNA profiling in single cells. Science 348, aaa6090 (2015).

pubmed: 25858977 pmcid: 4662681 doi: 10.1126/science.aaa6090

Lee, J. H. et al. Highly multiplexed subcellular RNA sequencing in situ. Science 343, 1360–1363 (2014).

pubmed: 24578530 pmcid: 4140943 doi: 10.1126/science.1250212

Merritt, C. R. et al. Multiplex digital spatial profiling of proteins and RNA in fixed tissue. Nat. Biotechnol. 38, 586–599 (2020).

pubmed: 32393914 doi: 10.1038/s41587-020-0472-9

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).

pubmed: 25962338 doi: 10.1038/ncomms8147

Kishi, J. Y. et al. SABER amplifies FISH: enhanced multiplexed imaging of RNA and DNA in cells and tissues. Nat. Methods 16, 533–544 (2019).

pubmed: 31110282 pmcid: 6544483 doi: 10.1038/s41592-019-0404-0

Payne, A. C. et al. In situ genome sequencing resolves DNA sequence and structure in intact biological samples. Science 371, eaay3446 (2021).

pubmed: 33384301 doi: 10.1126/science.aay3446

Mateo, L. J. et al. Visualizing DNA folding and RNA in embryos at single-cell resolution. Nature 568, 49–54 (2019).

pubmed: 30886393 pmcid: 6556380 doi: 10.1038/s41586-019-1035-4

Mateo, L. J., Sinnott-Armstrong, N. & Boettiger, A. N. Tracing DNA paths and RNA profiles in cultured cells and tissues with ORCA. Nat. Protoc. 16, 1647–1713 (2021).

pubmed: 33619390 pmcid: 8525907 doi: 10.1038/s41596-020-00478-x

Cardozo Gizzi, A. M. et al. Microscopy-based chromosome conformation capture enables simultaneous visualization of genome organization and transcription in intact organisms. Mol. Cell 74, 212–222.e5 (2019).

pubmed: 30795893 doi: 10.1016/j.molcel.2019.01.011

Cardozo Gizzi, A. M. et al. Direct and simultaneous observation of transcription and chromosome architecture in single cells with Hi-M. Nat. Protoc. 15, 840–876 (2020).

pubmed: 31969721 doi: 10.1038/s41596-019-0269-9

Rao, S. S. et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell 159, 1665–1680 (2014).

pubmed: 25497547 pmcid: 5635824 doi: 10.1016/j.cell.2014.11.021

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

pubmed: 11847345 doi: 10.1126/science.1067799

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).

pubmed: 16954542 pmcid: 1581439 doi: 10.1101/gr.5571506

Simonis, M. et al. Nuclear organization of active and inactive chromatin domains uncovered by chromosome conformation capture-on-chip (4C). Nat. Genet. 38, 1348–1354 (2006).

pubmed: 17033623 doi: 10.1038/ng1896

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

pubmed: 19815776 pmcid: 2858594 doi: 10.1126/science.1181369

Fiorillo, L. et al. Comparison of the Hi-C, GAM and SPRITE methods using polymer models of chromatin. Nat. Methods 18, 482–490 (2021).

pubmed: 33963348 pmcid: 8416658 doi: 10.1038/s41592-021-01135-1

Belaghzal, H., Dekker, J. & Gibcus, J. H. Hi-C 2.0: an optimized Hi-C procedure for high-resolution genome-wide mapping of chromosome conformation. Methods 123, 56–65 (2017).

pubmed: 28435001 pmcid: 5522765 doi: 10.1016/j.ymeth.2017.04.004

Krietenstein, N. et al. Ultrastructural details of mammalian chromosome architecture. Mol. Cell 78, 554–565.e7 (2020).

pubmed: 32213324 pmcid: 7222625 doi: 10.1016/j.molcel.2020.03.003

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.e13 (2019).

pubmed: 31522987 pmcid: 7251926 doi: 10.1016/j.molcel.2019.07.039

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

pubmed: 29887377 pmcid: 6548320 doi: 10.1016/j.cell.2018.05.024

Ramani, V. et al. Massively multiplex single-cell Hi-C. Nat. Methods 14, 263–266 (2017).

pubmed: 28135255 pmcid: 5330809 doi: 10.1038/nmeth.4155

Nagano, T. et al. Single-cell Hi-C for genome-wide detection of chromatin interactions that occur simultaneously in a single cell. Nat. Protoc. 10, 1986–2003 (2015).

pubmed: 26540590 doi: 10.1038/nprot.2015.127

Liu, Z. et al. Linking genome structures to functions by simultaneous single-cell Hi-C and RNA-seq. Science 380, 1070–1076 (2023).

pubmed: 37289875 doi: 10.1126/science.adg3797

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

pubmed: 28273065 pmcid: 5366070 doi: 10.1038/nature21411

Gross, D. S. & Garrard, W. T. Nuclease hypersensitive sites in chromatin. Annu. Rev. Biochem. 57, 159–197 (1988).

pubmed: 3052270 doi: 10.1146/annurev.bi.57.070188.001111

Thurman, R. E. et al. The accessible chromatin landscape of the human genome. Nature 489, 75–82 (2012).

pubmed: 22955617 pmcid: 3721348 doi: 10.1038/nature11232

Chen, X. et al. ATAC-see reveals the accessible genome by transposase-mediated imaging and sequencing. Nat. Methods 13, 1013–1020 (2016).

pubmed: 27749837 pmcid: 5509561 doi: 10.1038/nmeth.4031

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

pubmed: 21321607 pmcid: 3193420 doi: 10.1038/cr.2011.22

Hebbes, T. R., Thorne, A. W. & Crane-Robinson, C. A direct link between core histone acetylation and transcriptionally active chromatin. EMBO J. 7, 1395–1402 (1988).

pubmed: 3409869 pmcid: 458389 doi: 10.1002/j.1460-2075.1988.tb02956.x

Tessarz, P. & Kouzarides, T. Histone core modifications regulating nucleosome structure and dynamics. Nat. Rev. Mol. Cell Biol. 15, 703–708 (2014).

pubmed: 25315270 doi: 10.1038/nrm3890

Zhang, X. et al. The loss of heterochromatin is associated with multiscale three-dimensional genome reorganization and aberrant transcription during cellular senescence. Genome Res. 31, 1121–1135 (2021).

pubmed: 34140314 pmcid: 8256869 doi: 10.1101/gr.275235.121

Padeken, J., Methot, S. P. & Gasser, S. M. Establishment of H3K9-methylated heterochromatin and its functions in tissue differentiation and maintenance. Nat. Rev. Mol. Cell Biol. 23, 623–640 (2022).

pubmed: 35562425 pmcid: 9099300 doi: 10.1038/s41580-022-00483-w

Albert, I. et al. Translational and rotational settings of H2A.Z nucleosomes across the Saccharomyces cerevisiae genome. Nature 446, 572–576 (2007).

pubmed: 17392789 doi: 10.1038/nature05632

Kaya-Okur, H. S. et al. CUT&Tag for efficient epigenomic profiling of small samples and single cells. Nat. Commun. 10, 1930 (2019).

pubmed: 31036827 pmcid: 6488672 doi: 10.1038/s41467-019-09982-5

Skene, P. J. & Henikoff, S. An efficient targeted nuclease strategy for high-resolution mapping of DNA binding sites. Elife 6, e21856 (2017).

pubmed: 28079019 pmcid: 5310842 doi: 10.7554/eLife.21856

Fullwood, M. J. et al. An oestrogen-receptor-α-bound human chromatin interactome. Nature 462, 58–64 (2009).

pubmed: 19890323 pmcid: 2774924 doi: 10.1038/nature08497

Greil, F., Moorman, C. & van Steensel, B. DamID: mapping of in vivo protein-genome interactions using tethered DNA adenine methyltransferase. Methods Enzymol. 410, 342–359 (2006).

pubmed: 16938559 doi: 10.1016/S0076-6879(06)10016-6

Guelen, L. et al. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453, 948–951 (2008).

pubmed: 18463634 doi: 10.1038/nature06947

Bersaglieri, C. et al. Genome-wide maps of nucleolus interactions reveal distinct layers of repressive chromatin domains. Nat. Commun. 13, 1483 (2022).

pubmed: 35304483 pmcid: 8933459 doi: 10.1038/s41467-022-29146-2

Wu, W. et al. Mapping RNA-chromatin interactions by sequencing with iMARGI. Nat. Protoc. 14, 3243–3272 (2019).

pubmed: 31619811 pmcid: 7314528 doi: 10.1038/s41596-019-0229-4

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).

pubmed: 30154186 pmcid: 6219710 doi: 10.1083/jcb.201807108

Tsue, A. F. et al. Oligonucleotide-directed proximity-interactome mapping (O-MAP): a unified method for discovering RNA-interacting proteins, transcripts and genomic loci in situ. Preprint at bioRxiv https://doi.org/10.1101/2023.01.19.524825 (2023).

Belaghzal, H. et al. Liquid chromatin Hi-C characterizes compartment-dependent chromatin interaction dynamics. Nat. Genet. 53, 367–378 (2021).

pubmed: 33574602 pmcid: 7946813 doi: 10.1038/s41588-021-00784-4

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

pubmed: 24067610 doi: 10.1038/nature12593

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

pubmed: 28355183 pmcid: 5639698 doi: 10.1038/nature21711

Tan, L., Xing, D., Chang, C. H., Li, H. & Xie, X. S. Three-dimensional genome structures of single diploid human cells. Science 361, 924–928 (2018).

pubmed: 30166492 pmcid: 6360088 doi: 10.1126/science.aat5641

Zhang, C. et al. tagHi-C reveals 3D chromatin architecture dynamics during mouse hematopoiesis. Cell Rep. 32, 108206 (2020).

pubmed: 32997998 doi: 10.1016/j.celrep.2020.108206

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

pubmed: 28682332 pmcid: 5567812 doi: 10.1038/nature23001

Kirschenbaum, D. et al. Time-resolved single-cell transcriptomics defines immune trajectories in glioblastoma. Cell 187, 149–165.e23 (2024).

pubmed: 38134933 doi: 10.1016/j.cell.2023.11.032

Ma, H., Reyes-Gutierrez, P. & Pederson, T. Visualization of repetitive DNA sequences in human chromosomes with transcription activator-like effectors. Proc. Natl Acad. Sci. USA 110, 21048–21053 (2013).

pubmed: 24324157 pmcid: 3876203 doi: 10.1073/pnas.1319097110

Miyanari, Y., Ziegler-Birling, C. & Torres-Padilla, M. E. Live visualization of chromatin dynamics with fluorescent TALEs. Nat. Struct. Mol. Biol. 20, 1321–1324 (2013).

pubmed: 24096363 doi: 10.1038/nsmb.2680

Fu, Y. et al. CRISPR-dCas9 and sgRNA scaffolds enable dual-colour live imaging of satellite sequences and repeat-enriched individual loci. Nat. Commun. 7, 11707 (2016).

pubmed: 27222091 pmcid: 4894952 doi: 10.1038/ncomms11707

Chen, B. et al. Dynamic imaging of genomic loci in living human cells by an optimized CRISPR/Cas system. Cell 155, 1479–1491 (2013).

pubmed: 24360272 pmcid: 3918502 doi: 10.1016/j.cell.2013.12.001

Ma, H. et al. Multicolor CRISPR labeling of chromosomal loci in human cells. Proc. Natl Acad. Sci. USA 112, 3002–3007 (2015).

pubmed: 25713381 pmcid: 4364232 doi: 10.1073/pnas.1420024112

Ma, H. et al. Multiplexed labeling of genomic loci with dCas9 and engineered sgRNAs using CRISPRainbow. Nat. Biotechnol. 34, 528–530 (2016).

pubmed: 27088723 pmcid: 4864854 doi: 10.1038/nbt.3526

Chen, B. et al. Expanding the CRISPR imaging toolset with Staphylococcus aureus Cas9 for simultaneous imaging of multiple genomic loci. Nucleic Acids Res. 44, e75 (2016).

pubmed: 26740581 pmcid: 4856973 doi: 10.1093/nar/gkv1533

Gu, B. et al. Transcription-coupled changes in nuclear mobility of mammalian cis-regulatory elements. Science 359, 1050–1055 (2018).

pubmed: 29371426 pmcid: 6590518 doi: 10.1126/science.aao3136

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).

pubmed: 28828720 pmcid: 5676592 doi: 10.1007/s13238-017-0460-0

Tanenbaum, M. E., Gilbert, L. A., Qi, L. S., Weissman, J. S. & Vale, R. D. A protein-tagging system for signal amplification in gene expression and fluorescence imaging. Cell 159, 635–646 (2014).

pubmed: 25307933 pmcid: 4252608 doi: 10.1016/j.cell.2014.09.039

Janicki, S. M. et al. From silencing to gene expression: real-time analysis in single cells. Cell 116, 683–698 (2004).

pubmed: 15006351 pmcid: 4942132 doi: 10.1016/S0092-8674(04)00171-0

Shav-Tal, Y. et al. Dynamics of single mRNPs in nuclei of living cells. Science 304, 1797–1800 (2004).

pubmed: 15205532 pmcid: 4765737 doi: 10.1126/science.1099754

Park, H. Y. et al. Visualization of dynamics of single endogenous mRNA labeled in live mouse. Science 343, 422–424 (2014).

pubmed: 24458643 pmcid: 4111226 doi: 10.1126/science.1239200

Duan, J. et al. Live imaging and tracking of genome regions in CRISPR/dCas9 knock-in mice. Genome Biol. 19, 192 (2018).

pubmed: 30409154 pmcid: 6225728 doi: 10.1186/s13059-018-1530-1

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

pubmed: 23473598 pmcid: 3741673 doi: 10.1016/j.molcel.2013.02.011

Gilbert, N., Gilchrist, S. & Bickmore, W. A. Chromatin organization in the mammalian nucleus. Int. Rev. Cytol. 242, 283–336 (2005).

pubmed: 15598472 doi: 10.1016/S0074-7696(04)42007-5

Carter, C. W. Jr. Histone packing in the nucleosome core particle of chromatin. Proc. Natl Acad. Sci. USA 75, 3649–3653 (1978).

pubmed: 278980 pmcid: 392843 doi: 10.1073/pnas.75.8.3649

Stein, A., Bina-Stein, M. & Simpson, R. T. Crosslinked histone octamer as a model of the nucleosome core. Proc. Natl Acad. Sci. USA 74, 2780–2784 (1977).

pubmed: 197520 pmcid: 431287 doi: 10.1073/pnas.74.7.2780

Leffak, I. M., Grainger, R. & Weintraub, H. Conservative assembly and segregation of nucleosomal histones. Cell 12, 837–845 (1977).

pubmed: 562720 doi: 10.1016/0092-8674(77)90282-3

Oudet, P., Gross-Bellard, M. & Chambon, P. Electron microscopic and biochemical evidence that chromatin structure is a repeating unit. Cell 4, 281–300 (1975).

pubmed: 1122558 doi: 10.1016/0092-8674(75)90149-X

Axel, R., Melchior, W. Jr., Sollner-Webb, B. & Felsenfeld, G. Specific sites of interaction between histones and DNA in chromatin. Proc. Natl Acad. Sci. USA 71, 4101–4105 (1974).

pubmed: 4530287 pmcid: 434336 doi: 10.1073/pnas.71.10.4101

Kornberg, R. D. Chromatin structure: a repeating unit of histones and DNA. Science 184, 868–871 (1974).

pubmed: 4825889 doi: 10.1126/science.184.4139.868

Baldwin, J. P., Boseley, P. G., Bradbury, E. M. & Ibel, K. The subunit structure of the eukaryotic chromosome. Nature 253, 245–249 (1975).

pubmed: 1167623 doi: 10.1038/253245a0

Oosterhof, D. K., Hozier, J. C. & Rill, R. L. Nucleas action on chromatin: evidence for discrete, repeated nucleoprotein units along chromatin fibrils. Proc. Natl Acad. Sci. USA 72, 633–637 (1975).

pubmed: 1054845 pmcid: 432368 doi: 10.1073/pnas.72.2.633

Allan, J., Hartman, P. G., Crane-Robinson, C. & Aviles, F. X. The structure of histone H1 and its location in chromatin. Nature 288, 675–679 (1980).

pubmed: 7453800 doi: 10.1038/288675a0

Davidson, I. F. et al. DNA loop extrusion by human cohesin. Science 366, 1338–1345 (2019).

pubmed: 31753851 doi: 10.1126/science.aaz3418

Hao, X. et al. Super-resolution visualization and modeling of human chromosomal regions reveals cohesin-dependent loop structures. Genome Biol. 22, 150 (2021).

pubmed: 33975635 pmcid: 8111965 doi: 10.1186/s13059-021-02343-w

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).

pubmed: 26499245 pmcid: 4664323 doi: 10.1073/pnas.1518552112

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

pubmed: 22495300 pmcid: 3356448 doi: 10.1038/nature11082

Zhang, Y. et al. Spatial organization of the mouse genome and its role in recurrent chromosomal translocations. Cell 148, 908–921 (2012).

pubmed: 22341456 pmcid: 3320767 doi: 10.1016/j.cell.2012.02.002

Hafner, A. et al. Loop stacking organizes genome folding from TADs to chromosomes. Mol. Cell 83, 1377–1392 e1376 (2023).

pubmed: 37146570 pmcid: 10167645 doi: 10.1016/j.molcel.2023.04.008

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

pubmed: 27445307 pmcid: 4991974 doi: 10.1126/science.aaf8084

Falk, M. et al. Heterochromatin drives compartmentalization of inverted and conventional nuclei. Nature 570, 395–399 (2019).

pubmed: 31168090 pmcid: 7206897 doi: 10.1038/s41586-019-1275-3

Manuelidis, L. Individual interphase chromosome domains revealed by in situ hybridization. Hum. Genet. 71, 288–293 (1985).

pubmed: 3908288 doi: 10.1007/BF00388453

Lichter, P., Cremer, T., Borden, J., Manuelidis, L. & Ward, D. C. Delineation of individual human chromosomes in metaphase and interphase cells by in situ suppression hybridization using recombinant DNA libraries. Hum. Genet. 80, 224–234 (1988).

pubmed: 3192212 doi: 10.1007/BF01790090

Zink, D. et al. Structure and dynamics of human interphase chromosome territories in vivo. Hum. Genet. 102, 241–251 (1998).

pubmed: 9521598 doi: 10.1007/s004390050686

Cremer, T. & Cremer, C. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat. Rev. Genet. 2, 292–301 (2001).

pubmed: 11283701 doi: 10.1038/35066075

Gasser, S. M. Visualizing chromatin dynamics in interphase nuclei. Science 296, 1412–1416 (2002).

pubmed: 12029120 doi: 10.1126/science.1067703

Pickersgill, H. et al. Characterization of the Drosophila melanogaster genome at the nuclear lamina. Nat. Genet. 38, 1005–1014 (2006).

pubmed: 16878134 doi: 10.1038/ng1852

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).

pubmed: 18272965 doi: 10.1038/nature06727

Peric-Hupkes, D. et al. Molecular maps of the reorganization of genome-nuclear lamina interactions during differentiation. Mol. Cell 38, 603–613 (2010).

pubmed: 20513434 pmcid: 5975946 doi: 10.1016/j.molcel.2010.03.016

Peric-Hupkes, D. & van Steensel, B. Role of the nuclear lamina in genome organization and gene expression. Cold Spring Harb. Symp. Quant. Biol. 75, 517–524 (2010).

pubmed: 21209388 doi: 10.1101/sqb.2010.75.014

Pliss, A. et al. Spatio-temporal dynamics at rDNA foci: global switching between DNA replication and transcription. J. Cell Biochem. 94, 554–565 (2005).

pubmed: 15543556 doi: 10.1002/jcb.20317

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).

pubmed: 20826608 pmcid: 2965689 doi: 10.1091/mbc.e10-06-0508

Vertii, A. et al. Two contrasting classes of nucleolus-associated domains in mouse fibroblast heterochromatin. Genome Res. 29, 1235–1249 (2019).

pubmed: 31201210 pmcid: 6673712 doi: 10.1101/gr.247072.118

Spector, D. L. & Lamond, A. I. Nuclear speckles. Cold Spring Harb. Perspect. Biol. 3, a000646 (2011).

pubmed: 20926517 pmcid: 3039535 doi: 10.1101/cshperspect.a000646

Berry, J., Weber, S. C., Vaidya, N., Haataja, M. & Brangwynne, C. P. RNA transcription modulates phase transition-driven nuclear body assembly. Proc. Natl Acad. Sci. USA 112, E5237–E5245 (2015).

pubmed: 26351690 pmcid: 4586886 doi: 10.1073/pnas.1509317112

Feric, M. et al. Coexisting liquid phases underlie nucleolar subcompartments. Cell 165, 1686–1697 (2016).

pubmed: 27212236 pmcid: 5127388 doi: 10.1016/j.cell.2016.04.047

Riback, J. A. et al. Composition-dependent thermodynamics of intracellular phase separation. Nature 581, 209–214 (2020).

pubmed: 32405004 pmcid: 7733533 doi: 10.1038/s41586-020-2256-2

Osborne, C. S. et al. Active genes dynamically colocalize to shared sites of ongoing transcription. Nat. Genet. 36, 1065–1071 (2004).

pubmed: 15361872 doi: 10.1038/ng1423

Carter, K. C. et al. A three-dimensional view of precursor messenger RNA metabolism within the mammalian nucleus. Science 259, 1330–1335 (1993).

pubmed: 8446902 doi: 10.1126/science.8446902

Dillinger, S., Straub, T. & Nemeth, A. Nucleolus association of chromosomal domains is largely maintained in cellular senescence despite massive nuclear reorganisation. PLoS ONE 12, e0178821 (2017).

pubmed: 28575119 pmcid: 5456395 doi: 10.1371/journal.pone.0178821

Croft, J. A. et al. Differences in the localization and morphology of chromosomes in the human nucleus. J. Cell Biol. 145, 1119–1131 (1999).

pubmed: 10366586 pmcid: 2133153 doi: 10.1083/jcb.145.6.1119

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

pubmed: 28703188 doi: 10.1038/nature23263

Bonev, B. et al. Multiscale 3D genome rewiring during mouse neural development. Cell 171, 557–572.e24 (2017).

pubmed: 29053968 pmcid: 5651218 doi: 10.1016/j.cell.2017.09.043

Phanstiel, D. H. et al. Static and dynamic DNA loops form AP-1-bound activation hubs during macrophage development. Mol. Cell 67, 1037–1048.e6 (2017).

pubmed: 28890333 pmcid: 5610110 doi: 10.1016/j.molcel.2017.08.006

Zhang, Y. et al. Dynamic epigenomic landscapes during early lineage specification in mouse embryos. Nat. Genet. 50, 96–105 (2018).

pubmed: 29203909 doi: 10.1038/s41588-017-0003-x

Guan, Y. et al. Senescence-activated enhancer landscape orchestrates the senescence-associated secretory phenotype in murine fibroblasts. Nucleic Acids Res. 48, 10909–10923 (2020).

pubmed: 33045748 pmcid: 7641768 doi: 10.1093/nar/gkaa858

Tomimatsu, K. et al. Locus-specific induction of gene expression from heterochromatin loci during cellular senescence. Nat. Aging 2, 31–45 (2022).

pubmed: 37118356 doi: 10.1038/s43587-021-00147-y

Zirkel, A. et al. HMGB2 loss upon senescence entry disrupts genomic organization and induces CTCF clustering across cell types. Mol. Cell 70, 730–744.e6 (2018).

pubmed: 29706538 doi: 10.1016/j.molcel.2018.03.030

Liu, G. H. et al. Recapitulation of premature ageing with iPSCs from Hutchinson–Gilford progeria syndrome. Nature 472, 221–225 (2011).

pubmed: 21346760 pmcid: 3088088 doi: 10.1038/nature09879

Tavares-Cadete, F., Norouzi, D., Dekker, B., Liu, Y. & Dekker, J. Multi-contact 3C reveals that the human genome during interphase is largely not entangled. Nat. Struct. Mol. Biol. 27, 1105–1114 (2020).

pubmed: 32929283 pmcid: 7718335 doi: 10.1038/s41594-020-0506-5

Hansen, R. S. et al. Sequencing newly replicated DNA reveals widespread plasticity in human replication timing. Proc. Natl Acad. Sci. USA 107, 139–144 (2010).

pubmed: 19966280 doi: 10.1073/pnas.0912402107

Liang, Z. et al. Chromosomes progress to metaphase in multiple discrete steps via global compaction/expansion cycles. Cell 161, 1124–1137 (2015).

pubmed: 26000485 pmcid: 4448932 doi: 10.1016/j.cell.2015.04.030

Kschonsak, M. et al. Structural basis for a safety-belt mechanism that anchors condensin to chromosomes. Cell 171, 588–600 e524 (2017).

pubmed: 28988770 pmcid: 5651216 doi: 10.1016/j.cell.2017.09.008

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

pubmed: 29348367 pmcid: 5924687 doi: 10.1126/science.aao6135

Golfier, S., Quail, T., Kimura, H. & Brugues, J. Cohesin and condensin extrude DNA loops in a cell cycle-dependent manner. Elife 9, e53885 (2020).

pubmed: 32396063 pmcid: 7316503 doi: 10.7554/eLife.53885

Martinez-Balbas, M. A., Dey, A., Rabindran, S. K., Ozato, K. & Wu, C. Displacement of sequence-specific transcription factors from mitotic chromatin. Cell 83, 29–38 (1995).

pubmed: 7553870 doi: 10.1016/0092-8674(95)90231-7

Palozola, K. C. et al. Mitotic transcription and waves of gene reactivation during mitotic exit. Science 358, 119–122 (2017).

pubmed: 28912132 pmcid: 5727891 doi: 10.1126/science.aal4671

van Schaik, T., Vos, M., Peric-Hupkes, D., Hn Celie, P. & van Steensel, B. Cell cycle dynamics of lamina-associated DNA. EMBO Rep. 21, e50636 (2020).

pubmed: 32893442 pmcid: 7645246 doi: 10.15252/embr.202050636

Caravaca, J. M. et al. Bookmarking by specific and nonspecific binding of FoxA1 pioneer factor to mitotic chromosomes. Genes. Dev. 27, 251–260 (2013).

pubmed: 23355396 pmcid: 3576511 doi: 10.1101/gad.206458.112

Deluz, C. et al. A role for mitotic bookmarking of SOX2 in pluripotency and differentiation. Genes. Dev. 30, 2538–2550 (2016).

pubmed: 27920086 pmcid: 5159668 doi: 10.1101/gad.289256.116

Festuccia, N. et al. Mitotic binding of Esrrb marks key regulatory regions of the pluripotency network. Nat. Cell Biol. 18, 1139–1148 (2016).

pubmed: 27723719 doi: 10.1038/ncb3418

Raccaud, M. et al. Mitotic chromosome binding predicts transcription factor properties in interphase. Nat. Commun. 10, 487 (2019).

pubmed: 30700703 pmcid: 6353955 doi: 10.1038/s41467-019-08417-5

Young, D. W. et al. Mitotic retention of gene expression patterns by the cell fate-determining transcription factor Runx2. Proc. Natl Acad. Sci. USA 104, 3189–3194 (2007).

pubmed: 17360627 pmcid: 1805558 doi: 10.1073/pnas.0611419104

Abramo, K. et al. A chromosome folding intermediate at the condensin-to-cohesin transition during telophase. Nat. Cell Biol. 21, 1393–1402 (2019).

pubmed: 31685986 pmcid: 6858582 doi: 10.1038/s41556-019-0406-2

Zhang, H. et al. Chromatin structure dynamics during the mitosis-to-G1 phase transition. Nature 576, 158–162 (2019).

pubmed: 31776509 pmcid: 6895436 doi: 10.1038/s41586-019-1778-y

Paul, J. & Gilmour, R. S. Organ-specific restriction of transcription in mammalian chromatin. J. Mol. Biol. 34, 305–316 (1968).

pubmed: 5761390 doi: 10.1016/0022-2836(68)90255-6

Balsalobre, A. & Drouin, J. Pioneer factors as master regulators of the epigenome and cell fate. Nat. Rev. Mol. Cell Biol. 23, 449–464 (2022).

pubmed: 35264768 doi: 10.1038/s41580-022-00464-z

Iwafuchi-Doi, M. & Zaret, K. S. Cell fate control by pioneer transcription factors. Development 143, 1833–1837 (2016).

pubmed: 27246709 pmcid: 6514407 doi: 10.1242/dev.133900

Vos, S. M. Understanding transcription across scales: from base pairs to chromosomes. Mol. Cell 81, 1601–1616 (2021).

pubmed: 33770487 doi: 10.1016/j.molcel.2021.03.002

Consortium, E. P. An integrated encyclopedia of DNA elements in the human genome. Nature 489, 57–74 (2012).

doi: 10.1038/nature11247

Buenrostro, J. D., Giresi, P. G., Zaba, L. C., Chang, H. Y. & Greenleaf, W. J. Transposition of native chromatin for fast and sensitive epigenomic profiling of open chromatin, DNA-binding proteins and nucleosome position. Nat. Methods 10, 1213–1218 (2013).

pubmed: 24097267 pmcid: 3959825 doi: 10.1038/nmeth.2688

Neph, S. et al. An expansive human regulatory lexicon encoded in transcription factor footprints. Nature 489, 83–90 (2012).

pubmed: 22955618 pmcid: 3736582 doi: 10.1038/nature11212

Bulger, M. & Groudine, M. Functional and mechanistic diversity of distal transcription enhancers. Cell 144, 327–339 (2011).

pubmed: 21295696 pmcid: 3742076 doi: 10.1016/j.cell.2011.01.024

Ghirlando, R. et al. Chromatin domains, insulators, and the regulation of gene expression. Biochim. Biophys. Acta 1819, 644–651 (2012).

pubmed: 22326678 pmcid: 3484685 doi: 10.1016/j.bbagrm.2012.01.016

Wu, C. The 5’ ends of Drosophila heat shock genes in chromatin are hypersensitive to DNase I. Nature 286, 854–860 (1980).

pubmed: 6774262 doi: 10.1038/286854a0

Keene, M. A., Corces, V., Lowenhaupt, K. & Elgin, S. C. DNase I hypersensitive sites in Drosophila chromatin occur at the 5’ ends of regions of transcription. Proc. Natl Acad. Sci. USA 78, 143–146 (1981).

pubmed: 6264428 pmcid: 319007 doi: 10.1073/pnas.78.1.143

Shermoen, A. W. & Beckendorf, S. K. A complex of interacting DNAase I-hypersensitive sites near the Drosophila glue protein gene, Sgs4. Cell 29, 601–607 (1982).

pubmed: 6214314 doi: 10.1016/0092-8674(82)90176-3

Muskavitch, M. A. & Hogness, D. S. An expandable gene that encodes a Drosophila glue protein is not expressed in variants lacking remote upstream sequences. Cell 29, 1041–1051 (1982).

pubmed: 6817924 doi: 10.1016/0092-8674(82)90467-6

McGinnis, W., Shermoen, A. W., Heemskerk, J. & Beckendorf, S. K. DNA sequence changes in an upstream DNase I-hypersensitive region are correlated with reduced gene expression. Proc. Natl Acad. Sci. USA 80, 1063–1067 (1983).

pubmed: 6405379 pmcid: 393528 doi: 10.1073/pnas.80.4.1063

McGinnis, W., Shermoen, A. W. & Beckendorf, S. K. A transposable element inserted just 5’ to a Drosophila glue protein gene alters gene expression and chromatin structure. Cell 34, 75–84 (1983).

pubmed: 6309414 doi: 10.1016/0092-8674(83)90137-X

Ptashne, M. Gene regulation by proteins acting nearby and at a distance. Nature 322, 697–701 (1986).

pubmed: 3018583 doi: 10.1038/322697a0

Narlikar, G. J., Fan, H. Y. & Kingston, R. E. Cooperation between complexes that regulate chromatin structure and transcription. Cell 108, 475–487 (2002).

pubmed: 11909519 doi: 10.1016/S0092-8674(02)00654-2

Gorkin, D. U. et al. An atlas of dynamic chromatin landscapes in mouse fetal development. Nature 583, 744–751 (2020).

pubmed: 32728240 pmcid: 7398618 doi: 10.1038/s41586-020-2093-3

Banerji, J., Rusconi, S. & Schaffner, W. Expression of a β-globin gene is enhanced by remote SV40 DNA sequences. Cell 27, 299–308 (1981).

pubmed: 6277502 doi: 10.1016/0092-8674(81)90413-X

Bulger, M. & Groudine, M. Looping versus linking: toward a model for long-distance gene activation. Genes. Dev. 13, 2465–2477 (1999).

pubmed: 10521391 doi: 10.1101/gad.13.19.2465

Cullen, K. E., Kladde, M. P. & Seyfred, M. A. Interaction between transcription regulatory regions of prolactin chromatin. Science 261, 203–206 (1993).

pubmed: 8327891 doi: 10.1126/science.8327891

Morgan, G. T. Imaging the dynamics of transcription loops in living chromosomes. Chromosoma 127, 361–374 (2018).

pubmed: 29610944 pmcid: 6096578 doi: 10.1007/s00412-018-0667-8

Chen, H. et al. Dynamic interplay between enhancer-promoter topology and gene activity. Nat. Genet. 50, 1296–1303 (2018).

pubmed: 30038397 pmcid: 6119122 doi: 10.1038/s41588-018-0175-z

Deng, W. et al. Controlling long-range genomic interactions at a native locus by targeted tethering of a looping factor. Cell 149, 1233–1244 (2012).

pubmed: 22682246 pmcid: 3372860 doi: 10.1016/j.cell.2012.03.051

Bartman, C. R. et al. Transcriptional burst initiation and polymerase pause release are key control points of transcriptional regulation. Mol. Cell 73, 519–532.e4 (2019).

pubmed: 30554946 doi: 10.1016/j.molcel.2018.11.004

Jerabek, H. & Heermann, D. W. Expression-dependent folding of interphase chromatin. PLoS ONE 7, e37525 (2012).

pubmed: 22649534 pmcid: 3359300 doi: 10.1371/journal.pone.0037525

Racko, D., Benedetti, F., Dorier, J. & Stasiak, A. Transcription-induced supercoiling as the driving force of chromatin loop extrusion during formation of TADs in interphase chromosomes. Nucleic Acids Res. 46, 1648–1660 (2018).

pubmed: 29140466 doi: 10.1093/nar/gkx1123

Calandrelli, R. et al. Genome-wide analysis of the interplay between chromatin-associated RNA and 3D genome organization in human cells. Nat. Commun. 14, 6519 (2023).

pubmed: 37845234 pmcid: 10579264 doi: 10.1038/s41467-023-42274-7

Zhang, S., Ubelmesser, N., Barbieri, M. & Papantonis, A. Enhancer-promoter contact formation requires RNAPII and antagonizes loop extrusion. Nat. Genet. 55, 832–840 (2023).

pubmed: 37012454 doi: 10.1038/s41588-023-01364-4

Kagey, M. H. et al. Mediator and cohesin connect gene expression and chromatin architecture. Nature 467, 430–435 (2010).

pubmed: 20720539 pmcid: 2953795 doi: 10.1038/nature09380

Richter, W. F., Nayak, S., Iwasa, J. & Taatjes, D. J. The mediator complex as a master regulator of transcription by RNA polymerase II. Nat. Rev. Mol. Cell Biol. 23, 732–749 (2022).

pubmed: 35725906 pmcid: 9207880 doi: 10.1038/s41580-022-00498-3

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

pubmed: 29472443 pmcid: 6329450 doi: 10.1126/science.aar7831

Haarhuis, J. H. I. et al. The cohesin release factor WAPL restricts chromatin loop extension. Cell 169, 693–707 e614 (2017).

pubmed: 28475897 pmcid: 5422210 doi: 10.1016/j.cell.2017.04.013

Winick-Ng, W. et al. Cell-type specialization is encoded by specific chromatin topologies. Nature 599, 684–691 (2021).

pubmed: 34789882 pmcid: 8612935 doi: 10.1038/s41586-021-04081-2

Rao, S. S. P. et al. Cohesin loss eliminates all loop domains. Cell 171, 305–320 e324 (2017).

pubmed: 28985562 pmcid: 5846482 doi: 10.1016/j.cell.2017.09.026

Nora, E. P. et al. Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169, 930–944.e22 (2017).

pubmed: 28525758 pmcid: 5538188 doi: 10.1016/j.cell.2017.05.004

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

pubmed: 29094699 pmcid: 5687303 doi: 10.1038/nature24281

Gong, Y. et al. Stratification of TAD boundaries reveals preferential insulation of super-enhancers by strong boundaries. Nat. Commun. 9, 542 (2018).

pubmed: 29416042 pmcid: 5803259 doi: 10.1038/s41467-018-03017-1

Nanni, L., Ceri, S. & Logie, C. Spatial patterns of CTCF sites define the anatomy of TADs and their boundaries. Genome Biol. 21, 197 (2020).

pubmed: 32782014 pmcid: 7422557 doi: 10.1186/s13059-020-02108-x

Thiecke, M. J. et al. Cohesin-dependent and -independent mechanisms mediate chromosomal contacts between promoters and enhancers. Cell Rep. 32, 107929 (2020).

pubmed: 32698000 pmcid: 7383238 doi: 10.1016/j.celrep.2020.107929

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

pubmed: 28424523 pmcid: 6080695 doi: 10.1038/nature22063

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

pubmed: 30262496 pmcid: 6986801 doi: 10.1126/science.aau0320

Wang, D. et al. Reprogramming transcription by distinct classes of enhancers functionally defined by eRNA. Nature 474, 390–394 (2011).

pubmed: 21572438 pmcid: 3117022 doi: 10.1038/nature10006

Kim, T. K. et al. Widespread transcription at neuronal activity-regulated enhancers. Nature 465, 182–187 (2010).

pubmed: 20393465 pmcid: 3020079 doi: 10.1038/nature09033

De Santa, F. et al. A large fraction of extragenic RNA pol II transcription sites overlap enhancers. PLoS Biol. 8, e1000384 (2010).

pubmed: 20485488 pmcid: 2867938 doi: 10.1371/journal.pbio.1000384

Zoller, B., Little, S. C. & Gregor, T. Diverse spatial expression patterns emerge from unified kinetics of transcriptional bursting. Cell 175, 835–847 e825 (2018).

pubmed: 30340044 pmcid: 6779125 doi: 10.1016/j.cell.2018.09.056

Rodriguez, J. et al. Intrinsic dynamics of a human gene reveal the basis of expression heterogeneity. Cell 176, 213–226.e18 (2019).

pubmed: 30554876 doi: 10.1016/j.cell.2018.11.026

Hakim, O. et al. Diverse gene reprogramming events occur in the same spatial clusters of distal regulatory elements. Genome Res. 21, 697–706 (2011).

pubmed: 21471403 pmcid: 3083086 doi: 10.1101/gr.111153.110

D’Ippolito, A. M. et al. Pre-established chromatin interactions mediate the genomic response to glucocorticoids. Cell Syst. 7, 146–160.e7 (2018).

pubmed: 30031775 doi: 10.1016/j.cels.2018.06.007

Alexander, J. M. et al. Live-cell imaging reveals enhancer-dependent Sox2 transcription in the absence of enhancer proximity. Elife 8, e41769 (2019).

pubmed: 31124784 pmcid: 6534382 doi: 10.7554/eLife.41769

Fukaya, T., Lim, B. & Levine, M. Enhancer control of transcriptional bursting. Cell 166, 358–368 (2016).

pubmed: 27293191 pmcid: 4970759 doi: 10.1016/j.cell.2016.05.025

Goel, V. Y., Huseyin, M. K. & Hansen, A. S. Region capture micro-C reveals coalescence of enhancers and promoters into nested microcompartments. Nat. Genet. 55, 1048–1056 (2023).

pubmed: 37157000 pmcid: 10424778 doi: 10.1038/s41588-023-01391-1

Kawasaki, K. & Fukaya, T. Functional coordination between transcription factor clustering and gene activity. Mol. Cell 83, 1605–1622 e1609 (2023).

pubmed: 37207625 doi: 10.1016/j.molcel.2023.04.018

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

pubmed: 29930094 pmcid: 6543815 doi: 10.1126/science.aar4199

Stadhouders, R., Filion, G. J. & Graf, T. Transcription factors and 3D genome conformation in cell-fate decisions. Nature 569, 345–354 (2019).

pubmed: 31092938 doi: 10.1038/s41586-019-1182-7

Cardozo Gizzi, A. M. A shift in paradigms: spatial genomics approaches to reveal single-cell principles of genome organization. Front. Genet. 12, 780822 (2021).

pubmed: 34868269 pmcid: 8640135 doi: 10.3389/fgene.2021.780822

Finn, E. H. & Misteli, T. Molecular basis and biological function of variability in spatial genome organization. Science 365, eaaw9498 (2019).

pubmed: 31488662 pmcid: 7421438 doi: 10.1126/science.aaw9498

Bintu, B. et al. Super-resolution chromatin tracing reveals domains and cooperative interactions in single cells. Science 362, eaau1783 (2018).

pubmed: 30361340 pmcid: 6535145 doi: 10.1126/science.aau1783

Bintu, L. et al. Dynamics of epigenetic regulation at the single-cell level. Science 351, 720–724 (2016).

pubmed: 26912859 pmcid: 5108652 doi: 10.1126/science.aab2956

Vierstra, J. et al. Mouse regulatory DNA landscapes reveal global principles of cis-regulatory evolution. Science 346, 1007–1012 (2014).

pubmed: 25411453 pmcid: 4337786 doi: 10.1126/science.1246426

Cheng, Y. et al. Principles of regulatory information conservation between mouse and human. Nature 515, 371–375 (2014).

pubmed: 25409826 pmcid: 4343047 doi: 10.1038/nature13985

Stergachis, A. B. et al. Conservation of trans-acting circuitry during mammalian regulatory evolution. Nature 515, 365–370 (2014).

pubmed: 25409825 pmcid: 4405208 doi: 10.1038/nature13972

Franke, M. et al. Formation of new chromatin domains determines pathogenicity of genomic duplications. Nature 538, 265–269 (2016).

pubmed: 27706140 doi: 10.1038/nature19800

Lupianez, D. G. et al. Disruptions of topological chromatin domains cause pathogenic rewiring of gene-enhancer interactions. Cell 161, 1012–1025 (2015).

pubmed: 25959774 pmcid: 4791538 doi: 10.1016/j.cell.2015.04.004

Martinez, M. F., Martini, A. G., Sequeira-Lopez, M. L. S. & Gomez, R. A. Ctcf is required for renin expression and maintenance of the structural integrity of the kidney. Clin. Sci. 134, 1763–1774 (2020).

doi: 10.1042/CS20200184

Christov, M. et al. Inducible podocyte-specific deletion of CTCF drives progressive kidney disease and bone abnormalities. JCI Insight 3, e95091 (2018).

pubmed: 29467330 pmcid: 5916242 doi: 10.1172/jci.insight.95091

Alharbi, A. B. et al. Ctcf haploinsufficiency mediates intron retention in a tissue-specific manner. RNA Biol. 18, 93–103 (2021).

pubmed: 32816606 doi: 10.1080/15476286.2020.1796052

Moisan, S. et al. Novel long-range regulatory mechanisms controlling PKD2 gene expression. BMC Genomics 19, 515 (2018).

pubmed: 29986647 pmcid: 6038307 doi: 10.1186/s12864-018-4892-6

Livingston, S. et al. Cux1 regulation of the cyclin kinase inhibitor p27

pubmed: 34531000 doi: 10.1016/j.gene.2019.100007

Wang, H. et al. Glucocorticoid receptor wields chromatin interactions to tune transcription for cytoskeleton stabilization in podocytes. Commun. Biol. 4, 675 (2021).

pubmed: 34083716 pmcid: 8175753 doi: 10.1038/s42003-021-02209-8

Luo, Z. et al. NicE-C efficiently reveals open chromatin-associated chromosome interactions at high resolution. Genome Res. 32, 534–544 (2022).

pubmed: 35105668 pmcid: 8896462 doi: 10.1101/gr.275986.121

Yeung, J. et al. Transcription factor activity rhythms and tissue-specific chromatin interactions explain circadian gene expression across organs. Genome Res. 28, 182–191 (2018).

pubmed: 29254942 pmcid: 5793782 doi: 10.1101/gr.222430.117

Mermet, J. et al. Clock-dependent chromatin topology modulates circadian transcription and behavior. Genes. Dev. 32, 347–358 (2018).

pubmed: 29572261 pmcid: 5900709 doi: 10.1101/gad.312397.118

Zhang, Y. et al. Transcriptionally active HERV-H retrotransposons demarcate topologically associating domains in human pluripotent stem cells. Nat. Genet. 51, 1380–1388 (2019).

pubmed: 31427791 pmcid: 6722002 doi: 10.1038/s41588-019-0479-7

Dhillon, P. et al. Increased levels of endogenous retroviruses trigger fibroinflammation and play a role in kidney disease development. Nat. Commun. 14, 559 (2023).

pubmed: 36732547 pmcid: 9895454 doi: 10.1038/s41467-023-36212-w

Siebenthall, K. T. et al. Integrated epigenomic profiling reveals endogenous retrovirus reactivation in renal cell carcinoma. EBioMedicine 41, 427–442 (2019).

pubmed: 30827930 pmcid: 6441874 doi: 10.1016/j.ebiom.2019.01.063

Gillies, C. E. et al. An eQTL landscape of kidney tissue in human nephrotic syndrome. Am. J. Hum. Genet. 103, 232–244 (2018).

pubmed: 30057032 pmcid: 6081280 doi: 10.1016/j.ajhg.2018.07.004

Ko, Y. A. et al. Genetic-variation-driven gene-expression changes highlight genes with important functions for kidney disease. Am. J. Hum. Genet. 100, 940–953 (2017).

pubmed: 28575649 pmcid: 5473735 doi: 10.1016/j.ajhg.2017.05.004

Qiu, C. et al. Renal compartment-specific genetic variation analyses identify new pathways in chronic kidney disease. Nat. Med. 24, 1721–1731 (2018).

pubmed: 30275566 pmcid: 6301011 doi: 10.1038/s41591-018-0194-4

Gorkin, D. U. et al. Common DNA sequence variation influences 3-dimensional conformation of the human genome. Genome Biol. 20, 255 (2019).

pubmed: 31779666 pmcid: 6883528 doi: 10.1186/s13059-019-1855-4

Chandra, V. et al. Promoter-interacting expression quantitative trait loci are enriched for functional genetic variants. Nat. Genet. 53, 110–119 (2021).

pubmed: 33349701 doi: 10.1038/s41588-020-00745-3

Duan, A. et al. Chromatin architecture reveals cell type-specific target genes for kidney disease risk variants. BMC Biol. 19, 38 (2021).

pubmed: 33627123 pmcid: 7905576 doi: 10.1186/s12915-021-00977-7

Sieber, K. B. et al. Integrated functional genomic analysis enables annotation of kidney genome-wide association study loci. J. Am. Soc. Nephrol. 30, 421–441 (2019).

pubmed: 30760496 pmcid: 6405142 doi: 10.1681/ASN.2018030309

Maurano, M. T. et al. Large-scale identification of sequence variants influencing human transcription factor occupancy in vivo. Nat. Genet. 47, 1393–1401 (2015).

pubmed: 26502339 pmcid: 4666772 doi: 10.1038/ng.3432

Javierre, B. M. et al. Lineage-specific genome architecture links enhancers and non-coding disease variants to target gene promoters. Cell 167, 1369–1384.e19 (2016).

pubmed: 27863249 pmcid: 5123897 doi: 10.1016/j.cell.2016.09.037

Muto, Y. et al. Single cell transcriptional and chromatin accessibility profiling redefine cellular heterogeneity in the adult human kidney. Nat. Commun. 12, 2190 (2021).

pubmed: 33850129 pmcid: 8044133 doi: 10.1038/s41467-021-22368-w

Haug, S. et al. Multi-omic analysis of human kidney tissue identified medulla-specific gene expression patterns. Kidney Int. 105, 293–311 (2024).

pubmed: 37995909 doi: 10.1016/j.kint.2023.10.024

Claussnitzer, M. et al. FTO obesity variant circuitry and adipocyte browning in humans. N. Engl. J. Med. 373, 895–907 (2015).

pubmed: 26287746 pmcid: 4959911 doi: 10.1056/NEJMoa1502214

Mukhi, D. et al. ACSS2 gene variants determine kidney disease risk by controlling de novo lipogenesis in kidney tubules. J. Clin. Invest. 134, e172963 (2023).

pubmed: 38051585 pmcid: 10866669 doi: 10.1172/JCI172963

Mumbach, M. R. et al. Enhancer connectome in primary human cells identifies target genes of disease-associated DNA elements. Nat. Genet. 49, 1602–1612 (2017).

pubmed: 28945252 pmcid: 5805393 doi: 10.1038/ng.3963

Lidberg, K. A. et al. Serum protein exposure activates a core regulatory program driving human proximal tubule injury. J. Am. Soc. Nephrol. 33, 949–965 (2022).

pubmed: 35197326 pmcid: 9063895 doi: 10.1681/ASN.2021060751

Gisch, D. L. et al. The chromatin landscape of healthy and injured cell types in the human kidney. Nat. Commun. 15, 433 (2024).

pubmed: 38199997 pmcid: 10781985 doi: 10.1038/s41467-023-44467-6

Eun, M. et al. Chromatin accessibility analysis and architectural profiling of human kidneys reveal key cell types and a regulator of diabetic kidney disease. Kidney Int. 105, 150–164 (2024).

pubmed: 37925023 doi: 10.1016/j.kint.2023.09.030

Combes, A. N. et al. Haploinsufficiency for the Six2 gene increases nephron progenitor proliferation promoting branching and nephron number. Kidney Int. 93, 589–598 (2018).

pubmed: 29217079 doi: 10.1016/j.kint.2017.09.015

Georgas, K. et al. Analysis of early nephron patterning reveals a role for distal RV proliferation in fusion to the ureteric tip via a cap mesenchyme-derived connecting segment. Dev. Biol. 332, 273–286 (2009).

pubmed: 19501082 doi: 10.1016/j.ydbio.2009.05.578

Kobayashi, A. et al. Six2 defines and regulates a multipotent self-renewing nephron progenitor population throughout mammalian kidney development. Cell Stem Cell 3, 169–181 (2008).

pubmed: 18682239 pmcid: 2561900 doi: 10.1016/j.stem.2008.05.020

Perl, A. J. et al. Reduced nephron endowment in Six2-TGCtg mice is due to Six3 misexpression by aberrant enhancer-promoter interactions in the transgene. J. Am. Soc. Nephrol. 35, 566–577 (2024).

pubmed: 38447671 doi: 10.1681/ASN.0000000000000324

O’Brien, L. L. et al. Transcriptional regulatory control of mammalian nephron progenitors revealed by multi-factor cistromic analysis and genetic studies. PLoS Genet. 14, e1007181 (2018).

pubmed: 29377931 pmcid: 5805373 doi: 10.1371/journal.pgen.1007181

Ichimura, T. et al. Kidney injury molecule-1 (KIM-1), a putative epithelial cell adhesion molecule containing a novel immunoglobulin domain, is up-regulated in renal cells after injury. J. Biol. Chem. 273, 4135–4142 (1998).

pubmed: 9461608 doi: 10.1074/jbc.273.7.4135

Zhang, Z., Humphreys, B. D. & Bonventre, J. V. Shedding of the urinary biomarker kidney injury molecule-1 (KIM-1) is regulated by MAP kinases and juxtamembrane region. J. Am. Soc. Nephrol. 18, 2704–2714 (2007).

pubmed: 17898236 doi: 10.1681/ASN.2007030325

Wilson, P. C. et al. Mosaic loss of Y chromosome is associated with aging and epithelial injury in chronic kidney disease. Genome Biol. 25, 36 (2024).

pubmed: 38287344 pmcid: 10823641 doi: 10.1186/s13059-024-03173-2

Langelueddecke, C. et al. Lipocalin-2 (24p3/neutrophil gelatinase-associated lipocalin (NGAL)) receptor is expressed in distal nephron and mediates protein endocytosis. J. Biol. Chem. 287, 159–169 (2012).

pubmed: 22084236 doi: 10.1074/jbc.M111.308296

Chen, X. et al. Mapping disease regulatory circuits at cell-type resolution from single-cell multiomics data. Nat. Comput. Sci. 3, 644–657 (2023).

pubmed: 37974651 pmcid: 10653299 doi: 10.1038/s43588-023-00476-5

Wang, R. et al. SARS-CoV-2 restructures host chromatin architecture. Nat. Microbiol. 8, 679–694 (2023).

pubmed: 36959507 pmcid: 10116496 doi: 10.1038/s41564-023-01344-8

Chiariello, A. M. et al. Multiscale modelling of chromatin 4D organization in SARS-CoV-2 infected cells. Nat. Commun. 15, 4014 (2024).

pubmed: 38740770 pmcid: 11091192 doi: 10.1038/s41467-024-48370-6

Akilesh, S. et al. Multicenter clinicopathologic correlation of kidney biopsies performed in COVID-19 patients presenting with acute kidney injury or proteinuria. Am. J. Kidney Dis. 77, 82–93.e1 (2021).

pubmed: 33045255 doi: 10.1053/j.ajkd.2020.10.001

Nicosia, R. F., Ligresti, G., Caporarello, N., Akilesh, S. & Ribatti, D. COVID-19 vasculopathy: mounting evidence for an indirect mechanism of endothelial injury. Am. J. Pathol. 191, 1374–1384 (2021).

pubmed: 34033751 pmcid: 8141344 doi: 10.1016/j.ajpath.2021.05.007

Smith, K. D. & Akilesh, S. Pathogenesis of coronavirus disease 2019-associated kidney injury. Curr. Opin. Nephrol. Hypertens. 30, 324–331 (2021).

pubmed: 33767060 doi: 10.1097/MNH.0000000000000708

Davis, M. A. et al. A C57BL/6 mouse model of SARS-CoV-2 infection recapitulates age- and sex-based differences in human COVID-19 disease and recovery. Vaccines https://doi.org/10.3390/vaccines11010047 (2022).

Zazhytska, M. et al. Non-cell-autonomous disruption of nuclear architecture as a potential cause of COVID-19-induced anosmia. Cell 185, 1052–1064.e12 (2022).

pubmed: 35180380 pmcid: 8808699 doi: 10.1016/j.cell.2022.01.024

Zaidman, N. A. & Pluznick, J. L. Understudied G protein-coupled receptors in the kidney. Nephron 146, 278–281 (2022).

pubmed: 34261071 doi: 10.1159/000517355

Halperin Kuhns, V. L. et al. Characterizing novel olfactory receptors expressed in the murine renal cortex. Am. J. Physiol. Renal Physiol. 317, F172–F186 (2019).

pubmed: 31042061 pmcid: 6692720 doi: 10.1152/ajprenal.00624.2018

Shepard, B. D., Koepsell, H. & Pluznick, J. L. Renal olfactory receptor 1393 contributes to the progression of type 2 diabetes in a diet-induced obesity model. Am. J. Physiol. Renal Physiol. 316, F372–F381 (2019).

pubmed: 30484350 doi: 10.1152/ajprenal.00069.2018

Pluznick, J. L. et al. Functional expression of the olfactory signaling system in the kidney. Proc. Natl Acad. Sci. USA 106, 2059–2064 (2009).

pubmed: 19174512 pmcid: 2644163 doi: 10.1073/pnas.0812859106

Correa-Costa, M. et al. Transcriptome analysis of renal ischemia/reperfusion injury and its modulation by ischemic pre-conditioning or hemin treatment. PLoS ONE 7, e49569 (2012).

pubmed: 23166714 pmcid: 3498198 doi: 10.1371/journal.pone.0049569

A guide to studying 3D genome structure and dynamics in the kidney.

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Brian J Beliveau (BJ)

Shreeram Akilesh (S)

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