{"paper_id":"a299faeb-ac52-4248-bbe8-e35a0de7775c","body_text":"1 \n \nT\nitle: Quantitative imaging of loop extruders  \nrebuilding interphase genome architecture after mitosis \n \nTitle (short): Analysis of genome refolding during mitotic exit \n \nAuthors: Andreas Brunner1,2, Natalia Rosalía Morero1, Wanlu Zhang1, M. Julius Hossain3, Marko \nLampe4, Hannah Pflaumer1, Aliaksandr Halavatyi4, Jan-Michael Peters5, Kai S. Beckwith1,6, Jan \nEllenberg1 \n \nAffiliations: \n1 Cell Biology and Biophysics Unit, European Molecular Biology Laboratory (EMBL), 69117 Heidelberg, \nGermany \n2 Collaboration for Joint PhD Degree between EMBL and Heidelberg University, Faculty of \nBiosciences, Heidelberg, Germany \n3 Centre for Cancer Immunology, University of Southampton, SO17 1BJ Southampton, United \nKingdom \n4 Advanced Light Microscopy Facility, European Molecular Biology Laboratory (EMBL), 69117 \nHeidelberg, Germany \n5 Research Institute of Molecular Pathology, Vienna BioCenter 1030 Vienna, Austria \n6 Department of Biomedical Laboratory Science, Norwegian University of Science and Technology \n(NTNU), N-7491 Trondheim, Norway \n \nSummary: \nHow cells establish the interphase genome organiza tion after mitosis is incompletely understood. \nUsing quantitative and super-resolution microscopy, we show that the transition from a Condensin to \na Cohesin-based genome organization occurs dynamically over two hours. While a significant fraction \nof Condensins remains chromatin-bound until early G1, Cohesin-STAG1 and its boundary factor CTCF \nare rapidly imported into daughter nuclei in telophase, immediately bind chromosomes as individual \ncomplexes and are sufficient to build the first in terphase TAD structures. By contrast, the more \nabundant Cohesin-STAG2 accumulates on chromosomes only gradually later in G1, is responsible for \ncompaction inside TAD structures and forms paire d complexes upon compl eted nuclear import. Our \nquantitative time-resolved mapping of mitotic and interphase loop extruders in single cells reveals \nthat the nested loop architecture formed by sequ ential action of two Condensins in mitosis is \nseamlessly replaced by a less compact, but conceptually similar hierarchically nested loop architecture \ndriven by sequential action of two Cohesins.   \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n2 \n \nI\nntroduction \nDNA loop extrusion by SMC complexes (structural maintenance of chromosomes) has emerged as a \nkey principle in the spatial organization of chromosomes during interphase and mitosis (Yatskevich et \nal., 2019; Davidson & Peters, 2021). In mitosis, the two pentameric ring-like Condensin complexes I & \nII, consisting of two shared coiled-coil subunits (SMC2 and SMC4) and three isoform-specific subunits \n(the kleisin CAP-H or CAP-H2 and two HAWK proteins CAP-D2/3 and CAP-G/2, Hirano & Mitchison, \n1994; Hirano et al., 1997) have been shown to be capable of processive DNA loop extrusion (Ganji et \nal., 2018). Both Condensin I, activated through mitotic phosphorylation and KIF4A (Kimura et al., 1998; \nBazile et al., 2010; Tane et al., 2022; Cutts et al., 2024 Preprint), and Condensin II, deactivated during \ninterphase by MCPH1 (Houlard et al., 2021) and as sociating with chromosomes through M18BP1 in \nmitosis (Borsellini et al., 2024 Preprint), localize to the longitudinal axis of mitotic chromosomes (Ono \net al., 2003; Hirota et al., 2004). Condensin I & II impact the shape of mitotic chromosomes distinctly, \nwith Condensin II compacting chromosomes axially from prophase onward, and Condensin I \ncompacting chromosomes laterally once it gains access to DNA during prometaphase (Ono et al., 2003, \n2004; Hirota et al., 2004; Shintomi & Hirano, 2011; Green et al., 2012). Through their sequential action, \nthe Condensins shape mitotic chromosomes into rod-shaped entities and provide mechanical rigidity \n(Houlard et al., 2015) to ensure the faithful segregation of sister chromatids by spindle forces. Based \non quantitative and super-resolution imaging, as we ll as HiC and polymer modelling, it has recently \nbeen proposed that Condensins organize mitotic chromosomes into  nested loops, with the less \nabundant and stably binding Condensin II extruding big DNA loops (~450 kb) already during prophase \nthat are subsequently nested into smaller sub-loops (~90 kb) by the more abundant and more \ndynamically associating Condensin I complex after nuclear envelope breakdown (Walther et al., 2018; \nGibcus et al., 2018). These Condensin-driven loops are randomly generated across the linear \nchromosomal DNA molecules, thereby erasing sequence  specific interphase structures (Naumova et \nal., 2013). \nIn interphase, the two closely related Cohesin co mplexes Cohesin-STAG1 and Cohesin-STAG2 govern \nthe loop extruder-based genome organization (Wutz et al ., 2017, 2020). Like  the Condensins, the \nCohesins are ring-like protein complexes consisting of two shared coiled-coil subunits (SMC1 and \nSMC3), a shared kleisin subunit (RAD21, also called SCC1) and one isoform-specific HEAT-repeat \nsubunit (STAG1 or STAG2, Losada et al., 1998, 2000 ; Sumara et al., 2000). In the presence of the \naccessory HEAT repeat protein NIPBL, Cohesin complexes can extrude DNA loops (Kim et al., 2019; \nDavidson et al., 2019) until they are being stalle d by the protein CTCF binding to the conserved \nessential surface of STAG1/2 (Li et al., 2020). CTCF is a zink-finger containing protein that is enriched \nat its asymmetric cognate binding sites in the genome (de Wit et al., 2015; Guo et al., 2015), yielding \nmost efficient stalling of loop ex truding Cohesin when arranged in a convergent orientation (Rao et \nal., 2014). The protein WAPL functions as an un-loader of Cohesin on chromatin, restricting its maximal \nresidence time on chromatin and thereby achieving a constant turnover of DNA loops (Kueng et al., \n2006; Wutz et al., 2017). The combined action of these proteins leads to the continuous and dynamic \ngeneration of sequence specifically positioned DNA loops in the genome (Rao et al., 2014; Sanborn et \nal., 2015; Fudenberg et al., 2016; Gabriele et al., 2022; Mach et al, 2022; Beckwith et al., 2023 \nPreprint), thereby creating more comp act domains in the genome termed topologically associated \ndomains (TADs, Nora et al., 2012; Dixon et al., 2012). While their functional role is still an active area \nof research, TADs have been implicated in the regulation of gene expression through active regulation \nof enhancer promoter contact frequency (Lupiáñez et al., 2015; Zuin et al., 2022).  \nSimilar to the two Condensin isoforms, the Cohesi n isoforms STAG1/2 display different expression \nlevels and chromatin residence times, with Cohe sin-STAG1 being the less abundant subunit with a \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n3 \n \nl\nong residence time, and Cohesin-STAG2 making up 75% of the total Cohesin pool and being more \ndynamically bound to chromatin (Losada et al., 2000; Holzmann et al., 2019; Wutz et al., 2020). While \nthe two isoforms share a large portion of common binding sites in the genome and display a certain \nfunctional redundancy in the generation of DNA loops, Cohesin-STAG1/2 also have unique binding \nsites, with Cohesin-STAG1 being preferentially enriched at CTCF binding sites and TAD boundaries, and \nCohesin-STAG2 being enriched at non-CTCF sites (Kojic et al., 2018).  \nWhile the bona-fide interphase organization and the formation of mitotic chromosomes have been \nsubject to thorough investigation, much less is known about how the interphase organization is rebuilt \nafter mitosis. Previously, the genome-wide reorga nization of chromatin has been studied using a \ncombination of HiC and ChiP-seq in cell populations fixed after pharmacological synchronization in a \nlong mitotic arrest. This revealed a slow and gradual transition of the mitotic to the interphase fold \nover the course of several hours, via an apparently unstructured folding intermediate during telophase \nthat is devoid of Condensin and Cohesin loop extruders, as well as a gradual build-up of TAD structures \nover the course of several hours during G1 (Abramo et al., 2019; Zhang et al., 2019).  \nHere, we set out to systematically quantify and map the actions of the Condensin and Cohesin loop \nextrusion machinery during  mitotic exit in single living cells,  aiming to characterize the dynamic \nmolecular processes underlying the reformation of  the loop-extrusion governed interphase genome \norganization after mitosis. We find that the switch from mitotic to interphase organization takes about \n2 hours in unsynchronized cells, passing a transition state during telophase during which a minimal set \nof 3 Condensins and Cohesins each are simultaneously bound per megabase of genomic DNA. We find \nthat Cohesin-STAG1 is rapidly imported into the newly formed daughter nuclei alongside CTCF, \ncapable of the formation of large, TAD-scale, loop s early after mitosis as a monomer. We find that \nCohesin-STAG2 likely also extrudes DNA loops as a monomer, but that it undergoes a concentration-\ndependent dimerization on chromatin upon its full import into the nucleus. Based on our quantitative \nimaging data, we can infer that this phenomenon is a result of the high occupancy of 8 chromatin-\nbound Cohesin-STAG2 per megabase in late G1, leading to frequent encounters of neighboring \ncomplexes that lead to a nested/stacked arrangement of extruded loops. Surprisingly, we also find \nthat CTCF is increasingly stabilized on chromatin throughout G1 due to its increasing interaction with \nthe two Cohesin complexes. Based on these data, we propose a double-hierarchical loop model to \ngenerate interphase genome architecture after mito sis, in which the two interphase Cohesin loop \nextruders sequentially build a nested arrangement of large and then small DNA loops, conceptually \nsimilar to how Condensins have been suggested to drive mitotic genomic organization. \n  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n4 \n \nR\nesults \nThe transition from mitotic to interphase lo o p extruders occurs over two hours after \nmitosis and requires nuclear import \nTo examine the time required to complete the sw i tch from mitotic to interphase loop extruder \ngenome organization (Fig. 1A), we made use of human HeLa Kyoto (HK) homozygous knock-in cell lines \nin which all alleles of the endogenous genes for the kleisin subunits of Condensin I (NCAPH), \nCondensin II (NCAPH2) and the HEAT-repeat subunits of Cohe sin-STAG1 (STAG1) and Cohesin-STAG2 \n(STAG2) have been tagged with GF P (Walther et al., 2018; Cai et al ., 2018). After a single S-phase \nsynchronization, we performed continuous FCS-cali brated 4D live-cell imaging (Politi et al., 2018, \nFig. 1B, Suppl. Fig. 1A) through two subsequent cell divisions with 10-minute time-resolution, using \nSiR-Hoechst and extracellular Dextran to label nucl ear and cell volumes, respectively (Fig. 1C). \nComputational 3D segmentation of these cellular landmarks (Cai et al., 2018, Suppl. Fig. 1B), combined \nwith automatic cell tracking allowed us to align single cell trajectories from one anaphase to the next, \nand calculate absolute protein concentrations and copy numbers throughout a full cell cycle (Fig. 1D, \nSuppl. Fig. 1C-G).  \nAs expected, both Condensin isoforms were concen trated on mitotic chromosomes and Condensin II \nmaintained a stable nuclear concentration after division (Fig. 1D). Surprisingly, we found that while \nthe high Condensin I concentration of 380 nM on chromosomes dropped sharply after segregation, it \ndid not become completely cytoplasmic but maintain ed a concentration of 150 nM in the two newly \nformed interphase nuclei, where it then became diluted slowly with nuclear growth (Fig. 1D). \nPhotobleaching of this nuclear Condensin I pool in  interphase revealed that it moves freely in the \nnucleus and does not exchange with the cytosolic pool  (Suppl. Fig. 1H). Quantitative full cell cycle \nimaging showed that the nuclear pools of NCAP H/2 could in principle form complete Condensin \ncomplexes with the shared Condensin subunit SMC4, which is present inside the nucleus in sufficient \nnumbers (Suppl. Fig. 1E).  \nConversely to the sharp reduction in chromosomal Condensin I, the two Cohesin isoforms STAG1/2 \nthat are key for interphase genome organization be came enriched inside the nucleus after anaphase \nand reached essentially constant nuclear concentrations throughout the entire interphase (Fig. 1D). \nThis was also found to be true for CTCF (Suppl . Fig. 1E), revealing homeostatic stable nuclear \nconcentrations of these factors and no doubling of interphase loop extruders with DNA replication, \nconsistent with previous reports that showed uncoupling of nuclear growth from DNA replication \n(Otsuka et al., 2016). We found that the Cohesi n isoform STAG1 only makes up 25% of the total \nCohesin pool, consistent with previous studies (Fig. 1D, Losada et al., 2000; Holzmann et al., 2019). \nInterestingly, Cohesin-STAG1 displayed rapid and complete nuclear localization shortly after mitosis, \nfollowed by an equilibration of its nuclear concentrat ion upon nuclear expansion (Fig. 1D, Suppl. Fig. \n1D). The 3-fold more abundant Cohesin-STAG2, however, reached stable nuclear concentrations only \nabout two hours after mitosis (Fig. 1D, Suppl. Fig. 1I-N).  \nHaving characterized the chromosomal/nuclear concentration changes of the four loop extruders and \nCTCF throughout the cell cycle, we next focused our analysis on the transition between Condensin and \nCohesin occupancy on the genome during the first 2 hours after mitosis, increasing the time-resolution \nof our FCS-calibrated 4D imaging to 2 minutes (Fig. 1E). This detailed kinetic analysis revealed that the \nnumber of Condensin I proteins associating with chromatin rapidly dropped after its peak during \nanaphase (Fig. 1F). This drop, however, ceased at the time of reformation of the nuclear envelope, \n6 minutes after AO (Suppl. Fig 1O), that creates a permeability barrier and apparently retains the \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n5 \n \nr\nemaining Condensin I molecules inside the newly formed nucleus. Cohesin-STAG2 started its nuclear \nenrichment precisely from the ti me of nuclear envelope assembly, and required 2 hours until \ncomplete nuclear enrichment (Fig. 1F, Suppl. Fig. 1N). To test if the Cohesin accumulation required \nnuclear import, we acutely degraded degron knocke d-in Nup153, an essential component of nuclear \npore complexes (Fig. 1G, Suppl. Fig. 1Q-S). We fo und that both Cohesin and CTCF levels inside the \nnucleus were significantly reduced in Nup153 depleted  cells early after mitosis (Fig. 1G-I, Suppl. Fig. \n1T), showing that both factors require functional nuclear pores to reach the genome. \n \nCondensins and Cohesins bind simultaneousl y, yet independently, to the early G1 \ngenome at 3 complexes per megabase DNA \nG\niven that a significant number of both Condensin complexes are still present inside the newly formed \ndaughter cell nuclei when Cohesins start to be impo rted (Fig. 1F, Table 1), we wanted to go beyond \nnuclear concentration and protein numbers and ask how much of the mitotic and interphase loop \nextruding complexes are bound to chromatin after mitosis and could thus be actively engaged in \nextrusion. To quantify binding, we used fluorescence recovery after photobleaching (FRAP, Suppl. Fig. \n2A) of Condensin I and II on the metaphase plate and in the newly formed nucleus. Half nuclear \nphotobleaching indicated that a significant fraction  of Condensins remains chromatin-bound in early \nG1 (Suppl. Fig. 2 B-D), while at the same time a large fraction of the newly imported Cohesins are \nalready bound (Suppl. Fig. 2F). To assay changes in the chromatin-bound fraction of Condensins and \nCohesins quantitatively and in a highly time-resolved manner during mitotic exit, we used a rapid spot-\nbleach assay monitoring fluorescence depletion from a femtoliter-sized chromatin volume during 30 \nsecond continuous illumination with a diffraction li mited focused laser beam (Fig. 2A, Suppl. Fig. 2G-\nI). In this assay, the chromatin-bound protein fraction is bleached, while the unbound fraction recovers \nfrom the excess soluble nuclear p ool outside the small bleach spot. This approach thus provides a \nrapid measure for the bound fraction of GFP-tagged proteins on chromatin that can be carried out \nrepeatedly in a single living cell without interfering with mitotic progression (see Supplementary Table \n1&2 for more detail and comparison to classical FRAP).  \nWe then used this assay to monitor changes in chromatin-binding of Condensin and Cohesin every 5 \nminutes after exit from mitosis. We found that while all Condensins (using the isoform-shared subunit \nSMC4-mEGFP) progressively dissociated from chromatin during telophase and early G1, they retained \na significant chromatin bound fraction of around 25% 15 minutes after AO (Fig. 2B). This reduction in \nbound fraction, also following nuclear envelope reformation, was consistent for both Condensin \nisoforms, as shown by time-resolved spot bleaching using isoform-specific NCAPH and NCAPH2 \nsubunits (Suppl. Fig. 2K). By contrast, we found that the fraction of bound Cohesins (using isoform \nshared subunit RAD21-EGFP) increases continuously following nuclear envelope reformation (Fig. 2B), \nreaching a bound fraction of about 40% 15 min after AO. Again, this increase in binding was consistent \nfor both Cohesin isoforms (using isoform specific STAG1/2 subunits (Suppl. Fig. 2K). \nOur quantitative real time analysis of chromatin binding in single dividing cells provides clear evidence \nfor co-occupancy of chromatin by Condensin and Cohesin complexes throughout telophase and \nearly G1. Combining the bound fraction measurements by FRAP with the protein numbers measured \nby FCS-calibrated imaging (e.g. Fig. 1F, Suppl. Fig. 1P) allows us to calculate the number of proteins \nbound to genomic DNA (Fig. 2C, Suppl. Fig. 2L). This analysis shows that in early G1, 15 minutes after \nAO, the same number of around three Condensin and Cohesin complexes are simultaneously bound \nper megabase of genomic DNA (Fig. 2C). Could this simultaneous binding of mitotic and interphase \nloop extruders be functionally interlinked? To test this, we probed if the chromatin localization of \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n6 \n \nC\nondensins and Cohesins in early G1 depends on each other’s presence, using AID-degron knock-in \ncell lines for the isoform-shared Condensin subuni t SMC4 (Schneider et al., 2022) and the Cohesin-\nchromatin-loader NIPBL (Mitter et al ., 2020). In these cells, we could acutely degrade the degron \ntagged proteins during mitosis (Suppl. Fig. 2J), and ask if they are required for the other complex to \nassociate with chromatin by subsequent immunofluorescence staining for the non-degraded \nCondensin or Cohesin complex. This analysis did not show major differences in chromatin association \nof Condensin after NIPBL or Cohesin after SMC4 de pletion, respectively (Fig . 2D&E). This suggested \nthat while mitotic and interphase loop extruders bind to chromatin simultaneously in very similar \nnumbers during G1, they do so independently.  \n \nCohesin-STAG1 and CTCF are simultaneously imported immediately after mitosis and \nsufficient to build the first interphase hallmarks in genome structure \nO\nur full cell cycle data showed clear differences in the time required for complete nuclear import of \nthe two Cohesin isoforms, with STAG1 reaching maxi mal nuclear concentration within only 10 mins, \nwhile STAG2 reached steady state only after over two hours (Fig. 1D, Suppl. Fig. 1N). To get a first \ninsight into which complex might functionally be more important for early G1 genome architecture, \nwe compared these kinetics with the boundary factor CTCF using an endogenous CTCF-EGFP knock-in \ncell line (Cai et al., 2018). Calibrated full cell cycle imaging showed a strikingly similar kinetic signature \nof its nuclear concentration changes compared to  Cohesin-STAG1, reaching an approximately 2.5 \ntimes higher steady state concen tration in interphase (Suppl. Fig 1E). We therefore compared the \nnuclear import kinetics of Cohesin-STAG1 and CTCF  relative to the slower accumulating Cohesin-\nSTAG2 with high time-resolution after mitotic ex it, using our FCS-calibrated 4D imaging setup. \nStrikingly, we found that CTCF displayed indistinguishable import kinetics as Cohesin-STAG1 while \nCohesin-STAG2 was imported at a much lower rate (Fig. 3A, Suppl. Fig. 3A).  \nThe simultaneous import of Cohesin STAG1 and CTCF is consistent with a functional interaction on \nchromatin immediately after nuclear reformation. To test if the two proteins are bound to chromatin, \nwe performed real time spot-bleach, as well as FRAP measurements of CTCF, to compare its binding \nto chromatin with Cohesin STAG1 early after mitotic exit (Suppl. Fig. 2E&L). This analysis revealed that \nwhen Cohesin-STAG1 and CTCF reach their maximum concentration, about 2 Cohesin-STAG1 and 5 \nCTCF molecules are bound per megabase of genomic DNA (Table 1). Two actively extruding Cohesin-\nSTAG1 complexes per megabase of genomic DNA would in principle explain the frequency of compact \ntopologically associated domain (TAD) structures that have been estimated at 1.5 TADs/Mb using \nbiochemical approaches previously (Wutz et al., 2020 ). To test directly, if Cohesin STAG1 without \nCohesin STAG2 is indeed sufficient to create the first more compactly folded G1 genome structures in \nsingle cells, we took advantage of our recently developed nanoscale DNA tracing method LoopTrace, \nenabling us to inspect individual 3D DNA folds as well as ensemble averages  with precise physical \ndistance measures (Beckwith et al., 2023 Preprint, Fig. 3B, Suppl. Fig. 3B). We traced three \nindependent 1.2 megabase long genomic regions pred icted to contain TADs, in 3D at 12 kb genomic \nand 20 nm spatial resolution (Fig. 3C, Suppl. Fig. 3E&F). Our single cell DNA traces could indeed readily \nidentify compact 3D DNA folds already in single early G1 cells (Fig. 3D). Depletion of Cohesin-STAG2 \nduring the prior mitosis (Suppl. Fig. 3B&C) did not influence the overall genomic size of these domains, \nbut led to some reduction in internal loop nesting and slight physical decompaction (Fig. 3E, Suppl. \nFig. 3D), which was also clear when comparing pairwise physical 3D distance maps of these regions \nfrom hundreds of control or STAG2 depleted cells (Fig. 3F, Suppl. Fig. 3G&H). We conclude that \nCohesin-STAG1 and CTCF are imported with identical kinetics rapidly after mitosis and are sufficient \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n7 \n \nt\no build the first compact looped interphase structures  in single G1 cells, equivalent to biochemically \ndetected TADs in cell populations.  \n \nCohesin-STAG1 and CTCF bec ome increasingly stably bound to the genome \nthroughout G1 \nT\no investigate the interplay of Cohesin-STAG1 and Cohesin-STAG2 at later times after mitosis, we \nperformed FRAP measurements during G1 (2-5h pa st AO) and compared them to our measurements \nshortly after mitosis (Fig. 4A). We found that the chromatin-bound fraction for both Cohesin isoforms \nas well as CTCF significantly increased in later G1 (Suppl. Fig. 4A). A single exponential function with \nan immobile fraction fit the fluorescence equilibration kinetics of all proteins well (Fig. 4B) and allowed \nus to determine the dynamically chromatin-bound protein fraction, its residence time, as well as the \nstably bound fraction that did not exchange dynamically during our measurement time (Suppl. Fig. \n4B&C, Tables 1&2). While the average residence time of the dynamically bound pool of Cohesin \nisoforms (STAG1: 4 min, STAG2: 2 min and CTCF: 2 min) remained unchanged from early to late G1 \n(Suppl. Fig. 4B), we measured a significant increase in the stably chromatin-bound fraction for \nCohesin-STAG1 and CTCF, reaching up 30-40% of the total protein (Fig. 4C&E, Suppl. Fig. 4C). Cohesin-\nSTAG2 also displayed a significant increase in its stably chromatin-bound fraction, however reaching \nless than 10% of the total protein pool (Fig. 4D). While it has been previously reported that Cohesin-\nSTAG1 chromatin binding can be stabilized by CTCF (Wutz et al., 2020), whether CTCF’s own binding \nis reciprocally affected by the presence of Cohe sin has not been investigated. To test if CTCF’s \nincreasingly stable binding in G1 depends on Cohesin, we acutely depleted the isoform-shared subunit \nRAD21 (Suppl. Fig. 4D-G), which resulted in a significant reduction of stably chromatin-bound CTCF, \nwhich could be rescued by RAD21 overexpression (Fig. 4F). This shows that Cohesin is necessary and \nsufficient to stabilize CTCF’s interaction with chromatin in G1. \n \nCohesin-STAG2 completes  its nuclear import after 2 hours and exhibits concentration \ndependent dimerization on the genome in G1 \nT\no test directly whether the observed interdependent  increase in stable binding of both CTCF and \nCohesins is due to increased complex formation between these proteins on chromatin, we performed \nSTED super-resolution imaging of CTCF and the Cohesin isoform specific subunits STAG1 and STAG2 \nduring early and late G1. To achieve high and comparable labeling efficiency of the different Cohesin \nisoforms, we used our homozygous knock-in cell li nes for STAG1/2-EGFP and detected both with the \nsame GFP-nanobody, while using a specific antibody to detect endogenous CTCF as a reference (Fig. \n5A&B, Suppl. Fig. 5A-C). Having calculated the number of chroma tin complexes from our combined \nconcentration imaging and FRAP data allowed us to  estimate the labeling efficiency of our super-\nresolution imaging by counting the individual labeled fluorescent spots to about 60-80% for the two \nCohesin-isoforms (Suppl. Fig. 5D-F), very similar to our previous labeling efficiencies of this GFP \nnanobody (Thevathasan et al., 2019). Given that we could resolve the expected number of Cohesin \ncomplexes as individual fluorescent spots, the large majority of the labelled STAG1/2 proteins in early \nG1 therefore most likely represent monomeric Cohesin complexes.  \nColocalization of either STAG1 or STAG2 with CTCF resulted in comparable spatial correlations that in \nboth cases increased slightly but significantly from early to late G1, indicating an increase in Cohesin-\nCTCF complex formation for both isoforms (Fig. 5C).  This increased colocalization was not due to the \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n8 \n \ns\ntill ongoing accumulation of STAG2 in the nucleus, as shown by image simulations with random \nprotein distributions at realistic densities (Suppl. Fig. 5H&I). Our data thus suggests that CTCF \nassociates with both STAG1, that enters the nucleus early, and STAG2, that completes its import later \nand eventually becomes the more abundant Cohesin isoform. This finding is also in line with the fact \nthat also Cohesin-STAG2 becomes more stably bound to chromatin in late G1 (Fig. 4D).  \nThe ability to detect both Cohesin isoforms at the single complex level with high labeling efficiency in \nearly G1 also put us in a position to use the in tensity of Cohesin spots to ask if we can detect \nmultimerization of the Cohesins as the cell cycl e progresses, which would be expected with the \nformation of closely stacked or nested loops. Inte restingly, this analysis revealed that while the \naverage spot intensity and total number of spots detected did not change for Cohesin-STAG1 between \nearly and late G1, the STAG2 spot intensity increased about 2-fold between early and late G1 (Fig. 5D, \nSuppl. Fig. 5G) and correlated with a 2-fold drop in the number of STAG2 spots detected compared to \nour expectation from quantitative live imaging (Sup pl. Fig. 5F). When it has reached its maximum \nconcentration in late G1, Cohesin-STAG2 thus appe ars to associate with the genome in pairs of \nmolecules that are no longer resolvable individually  by STED microscopy, that has a lateral precision \nof around 60 nm. With an estimated extension of single Cohesin complexes of 50 nm, they must \ntherefore be very closely adjacent to each other or form dimers to result in single STED spots with \ndoubled intensity.  \nWhy might Cohesin-STAG2 form closely adjacent pairs of  complexes only at the end of G1? To test if \nits self-association is concentration dependent, we performed partial depletion of degron tagged \nCohesin-STAG2. This indeed shifted the average spot brightness in late G1 back to a value of nanobody \nmonomers (preliminary data, not shown), supporting a concentration rather than for example a cell \ncycle driven dimerization of Cohesin-STAG2 on DNA. In fact, our quantitative imaging data of the \nincreasing numbers of chromatin-bound Cohesins after mitosis provides a quantitative explanation \nfor the concentration-dependent dimerization of Cohesin-STAG2. During early G1, we found three \nCohesin-STAG2 molecules to be on average bound for 120 seconds per megabase of DNA. Assuming \nthey extrude loops with the estimated rate of 1 kb/s (Kim et al., 2019; Davidson et al., 2019), they \nwould form 120 kb large loops and would thus be relatively unlikely to encounter each other within \none megabase (Table 1). In late G1 however, about 8 Cohesin-STAG2 complexes are bound per \nmegabase with a similar residence time (Table 2), making Cohesin-STAG2 encounters between eight \n120 kb sized loops within one megabase much more likely. The fact that we observe a quantitative \nshift in the intensities of the Cohesin-STAG2 spot distribution from early to late G1 (Fig. 5D) in fact \nsuggests that Cohesin complexes not only encounter each other transiently, but potentially stay \nassociated with each other when they meet, which would induce stacking and nesting of loops. To test \nif such nested and stacked loops indeed form in late G1 in single cells in a Cohesin-STAG2 dependent \nmanner, we again made use of our nanoscale DNA tracing of interphase cells targeting the same three \n1.2 megabase TAD regions as before. The 3D folds of  these regions indeed revealed stronger nesting \nand stronger compaction of these regions compared to early G1, and again showed that this is largely \ndependent on Cohesin-STAG2 (Fig. 5E-H, Suppl. Fig. 5J-O). In conclusion, due to its continuous nuclear \nimport, Cohesin-STAG2 crosses a critical occupancy threshold on the genome within the first 1-2 hours \nafter mitosis that leads to a high probability of encounters between Cohesin-STAG2 complexes, \naccompanied by increased formation of nested loops inside TAD-scale compact domains of the \ninterphase genome.  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n9 \n \nA double hierarchical loop model  quantitatively explains the transition from mitotic to \ninterphase loop extruder driven genome organization \nOur systematic, quantitative time-resolved mapping o f mitotic and interphase loop extruders in single \ncells shows that the interphase genome is sequentially organized into compact TAD-scale regions \nwhich then compact further by internal stacking of ne sted loops. This is highly reminiscent of the \npreviously proposed nested loop organization in mitosis (Walther et al., 2018), in which chromosomes \nare organized by the sequential action of two Cond ensin complexes (Walther et al., 2018; Gibcus et \nal., 2018). In this model of establishing mitotic architecture, the less abundant Condensin II loop \nextrusion motor first forms large DNA loops during prophase that become subsequently nested by the \nmore abundant Condensin I, once it gains access to chromosomes during prometaphase.  \nOur study now shows, that following chromosome segregation and nuclear envelope reformation, \nsome Condensins are still bound to chromatin, while Cohesins and CTCF are rapidly imported into the \nnewly formed nucleus, leading to a co-occupancy of the genome by 3 Condensins and 3 Cohesins per \nmegabase of DNA in early G1. Very interestingly, wh en the interphase loop extruders Cohesins start \nbinding the genome, they do so independently of Condensins, but like Condensins during mitotic entry \nalso in a sequential manner during mitotic exit. First, the rapid and synchronous nuclear import of \nCohesin-STAG1 and CTCF (completed within 10 mi nutes after AO) and their immediate chromatin-\nbinding at relatively low abundance (3 complexes bound per megabase) with a long residence time (4 \nminutes) builds up the first compact interphase stru ctures even in the absence of Cohesin-STAG2. In \na second step, the slowly imported Cohesin-STAG2 (complete only 2 hours after mitosis) then binds in \nhigher abundance (8 complexes bound per megabase) and with a shorter residence time (2 minutes), \nleading to the generation of many smaller loops (~120 kb), and frequent encounters and likely stalling \nwith neighboring Cohesin-STAG2 complexes, leading to stacking of nested loops inside the larger \nSTAG1 defined domains. We therefore propose a double hierarchical loop model for the transition \nfrom mitotic to interphase loop extruder driven genome architecture, in which the Condensin-based, \nrandomly positioned nested loop architecture establ ished during mitotic entry is replaced by a less \ncompact, but conceptually similar Cohesin-driven nested loop architecture, positioned by CTCF, from \nmitotic exit to early G1 (Fig. 6). \n  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n10 \n \nD\niscussion \nThis study provides comprehensive quantitative an d time-resolved data on the chromatin-binding of \nCondensins and Cohesins throughout mitotic exit and G1. In addition to the new data it provides, it \nfurthermore allows to integrate many previous more  qualitative and individual observations into an \noverall, internally consistent and quantitative  model of how the loop extruder-based genome \norganization is handed over from mitosis to interphase.  \nA role for chromatin-bound Condensin during telophase? \nThe Condensin-driven mitotic chromosome organization, previously proposed to be best explained by \nan axial arrangement of nested DNA loops (Gibcus et al., 2018; Walther et al., 2018), is rapidly lost \nduring telophase when 75% of Condensins unbind DNA. Consistent with a previous report (Abramo et \nal., 2019), we find that this rapid removal of Cond ensins is followed by import of Cohesin and CTCF \ninto the newly forming nuclei, leading to a co-occupancy of only 3 Condensins and Cohesins per \nmegabase during early G1, which we show is the lowest number of genome-associated loop extrusion \ncomplexes at any time during the cell cycle. Nonetheless a significant fraction of Condensins remains \nchromatin bound during telophase and early G1, leading to a so far unappreciated pool of Condensin I \nto be retained in the nucleus during interphase. However, in interphase nuclear Condensin I is unlikely \nto be actively engaged in processive loop extrusion due to its mitosis-specific loading onto \nchromosomes (Hirano et al., 1997) regulated via ph osphorylation of the NCAPH N-terminal tail (Tane \net al., 2022) and mitotic activation by KIF4A (Cutts et al., 2024). However, it could be that the retained \nfraction of Condensins has a transient role during telophase and early G1 to facilitate the removal of \nintra-chromosomal catenations as suggested recently (Hildebrand et al., 2024).  \nThe two Cohesin isoforms bind sequentially and likely have different structural roles \nConsistent with previous systems analysis of mitotic protein networks (Cai et al., 2018), we found that \nCohesin-STAG1 and CTCF are rapidly imported into the newly formed daughter cell nuclei after cell \ndivision and are sufficient in numbers and loop extrusion processivity to form the first interphase TAD-\nscale loops shortly after mitosis. By contrast, we found that the more abundant second Cohesin \nisoform STAG2 is imported slowly over the course of 2 hours and is dispensable for the generation of \nthese TAD-scale compact structures in early G1 as well as later in interphase. Due to its later binding, \nits higher abundance and its short residence time on  chromatin, Cohesin-STAG2 leads to shorter and \nnested loops within the already established larger Cohesin-STAG1 loops. We speculate that this \nSTAG2-dependent highly dynamic sub-structuring of the more stable TAD loops could promote cell-\ntype-specific intra-TAD contacts between enhancers and promoters independently of CTCF, explaining \ncell-type dependent effects of Cohesin-STAG2 mutations or depletion (Kojic et al., 2018; Viny et al., \n2019). While we found that the overall fraction of chromatin-bound proteins increases for both \nCohesin isoforms and CTCF similarly from early to later G1, Cohesin-STAG1 and CTCF became \nspecifically stabilized on chromatin during G1. Our observations are consistent with recent reports \nthat Cohesin-STAG1 is stabilized through CTCF binding and acetylation of its SMC3 subunit by Esco1 \n(Wutz et al., 2020), suggesting a continued role of Cohesin-STAG1 in the generation and maintenance \nof long-range loops during interphase that are further sub-structured by Cohesin-STAG2.  \nChromatin binding of CTCF is stabilized by Cohesin \nOur FRAP analysis early af ter mitosis and later during G1 enabled us to see a clear increase in the \nfraction of stably chromatin bound CTCF. Interestingly, this stabilization was dependent on the \npresence on Cohesin, and occurred progressivel y throughout G1 when we found both Cohesin \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n11 \n \ni\nsoforms to increasingly co-localize with CTCF. It is therefore likely that CTCF’s stabilization is due to \ninteraction with one or both chromatin-bound Cohesin isoforms, which may serve as additional \nanchors at CTCF-sites. Combined with the fact that  Cohesin-STAG1 is preferentially associating with \nCTCF (Kojic et al., 2018) and is stabilized in part through CTCF (Wutz et al., 2020) our data suggests \nthat Cohesin-STAG1 and CTCF mutually stabilize each other on chromatin, which may be important to \nstabilize longer lived loop structures in interphase, equivalent of TADs.  \nThe oligomerization state of Cohesin \nUsing structured illumination mi croscopy of the isoform-shared Cohesin subunit RAD21, it was \nrecently reported that the majority of the loop-ext ruding Cohesin is present in dimers or multimers \n(Ochs et al., 2024). Consistent with this finding, our STED super-resolution imaging of isoform-specific \nCohesin subunits revealed that the less abundant Co hesin-STAG1 is present as a monomer, but that \nthe more abundant isoform Cohesin-STAG2 undergoes dimerization on chromatin in later G1 in a \nconcentration dependent manner. In addition, we found Cohesin-STAG2 to be on average bound to \nchromatin for 120 seconds and increasing its occupancy from 3 to 8 complexes per megabase from \nearly to late G1. Our data thus quantitatively explains how Cohesin STAG2 dimers form, if we assume \nthe bound complexes extrude loops with the repo rted speeds (i.e. 0.5-2 kb/s, Kim et al., 2019; \nDavidson et al., 2019): While 3 randomly loaded Cohesin-STAG2 complexes per megabase DNA are \nvery unlikely to encounter each other during early G1 due to the relatively small loops they can make \nduring 2 min (around 120 kb), encounters and potential stalling between loops of this size become \nmuch more likely when Cohesin-STAG2 is fully imported and present at 8 copies per megabase in late \nG1. While we cannot exclude that Cohesin-STAG2 di mers continue active loop extrusion, our data \nwould be consistent with the view that the default state of Cohesin complexes in loop extrusion is \nmonomeric and that dimers result from encounters and potential stalling events.  \nA comprehensive and quantitative dataset to constrain next generation polymer models \nIn summary, our systematic and quantitative assessment of Condensin and Cohesin loop extruder \ndynamics on chromatin provides a comprehensive an d integrated view of the transition from mitotic \nto interphase genome organization. Given the sequential import of Cohesin isoforms, their chromatin \nbinding dynamics, their different abundance as well as impact on chromatin upon depletion, we \npropose a hierarchical nested loop model fo r the establishment of the interphase genome \norganization by the Cohesin loop extruders after mi tosis. Our model is conceptually similar to the \nhierarchical nested loop architecture proposed for the establishment of Condensin driven mitotic \norganization (Gibcus et al., 2018; Walther et al., 2018). While the sub-structuring of large Condensin \nII loops in mitosis by Condensin I serves to laterally compact mitotic chromatin and confers additional \nmechanical rigidity to chromosomes (Ono et al., 2003; Shintomi and Hirano, 2011; Green et al., 2012; \nHoulard et al., 2015), we think that the sub-structuring of large STAG1 loops (Kojic et al., 2018; Wutz \net al., 2020) by STAG2 aids TAD-scale compaction  and specific intra-TAD contact enrichment, \npotentially in a cell type and species-specific manner (Dixon et al., 2012; Phillips-Cremins et al., 2013; \nRao et al., 2014). The quantitative data and understanding provided by our study should provide a \ncomprehensive quantitative basis for next generation predictive and mechanistically explanatory \nmodels of genome organization.  \n \n  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n12 \n \nA\ncknowledgements \nWe thank the EMBL Advanced Light Microscopy Facility (ALMF) and the EMBL Imaging Center for \nmicroscope support as well as the Centre for Bioimage Analysis for support related to image analysis. \nWe thank Gordana Wutz for valuable discussions and supply with antibodies and cell lines related to \nCohesin and CTCF. This work was supported by grants from the National Institutes of Health Common \nFund 4D Nucleome Program (Grant U01 EB021223 / U01 DA047728) to J.E., as well as by the The Paul \nG. Allen Frontiers Group through the Allen Distinguished Investigator Program to J.E.. A.B. has received \na PhD fellowship from the Boehringer Ingelheim Fonds and K.S.B. was supported by the Alexander von \nHumboldt foundation. Work in the laboratory of  J.-M.P. has received funding from Boehringer \nIngelheim, the Austrian Research  Promotion Agency (Headquarter grant FFG-852936), the European \nResearch Council (ERC) under the European Union’s Horizon 2020 research and innovation \nprogramme (grant agreements No 693949 and No 101020558), the Human Frontier Science Program \n(grant RGP0057/2018) and the Vienna Science and Technology Fund (grant LS19- 029).  \n \nAuthor contributions \nA.B. and J.E. conceived the project. K.S.B and J.E. supervised the project. A.B. designed, performed \nand analyzed all experiments with technical support from N.R.M. related to spot-bleach data \nacquisition, from K.S.B. related to FCS-calibrated imaging, FRAP and chromatin trace analysis, from \nM.J.H. related to the se gmentation of live-cell imaging data, from M.L. related to STED image \nacquisition, from H.P. related to protein degradat ion, and from A.H. related to automated live-cell \nimaging. N.R.M. and W.Z.  generated and validated genome-edited cell lines created within this study. \nA.B. and J.E. wrote the manuscript with reviewing and editing performed by all authors. Funding was \nacquired by A.B., K.S.B., J.E. and J.-M.P. \n  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n13 \n \nFigure 1 \nFigure 1. FCS-calibrated imaging of Cohesin isoforms shows that re-organization of loop extrusion during \nmitotic exit takes about 2h after anaphase onset.  \nA) Schematic of current loop-extrusion based models of mitotic and interphase genome organization. \nCondensin I and II build nested mitotic loops. Cohesins build topologically associating domains delimited by \nthe boundary factor CTCF during interphase.  \nB) Fluorescence intensity calibration using fluorescence correlation spectroscopy (FCS). Fluorescence \nintensity and photon count fluctuation measurements are performed in cells expressing varying amounts of \nmonomeric EGFP. An autocorrelation function simulating particle diffusion through the effective detection \nvolume is fit to the autocorrelated photon count signal, enabling estimation of protein number in the \neffective detection volume and therefore the calibration of fluorescent intensities to absolute protein \nconcentrations (Politi et al., 2018).  \nC) Imaging of genome-edited HK cells with homozygously EGFP-tagged Cohesin-STAG2 throughout 2 \nconsecutive mitoses. Fluorescent intensities (FI) measured in the second mitosis were set to the average \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n14 \n \np\nrotein concentration (C[nM]) during metaphase as measured by FCS-calibrated imaging of the same cell \nline during mitotic exit imaging (see E)). Protein concentrations of all other timepoints were adjusted based \non relative differences of measured fluorescence intensities.  \nD) Nuclear concentration throughout an entire cell cycle ranging from one anaphase to the next displayed \nfor genome-edited HK cells with homozygously (m)EGFP-tagged Cohesin-STAG1 (n = 20 cells) and Cohesin-\nSTAG2 (n = 13 cells), as well as Condensin I (NCPAH, n = 8 cells) and Condensin II (NCAPH2, n = 21 cells). \nInset shows focus of imaging on first two hours after mitosis performed with higher temporal resolution. \nError bands represent 95% confidence interval. \nE) FCS-calibrated imaging of genome-edited HK cells  with homozygously EGFP-tagged Cohesin-STAG2 \nthroughout mitotic exit. A total of 75 3D stacks with 2-minute intervals is triggered after successful \nautomated identification of metaphase cells based on SiR-Hoechst staining. Fluorescence intensity \ncalibration by FCS allows for the conversion of measured fluorescent intensities (FI) to absolute protein \nconcentrations and protein numbers (N) per unit volume. \nF) Absolute protein numbers co-localizing with chromatin/the two daughter nuclei displayed for genome-\nedited HK cells with homozygously (m)EGFP-tagged Cohesin-STAG2 (n = 11 cells) and Condensin I (NCPAH, n \n= 14 cells). Reformation and full establishment of the nuclear envelope as determined by Lamin B receptor \n(Suppl. Fig. 1N) is indicated through grey background. Error bands represent 95% confidence interval. \nG) Fluorescent micrographs of early G1 cells (~45 min past mitosis) stained with DAPI in WT or ΔNup153 \ncondition.  \nH) Average fluorescent intensity plots per 3D-segmented nucleus in grey (WT) or colored (ΔNup153). \nΔNup153 nuclei do not expand in size, show no residual Nup153 intensity and show a clear reduction in \nRAD21 intensity inside the nuclear lumen.  \nI) Average fluorescent intensity of early G1 cells in WT or ΔNup153 condition stained for RAD21. 50% drop \nin mean RAD21 intensity after 75 min release time. 33-48% reduction in average fluorescent intensity after \n45 min release time (not shown). Changes above/below 20% are considered a significant change.  \n \n \n \n \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n15 \n \nFigure 2 \nFigure 2. Condensins and Cohesins co-occupy chromatin during telophase and early G1, as revealed by \ntime-resolved bleaching.   \nA) Illustration of the spot-bleach assay. Genome edited HK cells homozygously expressing (m)EGFP tagged \nCondensin and Cohesin subunits are illuminated at a single spot on chromatin for a total duration of 30 \nseconds and the resulting fluorescent intensity is continuously measured. The chromatin-bound fraction of \na given protein of interest is calculated based on the mean fluorescent intensity of the first and last 500 \nmsec. Exemplary image and bleach data is shown for the common Condensin subunit SMC4.  \nB) The fraction of chromatin-bound Condensins (SMC4) and Cohesins (RAD21) determined using the spot-\nbleach assay at different timepoints during mitotic exit. Every bar plot represents at least 10 individual \ndatapoints measured in 10 separate cells.  \nC) Absolute number of proteins bound to chromatin were determined by multiplication of chromatin bound \nfractions shown in B with absolute protein numbers co-localizing with chromatin (n(SMC4) = 21 cells, \nn(RAD21) = 18 cells) as determined in Fig. 1E&F and displayed as per-megabase-count assuming an equal \ndistribution of the proteins on the entire 7.9 Mb HeLa genome (Landry et al., 2013). Grey background \nindicates reformation of nuclear envelope. Error bands represent 95% confidence interval. \nD) Fluorescent micrographs and quantification of early G1 cells in WT condition or after degradation of the \nisoform-shared Condensin subunit SMC4. Cells were pre-extracted for 1 minute prior to fixation and were \nstained for RAD21. SMC4 depletion caused a delay in cell division as well as major cell division errors (see \nmerged daughter nuclei in fluorescent micrograph indicated by arrow). Time of release from Nocodazole \nblock had to be increased to 60-70 minutes to fix cells in early G1 stage. Difference in mean fluorescence \nintensity: 8-12.5%. Changes above/below 20% are considered a significant change. \nE) Fluorescent micrographs and quantification of early G1 cells in WT condition or after degradation of the \nCohesin loader NIPBL. Cells were pre-extracted for 1 minute before fixation and were stained for SMC2. \nDifference in mean fluorescence intensity: ~15%. Changes above/below 20% are considered a significant \nchange. \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n16 \n \nFigure 3 \nFigure 3. Cohesin-STAG1 and CTCF cooperate to form interphase TAD structures after mitosis. \nA) FCS-calibrated protein numbers co-localizing with chromatin displayed for genome-edited HK cells with \nhomozygously EGFP-tagged Cohesin-STAG1 (n = 25 cells), Cohesin-STAG2 (n = 11 cells) and CTCF (n = 15 \ncells) relative to the measurement 2 hours after anaphase onset. Error bands represent 95% confidence \ninterval. \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n17 \n \nB\n) Scheme explaining LoopTrace chromatin tracing workflow. Fixed cells were subjected to single strand \nresection via exonuclease treatment (RASER) for maximal structure-preservation and subsequent \nhybridization with a tiled FISH library. Every FISH-probe contains a non-genome-complementary docking \nhandle that can be hybridized with a fluorescently labelled imager strand to read out the 3D location of a \ngenomic locus (Beckwith et al., 2023 Preprint). \nC) Overview of the traced 1.2 megabase locus on chromosome 14 with genes as well as ChIP-seq binding \nsites for RAD21 and CTCF (from the ENCODE portal (Sloan et al., 2016, https://www.encodeproject.org/) \nwith the following identifiers: ENCFF239FBO (RAD21), ENCFF111RWV (CTCF); CTCF directionality \nannotations from Rao et al., 2014). \nD-E) Exemplary chromatin traces of WT (E) or ΔSTAG2 (F) early G1 cells.  \nF) Distance and contact matrices of a 1.2 megabase region on chromosome 14 locus traced at a genomic \nresolution of 12 kb in early G1 cells with and without Cohesin-STAG2. Differences between WT and ΔSTAG2 \nare highlighted for distance and contact probability maps. \n \n \n \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n18 \n \nFigure 4 \nFigure 4. Fluorescence Recovery of Cohesin isoforms and CTCF. \nA) Fluorescence recovery after photobleaching (FRAP) was performed by bleaching half of a nucleus in early \nG1 cells (20-40 minutes after anaphase onset) or later G1 cells selected by nuclear volume.  \nB) FRAP shown for genome-edited HK cells with homozygously EGFP-tagged CTCF. Difference between the \nbleached an unbleached region is normalized by the maximal difference at time t=0 after bleaching. Black \nline indicates the data fit by a single-exponential function with immobile fraction. Single exponential \nfunctions with immobile fraction also fit the FRAP recovery of RAD21, STAG1/2 well. \nC-E) FRAP measurements using homozygous EGFP-knock-in HK cell lines in earlyG1 and G1 cells, \nrespectively. Bar plots display the fraction of protein that is stably bound to chromatin. Two-sample t-test \nwas used for calculating significance levels. Error bars show standard deviation.   \nC) Cohesin-STAG1 (early G1: n = 10 cells, G1: n = 9 cells) \nD) Cohesin-STAG2. (early G1: n = 10 cells, G1: n = 13 cells) \nE) CTCF. (early G1: n = 9 cells, G1: n = 10 cells) \nF) FRAP measurements of endogenous CTCF with WT levels of RAD21, after degradation of endogenous \nRAD21, and after rescue of RAD21 degradation by exogenous RAD21 expression for at least 24 hours. Bar \nplots display the fraction of protein that is stably bound to chromatin. (CTCF WT: n = 10 cells, CTCF dRAD21: \nn = 9 cells, CTCF dRAD21 rescue: n = 10 cells). Data from CTCF-EGFP knock-in line is used as WT reference as \nit displays WT expression levels of RAD21. The double-knock-in line Rad21-EGFP-AID CTCF-Halo-3xALFA #C7 \ndisplayed leaky degradation of RAD21, reducing CTCF-chromatin binding already in -IAA cells (see Suppl. Fig. \n4G and methods).  \n \n \n  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n19 \n \nFigure 5 \nFigure 5. STED super-resolution imaging of Cohesin isoforms and CTCF.  \nA-B) Exemplary STED images showing Cohesin-STAG1/2 (magenta) and CTCF (green) in early G1 (A) and G1 \n(B) nuclei (scalebar: 5 µm) and zoom-ins (scalebar: 1 µm).  \nC) Co-localization analysis of Cohesin-STAG1/2 with CTCF using the Pearson Correlation Coefficient of \nsegmented nuclei (n ≥ 17). Differences in STAG-CTCF co-localization in early G1 compared to G1 are \nsignificant as assessed by independent two-sample t-test.  \nD) Mean intensity of segmented STAG1/2 spots in STED images of replicate 3. Same results are observed in \nreplicate 1 and 2. Formal significance tests are meaningless due to large sample size. Median intensities of \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n20 \n \nthe mean spot intensity distributions are: STAG1 eG1: 12.02, STAG1 G1: 13.06, STAG2 eG1: 12.62, STAG2 \nG1: 21.02 (arbitrary intensity units).   \nE-F) Exemplary chromatin traces of WT (E) or ΔSTAG2 (F) interphase cells.  \nG) Scaling plot of Chr14 1.2 megabase region sampled at 12 kb resolution in WT or ΔSTAG2 interphase cells. \nTraces from ΔSTAG2 are on average less compact compared to WT.  \nH) Distance and contact matrices of a 1.2 megabase region on chromosome 14 locus traced at a genomic \nresolution of 12 kb in interphase cells with and without Cohesin-STAG2. Differences between WT and \nΔSTAG2 are highlighted for distance and contact probability maps. WT data from 1, ΔSTAG2 data from 2 \nindependent technical replicates (392 and 610 traces, respectively).  \n \n \n \n \nFigure 6 \nFigure 6. A new model for the mitosis-to-interphase transition of genome organization by loop extrusion. \nMitotic chromosomes are majorly organized by the abundant Condensin complexes (12/Mb, of those ~1.5 \nare Condensin II), only a small fraction of CTCF (about 14,000 proteins, ~ 10% of cellular pool) binds \nchromatin very transiently and a small fraction of the cellular Cohesin holds together sister chromatids until \nanaphase. During telophase and early G1, not all Condensins dissociate right away, leading to a co-\noccupancy of 3 Condensins and 3 Cohesins per megabase DNA and a significant pool of Condensin I that \nremains nuclear until the next mitosis starts. Cohesin-STAG1 and CTCF are fully imported into the newly \nformed nucleus within 10 minutes after anaphase onset and are sufficient to build interphase TAD \nstructures in the absence of Cohesin-STAG2. Cohesin-STAG2 only completes its import 2 hours after \nanaphase onset and – due to its abundance on chromatin upon full import (8 complexes per megabase) – \nfrequently crashes into neighboring loop extruders, leading to increased nesting of sub-TAD loops. While \nCohesin-STAG1 is stabilized through CTCF and SMC3 acetylation (Wutz et al., 2020), CTCF is increasingly \nstabilized on chromatin through its association with Cohesin on chromatin. After completed import of \nCohesins and CTCF, their nuclear concentrations remain stable irrespective of the DNA content of the \nnucleus. \n \n  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\nPOI e G1 (= 20 min past AO)\nnuclear, relative to 2h \npost AO\nabsolute number \nin nucleus\n% bound (spot-\nbleach, average) % bound (FRAP) absolute \nnumber, bound\nMean residence \ntime, \ndynamically \nbound pool (s)\nabsolute, \nchromatin \nbound, \nper Mb\nfraction long \nterm bound, \nper Mb\nabsolute, long \nterm bound, \nper Mb\nSMC4 104% 178000 0.21 0.19 33820 28 2.1 0.13 0.3\nNCAPH 103% 127000 0.15 0.11 13970 45 0.9 0.09 0.1\nNCAPH2 99% 25400 0.22 nan 5588\nOptimal \nParameters for \nFitting not found 0.4 nan nan\nCTCF 102% 125500 0.5 0.62 77810 125 4.9 0.32 1.6\nRAD21 63% 151000 0.41 0.53 80030 155 5.1 0.28 1.4\nSTAG1 100% 53500 0.37 0.62 33170 244 2.1 0.27 0.6\nSTAG2 56% 102000 0.37 0.47 47940 108 3.0 0.07 0.2\nPOI G 1 (= 2-4h past AO)\nnuclear, relative \nto 2h post AO\nabsolute number \nin nucleus\n%\n bound (spot-bleach, \naverage), interphase \ntime-point\n% bound (FRAP), G1 \ntimepoint\nResidence time, \ndynamically \nbound pool\nabsolute \nnumber, bound\nabsolute, \nchromatin \nbound, \nper Mb\nfraction long \nterm bound, \nper Mb\nabsolute, long \nterm bound, \nper Mb\nSMC4 100% 174000 0.1 17400\nNCAPH 100% 125000 0.09 11250\nNCAPH2 100% 25600 0.02 512\nCTCF 100% 124000 0.72 0.72 139 89280 5.7 0.488 2.8\nRAD21 100% 238000 0.59 0.77 172 183260 11.6 0.277 3.2\nSTAG1 100% 53500 0.59 0.76 276 40660 2.6 0.39 1.0\nSTAG2 100% 183000 0.5 0.71 126 129930 8.2 0.107 0.9\nTable 1. Quantitative data summary: early G1 \nTable 2. Quantitative data summary: G1 \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n21 \n \nR\neferences \nAbramo, K., A.L. Valton, S. V. Venev, H. Ozadam, A.N. Fox, and J. Dekker. 2019. A chromosome folding \nintermediate at the condensin-to-cohesin transition during telophase. Nat. Cell Biol. 21:1393–1402. \ndoi:10.1038/s41556-019-0406-2. \nBazile, F., J. St-Pierre, and D.D.’ Amours. 2010. Cell Cycle Three-step model for condensin activation during \nmitotic chromosome condensation Introduction: Tackling the Enigma. doi:10.4161/cc.9.16.12620. \nBeckwith, K.S., Ø. Ødegård-Fougner, N.R. Morero, C. Barton, F. Schueder, W. Tang, S. Alexander, J.-M. Peters, \nR. Jungmann, E. Birney, and J. Ellenberg. 2023. Nanoscale 3D DNA tracing in single human cells visualizes \nloop extrusion directly in situ. bioRxiv. 2021.04.12.439407. doi:10.1101/2021.04.12.439407. \nBorsellini, A., D. Conti, E. Cutts, R.J. Harris, K. Walstein, A. Graziadei, V. Cecatiello, T.F. Aarts, R. Xie, A. Mazouzi, \nS. Sen, C. Hoencamp, R. Pleuger, S. Ghetti, L. Oberste-Lehn, D. Pan, T. Bange, J.H.I. Haarhuis, A. Perrakis, \nT.R. Brummelkamp, B.D. Rowland, A. Musacchio, and A. Vannini. 2024. Condensin II activation by \nM18BP1. bioRxiv. 2024.05.02.592151. doi:10.1101/2024.05.02.592151. \nCai, Y., M.J. Hossain, J.K. Hériché, A.Z. Politi, N. Walther, B. Koch, M. Wachsmuth, B. Nijmeijer, M. Kueblbeck, \nM. Martinic-Kavur, R. Ladurner, S. Alexander, J.M. Peters, and J. Ellenberg. 2018. Experimental and \ncomputational framework for a dynamic protein atlas of human cell division. Nature. 561:411–415. \ndoi:10.1038/s41586-018-0518-z. \nCutts, E.E., D. Tetiker, E. Kim, and L. Aragon. 2024. Molecular Mechanism of Condensin I Activation by KIF4A. \nbioRxiv. 2024.03.19.585759. doi:10.1101/2024.03.19.585759. \nDavidson, I.F., B. Bauer, D. Goetz, W. Tang, G. Wutz, and J.-M. Peters. 2019. DNA loop extrusion by human \ncohesin. Science. 366:1338–1345. doi:10.1126/science.aaz3418. \nDavidson, I.F., and J.M. Peters. 2021. Genome folding through loop extrusion by SMC complexes. Nat. Rev. \nMol. Cell Biol. 2021 227. 22:445–464. doi:10.1038/s41580-021-00349-7. \nDixon, J.R., S. Selvaraj, F. Yue, A. Kim, Y. Li, Y. Shen, M. Hu, J.S. Liu, and B. Ren. 2012. Topological domains in \nmammalian genomes identified by analysis of chromatin interactions. Nature. 485:376–380. \ndoi:10.1038/nature11082. \nFudenberg, G., M. Imakaev, C. Lu, A. Goloborodko, N. Abdennur, and L.A. Mirny. 2016. Formation of \nChromosomal Domains by Loop Extrusion. Cell Rep. 15:2038–2049. doi:10.1016/j.celrep.2016.04.085. \nGabriele, M., H.B. Brandão, S. Grosse-Holz, A. Jha, G.M. Dailey, C. Cattoglio, T.H.S. Hsieh, L. Mirny, C. Zechner, \nand A.S. Hansen. 2022. Dynamics of CTCF- and cohesin-mediated chromatin looping revealed by live-cell \nimaging. Science (80-. ). 376:476–501. \ndoi:10.1126/SCIENCE.ABN6583/SUPPL_FILE/SCIENCE.ABN6583_MDAR_REPRODUCIBILITY_CHECKLIST.PD\nF. \nGanji, M., I.A. Shaltiel, S. Bisht, E. Kim, A. Kalichava, C.H. Haering, and C. Dekker. 2018. Real-time imaging of \nDNA loop extrusion by condensin. Science (80-. ). 360:102–105. doi:10.1126/science.aar7831. \nGibcus, J.H., K. Samejima, A. Goloborodko, I. Samejima, N. Naumova, J. Nuebler, M.T. Kanemaki, L. Xie, J.R. \nPaulson, W.C. Earnshaw, L.A. Mirny, and J. Dekker. 2018. A pathway for mitotic chromosome formation. \nScience (80-. ). 359. doi:10.1126/science.aao6135. \nGreen, L.C., P. Kalitsis, T.M. Chang, M. Cipetic, J.H. Kim, O. Marshall, L. Turnbull, C.B. Whitchurch, P. Vagnarelli, \nK. Samejima, W.C. Earnshaw, K.H.A. Choo, and D.F. Hudson. 2012. Contrasting roles of condensin I and \ncondensin II in mitotic chromosome formation. J. Cell Sci. 125:1591–1604. \ndoi:10.1242/JCS.097790/258379/AM/CONTRASTING-ROLES-OF-CONDENSIN-I-AND-II-IN-MITOTIC. \nGuo, Y., Q. Xu, D. Canzio, J. Shou, J. Li, D.U. Gorkin, I. Jung, H. Wu, Y. Zhai, Y. Tang, Y. Lu, Y. Wu, Z. Jia, W. Li, \nM.Q. Zhang, B. Ren, A.R. Krainer, T. Maniatis, and Q. Wu. 2015. CRISPR Inversion of CTCF Sites Alters \nGenome Topology and Enhancer/Promoter Function. Cell. 162:900–910. \ndoi:10.1016/J.CELL.2015.07.038. \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n22 \n \nH\nildebrand, E.M., K. Polovnikov, B. Dekker, Y. Liu, D.L. Lafontaine, A.N. Fox, Y. Li, S. V. Venev, L.A. Mirny, and J. \nDekker. 2024. Mitotic chromosomes are self-entangled and disentangle through a topoisomerase-II-\ndependent two-stage exit from mitosis. Mol. Cell. 84:1422-1441.e14. \ndoi:10.1016/J.MOLCEL.2024.02.025. \nHirano, T., R. Kobayashi, and M. Hirano. 1997. Condensins, chromosome condensation protein complexes \ncontaining XCAP- C, XCAP-E and a Xenopus homolog of the Drosophila Barren protein. Cell. 89:511–521. \ndoi:10.1016/S0092-8674(00)80233-0. \nHirano, T., and T.J. Mitchison. 1994. A heterodimeric coiled-coil protein required for mitotic chromosome \ncondensation in vitro. Cell. 79:449–458. doi:10.1016/0092-8674(94)90254-2. \nHirota, T., D. Gerlich, B. Koch, J. Ellenberg, and J.M. Peters. 2004. Distinct functions of condensin I and II in \nmitotic chromosome assembly. J. Cell Sci. 117:6435–6445. doi:10.1242/JCS.01604. \nHolzmann, J., A.Z. Politi, K. Nagasaka, M. Hantsche-Grininger, N. Walther, B. Koch, J. Fuchs, G. Dürnberger, W. \nTang, R. Ladurner, R.R. Stocsits, G.A. Busslinger, B. Novák, K. Mechtler, I.F. Davidson, J. Ellenberg, and J.-\nM. Peters. 2019. Absolute quantification of cohesin, CTCF and their regulators in human cells. Elife. 8. \ndoi:10.7554/eLife.46269. \nHoulard, M., E.E. Cutts, M.S. Shamim, J. Godwin, D. Weisz, A.P. Aiden, E.L. Aiden, L. Schermelleh, A. Vannini, \nand K. Nasmyth. 2021. MCPH1 inhibits Condensin II during interphase by regulating its SMC2-Kleisin \ninterface. Elife. 10. doi:10.7554/ELIFE.73348. \nHoulard, M., J. Godwin, J. Metson, J. Lee, T. Hirano, and K. Nasmyth. 2015. Condensin confers the longitudinal \nrigidity of chromosomes. Nat. Cell Biol. 17:771–781. doi:10.1038/NCB3167. \nKim, Y., Z. Shi, H. Zhang, I.J. Finkelstein, and H. Yu. 2019. Human cohesin compacts DNA by loop extrusion. \nScience (80-. ). 366:1345–1349. doi:10.1126/science.aaz4475. \nKimura, K., M. Hirano, R. Kobayashi, and T. Hirano. 1998. Phosphorylation and activation of 13S condensin by \nCdc2 in vitro. Science (80-. ). 282:487–490. \ndoi:10.1126/SCIENCE.282.5388.487/SUPPL_FILE/981578.XHTML. \nKojic, A., A. Cuadrado, M. De Koninck, D. Giménez-Llorente, M. Rodríguez-Corsino, G. Gómez-López, F. Le Dily, \nM.A. Marti-Renom, and A. Losada. 2018. Distinct roles of cohesin-SA1 and cohesin-SA2 in 3D \nchromosome organization. Nat. Struct. Mol. Biol. 2018 256. 25:496–504. doi:10.1038/s41594-018-0070-\n4. \nKueng, S., B. Hegemann, B.H. Peters, J.J. Lipp, A. Schleiffer, K. Mechtler, and J.M. Peters. 2006. Wapl Controls \nthe Dynamic Association of Cohesin with Chromatin. Cell. 127:955–967. \ndoi:10.1016/J.CELL.2006.09.040/ATTACHMENT/8DA5CD14-3CC1-4E70-A3D7-\nC9C3DDC10132/MMC1.PDF. \nLandry, J.J.M., P.T. Pyl, T. Rausch, T. Zichner, M.M. Tekkedil, A.M. Stütz, A. Jauch, R.S. Aiyar, G. Pau, N. \nDelhomme, J. Gagneur, J.O. Korbel, W. Huber, and L.M. Steinmetz. 2013. The genomic and \ntranscriptomic landscape of a hela cell line. G3 Genes, Genomes, Genet. 3:1213–1224. \ndoi:10.1534/g3.113.005777. \nLi, Y., J.H.I. Haarhuis, Á.S. Cacciatore, R. Oldenkamp, M.S. van Ruiten, L. Willems, H. Teunissen, K.W. Muir, E. de \nWit, B.D. Rowland, and D. Panne. 2020. The structural basis for cohesin-CTCF-anchored loops. Nature. \ndoi:10.1038/s41586-019-1910-z. \nLosada, A., M. Hirano, and T. Hirano. 1998. Identification of Xenopus SMC protein complexes required for \nsister chromatid cohesion. Genes Dev. 12:1986–1997. doi:10.1101/GAD.12.13.1986. \nLosada, A., T. Yokochi, R. Kobayashi, and T. Hirano. 2000. Identification and Characterization of SA/Scc3p \nSubunits in the Xenopus and Human Cohesin Complexes. J. Cell Biol. 150:405–416. \nLupiáñez, D.G., K. Kraft, V. Heinrich, P. Krawitz, F. Brancati, E. Klopocki, D. Horn, H. Kayserili, J.M. Opitz, R. \nLaxova, F. Santos-Simarro, B. Gilbert-Dussardier, L. Wittler, M. Borschiwer, S.A. Haas, M. Osterwalder, M. \nFranke, B. Timmermann, J. Hecht, M. Spielmann, A. Visel, and S. Mundlos. 2015. Disruptions of \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n23 \n \nt\nopological chromatin domains cause pathogenic rewiring of gene-enhancer interactions. Cell. 161:1012–\n1025. doi:10.1016/j.cell.2015.04.004. \nMach, P., P.I. Kos, Y. Zhan, J. Cramard, S. Gaudin, J. Tünnermann, E. Marchi, J. Eglinger, J. Zuin, M. \nKryzhanovska, S. Smallwood, L. Gelman, G. Roth, E.P. Nora, G. Tiana, and L. Giorgetti. 2022. Cohesin and \nCTCF control the dynamics of chromosome folding. Nat. Genet. 2022 5412. 54:1907–1918. \ndoi:10.1038/s41588-022-01232-7. \nMitter, M., C. Gasser, Z. Takacs, C.C.H. Langer, W. Tang, G. Jessberger, C.T. Beales, E. Neuner, S.L. Ameres, J.M. \nPeters, A. Goloborodko, R. Micura, and D.W. Gerlich. 2020. Conformation of sister chromatids in the \nreplicated human genome. Nat. 2020 5867827. 586:139–144. doi:10.1038/s41586-020-2744-4. \nNaumova, N., M. Imakaev, G. Fudenberg, Y. Zhan, B.R. Lajoie, L.A. Mirny, and J. Dekker. 2013. Organization of \nthe mitotic chromosome. Science (80-. ). 342:948–953. doi:10.1126/science.1236083. \nNora, E.P., B.R. Lajoie, E.G. Schulz, L. Giorgetti, I. Okamoto, N. Servant, T. Piolot, N.L. Van Berkum, J. Meisig, J. \nSedat, J. Gribnau, E. Barillot, N. Blüthgen, J. Dekker, and E. Heard. 2012. Spatial partitioning of the \nregulatory landscape of the X-inactivation centre. Nat. 2012 4857398. 485:381–385. \ndoi:10.1038/nature11049. \nOchs, F., C. Green, A.T. Szczurek, L. Pytowski, S. Kolesnikova, J. Brown, D.W. Gerlich, V. Buckle, L. Schermelleh, \nand K.A. Nasmyth. 2024. Sister chromatid cohesion is mediated by individual cohesin complexes. Science \n(80-. ). 383:1122–1130. doi:10.1126/SCIENCE.ADL4606. \nOno, T., Y. Fang, D.L. Spector, and T. Hirano. 2004. Spatial and temporal regulation of condensins I and II in \nmitotic chromosome assembly in human cells. Mol. Biol. Cell. 15:3296–3308. doi:10.1091/mbc.E04-03-\n0242. \nOno, T., A. Losada, M. Hirano, M.P. Myers, A.F. Neuwald, and T. Hirano. 2003. Differential contributions of \ncondensin I and condensin II to mitotic chromosome architecture in vertebrate cells. Cell. 115:109–121. \ndoi:10.1016/S0092-8674(03)00724-4. \nOtsuka, S., K.H. Bui, M. Schorb, M. Julius Hossain, A.Z. Politi, B. Koch, M. Eltsov, M. Beck, and J. Ellenberg. 2016. \nNuclear pore assembly proceeds by an inside-out extrusion of the nuclear envelope. Elife. 5. \ndoi:10.7554/ELIFE.19071. \nPhillips-Cremins, J.E., M.E.G. Sauria, A. Sanyal, T.I. Gerasimova, B.R. Lajoie, J.S.K. Bell, C.T. Ong, T.A. Hookway, \nC. Guo, Y. Sun, M.J. Bland, W. Wagstaff, S. Dalton, T.C. McDevitt, R. Sen, J. Dekker, J. Taylor, and V.G. \nCorces. 2013. Architectural Protein Subclasses Shape 3D Organization of Genomes during Lineage \nCommitment. Cell. 153:1281–1295. doi:10.1016/J.CELL.2013.04.053. \nPoliti, A.Z., Y. Cai, N. Walther, M.J. Hossain, B. Koch, M. Wachsmuth, and J. Ellenberg. 2018. Quantitative \nmapping of fluorescently tagged cellular proteins using FCS-calibrated four-dimensional imaging. Nat. \nProtoc. 13:1445–1464. doi:10.1038/nprot.2018.040. \nRao, S.S.P., M.H. Huntley, N.C. Durand, E.K. Stamenova, I.D. Bochkov, J.T. Robinson, A.L. Sanborn, I. Machol, \nA.D. Omer, E.S. Lander, and E.L. Aiden. 2014. A 3D map of the human genome at kilobase resolution \nreveals principles of chromatin looping. Cell. 159:1665–80. doi:10.1016/j.cell.2014.11.021. \nSanborn, A.L., S.S.P. Rao, S.C. Huang, N.C. Durand, M.H. Huntley, A.I. Jewett, I.D. Bochkov, D. Chinnappan, A. \nCutkosky, J. Li, K.P. Geeting, A. Gnirke, A. Melnikov, D. McKenna, E.K. Stamenova, E.S. Lander, and E.L. \nAiden. 2015. Chromatin extrusion explains key features of loop and domain formation in wild-type and \nengineered genomes. Proc. Natl. Acad. Sci. U. S. A. 112:E6456–E6465. doi:10.1073/pnas.1518552112. \nSchneider, M.W.G., B.A. Gibson, S. Otsuka, M.F.D. Spicer, M. Petrovic, C. Blaukopf, C.C.H. Langer, P. Batty, T. \nNagaraju, L.K. Doolittle, M.K. Rosen, and D.W. Gerlich. 2022. A mitotic chromatin phase transition \nprevents perforation by microtubules. Nat. 2022. 1–8. doi:10.1038/s41586-022-05027-y. \nShintomi, K., and T. Hirano. 2011. The relative ratio of condensin I to II determines chromosome shapes. Genes \nDev. 25:1464–1469. doi:10.1101/GAD.2060311. \nSloan, C.A., E.T. Chan, J.M. Davidson, V.S. Malladi, J.S. Strattan, B.C. Hitz, I. Gabdank, A.K. Narayanan, M. Ho, \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n24 \n \nB\n.T. Lee, L.D. Rowe, T.R. Dreszer, G. Roe, N.R. Podduturi, F. Tanaka, E.L. Hong, and J.M. Cherry. 2016. \nENCODE data at the ENCODE portal. Nucleic Acids Res. 44:D726–D732. doi:10.1093/NAR/GKV1160. \nSumara, I., E. Vorlaufer, C. Gieffers, B.H. Peters, and J.-M. Peters. 2000. Characterization of Vertebrate Cohesin \nComplexes and Their Regulation in Prophase. J. Cell Biol. 151:749–761. \nTane, S., K. Shintomi, K. Kinoshita, Y. Tsubota, M.M. Yoshida, T. Nishiyama, and T. Hirano. 2022. Cell cycle-\nspecific loading of condensin I is regulated by the N-terminal tail of its kleisin subunit. Elife. 11. \ndoi:10.7554/ELIFE.84694. \nThevathasan, J.V., M. Kahnwald, K. Cieśliński, P. Hoess, S.K. Peneti, M. Reitberger, D. Heid, K.C. Kasuba, S.J. \nHoerner, Y. Li, Y. Le Wu, M. Mund, U. Matti, P.M. Pereira, R. Henriques, B. Nijmeijer, M. Kueblbeck, V.J. \nSabinina, J. Ellenberg, and J. Ries. 2019. Nuclear pores as versatile reference standards for quantitative \nsuperresolution microscopy. Nat. Methods. 16:1045–1053. doi:10.1038/s41592-019-0574-9. \nViny, A.D., R.L. Bowman, Y. Liu, V.P. Lavallée, S.E. Eisman, W. Xiao, B.H. Durham, A. Navitski, J. Park, S. \nBraunstein, B. Alija, A. Karzai, I.S. Csete, M. Witkin, E. Azizi, T. Baslan, C.J. Ott, D. Pe’er, J. Dekker, R. \nKoche, and R.L. Levine. 2019. Cohesin Members Stag1 and Stag2 Display Distinct Roles in Chromatin \nAccessibility and Topological Control of HSC Self-Renewal and Differentiation. Cell Stem Cell. 25:682-\n696.e8. doi:10.1016/J.STEM.2019.08.003. \nWalther, N., M.J. Hossain, A.Z. Politi, B. Koch, M. Kueblbeck, Ø. Odegard-fougner, M. Lampe, and J. Ellenberg. \n2018. A quantitative map of human Condensins provides new insights into mitotic chromosome \narchitecture. J. Cell Biol. 217:2309–2328. doi:10.1083/jcb.201801048. \nde Wit, E., E.S.M. Vos, S.J.B. Holwerda, C. Valdes-Quezada, M.J.A.M. Verstegen, H. Teunissen, E. Splinter, P.J. \nWijchers, P.H.L. Krijger, and W. de Laat. 2015. CTCF Binding Polarity Determines Chromatin Looping. Mol. \nCell. 60:676–684. doi:10.1016/J.MOLCEL.2015.09.023. \nWutz, G., R. Ladurner, B.G. St Hilaire, R.R. Stocsits, K. Nagasaka, B. Pignard, A. Sanborn, W. Tang, C. Varnai, \nM.P. Ivanov, S. Schoenfelder, P. Van Der Lelij, X. Huang, G. Durnberger, E. Roitinger, K. Mechtler, I. Finley \nDavidson, P. Fraser, E. Lieberman-Aiden, and J.M. Peters. 2020. ESCO1 and CTCF enable formation of \nlong chromatin loops by protecting cohesinstag1 from WAPL. Elife. 9. doi:10.7554/eLife.52091. \nWutz, G., C. Várnai, K. Nagasaka, D.A. Cisneros, R.R. Stocsits, W. Tang, S. Schoenfelder, G. Jessberger, M. \nMuhar, M.J. Hossain, N. Walther, B. Koch, M. Kueblbeck, J. Ellenberg, J. Zuber, P. Fraser, and J. Peters. \n2017. Topologically associating domains and chromatin loops depend on cohesin and are regulated by \nCTCF, WAPL, and PDS5 proteins. EMBO J. 36:3573–3599. doi:10.15252/embj.201798004. \nYatskevich, S., J. Rhodes, and K. Nasmyth. 2019. Organization of Chromosomal DNA by SMC Complexes. Annu. \nRev. Genet. 53:445–482. doi:10.1146/annurev-genet-112618-043633. \nZhang, H., D.J. Emerson, T.G. Gilgenast, K.R. Titus, Y. Lan, P. Huang, D. Zhang, H. Wang, C.A. Keller, B. Giardine, \nR.C. Hardison, J.E. Phillips-Cremins, and G.A. Blobel. 2019. Chromatin structure dynamics during the \nmitosis-to-G1 phase transition. Nature. 576:158–162. doi:10.1038/s41586-019-1778-y. \nZuin, J., G. Roth, Y. Zhan, J. Cramard, J. Redolfi, E. Piskadlo, P. Mach, M. Kryzhanovska, G. Tihanyi, H. Kohler, M. \nEder, C. Leemans, B. van Steensel, P. Meister, S. Smallwood, and L. Giorgetti. 2022. Nonlinear control of \ntranscription through enhancer–promoter interactions. Nat. 2022 6047906. 604:571–577. \ndoi:10.1038/s41586-022-04570-y. \n \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n25 \n \nM\naterials and Methods \nCell Culture \nHeLa Kyoto cells (RRID: CVCL_1922) were obtained from S. Narumiya (Kyoto University, Kyoto, Japan) \nand cultured in high-glucose DMEM (41965-062, Thermo Fisher Scientific) supplemented with 10% \nFBS (10270-106, Lot. 42F2388K, Thermo Fisher Scientific ), 100 U/ml penicillin-streptomycin (15140-\n122, Thermo Fisher Scientific) and 1 mM sodium pyruvate (11360-039, Thermo Fisher Scientific) at \n37°C, 5% CO 2 unless otherwise stated. Cells were grown in cell culture dishes (Falcon) and passaged \nevery 2-3 days via trypsinization with 0.05% Trypsin-EDTA (25300-054, Thermo Fisher Scientific) at 80-\n90% confluency. Mycoplasma contamination was checked regularly and confirmed negative. \nFCS-calibrated confocal time-lapse imaging  \nCell samples for FCS-calibrated confocal time-lapse imaging were prepared according to Politi et al. \n(2018). Specifically, two days before the experiment, two 0.34 cm 2 wells of an 18-well chambered \ncoverglass (Ibidi µ-slide, 81817) were seeded with 3750 HK WT cells. 24 hours prior to the experiment, \none well of HK WT cells was transfected with a plasmid expressing monomeric EGFP and 2,000-4,000 \ngenome-edited cells expressing the protein of interest (POI) endogenously tagged with (m)EGFP were \nseeded in a third well. 1.5 hours prior to imaging, DMEM medium was exchanged to phenol-red free \nCO2-independent imaging medium based on Minimum Essential Medium (Sigma-Aldrich, M3024) \ncontaining 30 mM HEPES (pH 7.4), 10% FBS, 1X ME M non-essential amino-acids (11140-050, Thermo \nFisher Scientific) and 50-100 nM 5-SiR-Hoechst (gift from G. Lukinavi či us, Bucevičius et al., 2019). In \naddition, after 1.5 hours of DNA-labelling by 5-SiR-Hoechst, 500-kDa dextran-Dy481XL (Cai et al., 2018) \nwas added to the genome-edited cells to facilitate cell segmentation.  \nFluorescence Correlation Spectroscopy (FCS)-calibrated imaging was performed on Zeiss LSM780 \n(equipped with ConfoCor 3 unit, controlled by ZEN 2.3 Black software, Version 14.0.18.201, Zeiss) and \nLSM880 (controlled by ZEN 2.1 Black software, Versio n 14.0.9.201, Zeiss) laser-scanning microscopes \nwith an inverted Axio Observer microscope stand, equipped with an in-house constructed incubation \nchamber for temperature control set to 37°C (without CO 2 due to use of CO 2-independent imaging \nmedium) and using a C-Apochromat 40x/1.2 W Korr UV-Vis-IR water-immersion objective (421767-\n9971-711, Zeiss). Microscope calibration by FCS was performed as described by Politi et al. (2018), but \nusing 10 nM Atto488 carboxylic acid (AD 488-21, ATTO-TEC, Kapusta, 2010) in ddH2O instead of AF488 \ncoupled to a H2O-hydrolyzable NHS ester group to estimate the confocal volume in FCS \nmeasurements. This led to ~30% larger confocal volume estimates in better agreement with other \nmethods for confocal volume determination (Buschmann et al., 2009). This change resulted in a \nsystematic drop of the protein concentrations measured proportional to the change in confocal \nvolume size, compared to previous measurements using AF488-NHS (Politi et al., 2018). Ten FCS-\nmeasurements of 1 minute each were performed to estimate the effective confocal volume in the well \nwith Atto488 solution. FCS-measurements of 30  seconds were performed in the nucleus and \ncytoplasm in WT cells not expressing mEGFP to determine background fluorescence and photon \ncounts. Experiment-specific calibration factors were obtained from interphase cells expressing mEGFP \nby correlating measured fluorescence intensities and absolute mEGFP concentration calculated from \n30 seconds FCS-measurements (Politi et al., 2018).  \nCalibrated 4-dimensional confocal time-lapse imaging was performed on cells expressing the mEGFP-\ntagged protein of interest (POI) using a combin ation of MyPic macros for ZenBlack software \n(\nhttps://git.embl.de/grp-ellenberg/mypic), AutoMicTools library \n(https://git.embl.de/halavaty/AutoMicTools) for ImageJ (Schindelin et al., 2012) and ilastik ( Berg et \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n26 \n \na\nl., 2019). Specifically, metaphase cells were automatically identified in multiple pre-defined fields of \nview by low-resolution imaging of the DNA channe l (5-SiR-Hoechst). Subsequently, cells of interest \nwere imaged for the next 150 minutes with a time-resolution of 2 min to capture anaphase onset (AO) \nand 120 minutes of progression through mitotic exit with 31 z-slices with a voxel size of 250 nm in xy \nand 750 nm in z, covering a total of 75x75 µm in xy (300x300 pixels) and 22.5 µm in z, which was \nsufficient to cover the whole cell volume, in th e GFP ((m)EGFP-tagged POI), DNA, Dextran-Dy481XL \n(extracellular space), and transmission channels. A previously developed computational pipeline (Cai \net al., 2018) was adapted to track and segment dividing cells from high-zoom time lapses in 3D based \non the nuclear (SiR-Hoechst) and cellular (Dextran-Dy481XL) landmarks. The third eigenvalue of the \nsegmented chromatin mass, representing the thickn ess of the chromosomal volume, was utilized to \ndetect AO as chromosomes begin to be segregated to wards opposite cell poles. All mitotic exit time-\nseries were aligned to AO and set as the t=0 min timepoint. All individual aligned time-series displayed \na very consistent increase in chromatin volume over time, rendering any further alignment \ndispensable. \nEstimation of protein numbers from FCS-calibrated images \nFluorescence intensities in image voxels were converted to absolute protein concentrations and \nnumbers based on the experiment-specific calibration line (calibration factor (= slope) and background \nintensity) and the 3D binary masks of nucleus and the cell. The average protein concentration was \ncalculated by multiplying the calibration factor (slope of the calibration line) to the average \nbackground corrected fluorescent intensity in all nuclea r, cellular or cytosolic pixels (cytosol = within \nthe cell, but excluding the nucleus). The absolut e protein number inside each compartment was \nachieved by integrating all background-corrected fl uorescent intensities and multiplying them with \nthe calibration factor.  \nFull Cell Cycle Imaging \nAbout 750-1000 genome-edited cells expressing the POI endogenously tagged with EGFP were seeded \ntwo days before the experiment into a 0.34 cm 2 well of an 18-well chambered cover glass (Ibidi µ-\nslide, 81817) and incubated at 37°C, 5% CO2. 20 hours day later, cells were arrested in S-phase for 15-\n16 hours by changing the medium to DMEM supplemented with 2 mM thymidine (T1895, Sigma). Cells \nwere subsequently released from S-phase arrest by washing 3 times with DMEM. 4 hours after release, \nmedium was exchanged to phenol-red free, CO2-independent imaging medium (see above) containing \n50-100 nM 5-SiR-Hoechst and one hour later 500- kDa dextran-Dy481XL was added as a cell outline \nmarker (added later due to interference with efficient SiR-Hoechst staining ). Imaging was started 6 \nhours after release from S-phase, well before the firs t mitotic division. As a control of the effect of S-\nphase arrest, ~3750 asynchronous cells were seeded one day before imaging into a well of an 18-well \nIbidi µ-slide and imaging was carried out 1.5 hours after addition of imaging medium containing 5-SiR-\nHoechst and addition of 500-kDa dextran-Dy481XL. Imaging was carried out on a Zeiss LSM780 and \nLSM880 using a C-Apochromat 40x/1.2 W Korr UV-Vis-IR water-immersion objective (421767-9971-\n711, Zeiss) with a custom-made objective cap for automated water dispension , with a field of view \n(FOV) size of 177.12x177.12 µm covering a z -range of 22.5 µm with 253 nm pixel size in xy and 750 \nnm in z and a pixel dwell time of 0.76 µsec. 0.2% laser power of the 488 nm Argon laser line was used \nto ensure minimal bleaching and GFP fluorescence was recorded on the GaAsP detector (499 nm -553 \nnm range, gain set to 1100). 4 FOV were automa tically imaged every 10 minutes with an autofocus \nstep before every single 3D stack (based on peak reflection of 514 nm laser line at glass-sample \ninterface). Depending on the cell cycle length and whether synchronous or asynchronous cells were \nused, total imaging time varied from 25 to 40 hours, in order to capture two subsequent mitosis events \nfor most cells present in the FOV. Image data was processed using an adapted computational pipeline \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n27 \n \n(\nCai et al., 2018) performing 3D segmentation based on chromatin (5-SiR-Hoechst) and cellular \nlandmarks (500 kDa Dextran), as well as cell tracking of single cells using the 3D centroid of the \nchromatin mass. After manually filtering out duplicat e or poorly segmented si ngle cell tracks, single \ncell cycles were cropped out based on the cellular and nuclear volume information, resulting in a list \nof full cell cycle tracks ranging from one anaphase/telophase to the next. These full cell cycle tracks \nwere aligned to the first division and subsequently interpolated and fit to a common average cell cycle \ntiming. Calibration of the measured fluorescent intensities was performed not through direct FCS-\ncalibrated imaging, but by  setting the number of proteins inside a cell (N_cell) in the second mitosis \n(when the S-phase arrest effect has ceased) to the mean number of proteins inside a cell measured in \nasynchronous FCS-calibrated metaphase cells, resulting in a conversion factor that was used to \ntransform measured fluorescent intensities to ab solute protein numbers and concentrations at all \nother timepoints. While bleaching of GFP-tagged proteins was not tested over the course of an entire \ncell cycle, we assume it to be minimal due low laser exposure (488 nm: 0.2%, pixel dwell: 0.76 µsec, 1 \nstack every 10 min) and the fact that cellular concentrations of all proteins did not change from one \nmitosis to the next.  \n \nSimple Western \nProtein separation, immunodetection and quantificat ion from cell lysates was performed in a Jess \nAutomated Western Blot System (Bio-Techne), using 12-230 kDa and 66-440 kDa Fluorescence \nseparation capillary cartridges (SM-FL004-1, SM-FL 005-1, Bio-Techne). For this, total protein lysates \nwere prepared for each cell line and condition of  interest by growing cells in a 10-cm until ~80% \nconfluency, subsequently washing with PBS and resuspending cells in 500 μl of lysis buffer (RIPA buffer \n(R0278, Sigma-Aldrich), 1 mM PMSF (P7626, Sigma-Aldrich), cOmplete™ EDTA-free Protease Inhibitor \nCocktail (04693132001, Roche, 1 tablet/10 ml) an d PhosSTOP (4906845001, Roche, 1 tablet/10 ml)) \nwith the help of a cell scraper (on ice). Cells were then lysed by two cycles of freezing in liquid nitrogen \nand thawing at 37 °C. After centrifugation for 10 min at ~16,000xg, 4°C, the supernatant containing \nsoluble total protein extracts was separated and kept at -80°C until use. Total protein was quantified \nwith a Pierce BCA Protein Assay Ki t (23227, Thermo Fisher Scientific) and diluted to 0.4 µg/µL final \nconcentration including 1x Master Mix (from EZ St andard Pack 1 (PS-ST01EZ-8, Bio-Techne). Loading \nof samples and detection reagents into the Simple Western (SW) microplate was conducted following \nthe provider’s instructions. Detection was achieved by ECL using anti-rabbit and anti-mouse secondary \nHRP antibodies (042-206/ 042-205, Bio-Techne) and  Luminol-S/Peroxide solution (043-311/043-379, \nBio-Techne). Capillary electrophoresis run and an alysis was conducted wi th the Compass for SW \nsoftware (Bio-Techne) following the provider’s guidelines. \n \nPreparation of homozygous endogenous knock-in cell lines \nGenome-edited cell lines generated in this study (HK Rad21-EGFP-AID CTCF-Halo-3xALFA #C7 and HK \nNup153-mEGFP-FKBP12F36V #C10 (dTAG technology: Nabet et al., 2018) were obtained by C-terminal \ntagging of CTCF and Nup153 in HK RAD21-EGFP-AID (Davidson et al., 2016) or HK WT parental cell \nlines, respectively, using the CRISPR/Cas9 method. In brief, a linear DNA donor sequence encoding for \nthe tag of interest (and corresponding 50 base pair long homology arms) was electroporated into the \nparental cell line, together with the catalytic Cas9 /gRNA ribonucleoparticle complex, as previously \ndescribed (Koch et al., 2018; Kueblbeck et al., 2021 Preprint). For this, we used Alt-R™ S.p. HiFi Cas9 \nNuclease V3 (1081061, IDT) and single gRNAs (see Supplementary Information). Edited cells \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n28 \n \ne\nxpressing the tags of interest were selected by FACS sorting and the correct tagging of all target \ncopies was subsequently validated as described in (Kueblbeck et al., 2021). Expression of the tagged \nprotein of interest (POI) at endogenous levels was confirmed by simple western and confocal \nmicroscopy, the latter also indicating correct subcellular localization of the POI. Homozygous tagging \nof the POI was confirmed by PCR screening, simple western and digital PCR. Digital PCR (dPCR) allows \nto quantify the copy number of specific sequences of interest in a template genome, by partitioning \nthe amplification reaction (including a primer pair and an internal fluorescent probe, per region to be \nquantified) into thousands of nanodroplets, each containing 0-few DNA molecules. Upon amplification \nof the region of interest in a given droplet, the specific internal probe is released from the DNA and \nfluorescence is detected. The count of fluorescent vs non-fluorescent droplets is read out and used to \nquantify the absolute amount of template DNA. The triple-color dPCR assay used in this work allowed \nus to quantify: the total number of tags (“allGFP” or “allHalo”) integrated into the genome, the number \nof tags inserted at the intended target locus (“HD R”, homologous-directed repair after Cas9-directed \nDNA cut) and the copy number of a reference sequence located in the vicinity of the target locus. This \nsetup therefore allows to quantify how many endogenous alleles are tagged, as well as the detection \nof excess off-target tag integrations within the re cipient genome. Finally, the correct sequence and \npositioning of the integrated tags was corroborated by PCR-amplification and sequencing of the edited \ngenomic regions.  \n \nFluorescence recovery after photobleaching \nCells for FRAP measurements were  seeded at a density of 2.5x10 5 cells/ml into Ibidi glass bottom \nµ-Slide channels (80607, Ibidi) one day prior to imaging. DMEM was replaced by CO 2-independent \nimaging medium (as above) containing 50-100 nM 5-SiR-Hoechst at least 1 hour before imaging.  FRAP \nexperiments were performed on a LSM880 laser-scanning microscope with an inverted Axio Observer \ncontrolled by ZEN 2.1 Black software (Version 14.0.9.201, Zeiss), equipped with an in-house \nconstructed incubation chamber for temperature control set to 37°C and using a C-Apochromat \n40x/1.2 W Korr UV-Vis-IR water-immersion objective (421767-9971-711, Zeiss).\n Cells in metaphase \na\nnd early G1 were selected manually based on thei r chromatin staining and FRAP of metaphase cells \nwas performed as described previously (Walther et al., 2018). Cells in G1 stage were selected manually \nbased on nuclear size and filtered out computationally based on a nuclear size threshold of less than \n1050 µm3 corresponding to the size of cells about 5 hours into the cell cycle according to full cell cycle \ndata of asynchronous cells (exact nuclear size was derived from a 3D stack covering the whole \nchromatin mass, segmented with a previously developed script (Cattoglio et al., 2019). A single image \nwas recorded prior to bleaching, recording 5 z-planes in metaphase and early G1, 3 z-planes in G1 with \na pixel size of 213×213×750 nm, pixel dwell 1.7 µsec and a FOV size of 27.25x27.25 µm for metaphase \nand G1 cells and of 42.5x42.5 µm for early G1 cells, respectively in the EGFP (488 nm argon laser line, \nexcitation power: 1%, Ga AsP detection range set to 499 nm - 562 nm, gain set to 1000) and SiR-\nHoechst channels (633 nm diode laser, excitation power 0.2-0.4%, GaAsP detection range set to 641 \nnm - 696 nm, gain set to 1000). Subsequently, a square region covering half of the chromatin / nucleus \narea in the middle z-plane was bleached using si milar laser power for metaphase, early G1 and G1 \ncells (488 nm laser power: 100%). While metaphase pl ates were bleached with one bleach step (45 × \n35 pixels, 150 repetitions), early G1 and G1 cells were bleached 3 times within 30 seconds to \ncompletely bleach the freely diffusion soluble pool (45 × 35 pixels for eG1, 60 x 50 pixels for G1, 3x 50 \nrepetitions), enabling the determination of chromatin-bound fractions. The fluorescent recovery was \nrecorded by time-lapse imaging every 20 seconds for another 30 frames with the settings described \nfor the pre-bleach image, resulting in minimal bleaching throughout the imaging period (<10%).  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n29 \n \nF\nRAP image analysis was performed using a previously developed custom-written ImageJ script \n(Walther et al., 2018), adapted to enable the analysis of metaphase, early G1 and G1 cells at the same \ntime, as well as an R-script for downstream data processing (Walther et al., 2018). In brief, this analysis \nscript aggregates the (m)EGFP-POI and SiR-Hoechst fluorescence intensity data along the major 2D \nchromatin axis (segmented using SiR-Hoechst channel) into a 1D profile. Using a gap of 14 pixels in the \ncenter of the 1D profile, the border of the bleach ing ROI was omitted to avoid boundary effects. The \nweighted mean fluorescence intensi ties (using SiR-Hoechst) in the unbleached and bleached regions \nwere computed as described in (Walther et al., 2018). As in (Gerlich et al., 2006; Walther et al., 2018), \nthe weighted normalized difference between the unbleached and bleached region \n𝐹௨௕ሺ𝑡ሻ െ  𝐹௕ሺ𝑡ሻ\n𝐹௨௕ሺ0ሻ െ  𝐹௕ሺ0ሻ \nw\nas used as a readout for the residence time and immobile fraction. A single exponential function  \n𝑎 ൅ ሺ1 െ𝑎 ሻ 𝑒ିሺ௞೚೑೑ሻ௧ \nwas employed to fit the normalized fluo rescence recovery data. The parameter a represents the \nimmobile fraction and koff is the unbinding rate constant.  \nFRAP to investigate Cohesin-dependence of CTCF chromatin association \nFRAP measurements of CTCF after depletion of RAD21 were carried out in G1 cells of genome-edited \nHK cells in which all alleles of RAD21 were tagged with an AID degron and EGFP and all alleles of CTCF \nwere tagged with Halo (see above). G1 cells were selected based on nuclear volume, but no stringent \nsize filter was applied since the variance of individual measurements was found to be minimal and not \ndependent on nuclear volume. Complete depletion of RAD21 in these genome edited cells was \nachieved by incubation with Inole-3-acetic acid (IAA, I5148, Sigma) for at least 1.5 hours. For rescue of \nRAD21 depletion, exogenous RAD21-EGFP was overexpressed for at least 24 hours prior to the start \nof the experiment. FRAP measurements were carried out as described above, however bleaching and \nimaging of fluorescence recovery was performed us ing 561 nm excitation of the Halo-TMR (G8252, \nPromega) ligand coupled to endogenous CTCF-Hal o (excitation power: 0.7%, GaAsP detection range \nset to 570-624, gain set to 1000) after 10 minutes of labelling with Halo-TMR  at a concentration of \n100 nM at 37°C in imaging medium. Interestingly, we found that CTCF-Halo displayed a reduced \nchromatin residence time and immobile fraction in the absence of IAA, unlike CTCF-EGFP \nendogenously tagged in a different cell line. We found that this correlated with a leaky degradation of \nRAD21 in the RAD21-EGFP-AID CTCF-Halo cell line, reducing RAD21 levels about 40% relative to our \nCTCF-EGFP line (using Simple Western of asynchronous cell lysates, RAD21 detected via anti-RAD21 \nantibody (05-908, Merck Millipore, 1:50, Suppl. Fig. 4G). Overexpression of RAD21 rescued this effect, \nbringing CTCF-Halo residence time and bound fraction almost back to WT levels (data not shown). For \ncomparison with our ΔRAD21 and ΔRAD21+rescue conditions, we therefore decided to use our CTCF-\nmeasurements as WT reference condition.  \n \nCell synchronization by mitotic shake-off \nTo synchronize HK cells in mitosis for subsequent protein degradation or timed release into early G1 \nor G1, we used a combination of Nocodazole treatm ent and a mitotic shake-off. In brief, cells were \nregularly passaged (every second day) and seeded into a T-175 flask (353112, Corning)\n to reach a \nc\nonfluency of around 80% after 16-24 hours of incubation. One hour prior to mitotic shake-off, cells \nwere incubated in 12 mL of DMEM complete me dium supplemented with 82 nM Nocodazole \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n30 \n \n(\nSML1665, Sigma-Aldrich) to enrich mitotic cells.  The mitotic shake-off was conducted by banging 5 \ntimes the cell culture flask on a table covered with ~5 paper tissues. After confirming the detachment \nof most mitotic cells by inspection on a microscope , the mitotic cell suspension was transferred to a \n15 mL Falcon tube and centrifuged for 3 minutes at 90xg. The resulting cell pellet was resuspended in \n150 µL DMEM + 82 nM Nocodazole and the cell density was counted. 35 µL of cells at a desired density \n(between 1.2x106 cells/ml and 2.5x10 6 cells/ml) were seeded into an Ibidi µ-Slide glass bottom slide \n(80607, Ibidi) with channels pre-coated for 15 minutes with poly-L-lysine (P8920, Sigma). Ibidi slides \nwere incubated for 15 minutes at 37°C, 5% CO 2 to allow cells to attach. 100 µL of DMEM complete \nmedium supplemented with 82 nM Nocodazole was added to cells in every Ibidi µ-Slide channel prior \nto any further treatment.  \n \nImmunofluorescence \nFixed cells were prepared for immunostaining by permeabilization with 0.25% Tergitol (15S9, Sigma) \nin PBS for 15 minutes and subsequent incubation in blocking buffer (2% BSA, 0.05% Tergitol in PBS) \nfor at least 30 minutes at room temperature (RT, 20-25°C in this work). Primary antibody incubation \nwas performed in blocking buffer at 4°C in a humidified chamber overnight (16-24 hours), followed by \nwashing with blocking buffer (3 times, 5 min). Secondary antibody hybridization was performed in \nblocking buffer for 1h at RT. After washing with PBS (3 times, 5 min), samples were post-fixed with \n2.4% PFA (15710, EMS) in PBS for 15 minutes, quenched with 100 mM NH 4Cl in PBS for 10 minutes \nand washed in PBS. Samples used for LoopTrace-based chromatin tracing were permeabilized with \nTriton X-100 instead of Tergitol at the same concentration for consistency with previous experiments.   \n \nProtein depletion during mitosis \nFor the degradation of Nup153, SMC4, RAD21 and CTCF during mitosis, we used genome-edited HK \ncells in which all copies of the POI were endogenously tagged with a dTAG degron system (Nup153-\nmEGFP-FKBP12F36V, Nabet et al., 2018, 2020), or an Auxin-inducible degron tag (SMC4-Halo-mAID \n(Schneider et al., 2022), RAD21-EGFP-AID (Davidso n et al., 2016), CTCF-mEGFP-AID (Wutz et al., \n2017)). Nocodazole-assisted mitotic shake-off was conducted as described above and 35 µL of mitotic \ncells were seeded into Ibidi glass bottom µ-slides (80607, Ibidi) pre-coated  with poly-L-lysine (15 \nminutes) at a density of 2-2.5x10 6 cells/ml. Cells were allowed to attach for 15 minutes at 37°C, 5% \nCO2. Subsequently, the depletion of degron-tagged proteins was conducted for 1.5 hours in the \npresence of 82.5 nM Nocodazole and each specific degradation-triggering ligand (Nup153: 250 nM \ndTAG-13 (SML2601, Sigma) & 500 nM dTAG V-1 (6914, Tocris); SMC4: 1 uM 5-Ph-IAA (30-003, \nBioAcademia); RAD21 & CTCF: 500 uM  IAA). Afterwards, cells were re leased into mitotic exit by \nwashing out Nocodazole through cell incubation for 45-90 minutes in fresh medium supplemented \nwith dTAGs, 5-Ph-IAA or IAA, respectively. Then, cells were either pre-extracted by washing in PBS and \nthen incubating with 0.25% Tergitol in PBS for 1 minute followed by PFA-fixation (ΔSMC4, ΔNIPBL), or \nfixed directly with 2.4% PFA in PBS for 15 minutes (ΔNup153), followed by quenching of PFA with 100 \nmM NH 4Cl in PBS and washing with PBS. Immunofluorescence was performed as described above, \nusing the following primary antibodies: mouse anti-RAD21 (05-908, Merck Millipore, 1:500), rabbit \nanti-SMC2 (ab10412, Abcam, 1:1000); rabbit anti-CT CF (07-729, Merck Millipore, 1:2000) or rabbit-\nanti CTCF (Wutz et al., 2020, Glycine Elution, 1;3000). Secondary hybridization was performed using \nfluorescently tagged antibodies: AF647 goat anti-rabbit (A21245, Invitrogen, 1:1000), AF594 goat anti-\nrabbit (A11037, Life Technologies, 1:1000), AF55 5 goat anti-mouse (A28180, Invitrogen, 1:1000) or \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n31 \n \nA\nF594 goat anti-mouse (A11005, Life Technologies, 1:1000). Stained and post-fixed cells were imaged \non a Nikon TI-E2 equipped with a Lasercombiner, a 60X SR P-Apochromat IR AC 60x 1.27 NA water \nimmersion objective, a CSU-W1 SoRa spinning disk unit and an Orca Fusion CMOS camera in spinning \ndisk mode, operated using NIS Elements 5.2.02 (Nikon). Per condition (WT / ΔPOI), at least 5 z-stacks \ncovering a ROI size of 261.46x261.46x21 µm were ac quired in the DAPI channel (405 nm excitation), \nGFP channel (488 nm excitation, degradation control), and immunofluorescence channels (561 or 640 \nexcitation) with a pixel size of about 227 nm in xy and 500 nm in z. \nImage Analysis of mitotic exit degradation samples \nImage analysis of 3D stacks of stained mitotic exit  cells was performed with a custom-written Python \nscript. In brief, after a mild gaussian blur, the DAPI channel was converted to a 3D binary mask of \nnuclei used for 3D segmentation (method = triangle). Small objects and cropped nuclei at the image \nborders were removed automatically, and furthe r quality control to remove poorly segmented, \nmultinucleate or dead cells were removed manually  using napari. Interactive viewing of the nuclei \nimages and binary masks via napari was also used to classify cells as “mitosis” or “interphase” \n(representing all nuclei past anaphase). After classification, the nuclei mask and labels were used to \nextract fluorescent intensities of the endogenous POI-GFP, as well as stained proteins in the \nunprocessed 488 nm, 561 nm and 647 nm (if applicable). Image background from regions devoid of \ncells was subtracted from mean nuclear pixel intensities in every image channel. \n \nSpot-bleach assay and analysis \nCells for spot-bleach measurements were seeded at a density of 2.5x10 5 cells/ml into Ibidi glass \nbottom µ-Slide channels (80607, Ibidi) and grown for 16-24 hours. One hour before imaging, DMEM \nwas replaced by CO 2-independent imaging medium (as above) containing 50-100 nM 5-SiR-Hoechst. \nFRAP experiments were performed on a LSM880 laser-scanning microscope with an inverted Axio \nObserver controlled by ZEN 2.1 Black software (Version 14.0.9.201, Zeiss), equipped with an in-house \nconstructed incubation chamber for temperature control set to 37°C and using a C-Apochromat \n40x/1.2 W Korr UV-Vis-IR water-immersion objective (421767-9971-711, Zeiss). Cells were screened \nat low-resolution live imaging in the SiR-Hoechst channel and image acquisition was started once a \ncell undergoing anaphase onset was identified. At 5, 10, 15, 20 and 30 minutes after anaphase onset, \nan image of the dividing cell in the GFP (488 nm emission) and DNA (SiR-Hoechst, 633 nm emission) \nwas acquired and used to place and initiate a 30 second continuous illumination with a diffraction \nlimited focused laser beam (488 nm, ~1.5 µW laser power, corresponding to 0.1% Argon laser power). \nThis resulted in a clear depletion of the chromatin-bound (m)EGFP-tagged protein pool and minor \nbleaching of the overall cellular pool that readily  replaced the bleached soluble fraction at the \nmeasured spot. Measurement timepoints were distributed between the two daughter cells to further \nminimize light exposure of a single cell. During the 30 seconds illumination, emitted fluorescence was \ncontinuously measured using the GaAsP detector in  photon counting mode. The mean of the first \n(prebleach) and last (postbleach) 500 milliseconds of the fluorescen ce depletion trace was used to \ncalculate the chromatin-bound fraction for each measurement based on the following formula:  \n𝐵𝑜𝑢𝑛𝑑 𝑓𝑟𝑎𝑐𝑡𝑖𝑜𝑛 ൌ  𝑝𝑟𝑒𝑏𝑙𝑒𝑎𝑐ℎ െ  𝑝𝑜𝑠𝑡𝑏𝑙𝑒𝑎𝑐ℎ\n𝑝𝑟𝑒𝑏𝑙𝑒𝑎𝑐ℎ\n 100 \nI\nn addition to the measurements shortly after mitosis, chromatin-bound fractions of each POI were \nmeasured in asynchronous interphase cells. Measured bound fractions were calibrated using \nexogenously H2B-EGFP (low expres sion level, positive control representing ~100% chromatin bound \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n32 \n \nf\nraction) and freely diffusing mEGFP (unbound control, representing 0% chromatin bound fraction) \nexpressed in a HK WT cell background and measured in asynchronous interphase cell nuclei. The \naverage calibrated chromatin-bound fractions of 10 spot-bleach measurements per protein per \ntimepoint was interpolated (the asynchronous in terphase measurements were set to 300 minutes \nafter anaphase onset for this purpose) and used to calculate the average number of chromatin-bound \nPOIs at each timepoint during mitotic exit using the FCS-calibrated protein number information from \nFig. 1EF, Suppl. Fig. 1P.  \n \nCell synchronization and immunofluorescence for chromatin tracing \nTo prepare HK cells expressing AID-EGFP-tagged Cohesin-STAG2 as well as HK WT cells for chromatin \ntracing in interphase, 120 µL of asynchronous AID-tagged and WT cells were seeded at a 1:1 ratio and \na total density of 5x105 cells/ml into PBS-washed channels of Ibidi µ-Slide glass bottom slides (80607, \nIbidi) and cultured for 20 hours at 37°C, 5% CO2 in DMEM supplemented with 40 μM BrdU/BrdC (ratio \n3:1, BrdU: B5002, Sigma-Aldrich, BrdC: sc-284555, Santa Cruz Biotech). Degradation of EGFP-AID-\nSTAG2 was induced by the addition of 500 µM Inole-3-acetic acid (IAA, I5148, Sigma-Aldrich) for 2 \nhours at 37°C, 5% CO 2 in DMEM. Cells were then fixed using 2.4% PFA (15710, EMS) in PBS for 15 \nminutes, followed by quenching of PFA with 100 mM NH4Cl in PBS (5 minutes) and washing with PBS. \nTo prepare cells in early G1, HK WT and STAG2-AID cells were grown for 20 hours in a T-175 flask \n(353112, Corning) in the presence of 40 μM BrdU/BrdC (ratio 3:1) to reach a confluency of around \n80% suitable for mitotic shake off. Nocodazole-arrest, mitotic shake-off and resuspension of mitotic \ncells was performed as described above. Enriched mitotic HK WT and STAG2-AID cells (Wutz et al., \n2020) were diluted to 2.5x10 6 cells/ml, mixed 1:1 and 35 μL of this cell suspension was seeded into \nIbidi µ-Slide glass bottom slides (80607, Ibidi) pre-coated with poly-L-lysine and incubated for 15 \nminutes at 37°C, 5% CO2 to allow cells to attach. Degradation of STAG2 was induced upon addition of \n500 µM IAA in the presence of Nocodazole, ensuring near-complete degradation within 45 minutes. \nRelease into mitotic exit was triggered by Noco dazole washout using DMEM containing 500 µM IAA. \nCells were fixed 80 minutes after release. Live imagin g of cells at this point showed that they are on \naverage about 45 minutes past anaphase. After fixation, early G1 and asynchronous interphase cells \nwere permeabilized for 15 minutes using 0.25% TritonX-100 (T8787, Sigma-Aldrich) in PBS and 0.1 µm \nTetraspec beads were added to the Ibidi channels (1:100 dilution from stock, T7279, Thermo Fisher) \nto be used as fiducials for drift correction. After bl ocking with 2% BSA in 0.05% TritonX-100 at RT for \nat least 30 minutes, primary labelling of STAG2 was performed overnight at 4°C in a humidified \nchamber (with rabbit-anti STAG2, Glycine Elution, 1:200, Sumara et al., 2000), followed by \nhybridization with an AF488-labelled secondary antibody (goat-anti-rabbit AF488, A-11034, Molecular \nProbes).  \n \nNon-denaturing FISH (RASER-FISH) \nNon-denaturing FISH (RASER-FISH) as well as FISH library design and amplification was performed as \ndescribed previously (Beckwith et al., 2023 Preprint & Beckwith, Brunner et al., in preparation). In \nbrief, cells were incubated with 0.5 ng/µl DAPI in PBS at RT for 15 minutes to sensitize DNA for UV-\ninduced single-strand nicking of the replicated strand containing BrdU/C. Subsequently, the cells were \nexposed (without Ibidi lid) to 254 nm UV light for 15 min (Stratalinker 2400 fitted with 15W 254 nm \nbulbs-part no G15T8). The nicked strand of DNA was then digested using Exonuclease (1U/ul, M0206, \nNEB) in NEB buffer 1 at 37 °C for 15 min in a humidified chamber. Cells were post-fixed using 5 mM \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n33 \n \nB\nis(NHS)PEG5 (803537, Sigma-Aldrich) in PBS for 30 minutes at RT to preserve cell fixation during \nprimary FISH library hybridization at 37°C. Hybridization of primary FISH probe libraries targeting 1.2 \nMb regions (Chr14 50.92-52.10 Mb, Chr5 149.50- 150.70 Mb, Chr2 191.11-192.31 Mb) with 12 kb \ngenomic resolution (one trace-spot = tiled set of ~150 FISH probes with common docking handle) was \nperformed by incubation with hybridization buffer (50% formamide (FA, AM9342, Thermo Fisher), \n10% (w/v) dextran sulfate (D8906, Sigma-Aldrich)\n in 2xSSC (AM9763, Thermo Fisher) containing the \nF\nISH probe libraries at a final concentration of 100-200 ng/µL DNA per library for 1-2 nights at 37°C in \na humidified chamber. After primary hybridization, channels were rinsed 3 times with 50% FA in \n2xSSC, washed again twice with 50% FA in 2xSSC for 5 min at RT and finally washed with 2xSSC \ncontaining 0.2% Tween. RNA-DNA hybrids were removed by incubating cells with 0.05 U/µL RNAse H \n(M0297S, NEB) for 20 min at 37 °C in RNAse H buffer (NEB). To image and segment whole 1.2 Mb \ntracing loci, secondary FISH probes serving as bridges between all primary probes of a whole 1.2 Mb \nlocus and a common imager strand were applied at a concentration of 100 nm in secondary \nhybridization buffer (20% Ethylene Carbonate (EC,  E26258, Sigma-Aldrich), 2xSSC) for 20 minutes at \nRT rocking. Secondary probes were then washed with 30% FA in 2XSSC at RT (3 washes, 5 minutes \neach) and 2 additional washes with  2xSSC. Prior to imaging, DNA was stained with 0.5 ng/µl DAPI in \nPBS for 5 minutes at RT. \n \nChromatin Tracing using LoopTrace \n3D DNA trace acquisition using a custom-built auto mated fluidics setup was performed as described \nin Beckwith et al., 2023 (Preprint) and in \nhttps://git.embl.de/grp-ellenberg/tracebot. In brief, 12-mer \nimager strands with 3’ or 5’-azide functionalit y  (Metabion) complementary to the docking handles \nemployed by the primary FISH probe library, as well as the bridged regional barcode probes added \nduring secondary hybridization, were fluorescentl y labelled with Cy3B-alkyne (AAT Bioquest) or \nAtto643-alkyne (Attotec) using click chemistry (ClickTech Oligo Link Kit, Baseclick GmbH) according to \nthe manufacturer’s instructions to enable dual-col or tracing. Fluorescently labelled 12-mer imagers \nwere diluted to a final concentration of 20 nM in 5% EC 2X SSC in a 96 well plate and placed on the \nstage of a custom-built automated fluidics setup based on a GRBL controlled CNC stage (Beckwith et \nal., 2023). Furthermore, a 3-well deep plate containing washing buffer (10 % FA, 2X SSC) and stripping \nbuffer (30% FA, 2XSSC) covered with parafilm, as we ll as a 24-well plate containing imaging buffer \n(0.2X Glucose Oxidase (G7141, Sigma-Aldrich), 1.5 mM TROLOX (238813, Sigma-Aldrich), 10% Glucose, \n50 mM Tris, 2X SSC pH 8.0) were placed on the stage of the automated fluidics setup. A syringe needle \nmounted in place of the CNC drill head was connected to the sample and a CPP1 peristaltic micropump \n(Jobst Technologies, Freiburg, Germany, flow rate of 1 mL/min at maximal speed) using 1 mm i.d. PEEK \nand silicone tubing (VWR), allowing to pull liquids out of the well plates and through the sample \nchannel in an automated manner. Imaging was performed on a Nikon TI-E2 microscope equipped with \na Lasercombiner, a 100X 1.35 NA silicon oil immersion objective, a CSU-W1 SoRa spinning disk unit \nand an Orca Fusion CMOS camera in spinning disk mode, operated using NIS Elements 5.2.02 (Nikon) \nin combination with custom-made Python software for synchronization with automated liquid \nhandling. Prior to sequential imaging, a 3D stack of DAPI-stained nuclei (405 nm excitation), STAG2-\nEGFP fluorescence (488 nm excitation) and the fiducial beads (561 or 640 excitation) was acquired as \na reference stack for cell classification with a pixel size of 130 nm in xy and 300 nm in z at a total size \nof 149.76x149.76 µm in xy and covering a z-rang e of 14.1 (interphase) - 18.3 µm (early G1). \nSubsequently, imager strands were sequentially hybridized for ~ 2 minutes at 20 nM concentration in \n5% EC 2X SSC, washed for 1 minute with washing buffer, imaged after addition of GLOX-based imaging \nbuffer as a 3D stack, stripped for ~2 minutes using stripping buffer and washed again for 1 minute. 3D \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n34 \n \ns\ntacks acquired during sequential imaging had eq ual pixel sizes and z-range as before, but were \nacquired only in the 561 nm or 640 nm channels (100% laser power, 100 msec exposure time, triggered \nacquisition mode), to image fiducial beads and Cy3B or Atto643-labelled imagers, respectively.  \n \n \nAnalysis of LoopTrace data \nProcessing of acquired tracing data was perfor med as described in Beckwith et al., 2023 (Preprint ) \nwith code available under \nhttps://git.embl.de/grp-ellenberg/looptrace. In brief, nd2 image files were \nconverted to OME-ZARR format. Images were drift-corr ected based on cross-correlation and sub-pixel \ndrift was corrected by fitting the fiducial bead signal to a 3D gaussian function and subsequent \ncorrection for calculated sub-pixel drift. Images were deconvolved using the experimental PSF \nextracted from fiducial beads. Identification of  tracing regions was performed based on regional \nbarcodes using an intensity threshold. Detected spot  masks were then used to extract regions of \ninterest for 3D-superlocalisation of individual trace-spots by fitting with a 3D gaussian. Finally, \nextracted traces were corrected for chromatic ab erration between the 561 and 642 image channels \nby affine transformation obtained by least squares fitting of the centroid of fiducial beads imaged in \nboth channels, and traces were assigned to nuclei classified as “interphase”, “early G1” or “mitosis”.  \nThe resulting interphase and early G1 DNA traces were grouped into “WT” or “ΔSTAG2” based on their \nAF488 intensity and the subsequent analysis was performed as described in Beckwith et al., 2023 \n(Preprint). In brief, all fits were quality-controlled for their signal to background ratio, standard \ndeviation of the fit and fit center distance to the regional barcode signal. Traces containing less than \n20 high-quality fitted positions were removed from further analysis. Median pairwise distances were \ncalculated for all 3D coordinates within a single trace and used to  display either pairwise-distance \nmaps or contact maps by calculating the frequency of contacts below a certain 3D distance (set to 120 \nnm). Difference matrices were achieved by subtraction of “dSTAG2” from “WT” pairwise distances. \nScaling plots were generated from pairwise distance metrices as well, essentially plotting all measured \n3D distances for every given genomic distance.  \n \n  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n35 \n \nS\nample preparation for STED microscopy \nTo prepare genome-edited HK cells expressing endogenously EGFP-tagged Cohesin-STAG1/2 for STED \nmicroscopy in early G1 or G1, ce lls were synchronized in mitosis and subsequently released into \nmitotic exit, pre-extracted, PFA-fixed and immuno-stained. Cell synchronization was performed by \nmitotic shake-off as described above. 35 µL of Nocodazole-arrested enriched mitotic cells were added \nat a density of 1.2x10 6 cells/ml (for G1) or 2.5x10 6 cells/ml (for early G1) to pre-washed and poly-L-\nlysine coated channels of Ibidi µ-Slide glass bottom slides (80607, Ibidi) and incubated for 15 minutes \nat 37°C, 5% CO2 to allow cells to attach. Subsequently, 3 washes with fresh DMEM were performed to \nwash out Nocodazole, and cells were allowed to exit mitosis for 45 minutes (for early G1 stage) or 4h \n(for G1 stage) at 37°C, 5% CO 2. Pre-extraction was performed by washing cells once in PBS and then \nadding 0.25% Tergitol in 1X PBS for a total of 1 minute. Cells were then immediately fixed using 2.4% \nPFA in PBS for 15 minutes, followed by quenching of PFA (15710, EMS) with 100 mM NH4Cl in PBS and \nwashing with PBS. Fixed cells were prepared for immuno-staining by an additional 15-minute \npermeabilization (standard IF protocol) in PBS with 0.25% Tergitol and subsequent blocking using \nblocking buffer (2% BSA in 0.05% Tergitol in PBS) for at least 30 minutes at RT. Incubation with the \nanti-GFP nanobody (FluoTag®-X4 anti-GFP conjugated  to Abberior® Star 635P, 1:250 dilution N0304-\nAb635P, NanoTag) and rabbit anti-CTCF antibody (Glycine-Elution, 1:3000, Wutz et. al 2020) was \nperformed in blocking buffer at 4°C in a humidified chamber overnight. Secondary hybridization using \nAF594-conjugated goat-anti-rabbit antibody (1:1000, A11037, Life Technologies) was performed for \n1h at RT. Samples were post-fixed for 15 minutes in 2.4% PFA in PBS, with subsequent quenching (100 \nmM NH4Cl in PBS) and PBS washing. Samples were imaged by STED super-resolution microscopy on \nthe same day.  \n \nSTED microscopy \n2D STED imaging was performed on a Leica Stel laris 8 STED Falcon FLIM microscope (Leica \nMicrosystems) controlled by the Leica LAS X software (4.7.0.28176). Samples were imaged at RT using \na HC PL APO 86x/1.2 W motCORR STED white water immersion objective. The microscope was \nequipped with the SuperK FIANIUM FIB-12 white light laser with laser pulse picker (440-790 nm, Leica \nMicrosystems/NKT), 592 nm continuous wave (cw), 660 nm cw and 775 nm pulsed lasers \n(MPB Communications) and the HyD S, HyD X and HyD R detectors. Diffraction-limited as well as STED \nimaging of CTCF (AF594) and STAG1/2-EGFP (Abberior Star 635P) was performed with excitation at \n590 nm and 645 nm using the white light laser (diffraction-limited/confocal: 3% each, STED: \n590 nm: 9%, 645 nm: 6%). Fluorescence was detecte d with two HyD X detectors using a 601-619 nm \nand a 655-750 nm detection window, respectively. Imaging was performed in xy line sequential mode. \nThe pinhole size was set to 1 airy unit and the pixel size set to 18.88x18.88 nm in xy, resulting in images \ncapturing a region of 19.31x19.31 µm with 1024x1024 pixels.  STED imaging was performed using a \n2D depletion doughnut and 50% power with a 775 nm  depletion laser for supe r-resolved imaging of \nCTCF-AF594 and at 12% excitation power at 775nm for imaging of STAG1/2-Abberior Star 635P. STED \nimages were acquired using 16 line accumulations with a scan-speed of 200 Hz resulting in a pixel \ndwell time of 3.85 µs. STED imaging was performed in FLIM mode and the images were post-processed \nusing tau-STED enhancement with background suppression activated and tau-strength set to 0%. \nCrosstalk between fluorescent channels was quanti fied with the settings described above and found \nto be less than 5% (maybe ref to supplementary figure, if space allows).   \n \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n36 \n \n \n \nS\nTED image analysis \nPre-processing of diffraction-limited and STED images was performed using Fiji using a custom-made \nscript. Nuclear masks were created by segmenting  the CTCF-AF594 diffract ion-limited image after \ngaussian-blurring and used to crop out nuclei in all image channels. In addition, STED images were \nbackground subtracted using the rolling ball algorithm set to a radius of 50 pixels.  \nCo-localization analysis, as well as spot segmentation was performed using custom python scripts. For \nco-localization analysis, Pearson correlation coefficient of CTCF and STAG1/2 was computed based on \ncropped nuclei in the respective STED channel. Spot segmentation was performed by first coarsely \nsegmenting spots inside the nuclear mask based on a common threshold (method: Otsu) after \napplying a mild gaussian blur (sigma = 1). Image noise resulting in excess tiny spots was filtered out \nthrough binary mask erosion and filtering, followed by binary dilation of correctly detected spots. \nCoarsely segmented spots often represent clusters of spot signals and were further segmented using \na combination of local peak finding & watershed. The resulting masks for individual spots were used \nto extract average pixel intensities in the STED and confocal images. Assuming a z-depth of about 500 \nnm, the number of detected spots per μm3 was compared to protein number estimates derived by \nFCS-calibrated imaging of early G1 or G1 cells to estimate the overall labelling efficiency.  \nSTED image simulation \nSTED images were simulated by generating a desired number of randomly localized spots (single \npixels) in an image representing 200 μm2 (or 100 μm3 assuming a z-depth of 500 nm), given the pixel \nsize of 18.88 nm in the images acquired as described above. The randomly distributed spots were \ngaussian blurred (sigma = 2.6) and their pixel intensity was enhanced 6-fold to be distinguishable \nabove a random background. Simulated images were analyzed for co-localization or segmented and \nanalyzed to read out their average spot intensities as described above.  \n \n  \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n37 \n \nR\neferences for Materials & Methods \nBeckwith, K.S., Ø. Ødegård-Fougner, N.R. Morero, C. Barton, F. Schueder, W. Tang, S. Alexander, J.-M. Peters, \nR. Jungmann, E. Birney, and J. Ellenberg. 2023. Nanoscale 3D DNA tracing in single human cells visualizes \nloop extrusion directly in situ. bioRxiv. 2021.04.12.439407. doi:10.1101/2021.04.12.439407. \nBerg, S., D. Kutra, T. Kroeger, C.N. Straehle, B.X. Kausler, C. Haubold, M. Schiegg, J. Ales, T. Beier, M. Rudy, K. \nEren, J.I. Cervantes, B. Xu, F. Beuttenmueller, A. Wolny, C. Zhang, U. Koethe, F.A. Hamprecht, and A. \nKreshuk. 2019. ilastik: interactive machine learning for (bio)image analysis. Nat. Methods 2019 1612. \n16:1226–1232. doi:10.1038/s41592-019-0582-9. \nBucevičius, J., J. Keller-Findeisen, T. Gilat, S.W. Hell, and G. Lukinavičius. 2019. Rhodamine-Hoechst positional \nisomers for highly efficient staining of heterochromatin. Chem. Sci. 10:1962–1970. \ndoi:10.1039/c8sc05082a. \nBuschmann, V., B. Krämer, F. Koberling, R. Macdonald, and S. Rüttinge. 2009. Quantitative FCS: Determination \nof the Confocal Volume by FCS and Bead Scanning with the MicroTime 200. AppNote Quant. FCS. 1–8. \nCai, Y., M.J. Hossain, J.K. Hériché, A.Z. Politi, N. Walther, B. Koch, M. Wachsmuth, B. Nijmeijer, M. Kueblbeck, \nM. Martinic-Kavur, R. Ladurner, S. Alexander, J.M. Peters, and J. Ellenberg. 2018. Experimental and \ncomputational framework for a dynamic protein atlas of human cell division. Nature. 561:411–415. \ndoi:10.1038/s41586-018-0518-z. \nCattoglio, C., I. Pustova, N. Walther, J.J. Ho, M. Hantsche-Grininger, C.J. Inouye, M.J. Hossain, G.M. Dailey, J. \nEllenberg, X. Darzacq, R. Tjian, and A.S. Hansen. 2019. Determining cellular CTCF and cohesin \nabundances to constrain 3D genome models. Elife. 8. doi:10.7554/eLife.40164.001. \nDavidson, I.F., D. Goetz, M.P. Zaczek, M.I. Molodtsov, P.J. Huis in ’t Veld, F. Weissmann, G. Litos, D.A. Cisneros, \nM. Ocampo-Hafalla, R. Ladurner, F. Uhlmann, A. Vaziri, and J. Peters. 2016. Rapid movement and \ntranscriptional re-localization of human cohesin on DNA. EMBO J. 35:2671–2685. \ndoi:10.15252/embj.201695402. \nGerlich, D., B. Koch, F. Dupeux, J.M. Peters, and J. Ellenberg. 2006. Live-Cell Imaging Reveals a Stable Cohesin-\nChromatin Interaction after but Not before DNA Replication. Curr. Biol. 16:1571–1578. \ndoi:10.1016/j.cub.2006.06.068. \nKapusta, P. 2010. Absolute Diffusion Coefficients: Compilation of Reference Data for FCS Calibration. \ndoi:10.1209/0295-5075/83/46001. \nKoch, B., B. Nijmeijer, M. Kueblbeck, Y. Cai, N. Walther, and J. Ellenberg. 2018. Generation and validation of \nhomozygous fluorescent knock-in cells using CRISPR-Cas9 genome editing. Nat. Protoc. 13:1465–1487. \ndoi:10.1038/nprot.2018.042. \nKueblbeck, M., A. Callegari, B. Serrano-Solano, and J. Ellenberg. 2021. Rapid generation of homozygous \nfluorescent knock-in human cells using CRISPR/Cas9 genome editing and validation by automated \nimaging and digital PCR screening. bioRxiv. 2021.06.23.449557. doi:10.1101/2021.06.23.449557. \nNabet, B., F.M. Ferguson, B.K.A. Seong, M. Kuljanin, A.L. Leggett, M.L. Mohardt, A. Robichaud, A.S. Conway, \nD.L. Buckley, J.D. Mancias, J.E. Bradner, K. Stegmaier, and N.S. Gray. 2020. Rapid and direct control of \ntarget protein levels with VHL-recruiting dTAG molecules. Nat. Commun. 2020 111. 11:1–8. \ndoi:10.1038/s41467-020-18377-w. \nNabet, B., J.M. Roberts, D.L. Buckley, J. Paulk, S. Dastjerdi, A. Yang, A.L. Leggett, M.A. Erb, M.A. Lawlor, A. \nSouza, T.G. Scott, S. Vittori, J.A. Perry, J. Qi, G.E. Winter, K.K. Wong, N.S. Gray, and J.E. Bradner. 2018. \nThe dTAG system for immediate and target-specific protein degradation. Nat. Chem. Biol. 2018 145. \n14:431–441. doi:10.1038/s41589-018-0021-8. \nPoliti, A.Z., Y. Cai, N. Walther, M.J. Hossain, B. Koch, M. Wachsmuth, and J. Ellenberg. 2018. Quantitative \nmapping of fluorescently tagged cellular proteins using FCS-calibrated four-dimensional imaging. Nat. \nProtoc. 13:1445–1464. doi:10.1038/nprot.2018.040. \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint \n\n38 \n \nS\nchindelin, J., I. Arganda-Carreras, E. Frise, V. Kaynig, M. Longair, T. Pietzsch, S. Preibisch, C. Rueden, S. \nSaalfeld, B. Schmid, J.Y. Tinevez, D.J. White, V. Hartenstein, K. Eliceiri, P. Tomancak, and A. Cardona. \n2012. Fiji: an open-source platform for biological-image analysis. Nat. Methods 2012 97. 9:676–682. \ndoi:10.1038/nmeth.2019. \nSchneider, M.W.G., B.A. Gibson, S. Otsuka, M.F.D. Spicer, M. Petrovic, C. Blaukopf, C.C.H. Langer, P. Batty, T. \nNagaraju, L.K. Doolittle, M.K. Rosen, and D.W. Gerlich. 2022. A mitotic chromatin phase transition \nprevents perforation by microtubules. Nat. 2022. 1–8. doi:10.1038/s41586-022-05027-y. \nSumara, I., E. Vorlaufer, C. Gieffers, B.H. Peters, and J.-M. Peters. 2000. Characterization of Vertebrate Cohesin \nComplexes and Their Regulation in Prophase. J. Cell Biol. 151:749–761. \nWalther, N., M.J. Hossain, A.Z. Politi, B. Koch, M. Kueblbeck, Ø. Odegard-fougner, M. Lampe, and J. Ellenberg. \n2018. A quantitative map of human Condensins provides new insights into mitotic chromosome \narchitecture. J. Cell Biol. 217:2309–2328. doi:10.1083/jcb.201801048. \nWutz, G., R. Ladurner, B.G. St Hilaire, R.R. Stocsits, K. Nagasaka, B. Pignard, A. Sanborn, W. Tang, C. Varnai, \nM.P. Ivanov, S. Schoenfelder, P. Van Der Lelij, X. Huang, G. Durnberger, E. Roitinger, K. Mechtler, I. Finley \nDavidson, P. Fraser, E. Lieberman-Aiden, and J.M. Peters. 2020. ESCO1 and CTCF enable formation of \nlong chromatin loops by protecting cohesinstag1 from WAPL. Elife. 9. doi:10.7554/eLife.52091. \nWutz, G., C. Várnai, K. Nagasaka, D.A. Cisneros, R.R. Stocsits, W. Tang, S. Schoenfelder, G. Jessberger, M. \nMuhar, M.J. Hossain, N. Walther, B. Koch, M. Kueblbeck, J. Ellenberg, J. Zuber, P. Fraser, and J. Peters. \n2017. Topologically associating domains and chromatin loops depend on cohesin and are regulated by \nCTCF, WAPL, and PDS5 proteins. EMBO J. 36:3573–3599. doi:10.15252/embj.201798004. \n \n \n \n.CC-BY 4.0 International licenseavailable under a \nwas not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made \nThe copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint","source_license":"CC-BY-4.0","license_restricted":false}