Chromatin organization and behavior in HRAS-transformed mouse fibroblasts

doi:10.21203/rs.3.rs-3631068/v1

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Chromatin organization and behavior in HRAS-transformed mouse fibroblasts

https://doi.org/10.21203/rs.3.rs-3631068/v1

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published 24 Feb, 2024

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In higher eukaryotic cells, a string of nucleosomes, where long genomic DNA is wrapped around core histones, are rather irregularly folded into a number of condensed chromatin domains, which have been revealed by super-resolution imaging and Hi-C technologies. Inside these domains, nucleosomes fluctuate and locally behave like a liquid. The behavior of chromatin may be highly related to DNA transaction activities such as transcription and repair, which are often upregulated in cancer cells. To investigate chromatin behavior in cancer cells and compare those of cancer and non-cancer cells, we focused on oncogenic-HRAS (Gly12Val)-transformed mouse fibroblasts CIRAS-3 cells and their parental 10T1/2 cells. CIRAS-3 cells are tumorigenic and highly metastatic. First, we found that HRAS-induced transformation altered not only chromosome structure, but also nuclear morphology in the cell. Using single-nucleosome imaging/tracking in live cells, we demonstrated that nucleosomes are locally more constrained in CIRAS-3 cells than in 10T1/2 cells. Consistently, heterochromatin marked with H3K9me3 and H3K27me3 was upregulated in CIRAS-3 cells. Finally, Hi-C analysis showed enriched interactions of the B-B compartment in CIRAS-3 cells, which likely represents transcriptionally inactive chromatin. Increased heterochromatin may play an important role in cell migration, as they have been reported to increase during metastasis. Our study also suggests that single-nucleosome imaging provides new insights into how local chromatin is structured in living cells.

Chromatin

nucleosome

cancer

RAS

single-molecule imaging

Hi-C

In higher eukaryotic cells, a long strand of genomic DNA is wrapped around core histone, forms nucleosomes (Koyama and Kurumizaka 2018; Luger et al. 1997; Olins and Olins 2003). Nucleosomes are irregularly folded into various-sized clusters, or chromatin domains (∼200 nm), within the cell (Maeshima et al. 2021; Misteli 2020) and fluctuate like a liquid (Ashwin et al. 2019; Maeshima et al. 2023; Nozaki et al. 2023). Notably, recent studies suggest that not only heterochromatin, which is generally silenced with recessive histone marks, but also euchromatic regions with active histone marks, form condensed chromatin organization (Maeshima et al. 2023; Miron et al. 2020; Nozaki et al. 2023). The size of euchromatin domains seems smaller than those of heterochromatin (Miron et al. 2020). Hi-C methods, which can generate a fine contact probability map of genomic DNA (Dekker and Heard 2015) revealed that the chromatin domains are often clustered as two distinct, megabase scale compartments (A and B), which likely represent transcriptionally active chromatin and inactive chromatin, respectively (Lieberman-Aiden et al. 2009).

The behavior of chromatin can be highly related to DNA transaction activities such as transcription (Germier et al. 2017; Nagashima et al. 2019) and DNA repair (Iida et al. 2022; Seeber et al. 2018): Nucleosome motion in euchromatin appears higher than that in heterochromatin (Nozaki et al. 2017; Shinkai et al. 2017) (Lerner et al. 2020; Nozaki et al. 2023). Since cancer cells often have upregulations of such activities (Mullen and Singh 2023), it would be intriguing to investigate chromatin behavior in cancer cells and compare those of cancer and non-cancer cells.

For this purpose, we used oncogenic-RAS (Gly12Val)-transformed mouse fibroblasts, CIRAS-3 cells, which are tumorigenic and highly metastatic (Fig. S1A) (Egan et al. 1987). RAS is a small GTPase protein and an important factor involved in many signaling pathways, such as cell cycle progression, cell migration, apoptosis, and senescence (Punekar et al. 2022; Pylayeva-Gupta et al. 2011). RAS is also one of the famous human oncoproteins (Punekar et al. 2022; Pylayeva-Gupta et al. 2011). Oncogenic RAS mutations, which are frequently found in about 25% of all cancer diseases, activate the MAPK pathway (RAS/RAF/MEK/ERK pathway), upregulate transcription, and contribute to tumorigenesis (Punekar et al. 2022; Pylayeva-Gupta et al. 2011).

We investigated chromatin organization and dynamics in CIRAS-3 cells and their parental mouse pre-transformed 10T1/2 cells(Fig. S1A) (Reznikoff et al. 1973). The 10T1/2 cell line derived from mouse embryo cells and is sensitive to post-confluence inhibition of division (Reznikoff et al. 1973). We observed oncogenic-RAS-induced transformation altered not only chromosome structure but also nuclear morphology. Using single-nucleosome imaging/tracking (Ide et al. 2022; Lakadamyali 2022), we found nucleosomes were more constrained in CIRAS-3 cells than in 10T1/2 cells. Consistently, CIRAS-3 cells had increased heterochromatin marks. Hi-C analysis showed enriched B-B compartment interactions in CIRAS-3 cells. These results suggest altered physical properties of chromatin in transformed cells, which are consistent with previous reports of increased H3K9me3 and K3K27me3, promoting cell motility and cancer metastasis.

Characterization of the mouse fibroblast CIRAS-3 and 10T1/2 cell lines.

We first characterized the mouse fibroblast CIRAS-3 and 10T1/2 cell lines (Fig. S1A). CIRAS-3 cells had a prolonged G1 phase (Fig. S1B and C) and proliferated more slowly than 10T1/2 cells (Fig. S1D). Consistent with the previous report (Dunn et al. 2009), chromosome spread analysis showed that both cell lines were aneuploidy, but CIRAS-3 cells had more complex karyotypes, including the Robertsonian-fusion (centromere to centromere fusion) (Fig. 1A-C) (Robertson 1916). CIRAS-3 cells grew in agarose, indicating their anchorage-independent growth, while 10T1/2 did not (Fig. S2). These results confirmed that CIRAS-3 cells have a metastatic potential, as shown in (Dunn et al. 2009; Egan et al. 1987). Interestingly, when we measured the nuclear volumes of each cell line by confocal laser scanning microscopy and image analysis (Fig. 1D and E), we found that CIRAS-3 had smaller nuclei than 10T1/2 (Fig. 1F).

Single-nucleosome imaging and tracking to study local chromatin motion in living transformed mouse cells.

We performed single-nucleosome imaging in living CIRAS-3 (Egan et al. 1987) and 10T1/2 cells to investigate how HRAS-transformation affected local chromatin behavior. As our previous studies with human RPE-1 cells (Nagashima et al. 2019), HeLa and HCT116 cells (Iida et al. 2022), we established mouse CIRAS-3 and 10T1/2 cells that stably express a core histone H2B labeled with HaloTag (H2B-Halo)(Fig. 1G), which can be visualized with a HaloTag ligand Tetramethylrhodamine (TMR) dye (Fig. 1H). H2B-Halo had genome-wide nucleosome incorporation, including euchromatic and heterochromatic regions (Fig. 1H), probably because the turnover of histone H2B occurred within a few hours (Kimura and Cook 2001).

We conducted single-nucleosome imaging and tracking (Ide et al. 2022; Iida et al. 2022; Lakadamyali 2022; Lerner et al. 2020; Nagashima et al. 2019) on CIRAS-3 and 10T1/2 cells using oblique illumination microscopy to illuminate a thin area within a single nucleus with reduced background noise (Fig. 2A) (Tokunaga et al. 2008). Very low concentrations of TMR were used to obtain sparse labeling of nucleosomes for single-nucleosome imaging (Fig. 2B). TMR-labeled nucleosomes were recorded at 50 ms/frame in asynchronous cells (∼100 frames, 5 s total) (Movies S1 and S2) and observed as clear dots (Fig. 2C). These dots showed a single-step photobleaching profile (Fig. 2D), which suggested that each dot represents a single H2B-Halo-TMR molecule in a single nucleosome. Notably, we only tracked the nucleosome-incorporated H2B-Halo-TMR at this frame rate since free H2B-Halo, which is not constrained in nucleosomes, diffused too fast to detect as dots and track. The individual dots were fitted with a 2D Gaussian function to estimate the precise position of the nucleosome (Movies S1 and S2) (Betzig et al. 2006; Rust et al. 2006). They were tracked using u-track software (Jaqaman et al. 2008) and the nucleosome trajectory data was obtained (e.g., Fig. 2E).

No significant differences in MSD plots of nucleosome motions between 10T1/2 and CIRAS-3 cells.

From the trajectory data, we calculated displacement distributions (Figs. 3A and S3) and mean square displacement (MSD) of individual nucleosomes in 10T1/2 and CIRAS-3 cells (Figs. 3B and S4A). MSD generally shows how molecules spatially move in a certain time period. MSD plots of nucleosomes in both live cells seemed sub-diffusive (solid lines, Fig. 3B) while those obtained from their formaldehyde (FA)-fixed cells were severely suppressed (broken lines).

As shown in Figs. 3B and S4A, single-nucleosome imaging revealed no significant differences in MSD plots of nucleosome motions between CIRAS-3 and 10T1/2 cells. We further explored local chromatin motions in CIRAS-3 and 10T1/2 cells. Log-log plots of their MSD data showed that their MSD exponents for the first 0.5 sec were also similar (i.e., 0.38 for CIRAS-3 and 0.39 for 10T1/2) (Fig. S4B).

Nucleosomes in CIRAS-3 are locally more constrained than those in 10T1/2 cells.

To further investigate nucleosome behaviors in CIRAS-3 and 10T1/2 cells, we performed the angle-distribution analysis (Iida et al. 2022; Izeddin et al. 2014) of nucleosome trajectories from CIRAS-3 and 10T1/2 cells (Fig. 4A and B). The nucleosome angle distributions showed that the moving angles of individual nucleosomes were somehow biased towards 180° (Fig. 4C) compared with control particles displaying Brownian motion (Fig. 4D) (Iida et al. 2022), suggesting nucleosomes are often pulled back to their original positions (lower, Fig. 4A). To quantify this tendency, we obtained the asymmetry coefficient (AC), which is calculated as the ratio between the frequency of forward angles (between − 30° and 30°) and backward angles (150°–210°) of nucleosomes (right, Fig. 4B). The ACs for nucleosome motions in CIRAS-3 and 10T1/2 cells were − 1.523 and − 1.384, respectively (Fig. 4C). These results suggested that nucleosomes in CIRAS-3 cells have a greater pull-back force to their original position. That is, nucleosomes in CIRAS-3 cells were more constrained than those in 10T1/2 cells. But why?

Recessive histone marks are enriched in CIRAS-3 cells.

We wondered whether CIRAS-3 cells have more heterochromatin and examined the amounts of repressive histone marks such as H3K9me3 and H3K27me3 (Allshire and Madhani 2018; Grewal and Jia 2007; Janssen et al. 2018; Padeken et al. 2022) in both cell lines by Western blot (Fig. 5A). We found that the signals of H3K9me3 and H3K27me3 were substantially increased in CIRAS-3 cells (Fig. 5B), suggesting that CIRAS-3 cells have more heterochromatin.

Hi-C analysis reveals an increase in B-B compartment interactions in CIRAS-3 cells.

To further investigate the heterochromatin organization in CIRAS-3 cells, we performed in situ Hi-C analysis (Rao et al. 2014) of CIRAS-3 and 10T1/2 cells and focused on their three-dimensional (3D) chromatin interactions. We used the GRCm38/mm10 mouse genome (http://www.ncbi.nlm.nih.gov/assembly/GCF_000001635.20) for analysis because reference genomes of both cell lines were not available. Thus, the obtained Hi-C results might not completely reflect the chromatin organization in the CIRAS-3 and 10T1/2 cells, but were still useful to test our notion.

We first characterized the A/B compartments (Lieberman-Aiden et al. 2009) in CIRAS-3 and 10T1/2 cells in 160-kb bins and the interactions between compartments (Fig. 6A and B). To quantitatively assess the strength of interactions between compartments, we performed a saddle plot analysis (Fig. 6C). We sorted the compartment scores of each row of contact maps from lowest to highest (B to A compartments) and divided them into quantiles. We then calculated interactions between these quantiles to create saddle plots (Fig. 6C). The interaction strengths are depicted in the lower left for B-B, the upper right for A-A, and the off-diagonal corners for B-A and A-B interactions. We found that B-B interactions increased in CIRAS-3 cells (Fig. 6C), consistent with our previous data on the upregulation of heterochromatin marks in CIRAS-3 cells (Fig. 5B).

Using single-nucleosome imaging, we found that nucleosomes in CIRAS-3 cells were more constrained than those in 10T1/2 cells (Fig. 4C), suggesting that the local chromatin environment changed upon oncogenic-HRAS-induced transformation. Consistently, heterochromatin marks (H3K9me3 and H3K27me3) (Fig. 5B) and interactions between B-B compartments (Fig. 6C) increased in CIRAS-3 cells. Our results suggest that oncogenic-HRAS-induced transformation altered not only chromosome structure (Fig. 1A and B) (Dunn et al. 2009), but also nuclear morphology (Fig. 1D and E) and local chromatin organization. Genomic and chromosomal instability are necessary for tumorigenesis (Jinesh et al. 2018; Negrini et al. 2010). In CIRAS-3 cells, telomere alterations might be induced when oncogenic-HRAS is overexpressed, leading to chromosomal instability and subsequent chromosome fusions (e.g., Robertsonian).

Interestingly, although MSD plots of nucleosomes in single-nucleosome imaging are similar between the CIRAS-3 and 10T1/2 cells, the angle distributions of their nucleosome trajectories showed that nucleosomes in CIRAS-3 cells were more constrained than those in 10T1/2 cells (Fig. 4C). Our results suggest that single-nucleosome imaging provides new insights into how local chromatin is structured in living cells. With new data analyses (Ashwin et al. 2019; Lerner et al. 2020; Nozaki et al. 2023), we will be able to extract more and more information on chromatin organization from the nucleosome motion data.

We demonstrated that local chromatin motion in CIRAS-3 cells was more constrained with enriched heterochromatin (Fig. 7). This might contribute to the slower growth of CIRAS-3 cells (Fig. S1D). On the other hand, genome chromatin in cancer cells must be protected from physical stress during metastasis. Chromatin works as a spring that resists nuclear deformation (Bustin and Misteli 2016; Furusawa et al. 2015; Maeshima et al. 2018; Shimamoto et al. 2017; Stephens et al. 2017). The smaller nucleus with more constrained chromatin in CIRAS-3 cells could benefit the metastasis process with nuclear deformation (Fig. 7). There is a significant body of literature supporting the hypothesis that heterochromatin formation promotes cell motility (reviewed in (Gerlitz 2020)). Heterochromatin marks increase when cells are forced to migrate through small pores in 3D (Seelbinder et al. 2021) and may protect against DNA damage that accompanies migration through small spaces (Shah et al. 2021), such as during metastasis (Maizels et al. 2017). Increased H3K9me3 and H3K27me3 are associated with increased motility and metastasis (Maizels et al. 2017; Yokoyama et al. 2013). Inhibitors of DNA methylation and histone deacetylase, both of which promote the dissolution of heterochromatin, reduce cell migration while overexpression of SUV39H1 (i.e., increasing H3K9me3) increases migration (Yokoyama et al. 2013). Indeed, nuclear stiffness correlates with migration efficiency (Seelbinder et al. 2021). Furthermore, a more rigid nucleus containing more heterochromatin acts as a piston when the cell is squeezed through a small space, resulting in forward propulsion (Gerlitz 2020; Petrie et al. 2014). Thus, increased heterochromatinization and enhanced nuclear stiffness may serve to promote cell metastasis.

On the other hand, more than 30 years have passed since CIRAS-3 cells were established via the introduction of oncogenic HRAS (Egan et al. 1987). The chromosomes were highly rearranged during its long maintenance process, generating complicated karyotypes (Fig. 1B) (Dunn et al. 2009). This may have caused difficulty in directly comparing chromatin organization/behavior between the two cell lines. Investigation using an inducible system of oncogenes to transform normal human cells would be an essential next step to understand how oncogenic cell transformation governs chromatin organization/behavior.

Cell lines and establishment of stable cell lines

Mouse 10T1/2 (Reznikoff et al. 1973) and CIRAS-3 cells (Egan et al. 1987) (a gift from Dr. James R. Davie at the University of Manitoba) were cultured at 37°C with 5% CO₂ in Dulbecco’s Modified Eagle’s medium (DMEM) (D5796-500ML, Sigma-Aldrich) supplemented with 10% fetal bovine serum (FBS) (FB-1061/500, Biosera).

To stably express H2B-Halo in 10T1/2 and CIRAS-3 cells, the transposon system was used. The constructed plasmid pPB-EF1a-IB-H2B-HaloTag (Nagashima et al. 2019) was cotransfected with pCMV-hyPBase (provided from the Sanger Institute with a materials transfer agreement) to 10T1/2 and CIRAS-3 cells with the Effectene Transfection Reagent kit (301425; QIAGEN). Transfected 10T1/2 cells were selected with blasticidin S (10 µg/ml) (029-18701, Wako). Transfected CIRAS-3 cells were selected with blasticidin S (10 µg/ml) and G418 (400 µg/ml) (16512-81, Nakalai).

Western blot

To check the expression level of H2B-HaloTag, transfected 10T1/2 and CIRAS-3 cells were lysed in Laemmli sample buffer (Laemmli 1970) supplemented with 10% 2-mercaptoethanol (133–1457; Wako) and incubated at 95°C for 10 min to denature their proteins. The cell lysates, equivalent 2x10⁵ 10T1/2 or CIRAS-3 cells per well, were subjected to SDS–polyacrylamide gel electrophoresis (PAGE) (14%) and subsequent Western blotting. For Western blot, the fractionated proteins in the gel were transferred to a polyvinylidene difluoride membrane (PVDF) (IPVH00010, Millipore) by a semidry blotter (BE-320, BIO CRAFT). After blocking with 10% or 3% skim milk (190-12865, FujiFilm), the membrane-bound proteins were probed by the anti-H2B rabbit (1:10,000 dilution; ab1790, Abcam), anti-HaloTag mouse (1:1000; G9211, Promega), anti-H3K9me3 mouse (1:1,000 dilution; provided by Prof. Hiroshi Kimura), or anti-H3K27me3 mouse (1:1000; provided by Prof. Hiroshi Kimura) antibody, followed by the appropriate secondary antibody, anti-rabbit (1:10,000 dilution; 170–6515, Bio-Rad) or anti-mouse (1:5000 dilution; 170–6516, Bio-Rad) horseradish peroxidase (HRP)–conjugated goat antibody. Chemiluminescence reactions were used (WBKLS0100, Millipore) and detected by EZ-Capture MG (AE-9300H-CSP, ATTO).

Imaging and quantification of HaloTag-labeled cells

To examine H2B-HaloTag localization in 10T1/2 and CIRAS-3 cells, cells grown on poly-l-lysine-coated (P1524-500MG, Sigma-Aldrich) coverslips (C018001, Matsunami) were treated with 5 nM HaloTag TMR ligand (8251, Promega) overnight at 37°C in 5% CO₂. The cells were fixed with 1.85% FA (064–00406, Wako) on coverslips at room temperature for 15 min followed by quenching with 50 mM glycine (077–00735, Wako) for 5 min, permeabilized with 0.5% Triton X-100 (T-9284, Sigma-Aldrich) for 5 min, washed with 1x HMK (20 mM HEPES pH 7.5, 1 mM MgCl₂, 100 mM KCl), and stained with 4′,6-diamidino-2- phenylindole (DAPI) (0.5 µg/ml) (10236276001, Roche) for 5 min, before being embedded in PPDI (20 mM HEPES (pH 7.4), 1 mM MgCl₂, 100 mM KCl, 78% glycerol, and paraphenylene diamine (1 mg/ml) (695106-1G, Sigma-Aldrich)). Optical sectioning images were recorded with a 0.2-µm step size using a DeltaVision Personal Microscope (Applied Precision) with an Olympus PlanApo N 60x objective (NA, 1.42) and a scientific complementary metal-oxide semiconductor (sCMOS) camera. The four-color standard filter set was also equipped. DeltaVision acquisition software, Softworx, was used to project deconvolved Z-stacks to cover the whole nucleus.

Chromosome spreads

To observe and count the chromosome numbers of 10T1/2 and CIRAS-3 cells, mitotic chromosome spreads were made. Cells were treated with colcemid (0.5 µg/ml) (045-16963, Wako) for 1 h at 37°C. The following steps were performed at room temperature except for the final fixation step. After washing with phosphate-buffered saline (PBS), cells were trypsinized and resuspended in hypotonic buffer (75 mM KCl) for 8 min. Cells were then gently fixed by repeating the following treatment three times: fixation buffer (methanol: acetic acid, 3:1) for 5 min, centrifuged at 850×g for 4 min, and then resuspended in new fixation buffer. Lastly, the cells were completely fixed by treatment with 200 µl of fixation buffer at − 30°C for at least 30 min. For imaging, 10 µl of the fixed cell suspension was dropped onto coverslips after mixing and completely dried by incubating at 60°C for 30 min. Dried cells were stained with DAPI (0.5 µg/ml) for 5 min, followed by PPDI mounting. Images of mitotic chromosome spreads were obtained using DeltaVision Personal Microscopy with an Olympus PlanApoN 60x objective (NA 1.42) and an sCMOS camera. The number of chromosomes in each cell was counted manually.

Nuclear volume analysis

Nuclear volumes were measured (Iida et al. 2022) as follows. The cells were grown on poly-l-lysine-coated (P1524-500MG, Sigma-Aldrich) coverslips (C018001, Matsunami). Subsequent processes were performed at room temperature. The cells were washed with 1x PBS, fixed with 1.85% FA for 15 min followed by quenching with 50 mM glycine (077–00735, Wako) for 5 min, permeabilized with 0.5% Triton X-100 for 5 min, washed with 1x HMK, stained with DAPI (0.5 µg/ml) for 5 min, and washed with 1x HMK for 3 min three times before being embedded in PPDI. Z-stack images (every 0.4 µm, 20 to 40 sections in total) of the cells were obtained using a FLUOVIEW FV1000 confocal laser scanning microscope (OLYMPUS) equipped with an Olympus UPLANSAPO 60x W objective (NA 1.20) at room temperature. Obtained Z-stack images were loaded to Imaris (Bitplane AG, Zurich, Switzerland) and converted to Imaris 3D image files (.ims). To calculate the nuclear volume, the tool “Surface” was used. A threshold value was automatically determined to include all DAPI signals. Note that only well-isolated nuclei were recorded and analyzed.

Cell growth and cell cycle

Cells were seeded in 6-well plates (1.0×10⁴ cells/ml). The number of proliferating cells was examined microscopically at several time points. The growth rates of 10T1/2 and CIRAS-3 cells were calculated from exponentially fitted growth curves.

Flow cytometry was performed to check the cell cycle profiles of 10T1/2 and CIRAS-3 cells. Collected cells were fixed in 70% ethanol at − 30°C for over 30 min. After fixation, cells were centrifuged at 603×g for 1 min, and the cell pellets were washed with PBS containing 1% BSA (bovine serum albumin; A9647-100G, Sigma-Aldrich). After centrifugation, cell pellets were resuspended in 600 µl of PBS containing 1% BSA, ribonuclease A (50 µg/ml) (10109169001, Sigma-Aldrich), and propidium iodide (40 µg/ml) (P4170-10MG, Sigma-Aldrich) and then further incubated for 30 min at 37°C with light protection. Lastly, cells were analyzed by BD Accuri C6 Plus (BD FACS, San Jose, CA). At least 20,000 cells were used for each analysis, and the obtained results were displayed as histograms. The percentage of cell cycle stage distributions in the G0/G1, S, and G2/M phases was analyzed by the ModFit LT software (Verity Software House, Topsham, ME).

Single-nucleosome imaging

CIRAS-3 and 10T1/2 cells stably expressing H2B-Halo were cultured on 35-mm dishes (153066, Nunc). H2B-Halo incorporated in nucleosomes was fluorescently labeled with 110 pM HaloTag TMR ligand (for 10T1/2 cells) and 100 pM HaloTag TMR ligand (for CIRAS-3 cells) for 20 min at 37°C in 5% CO₂ and then washed once with 1x PBS. Cells were trypsinized and resuspended in phenol red-free DMEM (21063-029, Thermo Fisher Scientific) supplemented with 10% FBS, and then seeded on fibronectin (354008, Corning) coated glass-base dishes (3970-035, IWAKI). A live-cell chamber (INU-TIZ-F1, Tokai Hit) and digital gas mixer (GM-8000, Tokai Hit) were used to maintain cell culture conditions (37°C, 5% CO₂, and humidity) during microscopy. Single nucleosomes were observed using an inverted Nikon Eclipse Ti microscope with a 100-mW Sapphire 561-nm laser (Coherent) and sCMOS ORCA-Flash 4.0 camera or ORCA-Fusion BT (Hamamatsu Photonics). Live cells labeled with H2B-Halo-TMR were excited by the 561-nm laser through an objective lens (100x PlanApo TIRF, NA 1.49; Nikon) and detected at 575 to 710 nm. An oblique illumination system with the TIRF unit (Nikon) was used to excite fluorescent nucleosome molecules within a limited thin area in the cell nucleus and reduce background noise (Fig. 2A). Sequential image frames were acquired using MetaMorph software (Molecular Devices) or NIS-Elements AR (Nikon) at a frame rate of 50 ms under continuous illumination.

Single-nucleosome tracking analysis

Image processing, single-nucleosome tracking, and single-nucleosome movement analysis were performed as previously described (Iida et al. 2022; Nozaki et al. 2023). Briefly, sequential images were converted to a 16-bit grayscale, and the background noise signals were subtracted with the rolling ball background subtraction (radius, 50 pixels) of Fiji/ImageJ (Schindelin et al. 2012). The nuclear regions in the images were manually extracted. Following this step, the position of each fluorescent dot in each image was determined and its trajectory was tracked with u-track ( (Jaqaman et al. 2008), MATLAB package). We previously calculated the SD of the 2D movement of immobilized nucleosomes per 50 ms in FA-fixed RPE-1 cells (n = 10 nucleosomes) to determine the accuracy of the position of H2B-Halo nucleosomes. The localization accuracy was 15.55 nm (Nagashima et al. 2019). For single-nucleosome movement analysis, the displacement distribution and the mean square displacement (MSD) of the fluorescent dots were calculated based on their trajectory using lab-made Python programs. The originally calculated MSD was in 2D. To obtain the 3D value, the 2D value was multiplied by 1.5 (4 to 6 Dt). Graphs and statistical analyses of the obtained single-nucleosome MSD between various conditions were performed using R. For chemical fixation, cells grown on poly-l-lysine-coated glass-based dishes were incubated with 2% FA in 1x HBSS at 37°C for 15 min and washed with 1x PBS at 4°C for at least 3 h.

Angle distribution analysis

Angle distribution analysis was performed as previously described (Iida et al. 2022; Izeddin et al. 2014). For the tracked consecutive points [(x0, y0), (x1, y1), ⋯, (xn, yn), ⋯] of a single nucleosome on the xy plane, we converted the data into a set of displacement vectors, Δrn = (xn + 1 – xn, yn + 1 – yn)t. Then, we calculated the angle between the two vectors Δrn and Δrn + 1. We carried out this procedure for all the points of each trajectory in our experiments. Lastly, we plotted the normalized polar histogram with our Python program. The angle distribution was normalized by 2π, and the values corresponded to the probability density.

Anchorage-independent cell growth analysis

Cell culture in a soft agar layer was performed as previously described (Xu et al. 2021; Zhao et al. 2015) with a 0.5% agar (010-08725, FujiFilm) base layer containing 10% FBS in DMEM with penicillin/streptomycin and a 0.35% agar growth layer containing 10% FBS in DMEM. Colonies that had three or more cells were scored at 21 days after the cell seeding.

In situ Hi-C library preparation

In situ Hi-C libraries in this study were prepared following the protocols described in (Kawaguchi and Tanaka 2023; Schloissnig et al. 2021), with some modifications. One million of 10T1/2 or CIRAS-3 cells were collected into 50 ml conical tubes with 10% FCS/DMEM and then fixed with 1% formaldehyde for 10 min at room temperature. The fixation was halted by adding ice-cold glycine (125 mM final concentration) and tubes were kept on ice for 15 min with occasional inversion for mixing. Cells were collected by centrifugation (350xg for 10 min at 4 ℃) and washed twice with ice-cold PBS (350xg for 10 min at 4 ℃). Cell pellets were then resuspended in ice-cold lysis buffer (10 mM Tris-HCl pH 7.5, 10 mM NaCl, 0.2% NP-40, protease inhibitor-EDTA free (P8340, SIGMA)) and incubated on ice for 30 min with occasional agitation. Afterward, the nuclei were collected by centrifugation (300xg for 10 min at 4 ℃) and washed with 500 µl of 1.25x NEBuffer3 at the same speed. The nuclei were resuspended in 1 ml of 1.25x NEBuffer3 and transferred to DNA low-binding tubes (500 µl each). SDS (0.4% final concentration) was added and the tubes were incubated with agitation (950 rpm) for 2 h at 37 ℃. Triton X-100 (3.3% final concentration) was added to quench SDS, and the nuclei were incubated with agitation (950 rpm) for 2 h at 37 ℃. Nuclei were pelleted with centrifugation (100xg for 1 min at room temperature) and the pellets were resuspended in 450 µl of 1x DpnII digestion buffer (R0543M, NEB). DpnII digestion (NEB; 100 U per 0.5 million cells) was performed with agitation (950 rpm) for 2 h at 37 ℃. The enzyme was then inactivated by heating the tubes to 65 ℃ for 20 min. Overhang filling was carried out using biotin-14-dATP, dCTP, dGTP, and dTTP, along with Klenow (NEB; 50 U per 0.5 million cells) for 1 h at 37 ℃ with repeated agitation (repeated cycles of 700 rpm for 10 sec and rest for 30 sec for 1 h in a thermal cycler). Ligation was performed overnight at 18 ℃ with 2000 U of T4 DNA ligase. De-crosslinking with Protease K in SDS buffer was done overnight at 65°C, followed by RNase A treatment for 2 h at 37 ℃. Subsequently, DNA extraction was performed with phenol/chloroform and EtOH precipitation steps, and the ligated gDNA products were eluted in 20 µl of TE buffer. The yield was determined using a Qubit3 device, and 5 µg of biotinylated DNA was used for the library preparation. To remove excess biotin, non-ligated fragment ends were then treated with T4 DNA polymerase for 30 min at 37°C, and the reaction was stopped by adding EDTA (10 mM final concentration). DNA was sonicated using Covaris S220 system to generate DNA fragments with the size peak around 400 bp (Covaris settings were duty factor: 10%; peak incident power: 5; cycles per burst: 200; time: 65 sec). After end repair with T4 DNA polymerase, Large (Klenow) Fragment, and T4 DNA polynucleotide kinase (all from NEB) in the presence of dNTPs in T4 DNA ligation buffer (for 30 min at room temperature), the DNA was purified with QIAGEN mini purification kit. A double-sized selection using AMPure XP beads (Beckman) was performed (see (Kawaguchi and Tanaka 2023; Schloissnig et al. 2021)). Firstly, the ratio of the AMPure XP bead volume to DNA sample volume was adjusted to 0.6:1, and then the second step was performed with the final ratio of 0.9:1. Following two washes with 80% EtOH, the DNA was eluted in Elution buffer (QIAGEN). From DNA pools, Hi-C ligation products were isolated using pre-washed MyOne Streptavidin C1 Dynabeads (# 65001, Life Technologies) on a magnet stand in binding buffer (5 mM Tris pH 8.0, 0.5 mM EDTA, 1 M NaCl) for 30 min at room temperature. dA-tailing was carried out on the beads. dATP was added with Klenow exo- (for 1 h at 37°C), then the enzyme was heat-inactivated (for 20 min at 65°C). After two washes in binding buffer and one wash in T4 DNA ligation buffer, Illumina adapters were ligated onto Hi-C ligation products bound to streptavidin beads in T4 DNA ligase while slowly rotating (for 2 h at room temperature). After washing twice with wash buffer (5 mM Tris pH 8.0, 0.5 mM EDTA, 1 M NaCl, 0.05% Tween-20) and once with binding buffer, the Hi-C product bound beads were resuspended in a final volume of 20 µl 1x NEBuffer2. Captured Hi-C DNA was amplified with PCR amplification cycles using NEBNext® High-Fidelity 2X PCR Master Mix (# M0541S, NEB) for 7 cycles. The final Hi-C libraries were purified with 1x volume of AMPure XP beads. The concentration of the Hi-C library was determined using Tapestation (Agilent), and the Hi-C libraries were sequenced with the PE150 setting on NovaSeq 6000. We repeated the above experiment for the reproducibility and Hi-C sequence data are available in DDBJ database as DRA017367.

Hi-C data analysis

Hi-C data were analyzed using the nf-core (Ewels et al. 2020) Hi-C pipeline (nf-core/hic: 2.0.0) (Servant et al. 2023), executed within a Docker configuration profile. The Mus musculus genome mm10 (http://www.ncbi.nlm.nih.gov/assembly/GCF_000001635.20), retrieved from Illumina’s iGenomes, was used for the reference genome. HiC-Pro (Servant et al. 2015) was employed, which performs sequence reads mapping with Bowtie2 (Langmead and Salzberg 2012) and identifies valid chromosomal interactions. Subsequently, Cooler (Abdennur and Mirny 2020) was used for constructing and normalizing contact maps through its balancing algorithm, and compartment callings were conducted with cooltools (Venev et al. 2022). To detect chromosomal translocations and breakpoints from the Hi-C data, HiNT (Wang et al. 2020) software (version 2.1.9) was run in the HiNT-TL mode with the default parameter settings. The visualization of contact maps and A/B compartment tracks in Fig. 6 were generated using CoolBox (v0.3.9) (Xu et al. 2021). To visualize the strength of interactions between compartments, the saddle plot analysis was performed using the R-package GENOVA (v1.0) (van der Weide et al. 2021).

Acknowledgments

We are grateful to Dr. K.M. Marshall for critical reading and editing of this manuscript. We thank Dr. S. Kuraku for supporting the Hi-C experiment, Dr. J.R. Davie for providing CIRAS-3 and 10T1/2 cells, Dr. H. Kimura for providing his antibodies, Dr. T. Yugawa for valuable comments, Dr. K. Hibino, Ms. S. Iida, Mr. M. A. Shimazoe, and Maeshima laboratory members for their helpful discussions and support. A.O. thanks Progress Committee members Dr. K. Saito, Dr. M. Kanemaki, Dr. J. Kitano, Dr. A. Kimura for their support. This work was supported by JSPS grants JP21H02453 (K.Maeshima), JP20H05936 (K.Maeshima), JP23K17398 (K.Maeshima and S.I.), JP23K05798 (A.K.), JP22H05606 (S.I.), JP21H02535 (S.I.), JP22H04925 (PAGS) (K.Maeshima), The Naito Foundation (A.K.), Takeda Science Foundation（K.Maeshima）. A.O. is a SOKENDAI Special Researcher (JST SPRING JPMJSP2104). K.Minami was a SOKENDAI Special Researcher and is currently a JSPS Fellow (23KJ0998). M.J.H. is supported by a Canada Research Chair.

Author contributions

A.O., K.Minami, and K.Maeshima designed the project. A.O. and S.T. created the cell lines with the help of S.I. A.O. and K.Minami performed single-nucleosome imaging, tracking, and analysis. A.O. performed most of the cell biology experiments with help of S.I. A.K. performed the Hi-C experiment. K.H., K.K., and A.O. analyzed Hi-C data. S.T. contributed to the scientific illustrations. M.H. provided essential resources. K.Maeshima, K.Minami, and A.O. wrote the manuscript with input from all other authors.

Competing interests

The authors declare that they have no competing interests, financial, or otherwise.

Data and materials availability

Raw sequencing datasets for Hi-C analysis generated in this study are available in the DDBJ Sequenced Read Archive under the accession numbers DRA017367.

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No competing interests reported.

FigureS1.tif
Fig. S1. Cell growth and cell-cycle profiles of 10T1/2 and CIRAS-3 cells. (A) Representative brightfield images of 10T1/2 (left) and CIRAS-3 (right) cells. (B) Cellular DNA contents in 10T1/2 (left) and CIRAS-3 (right) cells. Each histogram represents more than 20,000 cells. Black triangles on the x-axes show the genomic DNA amount corresponding to G1/G0 and G2/M phases. The proportion of G1/G0 and G2/M cells were quantified by the Gaussian fitting of corresponding peaks (shown in red). (C) The proportion of each stage was quantified from data in (B). 10T1/2 cells (left): 33.0% in G1/G0; 54.1% in S; 12.9% in G2/M. CIRAS-3 cells (right): 45.5% in G1/G0; 43.8% in S; 10.7% in G2/M. (D) Growth curves of 10T1/2 (black) and CIRAS-3 (red) cells cultured for indicated times. Mean ± SD among three independent replicates is shown. Doubling times calculated from the exponentially fitted curves were 16.4 h for 10T1/2 cells and 22.1 h for CIRAS-3 cells.
FigureS2.tif
Fig. S2. Anchorage-independent cell growth of CIRAS-3 cells. (A) Representative 10T1/2 (top) and CIRAS-3 (bottom) cells cultured on agar layers. The images of cells were captured on the day of seeding (day 0) and 21 days after culture. At day 0, cells are indicated by arrows. (B) Number of colonies that contain three or more cells at 21 days after culture. The horizontal bar shows the mean of triplicate experiments. **, p = 0.0018 by Welch’s two-sample t-test.
FigureS3.tif
Fig. S3. Displacement distributions of the tracked single nucleosomes. (A, B) Displacement (movement) distributions of nucleosomes in live cells for Δt = 100, 200, 300, 400, and 500 ms. (A) 10T1/2 cells (n = 20 cells). (B) CIRAS-3 cells (n = 20 cells). Mean ± SD of displacement is indicated in each panel.
FigureS4.tif
Fig. S4. MSD plots of nucleosome motion in a longer time range and log-log scale. (A) MSD plots (± SD among cells) of nucleosome motion in 10T1/2 and CIRAS-3 cells over a longer time range (0.05 to 1.5 s). N = 20 cells for each cell line. Not significant (N.S.) by Kolmogorov-Smirnov test for 10T1/2 versus CIRAS-3 (p = 0.1745). (B) Log-log plot of the MSD data from Figs. 3B and S4A. The indicated lines were fitted using the data from t = 0.05 to 0.5 s.
MovieS1.avi
Movie S1. Movie data (50 ms/frame) of single nucleosomes labeled with TMR in a living 10T1/2 cell recorded by sCMOS ORCA-Fusion BT camera (Hamamatsu Photonics) (left). Note that clear and well-separated dots are visualized with a single-step photobleaching profile after background subtraction. The tracked data by u-track (MATLAB package; (Jaqaman et al. 2008)) was overlaid on the original movie and marked with circles (right).
MovieS2.avi
Movie S2. Movie data (50 ms/frame) of single nucleosomes labeled with TMR in a living CIRAS-3 cell recorded by sCMOS ORCA-Fusion BT camera (Hamamatsu Photonics) (left). Note that clear and well-separated dots are visualized with a single-step photobleaching profile after background subtraction. The tracked data by u-track (MATLAB package; (Jaqaman et al. 2008)) was overlaid on the original movie and marked with circles (right).
FigureS5.tif

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Chromatin organization and behavior in HRAS-transformed mouse fibroblasts

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