Quantitative imaging of loop extruders rebuilding interphase genome architecture after mitosis

for confocal volume determination (Buschmann et al., 2009). This change resulted in a systematic drop of the protein concentrations measured proportional to the change in confocal volume size, compared to previous measurements using AF488-NHS (Politi et al., 2018). Ten FCS- measurements of 1 minute each were performed to estimate the effective confocal volume in the well with Atto488 solution. FCS-measurements of 30 seconds were performed in the nucleus and cytoplasm in WT cells not expressing mEGFP to determine background fluorescence and photon counts. Experiment-specific calibration factors were obtained from interphase cells expressing mEGFP by correlating measured fluorescence intensities and absolute mEGFP concentration calculated from 30 seconds FCS-measurements (Politi et al., 2018). Calibrated 4-dimensional confocal time-lapse imaging was performed on cells expressing the mEGFP- tagged protein of interest (POI) using a combin ation of MyPic macros for ZenBlack software ( https://git.embl.de/grp-ellenberg/mypic), AutoMicTools library (https://git.embl.de/halavaty/AutoMicTools) for ImageJ (Schindelin et al., 2012) and ilastik ( Berg et .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 26 a l., 2019). Specifically, metaphase cells were automatically identified in multiple pre-defined fields of view by low-resolution imaging of the DNA channe l (5-SiR-Hoechst). Subsequently, cells of interest were imaged for the next 150 minutes with a time-resolution of 2 min to capture anaphase onset (AO) and 120 minutes of progression through mitotic exit with 31 z-slices with a voxel size of 250 nm in xy and 750 nm in z, covering a total of 75x75 µm in xy (300x300 pixels) and 22.5 µm in z, which was sufficient to cover the whole cell volume, in th e GFP ((m)EGFP-tagged POI), DNA, Dextran-Dy481XL (extracellular space), and transmission channels. A previously developed computational pipeline (Cai et al., 2018) was adapted to track and segment dividing cells from high-zoom time lapses in 3D based on the nuclear (SiR-Hoechst) and cellular (Dextran-Dy481XL) landmarks. The third eigenvalue of the segmented chromatin mass, representing the thickn ess of the chromosomal volume, was utilized to detect AO as chromosomes begin to be segregated to wards opposite cell poles. All mitotic exit time- series were aligned to AO and set as the t=0 min timepoint. All individual aligned time-series displayed a very consistent increase in chromatin volume over time, rendering any further alignment dispensable. Estimation of protein numbers from FCS-calibrated images Fluorescence intensities in image voxels were converted to absolute protein concentrations and numbers based on the experiment-specific calibration line (calibration factor (= slope) and background intensity) and the 3D binary masks of nucleus and the cell. The average protein concentration was calculated by multiplying the calibration factor (slope of the calibration line) to the average

corrected fluorescent intensity in all nuclea r, cellular or cytosolic pixels (cytosol = within the cell, but excluding the nucleus). The absolut e protein number inside each compartment was achieved by integrating all background-corrected fl uorescent intensities and multiplying them with the calibration factor. Full Cell Cycle Imaging About 750-1000 genome-edited cells expressing the POI endogenously tagged with EGFP were seeded two days before the experiment into a 0.34 cm 2 well of an 18-well chambered cover glass (Ibidi µ- slide, 81817) and incubated at 37°C, 5% CO2. 20 hours day later, cells were arrested in S-phase for 15- 16 hours by changing the medium to DMEM supplemented with 2 mM thymidine (T1895, Sigma). Cells were subsequently released from S-phase arrest by washing 3 times with DMEM. 4 hours after release, medium was exchanged to phenol-red free, CO2-independent imaging medium (see above) containing 50-100 nM 5-SiR-Hoechst and one hour later 500- kDa dextran-Dy481XL was added as a cell outline marker (added later due to interference with efficient SiR-Hoechst staining ). Imaging was started 6 hours after release from S-phase, well before the firs t mitotic division. As a control of the effect of S- phase arrest, ~3750 asynchronous cells were seeded one day before imaging into a well of an 18-well Ibidi µ-slide and imaging was carried out 1.5 hours after addition of imaging medium containing 5-SiR- Hoechst and addition of 500-kDa dextran-Dy481XL. Imaging was carried out on a Zeiss LSM780 and LSM880 using a C-Apochromat 40x/1.2 W Korr UV-Vis-IR water-immersion objective (421767-9971- 711, Zeiss) with a custom-made objective cap for automated water dispension , with a field of view (FOV) size of 177.12x177.12 µm covering a z -range of 22.5 µm with 253 nm pixel size in xy and 750 nm in z and a pixel dwell time of 0.76 µsec. 0.2% laser power of the 488 nm Argon laser line was used to ensure minimal bleaching and GFP fluorescence was recorded on the GaAsP detector (499 nm -553 nm range, gain set to 1100). 4 FOV were automa tically imaged every 10 minutes with an autofocus step before every single 3D stack (based on peak reflection of 514 nm laser line at glass-sample interface). Depending on the cell cycle length and whether synchronous or asynchronous cells were used, total imaging time varied from 25 to 40 hours, in order to capture two subsequent mitosis events for most cells present in the FOV. Image data was processed using an adapted computational pipeline .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 27 ( Cai et al., 2018) performing 3D segmentation based on chromatin (5-SiR-Hoechst) and cellular landmarks (500 kDa Dextran), as well as cell tracking of single cells using the 3D centroid of the chromatin mass. After manually filtering out duplicat e or poorly segmented si ngle cell tracks, single cell cycles were cropped out based on the cellular and nuclear volume information, resulting in a list of full cell cycle tracks ranging from one anaphase/telophase to the next. These full cell cycle tracks were aligned to the first division and subsequently interpolated and fit to a common average cell cycle timing. Calibration of the measured fluorescent intensities was performed not through direct FCS- calibrated imaging, but by setting the number of proteins inside a cell (N_cell) in the second mitosis (when the S-phase arrest effect has ceased) to the mean number of proteins inside a cell measured in asynchronous FCS-calibrated metaphase cells, resulting in a conversion factor that was used to transform measured fluorescent intensities to ab solute protein numbers and concentrations at all other timepoints. While bleaching of GFP-tagged proteins was not tested over the course of an entire cell cycle, we assume it to be minimal due low laser exposure (488 nm: 0.2%, pixel dwell: 0.76 µsec, 1 stack every 10 min) and the fact that cellular concentrations of all proteins did not change from one mitosis to the next. Simple Western Protein separation, immunodetection and quantificat ion from cell lysates was performed in a Jess Automated Western Blot System (Bio-Techne), using 12-230 kDa and 66-440 kDa Fluorescence separation capillary cartridges (SM-FL004-1, SM-FL 005-1, Bio-Techne). For this, total protein lysates were prepared for each cell line and condition of interest by growing cells in a 10-cm until ~80% confluency, subsequently washing with PBS and resuspending cells in 500 μl of lysis buffer (RIPA buffer (R0278, Sigma-Aldrich), 1 mM PMSF (P7626, Sigma-Aldrich), cOmplete™ EDTA-free Protease Inhibitor Cocktail (04693132001, Roche, 1 tablet/10 ml) an d PhosSTOP (4906845001, Roche, 1 tablet/10 ml)) with the help of a cell scraper (on ice). Cells were then lysed by two cycles of freezing in liquid nitrogen and thawing at 37 °C. After centrifugation for 10 min at ~16,000xg, 4°C, the supernatant containing soluble total protein extracts was separated and kept at -80°C until use. Total protein was quantified with a Pierce BCA Protein Assay Ki t (23227, Thermo Fisher Scientific) and diluted to 0.4 µg/µL final concentration including 1x Master Mix (from EZ St andard Pack 1 (PS-ST01EZ-8, Bio-Techne). Loading of samples and detection reagents into the Simple Western (SW) microplate was conducted following the provider’s instructions. Detection was achieved by ECL using anti-rabbit and anti-mouse secondary HRP antibodies (042-206/ 042-205, Bio-Techne) and Luminol-S/Peroxide solution (043-311/043-379, Bio-Techne). Capillary electrophoresis run and an alysis was conducted wi th the Compass for SW software (Bio-Techne) following the provider’s guidelines. Preparation of homozygous endogenous knock-in cell lines Genome-edited cell lines generated in this study (HK Rad21-EGFP-AID CTCF-Halo-3xALFA #C7 and HK Nup153-mEGFP-FKBP12F36V #C10 (dTAG technology: Nabet et al., 2018) were obtained by C-terminal tagging of CTCF and Nup153 in HK RAD21-EGFP-AID (Davidson et al., 2016) or HK WT parental cell lines, respectively, using the CRISPR/Cas9 method. In brief, a linear DNA donor sequence encoding for the tag of interest (and corresponding 50 base pair long homology arms) was electroporated into the parental cell line, together with the catalytic Cas9 /gRNA ribonucleoparticle complex, as previously described (Koch et al., 2018; Kueblbeck et al., 2021 Preprint). For this, we used Alt-R™ S.p. HiFi Cas9 Nuclease V3 (1081061, IDT) and single gRNAs (see Supplementary Information). Edited cells .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 28 e xpressing the tags of interest were selected by FACS sorting and the correct tagging of all target copies was subsequently validated as described in (Kueblbeck et al., 2021). Expression of the tagged protein of interest (POI) at endogenous levels was confirmed by simple western and confocal microscopy, the latter also indicating correct subcellular localization of the POI. Homozygous tagging of the POI was confirmed by PCR screening, simple western and digital PCR. Digital PCR (dPCR) allows to quantify the copy number of specific sequences of interest in a template genome, by partitioning the amplification reaction (including a primer pair and an internal fluorescent probe, per region to be quantified) into thousands of nanodroplets, each containing 0-few DNA molecules. Upon amplification of the region of interest in a given droplet, the specific internal probe is released from the DNA and fluorescence is detected. The count of fluorescent vs non-fluorescent droplets is read out and used to quantify the absolute amount of template DNA. The triple-color dPCR assay used in this work allowed us to quantify: the total number of tags (“allGFP” or “allHalo”) integrated into the genome, the number of tags inserted at the intended target locus (“HD R”, homologous-directed repair after Cas9-directed DNA cut) and the copy number of a reference sequence located in the vicinity of the target locus. This setup therefore allows to quantify how many endogenous alleles are tagged, as well as the detection of excess off-target tag integrations within the re cipient genome. Finally, the correct sequence and positioning of the integrated tags was corroborated by PCR-amplification and sequencing of the edited genomic regions. Fluorescence recovery after photobleaching Cells for FRAP measurements were seeded at a density of 2.5x10 5 cells/ml into Ibidi glass bottom µ-Slide channels (80607, Ibidi) one day prior to imaging. DMEM was replaced by CO 2-independent imaging medium (as above) containing 50-100 nM 5-SiR-Hoechst at least 1 hour before imaging. FRAP experiments were performed on a LSM880 laser-scanning microscope with an inverted Axio Observer controlled by ZEN 2.1 Black software (Version 14.0.9.201, Zeiss), equipped with an in-house constructed incubation chamber for temperature control set to 37°C and using a C-Apochromat 40x/1.2 W Korr UV-Vis-IR water-immersion objective (421767-9971-711, Zeiss). Cells in metaphase a nd early G1 were selected manually based on thei r chromatin staining and FRAP of metaphase cells was performed as described previously (Walther et al., 2018). Cells in G1 stage were selected manually based on nuclear size and filtered out computationally based on a nuclear size threshold of less than 1050 µm3 corresponding to the size of cells about 5 hours into the cell cycle according to full cell cycle data of asynchronous cells (exact nuclear size was derived from a 3D stack covering the whole chromatin mass, segmented with a previously developed script (Cattoglio et al., 2019). A single image was recorded prior to bleaching, recording 5 z-planes in metaphase and early G1, 3 z-planes in G1 with a pixel size of 213×213×750 nm, pixel dwell 1.7 µsec and a FOV size of 27.25x27.25 µm for metaphase and G1 cells and of 42.5x42.5 µm for early G1 cells, respectively in the EGFP (488 nm argon laser line, excitation power: 1%, Ga AsP detection range set to 499 nm - 562 nm, gain set to 1000) and SiR- Hoechst channels (633 nm diode laser, excitation power 0.2-0.4%, GaAsP detection range set to 641 nm - 696 nm, gain set to 1000). Subsequently, a square region covering half of the chromatin / nucleus area in the middle z-plane was bleached using si milar laser power for metaphase, early G1 and G1 cells (488 nm laser power: 100%). While metaphase pl ates were bleached with one bleach step (45 × 35 pixels, 150 repetitions), early G1 and G1 cells were bleached 3 times within 30 seconds to completely bleach the freely diffusion soluble pool (45 × 35 pixels for eG1, 60 x 50 pixels for G1, 3x 50 repetitions), enabling the determination of chromatin-bound fractions. The fluorescent recovery was recorded by time-lapse imaging every 20 seconds for another 30 frames with the settings described for the pre-bleach image, resulting in minimal bleaching throughout the imaging period (<10%). .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 29 F RAP image analysis was performed using a previously developed custom-written ImageJ script (Walther et al., 2018), adapted to enable the analysis of metaphase, early G1 and G1 cells at the same time, as well as an R-script for downstream data processing (Walther et al., 2018). In brief, this analysis script aggregates the (m)EGFP-POI and SiR-Hoechst fluorescence intensity data along the major 2D chromatin axis (segmented using SiR-Hoechst channel) into a 1D profile. Using a gap of 14 pixels in the center of the 1D profile, the border of the bleach ing ROI was omitted to avoid boundary effects. The weighted mean fluorescence intensi ties (using SiR-Hoechst) in the unbleached and bleached regions were computed as described in (Walther et al., 2018). As in (Gerlich et al., 2006; Walther et al., 2018), the weighted normalized difference between the unbleached and bleached region 𝐹௨௕ሺ𝑡ሻ െ 𝐹௕ሺ𝑡ሻ 𝐹௨௕ሺ0ሻ െ 𝐹௕ሺ0ሻ w as used as a readout for the residence time and immobile fraction. A single exponential function 𝑎 ൅ ሺ1 െ𝑎 ሻ 𝑒ିሺ௞೚೑೑ሻ௧ was employed to fit the normalized fluo rescence recovery data. The parameter a represents the immobile fraction and koff is the unbinding rate constant. FRAP to investigate Cohesin-dependence of CTCF chromatin association FRAP measurements of CTCF after depletion of RAD21 were carried out in G1 cells of genome-edited HK cells in which all alleles of RAD21 were tagged with an AID degron and EGFP and all alleles of CTCF were tagged with Halo (see above). G1 cells were selected based on nuclear volume, but no stringent size filter was applied since the variance of individual measurements was found to be minimal and not dependent on nuclear volume. Complete depletion of RAD21 in these genome edited cells was achieved by incubation with Inole-3-acetic acid (IAA, I5148, Sigma) for at least 1.5 hours. For rescue of RAD21 depletion, exogenous RAD21-EGFP was overexpressed for at least 24 hours prior to the start of the experiment. FRAP measurements were carried out as described above, however bleaching and imaging of fluorescence recovery was performed us ing 561 nm excitation of the Halo-TMR (G8252, Promega) ligand coupled to endogenous CTCF-Hal o (excitation power: 0.7%, GaAsP detection range set to 570-624, gain set to 1000) after 10 minutes of labelling with Halo-TMR at a concentration of 100 nM at 37°C in imaging medium. Interestingly, we found that CTCF-Halo displayed a reduced chromatin residence time and immobile fraction in the absence of IAA, unlike CTCF-EGFP endogenously tagged in a different cell line. We found that this correlated with a leaky degradation of RAD21 in the RAD21-EGFP-AID CTCF-Halo cell line, reducing RAD21 levels about 40% relative to our CTCF-EGFP line (using Simple Western of asynchronous cell lysates, RAD21 detected via anti-RAD21 antibody (05-908, Merck Millipore, 1:50, Suppl. Fig. 4G). Overexpression of RAD21 rescued this effect, bringing CTCF-Halo residence time and bound fraction almost back to WT levels (data not shown). For comparison with our ΔRAD21 and ΔRAD21+rescue conditions, we therefore decided to use our CTCF- measurements as WT reference condition. Cell synchronization by mitotic shake-off To synchronize HK cells in mitosis for subsequent protein degradation or timed release into early G1 or G1, we used a combination of Nocodazole treatm ent and a mitotic shake-off. In brief, cells were regularly passaged (every second day) and seeded into a T-175 flask (353112, Corning) to reach a c onfluency of around 80% after 16-24 hours of incubation. One hour prior to mitotic shake-off, cells were incubated in 12 mL of DMEM complete me dium supplemented with 82 nM Nocodazole .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 30 ( SML1665, Sigma-Aldrich) to enrich mitotic cells. The mitotic shake-off was conducted by banging 5 times the cell culture flask on a table covered with ~5 paper tissues. After confirming the detachment of most mitotic cells by inspection on a microscope , the mitotic cell suspension was transferred to a 15 mL Falcon tube and centrifuged for 3 minutes at 90xg. The resulting cell pellet was resuspended in 150 µL DMEM + 82 nM Nocodazole and the cell density was counted. 35 µL of cells at a desired density (between 1.2x106 cells/ml and 2.5x10 6 cells/ml) were seeded into an Ibidi µ-Slide glass bottom slide (80607, Ibidi) with channels pre-coated for 15 minutes with poly-L-lysine (P8920, Sigma). Ibidi slides were incubated for 15 minutes at 37°C, 5% CO 2 to allow cells to attach. 100 µL of DMEM complete medium supplemented with 82 nM Nocodazole was added to cells in every Ibidi µ-Slide channel prior to any further treatment. Immunofluorescence Fixed cells were prepared for immunostaining by permeabilization with 0.25% Tergitol (15S9, Sigma) in PBS for 15 minutes and subsequent incubation in blocking buffer (2% BSA, 0.05% Tergitol in PBS) for at least 30 minutes at room temperature (RT, 20-25°C in this work). Primary antibody incubation was performed in blocking buffer at 4°C in a humidified chamber overnight (16-24 hours), followed by washing with blocking buffer (3 times, 5 min). Secondary antibody hybridization was performed in blocking buffer for 1h at RT. After washing with PBS (3 times, 5 min), samples were post-fixed with 2.4% PFA (15710, EMS) in PBS for 15 minutes, quenched with 100 mM NH 4Cl in PBS for 10 minutes and washed in PBS. Samples used for LoopTrace-based chromatin tracing were permeabilized with Triton X-100 instead of Tergitol at the same concentration for consistency with previous experiments. Protein depletion during mitosis For the degradation of Nup153, SMC4, RAD21 and CTCF during mitosis, we used genome-edited HK cells in which all copies of the POI were endogenously tagged with a dTAG degron system (Nup153- mEGFP-FKBP12F36V, Nabet et al., 2018, 2020), or an Auxin-inducible degron tag (SMC4-Halo-mAID (Schneider et al., 2022), RAD21-EGFP-AID (Davidso n et al., 2016), CTCF-mEGFP-AID (Wutz et al., 2017)). Nocodazole-assisted mitotic shake-off was conducted as described above and 35 µL of mitotic cells were seeded into Ibidi glass bottom µ-slides (80607, Ibidi) pre-coated with poly-L-lysine (15 minutes) at a density of 2-2.5x10 6 cells/ml. Cells were allowed to attach for 15 minutes at 37°C, 5% CO2. Subsequently, the depletion of degron-tagged proteins was conducted for 1.5 hours in the presence of 82.5 nM Nocodazole and each specific degradation-triggering ligand (Nup153: 250 nM dTAG-13 (SML2601, Sigma) & 500 nM dTAG V-1 (6914, Tocris); SMC4: 1 uM 5-Ph-IAA (30-003, BioAcademia); RAD21 & CTCF: 500 uM IAA). Afterwards, cells were re leased into mitotic exit by washing out Nocodazole through cell incubation for 45-90 minutes in fresh medium supplemented with dTAGs, 5-Ph-IAA or IAA, respectively. Then, cells were either pre-extracted by washing in PBS and then incubating with 0.25% Tergitol in PBS for 1 minute followed by PFA-fixation (ΔSMC4, ΔNIPBL), or fixed directly with 2.4% PFA in PBS for 15 minutes (ΔNup153), followed by quenching of PFA with 100 mM NH 4Cl in PBS and washing with PBS. Immunofluorescence was performed as described above, using the following primary antibodies: mouse anti-RAD21 (05-908, Merck Millipore, 1:500), rabbit anti-SMC2 (ab10412, Abcam, 1:1000); rabbit anti-CT CF (07-729, Merck Millipore, 1:2000) or rabbit- anti CTCF (Wutz et al., 2020, Glycine Elution, 1;3000). Secondary hybridization was performed using fluorescently tagged antibodies: AF647 goat anti-rabbit (A21245, Invitrogen, 1:1000), AF594 goat anti- rabbit (A11037, Life Technologies, 1:1000), AF55 5 goat anti-mouse (A28180, Invitrogen, 1:1000) or .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 31 A F594 goat anti-mouse (A11005, Life Technologies, 1:1000). Stained and post-fixed cells were imaged on a Nikon TI-E2 equipped with a Lasercombiner, a 60X SR P-Apochromat IR AC 60x 1.27 NA water immersion objective, a CSU-W1 SoRa spinning disk unit and an Orca Fusion CMOS camera in spinning disk mode, operated using NIS Elements 5.2.02 (Nikon). Per condition (WT / ΔPOI), at least 5 z-stacks covering a ROI size of 261.46x261.46x21 µm were ac quired in the DAPI channel (405 nm excitation), GFP channel (488 nm excitation, degradation control), and immunofluorescence channels (561 or 640 excitation) with a pixel size of about 227 nm in xy and 500 nm in z. Image Analysis of mitotic exit degradation samples Image analysis of 3D stacks of stained mitotic exit cells was performed with a custom-written Python script. In brief, after a mild gaussian blur, the DAPI channel was converted to a 3D binary mask of nuclei used for 3D segmentation (method = triangle). Small objects and cropped nuclei at the image borders were removed automatically, and furthe r quality control to remove poorly segmented, multinucleate or dead cells were removed manually using napari. Interactive viewing of the nuclei images and binary masks via napari was also used to classify cells as “mitosis” or “interphase” (representing all nuclei past anaphase). After classification, the nuclei mask and labels were used to extract fluorescent intensities of the endogenous POI-GFP, as well as stained proteins in the unprocessed 488 nm, 561 nm and 647 nm (if applicable). Image background from regions devoid of cells was subtracted from mean nuclear pixel intensities in every image channel. Spot-bleach assay and analysis Cells for spot-bleach measurements were seeded at a density of 2.5x10 5 cells/ml into Ibidi glass bottom µ-Slide channels (80607, Ibidi) and grown for 16-24 hours. One hour before imaging, DMEM was replaced by CO 2-independent imaging medium (as above) containing 50-100 nM 5-SiR-Hoechst. FRAP experiments were performed on a LSM880 laser-scanning microscope with an inverted Axio Observer controlled by ZEN 2.1 Black software (Version 14.0.9.201, Zeiss), equipped with an in-house constructed incubation chamber for temperature control set to 37°C and using a C-Apochromat 40x/1.2 W Korr UV-Vis-IR water-immersion objective (421767-9971-711, Zeiss). Cells were screened at low-resolution live imaging in the SiR-Hoechst channel and image acquisition was started once a cell undergoing anaphase onset was identified. At 5, 10, 15, 20 and 30 minutes after anaphase onset, an image of the dividing cell in the GFP (488 nm emission) and DNA (SiR-Hoechst, 633 nm emission) was acquired and used to place and initiate a 30 second continuous illumination with a diffraction limited focused laser beam (488 nm, ~1.5 µW laser power, corresponding to 0.1% Argon laser power). This resulted in a clear depletion of the chromatin-bound (m)EGFP-tagged protein pool and minor bleaching of the overall cellular pool that readily replaced the bleached soluble fraction at the measured spot. Measurement timepoints were distributed between the two daughter cells to further minimize light exposure of a single cell. During the 30 seconds illumination, emitted fluorescence was continuously measured using the GaAsP detector in photon counting mode. The mean of the first (prebleach) and last (postbleach) 500 milliseconds of the fluorescen ce depletion trace was used to calculate the chromatin-bound fraction for each measurement based on the following formula: 𝐵𝑜𝑢𝑛𝑑 𝑓𝑟𝑎𝑐𝑡𝑖𝑜𝑛 ൌ 𝑝𝑟𝑒𝑏𝑙𝑒𝑎𝑐ℎ െ 𝑝𝑜𝑠𝑡𝑏𝑙𝑒𝑎𝑐ℎ 𝑝𝑟𝑒𝑏𝑙𝑒𝑎𝑐ℎ 100 I n addition to the measurements shortly after mitosis, chromatin-bound fractions of each POI were measured in asynchronous interphase cells. Measured bound fractions were calibrated using exogenously H2B-EGFP (low expres sion level, positive control representing ~100% chromatin bound .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 32 f raction) and freely diffusing mEGFP (unbound control, representing 0% chromatin bound fraction) expressed in a HK WT cell background and measured in asynchronous interphase cell nuclei. The average calibrated chromatin-bound fractions of 10 spot-bleach measurements per protein per timepoint was interpolated (the asynchronous in terphase measurements were set to 300 minutes after anaphase onset for this purpose) and used to calculate the average number of chromatin-bound POIs at each timepoint during mitotic exit using the FCS-calibrated protein number information from Fig. 1EF, Suppl. Fig. 1P. Cell synchronization and immunofluorescence for chromatin tracing To prepare HK cells expressing AID-EGFP-tagged Cohesin-STAG2 as well as HK WT cells for chromatin tracing in interphase, 120 µL of asynchronous AID-tagged and WT cells were seeded at a 1:1 ratio and a total density of 5x105 cells/ml into PBS-washed channels of Ibidi µ-Slide glass bottom slides (80607, Ibidi) and cultured for 20 hours at 37°C, 5% CO2 in DMEM supplemented with 40 μM BrdU/BrdC (ratio 3:1, BrdU: B5002, Sigma-Aldrich, BrdC: sc-284555, Santa Cruz Biotech). Degradation of EGFP-AID- STAG2 was induced by the addition of 500 µM Inole-3-acetic acid (IAA, I5148, Sigma-Aldrich) for 2 hours at 37°C, 5% CO 2 in DMEM. Cells were then fixed using 2.4% PFA (15710, EMS) in PBS for 15 minutes, followed by quenching of PFA with 100 mM NH4Cl in PBS (5 minutes) and washing with PBS. To prepare cells in early G1, HK WT and STAG2-AID cells were grown for 20 hours in a T-175 flask (353112, Corning) in the presence of 40 μM BrdU/BrdC (ratio 3:1) to reach a confluency of around 80% suitable for mitotic shake off. Nocodazole-arrest, mitotic shake-off and resuspension of mitotic cells was performed as described above. Enriched mitotic HK WT and STAG2-AID cells (Wutz et al., 2020) were diluted to 2.5x10 6 cells/ml, mixed 1:1 and 35 μL of this cell suspension was seeded into Ibidi µ-Slide glass bottom slides (80607, Ibidi) pre-coated with poly-L-lysine and incubated for 15 minutes at 37°C, 5% CO2 to allow cells to attach. Degradation of STAG2 was induced upon addition of 500 µM IAA in the presence of Nocodazole, ensuring near-complete degradation within 45 minutes. Release into mitotic exit was triggered by Noco dazole washout using DMEM containing 500 µM IAA. Cells were fixed 80 minutes after release. Live imagin g of cells at this point showed that they are on average about 45 minutes past anaphase. After fixation, early G1 and asynchronous interphase cells were permeabilized for 15 minutes using 0.25% TritonX-100 (T8787, Sigma-Aldrich) in PBS and 0.1 µm Tetraspec beads were added to the Ibidi channels (1:100 dilution from stock, T7279, Thermo Fisher) to be used as fiducials for drift correction. After bl ocking with 2% BSA in 0.05% TritonX-100 at RT for at least 30 minutes, primary labelling of STAG2 was performed overnight at 4°C in a humidified chamber (with rabbit-anti STAG2, Glycine Elution, 1:200, Sumara et al., 2000), followed by hybridization with an AF488-labelled secondary antibody (goat-anti-rabbit AF488, A-11034, Molecular Probes). Non-denaturing FISH (RASER-FISH) Non-denaturing FISH (RASER-FISH) as well as FISH library design and amplification was performed as described previously (Beckwith et al., 2023 Preprint & Beckwith, Brunner et al., in preparation). In brief, cells were incubated with 0.5 ng/µl DAPI in PBS at RT for 15 minutes to sensitize DNA for UV- induced single-strand nicking of the replicated strand containing BrdU/C. Subsequently, the cells were exposed (without Ibidi lid) to 254 nm UV light for 15 min (Stratalinker 2400 fitted with 15W 254 nm bulbs-part no G15T8). The nicked strand of DNA was then digested using Exonuclease (1U/ul, M0206, NEB) in NEB buffer 1 at 37 °C for 15 min in a humidified chamber. Cells were post-fixed using 5 mM .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 33 B is(NHS)PEG5 (803537, Sigma-Aldrich) in PBS for 30 minutes at RT to preserve cell fixation during primary FISH library hybridization at 37°C. Hybridization of primary FISH probe libraries targeting 1.2 Mb regions (Chr14 50.92-52.10 Mb, Chr5 149.50- 150.70 Mb, Chr2 191.11-192.31 Mb) with 12 kb genomic resolution (one trace-spot = tiled set of ~150 FISH probes with common docking handle) was performed by incubation with hybridization buffer (50% formamide (FA, AM9342, Thermo Fisher), 10% (w/v) dextran sulfate (D8906, Sigma-Aldrich) in 2xSSC (AM9763, Thermo Fisher) containing the F ISH probe libraries at a final concentration of 100-200 ng/µL DNA per library for 1-2 nights at 37°C in a humidified chamber. After primary hybridization, channels were rinsed 3 times with 50% FA in 2xSSC, washed again twice with 50% FA in 2xSSC for 5 min at RT and finally washed with 2xSSC containing 0.2% Tween. RNA-DNA hybrids were removed by incubating cells with 0.05 U/µL RNAse H (M0297S, NEB) for 20 min at 37 °C in RNAse H buffer (NEB). To image and segment whole 1.2 Mb tracing loci, secondary FISH probes serving as bridges between all primary probes of a whole 1.2 Mb locus and a common imager strand were applied at a concentration of 100 nm in secondary hybridization buffer (20% Ethylene Carbonate (EC, E26258, Sigma-Aldrich), 2xSSC) for 20 minutes at RT rocking. Secondary probes were then washed with 30% FA in 2XSSC at RT (3 washes, 5 minutes each) and 2 additional washes with 2xSSC. Prior to imaging, DNA was stained with 0.5 ng/µl DAPI in PBS for 5 minutes at RT. Chromatin Tracing using LoopTrace 3D DNA trace acquisition using a custom-built auto mated fluidics setup was performed as described in Beckwith et al., 2023 (Preprint) and in https://git.embl.de/grp-ellenberg/tracebot. In brief, 12-mer imager strands with 3’ or 5’-azide functionalit y (Metabion) complementary to the docking handles employed by the primary FISH probe library, as well as the bridged regional barcode probes added during secondary hybridization, were fluorescentl y labelled with Cy3B-alkyne (AAT Bioquest) or Atto643-alkyne (Attotec) using click chemistry (ClickTech Oligo Link Kit, Baseclick GmbH) according to the manufacturer’s instructions to enable dual-col or tracing. Fluorescently labelled 12-mer imagers were diluted to a final concentration of 20 nM in 5% EC 2X SSC in a 96 well plate and placed on the stage of a custom-built automated fluidics setup based on a GRBL controlled CNC stage (Beckwith et al., 2023). Furthermore, a 3-well deep plate containing washing buffer (10 % FA, 2X SSC) and stripping buffer (30% FA, 2XSSC) covered with parafilm, as we ll as a 24-well plate containing imaging buffer (0.2X Glucose Oxidase (G7141, Sigma-Aldrich), 1.5 mM TROLOX (238813, Sigma-Aldrich), 10% Glucose, 50 mM Tris, 2X SSC pH 8.0) were placed on the stage of the automated fluidics setup. A syringe needle mounted in place of the CNC drill head was connected to the sample and a CPP1 peristaltic micropump (Jobst Technologies, Freiburg, Germany, flow rate of 1 mL/min at maximal speed) using 1 mm i.d. PEEK and silicone tubing (VWR), allowing to pull liquids out of the well plates and through the sample channel in an automated manner. Imaging was performed on a Nikon TI-E2 microscope equipped with a Lasercombiner, a 100X 1.35 NA silicon oil immersion objective, a CSU-W1 SoRa spinning disk unit and an Orca Fusion CMOS camera in spinning disk mode, operated using NIS Elements 5.2.02 (Nikon) in combination with custom-made Python software for synchronization with automated liquid handling. Prior to sequential imaging, a 3D stack of DAPI-stained nuclei (405 nm excitation), STAG2- EGFP fluorescence (488 nm excitation) and the fiducial beads (561 or 640 excitation) was acquired as a reference stack for cell classification with a pixel size of 130 nm in xy and 300 nm in z at a total size of 149.76x149.76 µm in xy and covering a z-rang e of 14.1 (interphase) - 18.3 µm (early G1). Subsequently, imager strands were sequentially hybridized for ~ 2 minutes at 20 nM concentration in 5% EC 2X SSC, washed for 1 minute with washing buffer, imaged after addition of GLOX-based imaging buffer as a 3D stack, stripped for ~2 minutes using stripping buffer and washed again for 1 minute. 3D .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 34 s tacks acquired during sequential imaging had eq ual pixel sizes and z-range as before, but were acquired only in the 561 nm or 640 nm channels (100% laser power, 100 msec exposure time, triggered acquisition mode), to image fiducial beads and Cy3B or Atto643-labelled imagers, respectively. Analysis of LoopTrace data Processing of acquired tracing data was perfor med as described in Beckwith et al., 2023 (Preprint ) with code available under https://git.embl.de/grp-ellenberg/looptrace. In brief, nd2 image files were converted to OME-ZARR format. Images were drift-corr ected based on cross-correlation and sub-pixel drift was corrected by fitting the fiducial bead signal to a 3D gaussian function and subsequent correction for calculated sub-pixel drift. Images were deconvolved using the experimental PSF extracted from fiducial beads. Identification of tracing regions was performed based on regional barcodes using an intensity threshold. Detected spot masks were then used to extract regions of interest for 3D-superlocalisation of individual trace-spots by fitting with a 3D gaussian. Finally, extracted traces were corrected for chromatic ab erration between the 561 and 642 image channels by affine transformation obtained by least squares fitting of the centroid of fiducial beads imaged in both channels, and traces were assigned to nuclei classified as “interphase”, “early G1” or “mitosis”. The resulting interphase and early G1 DNA traces were grouped into “WT” or “ΔSTAG2” based on their AF488 intensity and the subsequent analysis was performed as described in Beckwith et al., 2023 (Preprint). In brief, all fits were quality-controlled for their signal to background ratio, standard deviation of the fit and fit center distance to the regional barcode signal. Traces containing less than 20 high-quality fitted positions were removed from further analysis. Median pairwise distances were calculated for all 3D coordinates within a single trace and used to display either pairwise-distance maps or contact maps by calculating the frequency of contacts below a certain 3D distance (set to 120 nm). Difference matrices were achieved by subtraction of “dSTAG2” from “WT” pairwise distances. Scaling plots were generated from pairwise distance metrices as well, essentially plotting all measured 3D distances for every given genomic distance. .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 35 S ample preparation for STED microscopy To prepare genome-edited HK cells expressing endogenously EGFP-tagged Cohesin-STAG1/2 for STED microscopy in early G1 or G1, ce lls were synchronized in mitosis and subsequently released into mitotic exit, pre-extracted, PFA-fixed and immuno-stained. Cell synchronization was performed by mitotic shake-off as described above. 35 µL of Nocodazole-arrested enriched mitotic cells were added at 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- lysine coated channels of Ibidi µ-Slide glass bottom slides (80607, Ibidi) and incubated for 15 minutes at 37°C, 5% CO2 to allow cells to attach. Subsequently, 3 washes with fresh DMEM were performed to wash out Nocodazole, and cells were allowed to exit mitosis for 45 minutes (for early G1 stage) or 4h (for G1 stage) at 37°C, 5% CO 2. Pre-extraction was performed by washing cells once in PBS and then adding 0.25% Tergitol in 1X PBS for a total of 1 minute. Cells were then immediately fixed using 2.4% PFA in PBS for 15 minutes, followed by quenching of PFA (15710, EMS) with 100 mM NH4Cl in PBS and washing with PBS. Fixed cells were prepared for immuno-staining by an additional 15-minute permeabilization (standard IF protocol) in PBS with 0.25% Tergitol and subsequent blocking using blocking buffer (2% BSA in 0.05% Tergitol in PBS) for at least 30 minutes at RT. Incubation with the anti-GFP nanobody (FluoTag®-X4 anti-GFP conjugated to Abberior® Star 635P, 1:250 dilution N0304- Ab635P, NanoTag) and rabbit anti-CTCF antibody (Glycine-Elution, 1:3000, Wutz et. al 2020) was performed in blocking buffer at 4°C in a humidified chamber overnight. Secondary hybridization using AF594-conjugated goat-anti-rabbit antibody (1:1000, A11037, Life Technologies) was performed for 1h at RT. Samples were post-fixed for 15 minutes in 2.4% PFA in PBS, with subsequent quenching (100 mM NH4Cl in PBS) and PBS washing. Samples were imaged by STED super-resolution microscopy on the same day. STED microscopy 2D STED imaging was performed on a Leica Stel laris 8 STED Falcon FLIM microscope (Leica Microsystems) controlled by the Leica LAS X software (4.7.0.28176). Samples were imaged at RT using a HC PL APO 86x/1.2 W motCORR STED white water immersion objective. The microscope was equipped with the SuperK FIANIUM FIB-12 white light laser with laser pulse picker (440-790 nm, Leica Microsystems/NKT), 592 nm continuous wave (cw), 660 nm cw and 775 nm pulsed lasers (MPB Communications) and the HyD S, HyD X and HyD R detectors. Diffraction-limited as well as STED imaging of CTCF (AF594) and STAG1/2-EGFP (Abberior Star 635P) was performed with excitation at 590 nm and 645 nm using the white light laser (diffraction-limited/confocal: 3% each, STED: 590 nm: 9%, 645 nm: 6%). Fluorescence was detecte d with two HyD X detectors using a 601-619 nm and a 655-750 nm detection window, respectively. Imaging was performed in xy line sequential mode. The pinhole size was set to 1 airy unit and the pixel size set to 18.88x18.88 nm in xy, resulting in images capturing a region of 19.31x19.31 µm with 1024x1024 pixels. STED imaging was performed using a 2D depletion doughnut and 50% power with a 775 nm depletion laser for supe r-resolved imaging of CTCF-AF594 and at 12% excitation power at 775nm for imaging of STAG1/2-Abberior Star 635P. STED images were acquired using 16 line accumulations with a scan-speed of 200 Hz resulting in a pixel dwell time of 3.85 µs. STED imaging was performed in FLIM mode and the images were post-processed using tau-STED enhancement with background suppression activated and tau-strength set to 0%. Crosstalk between fluorescent channels was quanti fied with the settings described above and found to be less than 5% (maybe ref to supplementary figure, if space allows). .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made The copyright holder for this preprint (whichthis version posted May 30, 2024. ; https://doi.org/10.1101/2024.05.29.596439doi: bioRxiv preprint 36 S TED image analysis Pre-processing of diffraction-limited and STED images was performed using Fiji using a custom-made script. Nuclear masks were created by segmenting the CTCF-AF594 diffract ion-limited image after gaussian-blurring and used to crop out nuclei in all image channels. In addition, STED images were

subtracted using the rolling ball algorithm set to a radius of 50 pixels. Co-localization analysis, as well as spot segmentation was performed using custom python scripts. For co-localization analysis, Pearson correlation coefficient of CTCF and STAG1/2 was computed based on cropped nuclei in the respective STED channel. Spot segmentation was performed by first coarsely segmenting spots inside the nuclear mask based on a common threshold (method: Otsu) after applying a mild gaussian blur (sigma = 1). Image noise resulting in excess tiny spots was filtered out through binary mask erosion and filtering, followed by binary dilation of correctly detected spots. Coarsely segmented spots often represent clusters of spot signals and were further segmented using a combination of local peak finding & watershed. The resulting masks for individual spots were used to extract average pixel intensities in the STED and confocal images. Assuming a z-depth of about 500 nm, the number of detected spots per μm3 was compared to protein number estimates derived by FCS-calibrated imaging of early G1 or G1 cells to estimate the overall labelling efficiency. STED image simulation STED images were simulated by generating a desired number of randomly localized spots (single pixels) in an image representing 200 μm2 (or 100 μm3 assuming a z-depth of 500 nm), given the pixel size of 18.88 nm in the images acquired as described above. The randomly distributed spots were gaussian blurred (sigma = 2.6) and their pixel intensity was enhanced 6-fold to be distinguishable above a random background. Simulated images were analyzed for co-localization or segmented and analyzed to read out their average spot intensities as described above. .CC-BY 4.0 International licenseavailable under a was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. 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