Repeat-rich RNA guides repetitive genomic elements into biomolecular condensates for heterochromatin organization and muscle integrity
Jinmi Choi, Sangha Park, Nahee Kim, Soonho Kwon, Jinyoung Park, Somin Lee, Hongmin Lee, Keonjin Kang, Taewan Kim, Jaehoon Sul, Dong-Gyu Jo, Hong-Duk Youn, Eun-Jung Cho
TL;DR
A muscle-specific RNA called ChRO1 helps organize heterochromatin into condensates, maintaining chromatin structure and muscle integrity.
Contribution
ChRO1 is a novel repeat-rich RNA that drives heterochromatin condensation and protects muscle cells from atrophy.
Findings
ChRO1 promotes heterochromatin clustering and partitions heterochromatin proteins in muscle cells.
Disruption of ChRO1 leads to chromocenter disintegration and muscle atrophy.
ChRO1 mitigates atrophy in chemically induced models, highlighting its protective role.
Abstract
Biomolecular condensation is a pivotal mechanism in chromatin organization and nuclear compartmentalization. However, the molecular mechanism that drives heterochromatin organization and selectively partitions heterochromatin components in muscle cells remains unclear. Furthermore, its pathological implications remain unexplored. Here, we demonstrate that ChRO1, a muscle-specific RNA enriched with simple dinucleotide repeats, is associated with static heterochromatin foci containing similar repetitive elements in mouse muscle cells. Through its CU-repeat-rich region, ChRO1 promotes heterochromatin clustering and facilitates the selective partitioning of heterochromatin proteins, as shown in vitro and in C2C12 cells. Consequently, chromatin interaction stabilized at ChRO1-bound regions, reinforcing TAD boundaries and promoting inactive chromatin states. The enhanced intra- and…
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Figure 7- —National Research Foundation of Korea10.13039/501100003725
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Taxonomy
TopicsGenomics and Chromatin Dynamics · RNA Research and Splicing · Muscle Physiology and Disorders
Introduction
Recent advances in biomolecular condensate research have revealed the mechanisms that maintain high local concentrations of regulatory factors, ensuring the optimal efficiency of various biological processes [1–5]. Biomolecular condensates are dynamic compartments that concentrate proteins and nucleic acids to facilitate selective biochemical reactions. For example, transcription factors, including RNA polymerase II, MED1, BRD4, elongation factors, and splicing factors, generate nuclear condensates to promote efficient and selective transcription regulation [6–8]. Upon stimulation, factors involved in DNA damage repair and stress response enhance the formation of such condensates [9–11]. Similarly, HP1α and other heterochromatin-associated proteins organize into condensates to establish heterochromatin domains [12–14]. However, the mechanisms of dynamic and selective partitioning of biomolecular components into condensates under different circumstances remain unknown.
Biomolecular condensates and the spatial organization of chromatin are closely linked, reinforcing each other [15–17]. In particular, heterochromatin compartmentalization is critical for organizing nuclear structure, protecting DNA from damage, silencing repetitive DNA elements, preventing abnormal chromosomal rearrangement, and preserving genome function and stability [18, 19]. H3K9 trimethylation (H3K9me3) and HP1α [20–22] play important roles in the establishment and maintenance of heterochromatin. Two hypotheses have been proposed to explain how HP1α induces compact heterochromatin compartments. The bridging model [23, 24] suggests that HP1α forms bridges between nucleosomes containing H3K9me3 and collapses heterochromatin into globular-like domains with stiff and sturdy structures. Alternatively, the liquid–liquid phase separation (LLPS)-based model proposes that the local concentration of HP1α increases, resulting in condensate formation [12, 13]. Notably, heterochromatin compartments formed by these models would exhibit distinct physicochemical and functional characteristics.
The pericentromeric heterochromatin (PCH), characterized by major satellite repeats, represents a key silencing domain in most eukaryotic genomes. PCH regions from different chromosomes coalesce into a higher-order interchromosomal structure at the mesoscale, known as a chromocenter [25, 26]. This feature is highly cell-type-specific and often becomes prominent during cellular differentiation in muscle and neurons [27]. In C2C12 myoblast cells, a well-established model of myogenesis, PCH clusters into chromocenters where heterochromatin-associated proteins, such as HP1α, HP1β, HP1γ, ATRX, DAXX, and MeCP2, accumulate in high concentrations [28]. Interestingly, some heterochromatin proteins, such as HP1γ and DAXX, are excluded from PCH before differentiation but are specifically recruited into chromocenters during differentiation, contributing to chromocenter properties distinct from PCH. Additionally, distant genomic regions characterized by repetitive elements and repressive epigenetic marks spatially organize to the vicinity of PCH, potentially through LLPS-like mechanisms, as observed in Drosophila embryos [29]. Disruption of chromocenters leads to nuclear membrane instability, micronuclei formation, DNA damage, and cell death, underscoring their pivotal role in maintaining genome function [25, 27]. However, the mechanism driving chromocenter assembly, selective partitioning of components, and its pathological consequences remain largely unexplored.
In this study, we investigated the mechanism underlying myotube-specific chromocenter formation, selective partitioning of genomic and protein components, and functional and pathological significance of chromocenter organization. Our findings revealed that RNA-driven condensate organization plays a critical role in the formation of chromocenter compartments. Specifically, the low-complexity region of muscle-specific ChRO1 RNA promotes clustering of static PCH and genomic domains characterized by similarly low-complexity repeats, such as satellites and simple repeats. By enhancing condensation of heterochromatin-associated proteins, including H3K9me3, HP1α, or HP1β, ChRO1 increased the proximity of repeat-rich genomic loci and strengthened their interactions. Meanwhile, ChRO1 regulated the partitioning of DAXX and HP1γ into chromocenters, contributing to the peculiar chromocenter environment in myotubes. Furthermore, we demonstrated that chromocenter integrity is essential for preventing muscle atrophy. Overall, this study provides novel insights into RNA-mediated heterochromatin compartmentalization through dynamic condensate organization and selective partitioning.
Materials and methods
Reagents and materials
Reagents and materials, including plasmids and cell lines are listed in Supplementary Table S1.
Animals and ethics
The CRISPR/Cas9 technology was used by Macrogen Inc. (Seoul, Korea) to produce ChRO1 knockout (KO) mice. ChRO1 KO and littermate controls were housed in Sungkyunkwan University’s (SKKU) pathogen-free animal facility. Mice were kept on a 12-h light/dark cycle at room temperature (20°C–25°C, RT), with unrestricted access to food and drink. For the study, muscle tissues were obtained from ChRO1 KO and littermate control mice (n = 5 for each). All animal experiments were approved by the SKKU Institutional Animal Care and Use Committee (IACUC) and performed in accordance with the guidelines in the Guide for the Care and Use of Laboratory Animals (SKKUIACUC2022-03-09-1 and SKKUIACUC2022-05-68-1).
Cell lines and cell transfection
C2C12 murine myoblast cells and NIH3T3 mouse fibroblast cells were obtained from the American-type culture collection and grown in a growth medium (GM) consisting of Dulbecco’s modified Eagle medium (DMEM) with 10% (v/v) fetal bovine serum at 37°C and 5% CO_2_. GM was replaced with a differentiation medium (DM), which was composed of DMEM and 2% horse serum (Gibco), to convert C2C12 myoblasts to myotubes. To generate ChRO1a-expressing cell lines, TRE3G-ChRO1a, TRE3G-ChRO1a (1–413), or pBabe-puro-MS2-ChRO1a were transfected into C2C12 or NIH3T3 cells using NEON electroporation (pulse voltage, 1050 V; pulse width, 30 ms; pulse number, 2) (Thermo Fisher Scientific) or Lipofectamine 2000 (Thermo Fisher Scientific), followed by puromycin selection. Cells were treated with 1 µM doxycycline to induce ChRO1a.
The ChRO1 KO cell line was established using CRISPR/Cas9. C2C12 cells were transfected with the CRISPR/Cas9–guide RNA (gRNA) plasmid using Lipofectamine Stem (Invitrogen) according to the manufacturer’s protocol. Forty eight hours after transfection, cells were sorted by Green Fluorescent Protein (GFP) using FACS, followed by selection with 4 µg/ml puromycin for 48 h. KO efficiency was validated by genomic polymerase chain reaction (PCR), and loss of ChRO1 RNA expression was confirmed by quantitative real-time PCR (qRT-PCR).
Plasmids
Plasmids pcDNA 3.0-ChRO1a-mCherry, in either sense (S) or antisense (AS) orientation, were generated via subcloning ChRO1a from pBabe-ChRO1a (S) and pBabe-ChRO1a (AS) into EcoRI/BamHI sites of pcDNA 3.0-mCherry, respectively. pBabe-ChRO1a (S) and (AS) were described previously [30]. Various regions of ChRO1a were obtained via PCR and inserted between BamHI/EcoRI to generate pcDNA 3.0-ChRO1a (full length, 1–1338), ChRO1a (1–413), ChRO1a (1–824), ChRO1a (414–1338), and ChRO1a (825–1338). These constructs were used for in vitro transcription. For inducible expression of ChRO1a, ChRO1a full length and ChRO1a (1–413) were PCR-amplified and inserted between NcoI/BsrGI of the inducible CasPex expression plasmid (Addgene #97421) to generate TRE3G-ChRO1a (S) and TRE3G-ChRO1a (1–413), respectively. To study localization of overexpressed ChRO1, MS2-binding domain repeats from pSL-MS2-12x (Addgene #27119) were digested with BamHI/BglII, and inserted into pBabe-ChRO1a (S). The resulting MS2-ChRO1a was visualized using MS2 coat proteins (MCPs) fused to tdTomato (NLS-HA-tdMCP-tdTomato, Addgene #183938). For CRISPR/Cas9-mediated genome editing of ChRO1, gRNAs targeting regions −582 and 402 bp relative to the ChRO1 transcription start site were designed (Supplementary Table S1), annealed, and cloned into BbsI site of pSpCas9(BB)-2A-Puro (Addgene #48139) and pSpCas9(BB)-2A-GFP(Addgene #48138), respectively. To generate the 4xMajSat repeat construct, a 4xMajSat DNA was obtained by digesting pγSat (Addgene #39238) with BamHI/SalI and inserted between BamHI and XhoI of pcDNA3.0.
Bacterial expression plasmids, T7pET-6xHis-mEGFP-Med1 (IDR, 600–1581) and T7pET-6xHis-mEGFP-hHP1α, were provided by Dr Richard Young. A plasmid for 6xHis-mEGFP was generated by digestion of T7pET-6xHis-mEGFP-hHP1α with EcoRI/BamHI and religation after blunt end formation. Mouse HP1 complementary DNAs (cDNAs) were subcloned from pRSETA-mHP1α, -mHP1β, and -mHP1γ into the BamHI/EcoRI sites of T7pET-6xHis-mEGFP-hHP1α to generate T7pET-6xHis-mEGFP-mouse HP1α, HP1β, and HP1γ, respectively. To construct T7pET-6xHis-mCherry-mHP1α, mCherry was PCR-amplified from pEJS578_DD-dSpyCas9-mCherry-APEX2 (Addgene #108570), digested with EcoRV/BsrGI, and inserted into NcoI (blunt-ended)/BsrGI of T7pET-6xHis-mEGFP-mHP1α to replace mEGFP with mCherry. DAXX was PCR-amplified from Flag-DAXX (a gift from Dr Frank Funari), digested with EcoRI/BamHI, and ligated with a similarly digested T7pET-6xHis-mEGFP-mHP1α to replace mHP1α with DAXX. All plasmids were validated via sequencing. Primers used in cloning are listed in Supplementary Table S2.
RNA extraction, reverse transcription, and quantitative real-time PCR
Muscle tissues were extracted from mice and pulverized with liquid nitrogen using a mortar. Cells were collected through centrifugation, and total RNA was extracted using TRIzol (Thermo Fisher Scientific) as per the manufacturer’s instructions. The concentration and purity of the total RNA were determined using a NanoDrop spectrophotometer (Thermo Fisher Scientific) based on the OD_260_/OD_280_ ratio. An aliquot of 1.0 μg of total RNA and a cDNA synthesis kit (Cellsafe) were used for cDNA synthesis. qRT-PCR was performed using Thunderbird SYBR qPCR Mix (Toyobo) and measured using a CFX96 system (Bio-Rad). Relative RNA quantification was conducted using the 2^−ΔΔCT^ method, with Gapdh serving as an internal control for normalization, unless stated otherwise. The primers employed for qRT-PCR are listed in Supplementary Table S3.
Immunofluorescence staining of paraffin sections
For immunohistological analysis, five mice from each of the ChRO1 WT and KO groups were used [2-months-old male (n = 1), 5-months-old male (n = 1), 13-months-old male (n = 2), 12-months-old female (n = 1) from each group]. Muscle tissues (gastrocnemius, soleus, and tibialis anterior) were removed from mice, immediately fixed in 4% formaldehyde in phosphate-buffered saline (PBS) for 10 min at RT, and embedded in paraffin blocks. The samples were sliced into 5-μm-thick sections using a paraffin slicer. Immunostaining was performed by incubating the deparaffinized sections with wheat germ agglutinin conjugated with AlexaFluor 488 (Thermo Fisher Scientific) for 30 min at 37°C. Nuclei were counterstained with 50% glycerol containing 2 µg/ml DAPI. For the detection of DAXX and HP1α, antigen retrieval was performed by heating the tissue sections in 1× sodium citrate buffer (10 mM sodium citrate, 0.05% tween 20, pH 6.0) for 20 min. The tissue sections were then incubated with 0.5% Triton-X100 in DAKO Protein Block Serum-Free Ready-to-Use (Agilent Dako, X0909). Subsequently, DAXX (Santa Cruz, SC7152) and H3K9me3 (Abcam, ab8898) antibodies, diluted 1:100 in DAKO Antibody Diluent (Agilent Dako, S0809), were applied to the tissue sections and incubated overnight at 4°C. The following day, the tissue sections were briefly washed and incubated with Goat anti-rabbit IgG-AlexaFluor 488 (Invitrogen, A11008) and Goat anti-rabbit IgG-AlexaFluor 594 (Invitrogen, A11012) at a 1:200 dilution in DAKO Antibody Diluent for 2 h in the dark. After washing, the tissue sections were mounted with 50% glycerol containing 5 μg/ml of DAPI. The sections were examined using an LSM700 confocal microscope (Carl Zeiss). Image processing was performed using ImageJ software (National Institutes of Health, USA).
DNA recovery assay
Cells were cross-linked with 1% formaldehyde for 5 min at RT and quenched by 0.125 M glycine with shaking for 5 min at RT. After washing three times with PBS, cells were lysed with FAIRE buffer I [50 mM HEPES-KOH, pH 7.5, 140 mM NaCl, 1 mM ethylenediaminetetraacetic acid (EDTA), 10% glycerol, 0.5% NP-40, 0.25% triton X-100] for 10 min at 4°C. Following centrifugation at 5000 rpm for 5 min at 4°C, the supernatant was removed, and the pellet was resuspended in FAIRE buffer II [10 mM Tris–HCl, pH 8.0, 200 mM NaCl, 1 mM EDTA, 0.5 mM ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA)], incubated for 10 min at RT, then centrifuged at 13 000 rpm for 10 min at 4°C. The pellet was resuspended with FAIRE buffer III (10 mM Tris–HCl, pH 8.0, 100 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, 0.1% sodium deoxycholate, 0.5% N-lauroylsarcosine) and sonicated for 30 min at 0°C using Bioruptor UCD200-TM (Cosmo Bio). After centrifugation at 13 000 rpm for 10 min at 4°C, 10% of the supernatant was taken and decrosslinked with RNase A and proteinase K at 65°C overnight. DNA from the remaining cell lysate was purified by phenol, chloroform/isoamyl alcohol extraction, followed by decrosslinking. Final purification of DNA was performed after overnight decrosslinking. FAIRE enrichment was analyzed via qRT-PCR using primers as listed in Supplementary Table S3.
Immunofluorescence confocal microscopy
Cells plated on glass coverslips were used for immunostaining. The cells were fixed with 4% paraformaldehyde in PBS for 10 min at RT, followed by a wash with PBS. Subsequently, the cells were permeabilized with 0.1% triton X-100 in PBS for 10 min at RT, followed by another wash with PBS. Cells were treated with 2% bovine serum albumin (Gold Biotechnology) in PBS (PBA) for 10 min at RT for blocking and incubated with a primary antibody in 2% PBA for 1 h at RT or overnight at 4°C. After washing three times with 0.1% tween 20 in PBS, cells were incubated with secondary antibodies, such as Alexa 594- or Alexa 488-conjugated goat anti-rabbit IgG (Thermo Fisher Scientific) or anti-mouse IgG (Thermo Fisher Scientific), for 1 h at RT. Nuclear DNA was stained with DAPI (2 μg/ml, Molecular Probe). Immunofluorescence analysis was performed using an LSM 700 confocal microscope (Carl Zeiss), and image processing was conducted using the ZEN software (Carl Zeiss) or ImageJ (National Institutes of Health, USA) for quantifying the number of heterochromatin foci per nucleus and estimating foci overlap.
Live-cell imaging
Live-cell imaging was performed on differentiated C2C12 cells cultured on glass-bottom dishes (SPL). Nuclei were stained by incubating cells with 1 μg/ml Hoechst 33342 (Invitrogen) in DM for 10 min at 37°C in a humidified incubator. Subsequently, cells were washed once with PBS and returned to DM for the duration of the imaging session. Confocal micrographs were acquired using a Zeiss LSM 700 microscope (Carl Zeiss), and image processing was conducted using the Zen software (Carl Zeiss) or ImageJ (National Institutes of Health, USA). Cell morphology was monitored throughout image acquisition to ensure minimal phototoxicity and cell health.
RNA fluorescence in situ hybridization
Custom Stellaris RNA fluorescence in situ hybridization (RNA FISH) probe sets were designed using the Stellaris probe designer software (Biosearch Technologies), targeting ChRO1 or LacZ. Biotin-labeled probes were subsequently ordered from Biosearch Technologies according to the manufacturer’s instructions (Supplementary Table S4). Cells were fixed with methanol/acetic acid at RT for 10 min, followed by washing with wash buffer A (Biosearch technologies, SMF-WA1-60). Hybridization was performed overnight in hybridization buffer (Biosearch technologies, SMF-HB1-10) with 10% formamide. After hybridization, cells were washed with wash buffer A for 30 min at 37°C and then blocked for 30 min using DAKO Protein Block Serum-Free Ready-to-Use (Agilent Dako, X0909). The cells were then incubated with Fluorescein Avidin DCS (5 μg/ml, Vector Laboratories, A-2011) for 30 min at RT. Slides were washed twice for 3 min each with wash buffer B (Biosearch technologies, SMF-WB1-20), followed by incubation with Biotinylated Anti-Avidin (5 μg/ml, Vector Laboratories, BA-0300) for 30 min at RT. After washing again twice for 3 min each with wash buffer B, a second incubation with Fluorescein Avidin DCS (5 μg/ml) was performed for 30 min at RT. Finally, slides were washed twice for 5 min each with wash buffer B and mounted with DAPI. Imaging was performed using an LSM 700 confocal microscope (Carl Zeiss) equipped with a 6.5 Airy Unit pinhole to increase signal collection while compromising some optical sectioning. Images were processed using ZEN software (Carl Zeiss) or ImageJ (National Institutes of Health, USA).
Western blot analysis
Proteins were extracted from C2C12 cells that had been differentiated for 5 days. The concentration of protein extracts was measured using the Bradford assay, and 30 μg of each extract was loaded onto sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) gel. After gel electrophoresis, the proteins were transferred onto a PVDF or NC membrane. The membrane was then blocked with 5% skim milk. Subsequently, the membrane was incubated overnight at 4°C with H3K9me3 antibody (Abcam, ab8898), H3 antibody (Abcam, ab18521), MyHC antibody (Millipore #05-716), and GAPDH antibody (Santa Cruz, SC25778) at a dilution of 1:1000 in 1% BSA/Tris-buffered saline with Tween 20 (TBST). Following the primary antibody incubation, the membrane was washed with TBST and incubated with secondary antibodies (Bethyl A120-101P and A90-116P) at a dilution of 1:10000 in TBST for 1 h at RT. After three washes with TBST, the membrane was exposed to an ECL solution (Abfrontier, LF-QC0103) to generate a detectable signal. The signal was captured using X-ray films.
Recombinant protein purification
Protein purification was performed following the methods described by Klein et al. [31]. In brief, BL21 Escherichia coli cells were transformed with T7pET-6xHis-mEGFP DNA. A single colony was picked for the scale-up culture and grown at 37°C in 100 ml of LB medium with kanamycin. Protein expression was induced by isopropyl β-D-1-thiogalactopyranoside (IPTG) at a final concentration of 0.5–1 mM, followed by continued growth for 16–20 h at 18°C. After cell harvest, bacterial pellets were resuspended in lysis buffer (1 mg/ml lysozyme, 1 mM phenylmethylsulfonyl fluoride (PMSF), 1× protease inhibitor cocktail in PBS) and incubated for 30 min at 4°C under rotation. After sonication (5 min × 2 with cycles of 1.0 s on and 1.0 s off using VCX750, Sonics & Materials), lysates were centrifuged at 13 000 rpm at 4°C for 15 min. The supernatant was incubated with Ni-NTA agarose (QIAGEN) at 4°C for 2 h. The Ni-NTA agarose slurry was poured onto Poly-Prep Chromatography Columns (Bio-Rad) and washed with wash buffer (1 mM PMSF, 1× protease inhibitor cocktail, 10 mM imidazole, 300 mM NaCl in PBS). Proteins were eluted with an elution buffer containing increasing concentrations of imidazole. High-purity, high-concentration fractions were pooled, and the imidazole buffer was exchanged with storage buffer (50 mM Tris–HCl, pH 7.5, 125 mM NaCl, 1 mM DTT, 10% glycerol in DEPC-treated water) using an Amicon Ultra Centrifugal Filter (Millipore) until the desired concentration was reached.
In vitro transcription assay
Plasmids such as pcDNA3.0-ChRO1a (full length, 1–1338), pcDNA3.0-ChRO1a (AS), pcDNA3.0-ChRO1a (1–413), pcDNA3.0-ChRO1a (1–824), pcDNA3.0-ChRO1a (414–1338), and pcDNA3.0-ChRO1a (825–1338) were linearized via EcoRI digestion. DNA was purified via phenol/chloroform extraction and ethanol precipitation. One microgram of each DNA was used for in vitro transcription, either using the TranscriptAid T7 High Yield Transcription Kit (Thermo Fisher Scientific) or the High Yield T7 Cy5 RNA Labeling Kit (Jena Bioscience), following the manufacturer’s protocol.
In vitro phase separation assay
Recombinant proteins, including HP1s, were added to the droplet formation buffer (50 mM Tris–HCl, pH 7.5, 125 mM NaCl, 1 mM DTT, 10% glycerol, and 10% PEG-8000 in DEPC-treated water) at the indicated concentrations in low-binding microcentrifuge tubes (Eppendorf, 022431081). After mixing, the samples were transferred to a 384-well glass-bottom plate (Cellvis, P384-1.5H-N) and incubated at RT for at least an hour. Imaging was performed using the LSM 700 confocal microscope (Carl Zeiss). To assess the effect of RNA on droplet formation, RNAs such as ChRO1a, yeast tRNAs (Sigma), or 4xMajSat transcripts were combined with the proteins in the droplet formation buffer before loading onto the 384-well plates. Quantification of phase separation involved analyzing the number and area of droplets using ImageJ (National Institutes of Health, USA).
Fluorescence recovery after photobleaching assay
Regions of interest (ROI) were selected, and laser power was set at 100% with 405, 488, and 555 nm lasers for bleaching. Bleaching was initiated after the first scan and was repeated until the intensity dropped to 50%. Images were taken 50 times at 1.0 msec intervals. A control ROI without photobleaching was included in all experiments. The time point with the minimum fluorescence was considered a recovery time point of 0 s. The fluorescence during recovery was measured relative to the fluorescence at time point 0 s.
RNA immunoprecipitation
Cells were crosslinked with 1% formaldehyde for 10 min, followed by quenching with 0.125 M glycine. After washing with PBS twice, cells were lysed with RNA immunoprecipitation (RIP) lysis buffer (50 mM Tris–HCl, pH 8.0, 400 mM NaCl, RNase inhibitor). Lysates were sonicated for 15 cycles of 1 min on and 1 min off using Bioruptor UCD200-TM (Cosmo Bio) and centrifuged at 13 000 rpm for 10 min. Supernatants were incubated with the indicated antibodies overnight and then bound to agarose protein A/G beads for 3 h at 4°C. The bead-bound complex was washed with RIP buffer three times and decrosslinked at 70°C for 1 h. TRIzol (Invitrogen) was added to extract RNAs, which were then subjected to cDNA synthesis and qRT-PCR (refer to the RNA extraction section in the “Materials and methods” section).
Chromatin isolation by RNA purification–qRT-PCR
Cells were harvested by trypsinization, washed with PBS, and crosslinked with 1% glutaraldehyde for 10 min at RT (20 million cells per probe). After quenching with 0.125 M glycine, cells were pelleted, washed, and lysed in lysis buffer (50 mM Tris–HCl, pH 7, 10 mM EDTA, 1% SDS, proteinase inhibitors, and RNase inhibitors). Lysates were sonicated using a Bioruptor Pico (high setting, 30 s ON/30 s OFF pulses, Diagenode) for 45 min until DNA fragments ranged 100–500 bp, then cleared by centrifugation. Hybridization was performed using the same Stellaris probe sets targeting ChRO1 and LacZ as described in RNA FISH procedures and Supplemental Table S4. For hybridization, 1 ml sonicated chromatin was mixed with 2 ml hybridization buffer (50 mM Tris–HCl, pH 7, 10 mM EDTA, 1% SDS, 750 mM NaCl, 15% formamide, proteinase inhibitors, and RNase inhibitors), and 50 nM probe and incubated at 37°C overnight. The hybridization reaction was incubated with Streptavidin C-1 beads (Invitrogen, 65001) at 37°C for 30 min, followed by five washes at 37°C. Beads were split for RNA and DNA isolation. RNA was extracted by proteinase K treatment, TRIzol (Invitrogen)-chloroform extraction, ethanol precipitation, and DNase treatment, and then analyzed by qRT-PCR. DNA was eluted twice with elution buffer (50 mM NaHCO, 1% SDS, RNase A, RNase H). After proteinase K digestion, DNA was purified using QIAquick PCR Purification Kit (QIAGEN), and analyzed by qRT-PCR.
ChRO1 sequence analysis
To quantify the repetitiveness of human and mouse ChRO1 RNA sequences, we carried out k-mer analysis using RepLoc (k-mer = 4, 6, and 8) [32]. The centroid structure of ChRO1 with minimal base-pair distance to all other secondary structures in the Boltzmann ensemble was predicted using ViennaRNA Package 2.0 [33]. RNA–protein interaction prediction was carried out using PRIdictor, which provides scores at every position of the RNA sequence [34].
High-throughput chromatin immunoprecipitation combined with Hi-C
High-throughput chromatin immunoprecipitation combined with Hi-C (HiChIP) experiments were performed as previously described [35]. Briefly, cells were differentiated for 4 days, harvested by trypsinization, and crosslinked with 1% formaldehyde for 10 min. Crosslinking was quenched with 0.125 M glycine, followed by two washes with PBS. Cells were lysed in lysis buffer (10 mM Tris–HCl, pH 7.5, 10 mM NaCl, 0.2% NP-40, proteinase inhibitors). The pellet was incubated with 0.5% SDS at 62°C for 10 min, after which triton X-100 was added, and samples were incubated at 37°C for 15 min. Chromatin was digested overnight by DpnII (NEB, R0543) at 37°C. Biotinylation was performed using biotin-dATP (Jena, NU-835-BIO14), followed by ligation at RT for 6 h. The samples were then sonicated using a Bioruptor Pico (high setting, 30 s ON/30 s OFF pulses, Diagenode) for 3 min. Immunoprecipitation was carried out overnight at 4°C with an anti-H3K9me3 antibody (Abcam, ab8898). After bead binding and five washes, DNA was eluted with elution buffer (10 mM Tris–HCl, pH 8.0, 1% triton X-100, 5 mM EDTA, 300 mM NaCl, 0.4% SDS, proteinase K) at 65°C overnight, and then at 55°C for 1 h for a second elution. DNA was purified using AMPure beads (Beckman) and captured on Streptavidin C-1 beads (Invitrogen, 65001). Tagmentation was performed on beads with 1.5 μl of Tn5 transposase (Diagenode, C C01070012) at 55°C for 10 min, followed by inactivation with 50 mM EDTA, and three washes. Biotinylated DNA fragments were PCR-amplified for 10 cycles using KAPA HiFi HotStart Ready Mix (KAPA), and cleaned with AMPure beads (Beckman). Libraries were sequenced as paired-end 150 bp reads on a Novaseq X platform (Macrogen).
HiChIP analysis
Sequencing reads were preprocessed using Fastp (v1.0.1) [36] to remove adaptor sequences, filter out low-quality reads (Q < 20), and eliminate duplicate reads. After filtering, each sample yielded ~40 million paired-end reads. Reads were mapped to the mouse genome assembly mm10 using Bowtie2 [37, 38] with default parameters, and downstream processing was performed with HiC-Pro 3.1.0 [39]. Only valid interaction pairs were retained, excluding dangling ends, religated fragments, self-cycle products, and single-end reads, resulting in mapping efficiencies between 80% and 89%. Quality control was performed withiin HiC-Pro 3.1.0 [39], including contact map generation and normalization. Contact matrices were subsequently converted into multiresolution mcool format using cooltools [40] for visualization and downstream analyses. Specifically, the insulation score with 10 kb resolution, 20 kb window size.
Chromatin interaction peaks were inferred with FitHiChIP 11.0 [41] using a peak-to-all background model and a q-value cutoff of 0.05. Peak regions were extended by 5 kb from the summit. Replicates showed high consistency, with overlap rates of 69.1% (128,980/186,574 peaks) for wild type (WT1 versus WT2) and 86.45% (122,507/147,527 peaks) for KO (KO1 versus KO2) samples. Chromatin loop calling was performed using a bin size of 10 kb and q < 0.05. For differential loop analysis, replicates were processed separately in FitHiChIP [41] with significance thresholds of FDR < 0.05.
For visualization and integrative downstream analysis, HiCExperiment and HiContact packages [42] in R were employed to explore contact maps, loop structures, and replicate concordance.
ChIRP-seq data analysis
Chromatin isolation by RNA purification (ChIRP)-seq data were obtained from GSE94498 and GSE113248 [43, 44]. The data were mapped to the mouse genome assembly mm10 using Bowtie2 [37, 38]. Peaks were called using MACS2.4 [45]. For further analysis, we selected 4054 overlapping peaks from probes with even numbers and odd numbers, excluding peaks from LacZ. Coordinates and families of genomic repetitive elements were acquired from RepeatMasker [46]. For de novo motif analysis, MEME from the MEME suite 5.5.7 was used to find motifs within ±100 bp from the midpoint of ChIRP-seq peaks [47]. To define colocalized regions between ChRO1 peaks and satellite repeats, we used the findOverlaps() function from the GenomicRanges R package.
ATAC-seq data analysis
ATAC-seq data were obtained from GSE76010 [48]. The data were mapped to the mouse genome assembly mm10 using Bowtie2 [37, 38]. The data were normalized to the library size, and mean coverage was plotted on the mid-point of ChIRP-seq peaks or transcription start sites of nearby genes.
RNA-seq data analysis
RNA-seq count data were obtained from GSE113165 and GSE247438 [49, 50]. The data were normalized to the library size. For Fig. 6H, Pearson correlation coefficients and P-values for the fold change of ChRO1 and other genes (post-bedrest over pre-bedrest) were calculated. Genes with a P-value of <.1 were filtered for further analysis. Gene ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analyses were performed using DAVID [51, 52].
Hi-C data analysis
Hi-C raw data were obtained from PRJNA338854 [53]. The data analyses were performed using Homer’s analysis tools and workflow [54]. To be more specific, the data were mapped to the mouse genome assembly mm10 using Bowtie2 [29, 37]. Multimapping reads and PCR duplicates were removed, and only uniquely mapped read pairs were converted to Homer tag directory format [54]. Interactions, loops, and TADs (IR score threshold 1.5) in myoblast and myotube were obtained at 10 kb resolution. Compartmental states (A/B) were identified in 50 kb resolution using principal component analysis analysis on Hi-C data of myoblasts and myotubes. The correlation of Hi-C experiments from myoblasts and myotubes was calculated to identify genomic regions that switch compartmental states. Interaction heatmaps and a circos plot were generated using HiContacts and circlize packages in R [42, 55].
Quantification and statistics
Statistical analysis was performed using GraphPad Prism. Bar plots represent the mean ± SD. Box plots represent the interquartile range (box), median (line within the box), and maximum and minimum values excluding outliers (whiskers). Comparison between two groups was performed using an unpaired two-tailed Student’s t-test. P-values <.05, .01, .001, and .0001 were indicated by *, **, ***, and ****, respectively. Statistical details, including P-values and sample sizes (n), are described in the figure legends. Experiments were performed at least three times unless otherwise specified.
Results
CU-rich RNA promotes heterochromatin condensate organization during differentiation
During skeletal myogenesis, PCH consolidates into chromocenters, which contain a high concentration of heterochromatin proteins [30, 56]. To assess chromocenter dynamics, C2C12 myoblast cells were differentiated into myotubes and treated with 1,6-hexanediol (1,6-HD), a hydrophobic compound commonly used to study the physicochemical features of LLPS [57, 58]. Treatment with 1,6-HD (1.5%) dispersed chromocenters into separate foci without complete dissolution, as shown by the increased number and reduced size of heterochromatin foci in fixed cells stained with 4′,6-diamidino-2-phenylindole (DAPI) (Fig. 1A). This effect was specific to 1,6-HD, since 2,5-hexanediol (2,5-HD), a less hydrophobic molecule, did not affect chromocenter integrity (Fig. 1A). We next performed live-cell imaging with Hoechst 33342 to capture real-time chromocenter behavior during 1,6-HD treatment. While changes in foci number were modest, 1,6-HD treatment induced clear alterations in chromocenter intensity and integrity relative to untreated cells (Fig. 1B, Supplementary Fig. S1A and B, and Supplementary Movies S1 and S2). Furthermore, 1,6-HD treatment increased the extractability of pericentromeric and centromeric DNAs and release of chromocenter-associated RNAs (Supplementary Fig. S1C). ChRO1 expression was only marginally affected (Supplementary Fig. S1C). The dynamic and reversible nature of chromocenters was demonstrated by their recovery following 1,6-HD washout (Fig. 1C and Supplementary Fig. S1D) [3]. Notably, heterochromatin foci in myoblasts and myotubes exhibited distinct responses to 1,6-HD. Specifically, myoblast heterochromatin foci were highly resistant to 1,6-HD, while myotube chromocenters were more susceptible, suggesting that chromocenters in myotubes are more liquid-like compared to the stable and solid-like foci in myoblasts (Fig. 1D). Altogether, these results revealed that chromocenters are dynamic and reversible clusters composed of PCH. The incomplete dissolution of PCH foci upon 1,6-HD treatment distinguishes their physical state from canonical LLPS. To capture these unique organizational and dynamic properties, we hereafter refer to the clustering of PCH into dynamic chromocenters as condensation, and the resulting chromocenters as chromocenter condensates.
CU-rich RNA promotes heterochromatin condensate organization during differentiation. (A) Nuclei of C2C12 myotubes (MT) treated with 1.5% 1,6-hexanediol (1,6-HD) or 1.5% 2,5-hexanediol (2,5-HD). Left: Representative images of 4,6-diamidino-2-phenylindole (DAPI)-stained nuclei. Scale bar, 5 μm. Middle: Time-course quantification of the number of heterochromatin foci per nucleus. Right: Boxplots show foci area (μm2, y-axis) at 5, 10, and 15 min posttreatment (x-axis). n = 55 nuclei, three biological replicates. (B) Representative live-cell images of Hoechst 33342-stained MT nuclei before and after 1,6-HD treatment (0 and 15 min, respectively), taken from Supplementary Movies S1 and S2. Arrows indicate changes in heterochromatin foci intensity (pink, increased; blue, decreased), and the red arrow highlights an alteration in chromocenter integrity. (C) Number of heterochromatin foci per nucleus in MT following recovery from 1.5% 1,6-hexanediol (1,6-HD) treatment for 15 min, measured at indicated time points. n = 68 nuclei, three biological replicates. (D) Representative images of nuclei of myoblast (MB) and MT with or without 1,6-HD treatment (1.5%, 15 min). Right: Quantification of the number of heterochromatin foci per nucleus. n = 40 (MB), n = 60 (MT), three biological replicates. (E) Quantification of colocalization between indicated proteins and DAPI foci in MT with or without 1,6-HD treatment (1.5%, 15 min), by Pearson’s correlation coefficients. n = 60, three biological replicates. (F) Boxplot showing the distribution of Z-score of the interchromosomal interaction frequencies in MB and MT. (G) RNA FISH using ChRO1 and LacZ biotinylated probes. Biotin signal was detected by Fluorescein-conjugated Avidin DCS and amplified with biotinylated anti-Avidin and additional Fluorescein Avidin DCS. Right: Quantification of colocalization between biotin signal and DAPI foci. n = 50 nuclei. (H) Number of heterochromatin foci per nucleus of mouse fibroblast cells (NIH3T3) with or without doxycyline (Dox)-induced ChRO1a expression and/or 1,6-HD treatment (1.5%, 15 min). (EV; empty vector). n = 50, three biological replicates. (I) Number of heterochromatin foci per nucleus in MB with or without Dox-induced ChRO1a fragment (1–413, CUR) expression and/or 1,6-HD treatment (1.5%, 15 min). n = 75, three biological replicates. Statistical analyses and data presentation details are described in the “Materials and methods” section.
We next investigated whether chromocenter condensation affects the dynamic localization of heterochromatin proteins into chromocenters in a myotube-specific manner (Supplementary Fig. S1E and F) [30]. Using immunostaining, we examined their colocalization with DAPI foci after 1,6-HD treatment. Notably, H3K9me3, HP1α, HP1β, and ATRX remained associated with DAPI foci after 1,6-HD treatment, indicating their static interaction with heterochromatin (Fig. 1E and Supplementary Fig. S1G). In contrast, proteins such as HP1γ and DAXX, which translocate to chromocenters during myogenesis, were significantly delocalized by 1,6-HD (Fig. 1E and Supplementary Fig. S1G). This suggests that their myotube-specific enrichment at chromocenters depends on dynamic and selective condensation, resulting in heterochromatin protein composition distinct from that of myoblast heterochromatin foci.
We next asked whether chromocenter condensation is accompanied by mesoscale change of the 3D genome architecture. Hi-C analysis of myoblasts and myotubes [53] revealed that active compartments (A) in myoblasts shifted to more inactive states, while the inactive compartments (B) remained largely stable (Supplementary Fig. S2A). In parallel, TAD score analysis showed a more robust and stable TAD organization in myotubes (Supplementary Fig. S2B), exemplified by chromosome ends near the pericentromeric region (Supplementary Fig. S2C). Interchromosomal interactions were also increased, reflecting the multichromosomal composition of chromocenters (Fig. 1F). Collectively, these findings support heterochromatin reorganization with chromocenter condensation.
ChRO1 is a myotube-specific, chromatin-associated non-coding RNA (ncRNA) [30]. ChRO1 is enriched at chromocenters (Fig. 1G). However, its chromocenter localization and interaction with H3K9me3 were disrupted by 1,6-HD treatment (Fig. 1G and Supplementary Fig. S3A), indicating that ChRO1a is associated with H3K9me3-enriched heterochromatin and maintains chromocenters via condensation. To test whether ChRO1 is sufficient for heterochromatin condensation, ChRO1a, a short isoform containing part of intron 1, was expressed in myoblasts or NIH3T3 fibroblasts lacking endogenous ChRO1. ChRO1a colocalized with DAPI-stained heterochromatin (Supplementary Fig. S3B) and induced the clustering of heterochromatin foci in a 1,6-HD-sensitive manner, as evidenced by increased size and decreased number of DAPI foci (Fig. 1H and Supplementary Fig. S3C). We also confirmed that chromocenter deformation and impaired myogenic differentiation were rescued by exogenous expression of ChRO1a in ChRO1 KO C2C12 cells (Supplementary Fig. S3D–F).
To further understand how ChRO1 mediates global heterochromatin reorganization, we performed sequence analysis [32]. We found that simple CU dinucleotide repeats are particularly enriched within intron 1 of ChRO1, a feature that is also observed in human ChRO1 (Supplementary Fig. S3G). We therefore hypothesized that these repeats are crucial for ChRO1’s function in chromocenter organization. Indeed, the expression of the CU-rich region (CUR) of ChRO1a (1–413) was sufficient to induce chromocenter clustering in myoblasts in a 1,6-HD-sensitive manner (Fig. 1I and Supplementary Fig. S3H). Together, these results demonstrate that CU-rich ChRO1-driven condensation induces chromocenter formation and heterochromatin organization in myotubes, highlighting the role of repeat-rich RNA in mesoscale, cell-type-specific heterochromatin reorganization.
CU-rich ChRO1 associates with genomic low-complexity repeats
Given the established role of homotypic clustering of repetitive DNA elements and their transcripts in genome compartmentalization [59], we postulated that ChRO1 may specifically target low-complexity genomic regions via its simple dinucleotide repeats. To investigate this, we first characterized the global genomic binding profile of ChRO1. In line with our previous finding, ChRO1, also referred to as Charme in an independent study, has been shown to regulate myogenesis through cis interaction with the Nctc1 region [43]. Our analysis of published ChIRP-seq [43] further revealed that ChRO1 binds broadly across the genome, including the Nctc1 region on chromosome 7, supporting its role in both cis- and trans-regulation (Supplementary Fig. S4A). Remarkably, 91% of ChRO1 binding peaks overlapped with DNA repeats, compared to only 46% for MyoDeRNA used as a control (Supplementary Fig. S4B). De novo motif discovery revealed that ChRO1 preferentially binds to microsatellite repeats, particularly CA and CT dinucleotides, which resemble its own CU repeats (Supplementary Fig. S4C). Interestingly, metagene analysis revealed a significant enrichment of satellite and simple repeats at ChRO1 binding sites (Fig. 2A and Supplementary Fig. S4D), and conversely, these repeat classes showed the highest ChRO1 occupancy among repeat elements (Supplementary Fig. S4E and F). The specificity of ChRO1 occupancy was validated by examining multiple genomic regions, including Nctc1 region (Fig. 2B and Supplementary Fig. S4G and H). These data indicate that CU-rich ChRO1 associates with genomic regions enriched in similar low-complexity repeats.
CU-rich ChRO1 associates with genomic low-complexity repeats. (A) Relative repeat enrichment of indicated families of repeats within +/−500 bp of the center of ChRO1 peaks. Relative repeat content is calculated as the mean occurrence of repeats within ChRO1-bound sites normalized to total genomic repeat occurrence. (B) Normalized coverage tracks from ChRO1 or LacZ ChIRP-seq within 30 kb window surrounding satellite-bound ChRO1 peaks. The locus are indicated at the top of each panel. For each locus, ChIRP-seq coverage for odd and even probe sets displayed in orange and LacZ negative control is shown in gray. The red line marks the position of the ChRO1 peak overlapping with satellite repeats. (C) Coverage of satellite (left), SINE, and LINE (right) repeats across ChRO1-bound versus -unbound A/B compartments. Each column represents ChRO1-bound A compartment, ChRO1-bound B compartment, ChRO1-unbound A compartment, and ChRO1-unbound B compartment. X-axis: Relative position of repeats within the compartment, representing normalized coordinates from the start (1%) to the end (100%) of each compartment; Y-axis: individual A/B compartments, systematically ordered by their total SINE content (top to bottom). (D) Genome-wide distribution of ChRO1 binding peaks (ChRO1), satellite repeat regions (Satellite), and their overlap (ChRO1+satellite) across individual chromosomes. The ChRO1+satellite panel indicates genomic loci where ChRO1 peaks (ChIRP-seq) and satellite elements (RepeatMasker) preferentially colocalize at pericentromeric and telomeric regions. Arrows show the direction of peri-/centromeric region. (E) Histogram showing the distribution of ChRO1+satellite across normalized chromosome coordinates (5′ end at 3 Mb, 3′ chromosome end). Peaks from all chromosomes were combined with equal weighting per chromosome to account for differences in peak numbers across chromosomes. The dashed red line indicates chromosome ends at 5% and 95% of chromosome length. P-value was calculated using binomial test assuming a random distribution of peaks across normalized chromosomal positions (expected proportion at ends = 0.10). (F) Barplot showing the proportion of peaks localized to chromosome ends (normalized position < 0.05 or > 0.95) for three peak categories: ChRO1+satellite (16.4%), ChRO1 (11.3%), Satellite (11.3%). The dashed gray line indicates the expected proportion (10%) for random distribution. Three-way chi-square test revealed significant differences among the three peak groups (χ2 = 9.726, P-value = .0077). Pairwise comparisons with Bonferroni correction (α = 0 .0167) show that ChRO1+satellite is significantly more enriched at chromosome ends than ChRO1 and Satellite only. (G) Distribution of ChRO1 binding peaks (ChRO1), satellite repeat regions (Satellite), and their overlap (ChRO1+satellite) within 3 000 000–10 000 000 of chromosome 4, corresponding to the region shown in Fig. 2C. The ChRO1+satellite panel indicates genomic loci where ChRO1 peaks (ChIRP-seq) colocalizes with satellite elements (RepeatMasker). (H) Chromatin accessibility (ATAC-seq) at ChRO1 + Satellite regions in myoblasts (MB) and myotubes (MT). ATAC-seq signal intensity within +/−250 bp around ChRO1 + Satellite loci is compared between conditions. Statistical analyses and data presentation details are described in the “Materials and methods” section.
To understand how the repeat-specific targeting by ChRO1 influences heterochromatin compartmentalization into chromocenters (Fig. 1H and I), we examined the relationship between ChRO1 binding and genomic compartments. While SINEs and LINEs exhibited canonical compartmental distribution (A and B, respectively), satellite repeats were preferentially enriched within ChRO1-bound compartments regardless of A/B status (Fig. 2C). These results suggest that ChRO1 may promote the organization of satellite-rich regions into distinct compartments that are separable from the canonical A/B compartmental framework.
To further characterize these ChRO1-bound, satellite-enriched domains, we examined the chromosomal distribution of ChRO1-bound satellite repeats (hereafter ChRO1+satellite). These regions were markedly enriched at chromosomal ends, particularly near pericentromeric and telomeric regions (Fig. 2D and E). Although ChRO1 peaks and satellite repeats individually display relatively uniform distributions along chromosomes, sites of ChRO1+satellite show a strong bias toward chromosome ends (Fig. 2F). This is exemplified by the pericentromeric region of chromosome 4, where ChRO1+satellite sites align closely with TAD organization (Fig. 2G). In addition, repeat-bound ChRO1 sites were largely inaccessible and consistently enriched for H3K9me3 throughout myogenesis (Fig. 2H and Supplementary Fig. S4I). Together, these findings indicate that ChRO1 recognizes inaccessible, H3K9me3-enriched “repeat codes” to establish heterochromatin hubs at chromosomal ends.
ChRO1 augments the dynamic condensation of heterochromatin-associated proteins
Based on our findings that ChRO1 occupancy is enriched in repeat regions within a repressive environment, we examined whether ChRO1 can facilitate the condensation of heterochromatin-associated proteins, thereby orchestrating the reorganization of repeat-rich heterochromatin domains. To investigate this, we synthesized ChRO1a (~1.3 kb) in vitro and conducted an in vitro phase separation assay using GFP-labeled mouse HP1 proteins. Under the tested conditions, GFP–HP1 proteins readily formed condensates (see the “Materials and methods” section for details) (Fig. 3A). Remarkably, ChRO1a significantly increased the size of HP1 condensates, with a more pronounced effect on HP1α and HP1γ (Fig. 3A and Supplementary Fig. S5A). In contrast, ChRO1a had no impact on GFP–MED1–IDR, a domain crucial for the formation of transcriptionally active condensates, underscoring its specificity toward heterochromatin proteins (Supplementary Fig. S5B). ChRO1a RNA was significantly enriched within the condensates, indicating its direct involvement in enhancing the HP1 condensation (Supplementary Fig. S5C). ChRO1a’s specific action was further validated by comparing it to other nucleic acids, including antisense ChRO1a, nucleoplasmic RNAs, and their cDNAs, salmon sperm DNA, and yeast tRNAs. While these nucleic acids promoted condensation as reported [20], ChRO1a showed markedly stronger activity (Fig. 3B and Supplementary Fig. S5D). Additionally, ChRO1a was more efficient than major satellite repeat RNAs, pericentromeric transcripts implicated in HP1α-mediated heterochromatin compaction (Supplementary Fig. S5E) [60]. The effect of ChRO1a on HP1α condensate dynamics was further assessed using fluorescence recovery after photobleaching (FRAP). A higher recovery rate following photobleaching indicated that ChRO1a substantially increased HP1α dynamics and mobility (Fig. 3C and Supplementary Fig. S5F). This effect of ChRO1 is similar to the increased fluorescence recovery rate with higher HP1α concentration (Supplementary Fig. S5G), suggesting that ChRO1a enhances the local protein concentration within condensates by serving as a multivalent interaction platform.
ChRO1 augments dynamic condensation of heterochromatin-associated proteins and their co-partitioning. (A) Radius of droplets from in vitro phase separation assays using indicated HP1 proteins (HP1α, HP1β, and HP1γ) at various concentrations, in the presence or absence of in vitro transcribed ChRO1a. Droplet radii were quantified from three independent experiments per condition; the number of droplets ranges from 300 to 1000 depending on the phase separation capacity of each protein. Statistical significance was assessed using the Wilcoxon rank-sum test. (B) Radius of HP1α condensates in the presence of indicated nucleic acids (100 ng/μl for all except yeast tRNA, 200 ng/μl). n = 130 condensates from three biological replicates. (C) FRAP analysis of GFP–HP1α with or without ChRO1a. n > 5 for control without or with ChRO1, and n > 12 for bleached without or with ChRO1. (D) Diagram representing CU dinucleotide repeat-rich regions of ChRO1a and its fragments. (E) Radius of HP1α condensates in the presence of full-length or indicated fragments of ChRO1a. n = 120 condensates from three independent replicates. (F) Representative images of in vitro phase separation assays of mCherry-HP1α (12.5 μM) mixed with indicated proteins (12.5 μM for GFP–HP1 proteins, 3.125 μM for GFP–DAXX, 0.78 μM for GFP–MED1) with or without ChRO1a (100 ng/μl). (G) Scatterplot of mCherry and GFP signal intensity for the assays in (F). Condensates formed with or without ChRO1a are indicated by black and red dots, respectively. Pearson correlation coefficients of mCherry and GFP signals in the absence or presence of ChRO1a are shown. (H) Quantification of colocalization between indicated proteins and DAPI using Pearson’s correlation coefficients in MB with or without Dox-induced ChRO1a. n > 50 nuclei. Statistical analyses, box plot elements, and data normalization procedures are detailed in the “Materials and methods” section.
We next examined the potential role of ChRO1 as a multivalent interaction platform by analyzing its secondary structure [33]. Intriguingly, the low-complexity repeats in intron 1 are predicted to remain unpaired, forming a loop structure (Supplementary Fig. S5H), suggesting a role in mediating protein interactions (Supplementary Fig. S5I). To assess the importance of the CUR of ChRO1 in HP1 condensation, several ChRO1 segments were transcribed in vitro (Fig. 3D). Each fragment enhanced the in vitro condensation of mCherry–HP1α in proportion to its CU content, demonstrating that the CU-rich low-complexity regions of ChRO1 effectively promote HP1α condensation (Fig. 3E and Supplementary Fig. S5J). These findings align with the enhanced reorganization of heterochromatin foci marked by HP1α in C2C12 myoblast cells expressing the CUR of ChRO1a (1–413) (Fig. 1I). Overall, our findings suggest that ChRO1, particularly its low-complexity CUR, functions as a multivalent interaction platform that partitions HP1 proteins into chromocenters, thereby organizing complex heterochromatin condensates.
ChRO1 facilitates co-partitioning of heterochromatin-associated proteins
The three HP1 paralogs, HP1α, HP1β, and HP1γ, share a common overall domain structure but differ in their IDR hinge regions, resulting in varying propensities for condensate formation and DNA compaction [61–63]. Moreover, these proteins organize genomic regions differentially: HP1α and HP1β are primarily localized to genomic repeats for heterochromatin formation, whereas HP1γ is more associated with euchromatin and gene regulation [21, 64–66]. However, since all three HP1 proteins are present at chromocenters in myotubes, we speculated that ChRO1 may facilitate co-partitioning of HP1s, specifically concentrating HP1γ at chromocenters through dynamic condensation. To further investigate a potential mechanism for the colocalization of HP1 paralogs in overlapping genomic regions, we mixed mCherry-labeled HP1α with GFP-labeled HP1α, HP1β, or HP1γ in vitro with or without ChRO1a. As expected, HP1α condensates mixed fully within the range of their physiological concentrations (Fig. 3F) [67, 68]. Under identical conditions, HP1α and HP1β condensates largely intermingled, whereas HP1γ remained excluded from HP1α condensates, mimicking their localization in myoblast cells (Fig. 3F and G). However, ChRO1a significantly facilitated complete mixing of HP1 proteins within a condensate, recapitulating their colocalization in myotubes (Fig. 3F and G). Although high protein concentrations were required for droplet mixing (Supplementary Fig. S6A), ChRO1a substantially lowered this threshold, enabling efficient co-partitioning of HP1α and HP1γ at physiological concentrations. This mixing effect was more pronounced with ChRO1, compared to yeast tRNA (Supplementary Fig. S6B).
During myogenesis, DAXX relocates from small promyelocytic leukemia bodies to chromocenters [22]. As the subcellular localization of DAXX is often regulated by its IDR-mediated condensation [69], we performed an in vitro phase separation assay to examine how ChRO1a affects DAXX interactions with HP1α condensates. ChRO1a readily facilitated the co-partitioning of two proteins within the same droplets. In contrast, ChRO1a prevented the co-partitioning of HP1α and GFP–MED1–IDR, underscoring its selectivity for heterochromatin-associated components (Fig. 3F and G).
To investigate this further, we ectopically expressed ChRO1a in myoblasts and analyzed changes in the localization of heterochromatin-associated proteins. ChRO1a induction dramatically increased the translocation of HP1γ and DAXX to DAPI foci but did not affect the localization of HP1α, β, H3K9me3, and ATRX (Fig. 3H and Supplementary Fig. S6C). Similar results were observed with overexpression of ChRO1 CUR (1–413) (Supplementary Fig. S6D and E). These findings demonstrate that ChRO1 is essential for the myotube-specific partitioning of HP1γ and DAXX into chromocenters, consistent with previous observations that ChRO1 knockdown displaces DAXX, but not ATRX, from chromocenters [30]. Overall, our results indicate that ChRO1 regulates the permeability of heterochromatin compartments and controls the selective partitioning of chromocenter components.
ChRO1 establishes chromatin interaction hubs, securing heterochromatinization of non-muscle genes
Based on these findings, which demonstrate that ChRO1 facilitates heterochromatin remodeling via dynamic condensation of associated proteins, we next examined its effects on chromatin interactions and nearby gene regulation. Our Hi-C analysis [53] showed that chromatin interaction frequency and TAD scores were increased during myogenesis, preferentially at ChRO1-bound genomic regions compared to unbound regions (Fig. 4A and Supplementary Fig. S7A), supporting a role for ChRO1 in strengthening intra-TAD interactions and TAD borders. Moreover, PC1 correlation indicated increased compartmental reorganization of compartment A and sustained stability of compartment B at ChRO1-bound domains during myogenesis (Supplementary Fig. S7B). The ChRO1+satellite colocalization regions on chromosome 4 displayed enhanced chromatin interaction patterns accompanied by local changes in A/B compartmental features, indicative of ChRO1-bound regions as focal sites of chromatin reorganization (Fig. 4B). Additionally, interchromosomal interactions were markedly increased at ChRO1+satellite sites near pericentromeric regions, supporting their clustering and the formation of multichromosomal chromocenters (Fig. 4C and Supplementary Fig. S7C). These data suggest that ChRO1, by binding to satellite-enriched loci, contributes to the establishment of both intra- and interchromosomal interactions, important for chromocenter formation.
ChRO1 establishes chromatin interaction hubs, securing heterochromatinization of non-muscle genes. (A) Aggregated loop interaction scores of ChRO1-associated regions (±150 kb) in MB and MT. (B) Heatmap of chromatin interactions on chromosome 4 (Chr 4), showing MT (upper triangle) and MB (lower triangle). PC1 tracks indicate compartment polarity with compartment A (light blue) and compartment B (dark blue). Dots mark satellite repeats (dark red) and ChRO1 binding sites (orange). (C) Scatter plot of z-score differences (MT–MB) across chromosomal positions. Interactions at ChRO1 bound satellite repeats are in red. Differences in z-score was greater in ChRO1 bound satellite repeats than others (Wilcoxon rank-sum test, P-value = 1.5 × 10−8). (D–F) H3K9me3 HiChIP was performed WT and ChRO1 KO differentiated cells. Loops were defined as chromatin interactions containing at least one H3K9me3 peak at either anchor. (D) Aggregated loop interaction scores at ChRO1-bound loops present in WT differentiated cells but lost upon ChRO1 KO. Heatmaps display averaged interaction frequencies centered on loop anchors (±100 kb). (E) Loop span distribution of steady, lost, or gained H3K9me3 loops in KO cells. Distance was calculated as the difference between start coordinates of looped bins (Mb). Statistical significance between each pair of groups was assessed by Wilcoxon rank-sum test (P-value < 2.2 × 10−16). Blue dots indicate ChRO1-unbound loops, and red dots indicate ChRO1-bound loops in each group. (F) Heatmap of chromatin interactions from H3K9me3 HiChIP within Chr 4 [same region as in panel (B)], comparing WT (upper triangle) and KO (lower triangle). Dots mark satellite repeats (dark red) and ChRO1 binding sites (orange). Genomic tracks show H3K9me3 HiChIP read coverage in WT and KO cells (blue). (G) GO analysis of genes near ChRO1+satellite sites. (H) Heatmap of RNA-seq expression levels for genes located near ChRO1+satellite sites in MB and MT. Each column represents an independent biological replicate (n = 3 per condition). (I) Chromatin accessibility (ATAC-seq) ±500 bp around TSS of genes near ChRO1+satellite sites in MB and MT. (J) GO analysis of genes located within H3K9me3 loops lost in KO cells near ChRO1 binding sites. Statistical analyses, box plot elements, and data normalization procedures are detailed in the “Materials and methods” section. (K) Distance of genes in the indicated groups from ChRO1+satellite sites. Statistical significance was assessed by Wilcoxon rank-sum test: upregulated MT versus downregulated MT (P-value = .0026), upregulated MT versus nervous system development (GO:0007399) (P-value = .002). Data shown median distance (kb with quartiles).
Building on these genome-wide observations, we specifically investigated the requirement of ChRO1 for loop formation within heterochromatin domains. Given the high enrichment of ChRO1 at H3K9me3-marked heterochromatin (Fig. 1G and Supplementary Figs S3B, S2H, and S4I), and the observation that H3K9me3 levels remain unchanged in ChRO1 KO cells (Supplementary Fig. S7D), we performed H3K9me3 HiChIP in WT and ChRO1 KO differentiated C2C12 cells to obtain a direct and sensitive view of the heterochromatin-anchored interactome.
Of the merged 14 100 loops, defined as interactions with at least one anchor overlapping an H3K9me3-marked region, 2174 were differentially regulated (1188 lost, 986 gained). Notably, >74% of ChRO1-bound loops were lost in KO cells (Fisher’s exact test, P-value = 1.26 × 10^−5^; Fig. 4D and Supplementary Fig. S7E), particularly those with short-span interactions (Fig. 4E). This reduction was not limited to direct ChRO1 binding sites; ChRO1 KO was also associated with widespread weakening of unbound loops across H3K9me3-marked regions, particularly in the vicinity of ChRO1-bound lost loops (Supplementary Fig. S7F and G). This pattern suggests that ChRO1-mediated compaction may propagate locally to stabilize the broader heterochromatin environment and ensure structural coherence. In addition, ChRO1 KO cells exhibited higher insulation scores, indicative of weakened and disrupted local boundary strength throughout the H3K9me3 interactome (Supplementary Fig. S7H). Consistently, the chromosomal interactions of H3K9me3-marked domains were selectively weakened or lost in ChRO1 KO cells, whereas the H3K9me3 profile remained largely intact (Fig. 4F and Supplementary Fig. S7I). Together, these results suggest that ChRO1 plays a key role in organizing heterochromatin interaction hubs during differentiation, potentially by direct stabilization of ChRO1-bound loops and broader effects on the structural coherence of the surrounding H3K9me3-marked chromatin network.
Next, we asked how this heterochromatin organization influences the regulatory states of nearby genes. Remarkably, genes adjacent to ChRO1+satellite sites were enriched for developmental processes, including neurogenesis (Fig. 4G). Although the expression of these genes remained mostly unchanged during myogenesis, (Fig. 4H), their promoter accessibility shifted markedly to a more closed state, suggesting that ChRO1 reinforces a repressive environment for non-muscle lineage genes (Fig. 4I). Consistently, genes near loops lost in KO cells were enriched for neuronal functions, including regulators such as Nfix, Runx3, and Gprin1 (Fig. 4J). Moreover, genomic distance analysis revealed that myotube-specific downregulated genes were located significantly closer to ChRO1+satellite sites than upregulated genes (Fig. 4K). Notably, genes associated with nervous system development (GO:0007399) were also preferentially positioned close to ChRO1+satellite regions (Fig. 4K), suggesting that ChRO1-mediated heterochromatin formation occurs selectively within genomic neighborhoods containing non-muscle genes, whereas muscle-specific genes are largely insulated from these repressive compartments. Altogether, these results suggest that ChRO1 secures the suppression of alternative developmental genes by clustering satellite-marked genomic regions, enhancing both intra- and interchromosomal interactions, and ultimately establishing myotube-specific chromatin organization and gene expression.
ChRO1 deficiency and condensate disruption induce a muscle atrophic phenotype
Although biomolecular condensation is critical for diverse cellular processes [70, 71], its pathological implications for skeletal muscle health remain unexplored. To investigate the physiological importance of ChRO1-driven condensate reorganization into chromocenter, we examined the impact of ChRO1 deficiency on skeletal muscle tissue. Using CRISPR/Cas9, we generated ChRO1 KO mice by deleting an ∼1.6 kb genomic region encompassing the promoter, exon 1, and 5′ part of intron 1, including MyoD binding sites (Fig. 5A and Supplementary Fig. S8A). This deletion led to the loss of ChRO1 expression (Fig. 5B). We then assessed the effects of ChRO1 KO on muscle integrity. Remarkably, the gastrocnemius of 13-months-old male ChRO1 KO mice showed significant upregulation of atrophic markers, such as Atrogin1 and Murf1 (Fig. 5C). A notable reduction in muscle fiber cross-sectional area (CSA) was observed across various skeletal muscle types, consistently in different gender and age groups (Fig. 5D and Supplementary Fig. S8B), resembling the KO mice phenotype reported by Ballarino et al. using an alternate strategy [43]. To evaluate chromocenter integrity in ChRO1 KO mice, we performed DAPI staining on skeletal muscle tissues. Consistent with the results performed in ChRO1 KO C2C12 cells (Supplementary Fig. S3F), chromocenters were fragmented, and the average number of heterochromatin foci per nucleus increased compared to WT mice (Fig. 5E). Immunofluorescence analysis further revealed that DAXX was delocalized from chromocenters in ChRO1 KO mice, while H3K9me3 localization remained largely unchanged (Fig. 5F and Supplementary Fig. S8C). Overall, these findings suggest that ChRO1 deficiency partially disrupts chromocenters and alters the localization of heterochromatin proteins, potentially contributing to muscle atrophy in vivo.
ChRO1 deficiency and condensate disruption induce a muscle atrophic phenotype. (A) Strategy for generating ChRO1 KO (ChRO1−/−) mice using CRISPR–Cas9 genome editing targeting two gRNA sites to delete ChRO1 promoter, exon1, and part of intron1 region. (B) qRT-RCR quantification of ChRO1a and ChRO1b expression in gastrocnemius from 13-months-old-male mice WT and KO mice (ChRO1+/+ and ChRO1−/−). (C) qRT-PCR analysis of atrophic genes and satellite RNAs in gastrocnemius muscle from WT and ChRO1 KO mice. (D) Representative wheat germ agglutinin staining (left) and myofiber CSA (μm2) quantification (right) of indicated skeletal muscle tissues from WT and ChRO1 KO mice. n ≥ 500 fibers analyzed per tissue. Scale bars, 40 μm. (E) Representative images (left) and quantification (right) of heterochromatin foci in gastrocnemius muscle from WT and ChRO1 KO mice. Scale bars, 1 μm. n = 50 nuclei. (F) Immunostaining for DAXX and H3K9me3 in gastrocnemius muscle of WT and ChRO1 KO mice. n = 50 nuclei. Scale bars, 2 μm. (G) qRT-PCR analysis of Atrogin1 (left) and Murf1 (right) in C2C12 MT differentiated for 5 days followed by 1,6-HD treatment at indicated concentration (%) for varying durations (1.5, 5.5, or 24 h). Expression normalized to Gapdh and shown as fold change relative to untreated control for each time point. (H) Western blot analysis of MyHC protein levels in C2C12 MT treated with 1,6-HD at indicated concentration (%) for varying durations (1.5, 5.5, or 24 h). Immunofluorescence staining of MyHC (AF488, green) and nuclei (DAPI, blue) (I) and quantification of myotube diameters [(J), n = 100 cells] of C2C12 MT treated with 1,6-HD at indicated concentrations. (K) Quantification of myotube diameters of C2C12 MT treated with 2,5-HD or 1,6-HD at indicated concentration for 24 h. n = 45 cells. Statistical analyses, box plot elements, and data normalization procedures are detailed in the “Materials and methods” section.
To directly explore the impact of chromocenter condensate disruption, we differentiated C2C12 myoblasts into myotubes and treated them with 1,6-HD or 2,5-HD at varying concentrations (0.25%, 0.5%, and 1%) and durations (1.5, 5.5, and 24 h). Concentration- and time-dependent chromocenter scattering was observed without complete dissolution (Supplementary Fig. S8D), whereas 2,5-HD had minimal effects on chromocenter integrity under the same conditions (Supplementary Fig. S8E). Importantly, 1,6-HD treatment significantly induced the expression of atrophy marker genes, suggesting that disruption of chromocenter condensates contributes to muscle atrophy (Fig. 5G). Moreover, a concentration- and time-dependent reduction in myosin heavy chain (MyHC) proteins, a myotube marker, was observed (Fig. 5H). Measurements of myotube diameter after 1,6-HD treatment also showed a significant reduction in the median diameter (Fig. 5I and J). However, treatment with 2,5-HD did not exhibit any apparent effects on myotube diameter or morphology (Fig. 5K and Supplementary Fig. S8F). These results highlight the critical role of condensate organization and heterochromatin compartments in maintaining muscle integrity. Overall, our results provide evidence that impaired compartmentalization of PCH into chromocenters may undermine the maintenance of healthy myotubes and contribute to the development of the muscle atrophy phenotype.
Chromocenters are disintegrated in a skeletal muscle atrophy model
Conversely, to determine whether muscle atrophy affects chromocenter integrity, we induced muscle atrophy in C2C12 myotubes using dexamethasone (Dex), a synthetic glucocorticoid, or MK2206, an Akt inhibitor (Fig. 6A). Both agents are well-established inducers of muscle atrophy [72–74]. Treatment with Dex caused noticeable myotube degeneration (Fig. 6B), and increased mRNA expression of Atrogin1 and Murf1, while reducing the expression of myogenic genes MyHC and muscle creatine kinase (Fig. 6C). Under these atrophic conditions, chromocenters disintegrated into smaller, discrete DAPI foci of PCH, as evidenced by an increase in their number and a decrease in their size (Fig. 6D). Additionally, ChRO1 expression was reduced while major and minor satellite RNA levels were elevated (Fig. 6E and Supplementary Fig. S9A). We also examined the localization of chromocenter-associated proteins. Notably, HP1α, HP1β, and H3K9me3 remained localized to chromocenters, whereas DAXX was displaced from DAPI foci after Dex treatment. HP1γ also showed a modest but consistent reduction in colocalization with DAPI foci following Dex treatment (Fig. 6F and Supplementary Fig. S9B). Similar effects were observed in myotubes treated with MK2206, except for ATRX, whose localization varied depending on the specific atrophy-inducing agents (Supplementary Fig. S9C–G). Overall, atrophic stimulation reduced ChRO1 levels, leading to chromocenter disintegration and delocalization of associated proteins, similar to the effects observed with 1,6-HD treatment (Fig. 1). These findings suggest that muscle remodeling upon atrophic stimulation is partly due to impaired condensation of chromocenters and disrupted heterochromatin compartmentalization, potentially driven by ChRO1 downregulation.
Chromocenters are disintegrated in the skeletal muscle atrophy model. (A) Schematic diagram of experiment design. After 4 days of differentiation, MT was treated with 100 μM Dex for 48 h. Representative immunofluorescence images of MyHC (green) and DAPI (blue) staining (B) and relative mRNA expression levels (C) after 48 h of Dex treatment following 4 days of differentiation. (D) Representative image (left) and quantification (right) of heterochromatin foci per MT nucleus with or without Dex treatment after 4 days of differentiation. n = 80 nuclei. (E) Relative expression of ChRO1a measured by qRT-PCR following 48 h of Dex treatment after 4 days of differentiation. (F) Quantification of colocalization between DAPI and indicated proteins in MT with or without Dex treatment using Pearson’s correlation coefficients. n > 75 nuclei. (G) Fold change (post/pre) of RNA expression after 5 days of bedrest, with genes sorted by fold change of Atrogin1. Each column represents a different individual. (H) Pearson correlation coefficients and negative logarithm of P-values for fold changes (post/pre) of each gene compared with the fold change of ChRO1. Only genes with P-values < .1 are plotted. (I) KEGG pathway enrichment analysis of genes showing significant negative or positive correlation with ChRO1. Statistical analyses, box plot elements, and data normalization procedures are detailed in the “Materials and methods” section.
To explore the clinical relevance, we examined previously published RNA-seq data from human vastus lateralis muscle biopsies (n = 18, ages 60–79) collected before (pre) and after (post) a 5-day bedrest period [75]. Although the individual responses to acute bedrest varied, Atrogin1 and Murf1 RNA levels were generally elevated, whereas ChRO1 levels were decreased (Fig. 6G). To further investigate this, we identified genes whose changes correlated with alterations in ChRO1 levels by calculating correlation coefficients of fold changes. Genes negatively correlated with ChRO1 included Atrogin1 and Murf1, whereas those positively correlated with ChRO1 included the myogenic transcription factor MEF2D, JUND, and NFATs (Fig. 6H). We then performed KEGG pathway analysis on the gene sets correlated with ChRO1. Genes negatively correlated with ChRO1 were commonly involved in proteins or organelle homeostasis, including proteasome ubiquitin receptor (ADRM1), proteasome subunits (PSMAs, PSMBs, PSMCs, and PSMDs), ubiquitin-conjugating enzymes (UBE2A, UBE2D3, and UBE2W), and autophagy-related proteins (SQSTM1, Beclin1, ATGs, and CHMPs). In contrast, genes positively correlated with ChRO1 were highly enriched in pathways related to Notch and Wnt signaling (NOTCH, JAG2, GSK3B, FZD7), transcription activation (BRD4, MED12, and EP300), and ion transport (CACNA1A and ATP1A2) (Fig. 6I). Activation of these pathways supports that ChRO1 expression is positively correlated with satellite cell proliferation, muscle differentiation, and reinforcement of physiological function. Overall, our findings suggest that atrophic stimulation reduces ChRO1 levels, compromising chromocenter integrity and selective protein partitioning. Additionally, our data indicate that ChRO1 may play a protective role by preventing protein degradation and potentially mitigating muscle atrophy.
ChRO1a alleviates muscle atrophy by maintaining chromocenter clustering
Next, to determine whether ChRO1 overexpression has a protective effect against muscle atrophy induced by pharmacological stimulation, we differentiated C2C12 cells into myotubes for 4 days and induced atrophy by Dex or MK2206 for 48 h, with or without ChRO1a induction (+/−Dox) (Supplementary Fig. S10A). Remarkably, ChRO1a overexpression prevented chromocenter scattering, partially restoring the number of heterochromatin foci (Fig. 7A and Supplementary Fig. S10B and C). Furthermore, ChRO1 overexpression significantly suppressed the upregulation of Atrogin1 and Murf1 induced by Dex or MK2206 compared to control (Fig. 7B and Supplementary Fig. S10D). Immunostaining for MyHC and myotube diameter analysis confirmed that ChRO1a overexpression significantly alleviated the degenerative muscle phenotype caused by Dex or MK2206 (Fig. 7C and Supplementary Fig. S10E). Given that the CUR (1–413) region of ChRO1a is sufficient to promote condensation of chromocenter proteins, we tested whether CUR (1–413) could also protect the myotubes from atrophic stimulation. As expected, our results showed that CUR (1–413) overexpression mitigated Dex-induced atrophic phenotypes (Supplementary Fig. S10F–H). To further evaluate whether ChRO1a’s protective effects were linked to its condensation-promoting activity, ChRO1a was induced on day 4 of differentiation for 48 h, followed by 5h treatment with 1% 1,6-HD (Supplementary Fig. S10I). In the presence of 1,6-HD, ChRO1a failed to prevent chromocenter scattering (Fig. 7D and Supplementary Fig. S10J) and was unable to suppress Atrogin1 and Murf1 expression (Fig. 7E). Moreover, ChRO1a was ineffective in reducing myotube thinning in the presence of 1,6-HD (Fig. 7F). Overall, these findings support a causal link between ChRO1-driven condensation of chromocenters and muscle integrity. By marking genomic repeat elements, CU-rich ChRO1 clusters heterochromatin into chromocenter condensates, thereby suppressing the expression of atrophic genes, and protecting muscle from atrophic stimuli.
ChRO1a alleviates muscle atrophy by maintaining chromocenter clustering. Number of heterochromatin foci per MT nucleus following Dex [(A), n = 90 nuclei] or 1,6-HD [(D), n = 70 nuclei] treatment, with or without Dox-induced ChRO1a expression. Relative mRNA expression levels after Dex (B) or 1,6-HD (E) treatment, with or without Dox-induced ChRO1a. Representative immunofluorescence images for MyHC (green) and DAPI (blue) staining (left), and quantification of MT diameters (right) following Dex (C) or 1,6-HD (F) treatment, with or without Dox-induced ChRO1a. (G) Associations of CU-rich ChRO1 with similar low-complexity genomic repeats confer selectivity as genome markers, enabling cell-type-specific chromatin interactions. Repetitive genome elements are represented by boxes in the upper panel; black for PCH enriched with major satellite repeats, red for low-complexity repeats bound by ChRO1, and white for low-complexity repeats absent of ChRO1. ChRO1 induces heterochromatin condensation, and its stably associated proteins into dynamic chromocenters. The distinct structural environment within chomocenters facilitates the co-partitioning of heterochromatin-associated proteins, establishing chromatin-regulatory network, and reinforcing heterochromatinization of non-muscle compartments. Chromocenter integrity is critical for muscle maintenance, protecting against atrophic stimuli and stress. Statistical analyses, box plot elements, and data normalization procedures are detailed in the “Materials and methods” section.
Discussion
Cell-type-specific heterochromatin reorganization is essential for regulating gene expression, chromatin organization, and nuclear structure. However, the mechanisms underlying its temporal and spatial specificity remain unclear. Here, our study demonstrates that repeat-rich RNA clusters DNA regions enriched with similar low-complexity repeats, thereby promoting chromocenter condensation and the selective co-partitioning of heterochromatin-associated proteins to establish chromocenters distinct from PCH. This clustering enhances chromatin interactions, promoting repressive compartmental states and heterochromatinization of adjacent genes. Importantly, disruption of chromocenters by ChRO1 depletion or 1,6-HD induced a muscle atrophic phenotype. Conversely, chemically induced muscle atrophy results in chromocenter disintegration and muscle thinning, which can be protected by ChRO1a overexpression. These findings deepen our understanding of heterochromatin condensation into dynamic chromocenters and its pathological consequences in muscle health and disease.
Despite HP1α’s intrinsic ability to undergo LLPS, whether HP1α-associated heterochromatin represents bona fide LLPS-like condensates has remained contentious. Our findings indicate that PCH foci in myoblasts are resistant to 1,6-HD and exhibit solid-like properties, suggesting that their structure, at this stage, is maintained by more static interactions, such as HP1α-mediated nucleosome bridging or the formation of collapsed globule states [12, 24, 56, 76, 77] (Fig. 7G). In contrast, during myogenesis, heterochromatin reorganizes into chromocenters with distinct physicochemical properties indicative of a more dynamic and reversible architecture, facilitated by ChRO1. These findings reveal that HP1α-associated heterochromatin can adopt distinct chromatin states in an RNA-dependent, cell-context-specific manner.
Our study highlights the pivotal role of chromocenters in shaping and maintaining cell-type-specific genome organization. This is primarily achieved through the compartmentalization of inactive chromatin regions marked by H3K9me3 (Fig. 2). By pooling inactive chromatin, chromocenters effectively sequester these regions from chromatin opening, DNA rearrangement, and active transcription of muscle genes, thereby preventing inappropriate activation of non-muscle genes (Fig. 4). Additionally, chromocenters provide specialized nuclear environments that facilitate precise cellular regulation [78]. Considering that chromocenter condensates are built upon PCH, their formation relies on both specific (solid-like) and transient (liquid-like) interactions among macromolecular components (Fig. 1). This architectural duality enables chromocenters to accommodate multiple molecular components with disparate physicochemical properties [30, 31, 66, 70]. In this context, intron 1 of ChRO1, enriched in CU-repeats, serves as a multivalent interaction scaffold that recruits diverse biomolecules with distinct functions [60, 79–83].
Although not strictly required for condensation, the preferential association of ChRO1 with CA- and CT-rich repetitive regions, possibly via hybridization with GA-rich strands of repeat DNA, may provide a spatial cue that guides heterochromatin condensation. Furthermore, multivalent contacts involving RNA and repeat-binding heterochromatin proteins, such as HP1γ and DAXX, can promote chromocenter assembly and stabilization [84, 85]. Their incorporation, along with the co-partitioning of MeCP2, SUV420h2, KDM4A, cohesin, and TET1, may efficiently establish cooperative regulatory networks linking DNA, histone modifications, and chromatin–chromatin interactions [86–90]. Consequently, diverse chromatin-regulatory complexes become highly concentrated within chromocenters, enhancing local interaction networks and heterochromatin propagation, and thereby contributing to chromocenter stabilization downstream of ChRO1-mediated initiation. Moreover, chromocenters spatially restrict heterochromatin-associated factors from euchromatin, thereby enabling efficient transcription of muscle genes within a permissive environment.
DNA repetitive elements have accumulated throughout genomic evolution, now constituting over 50% of the human genome. Once regarded as “junk DNA,” these elements are increasingly recognized for their crucial roles in chromatin organization, nuclear structure, and integrity. Their transcripts mediate condensation of chromatin-associated proteins through RNA:DNA triplex formation [91, 92]. A key function of repetitive elements is chromatin compartmentalization. For instance, A and B compartments are established by homotypic interactions of SINEs and LINEs, respectively [59]. DNA repeat families also serve as “genome markers,” barcoding genes with unique functions, such as development [93]. While repetitive elements alone do not inherently define cell-type-specific genes, our findings reveal that low-complexity DNA repeats with ChRO1 occupancy selectively barcode non-muscle genes, designating specific compartments for heterochromatinization.
Chromatin compartments are further stabilized by their attachment to subnuclear structures such as nucleoli and nuclear speckles [94]. In light of this, we propose that ChRO1-satellite-marked chromatin domains may gain stability through tethering to chromocenters, reorganizing into pericentromeric-associated domains [90]. This stability may be further supported by basal expression of major satellite RNAs and heterochromatin proteins, which can partially compensate for one another in heterochromatin organization. Together, these mechanisms help preserve the integrity of peripheral heterochromatin and chromocenters, which likely contribute to maintaining nuclear architecture and providing resistance to external stimuli, including atrophy and mechanostress [19].
Skeletal muscle atrophy is often accompanied by physical immobilization, cancer, aging, and muscular diseases. Patients with tumors or sarcopenia are exposed to a greater risk of injuries and death compared to those without muscle reduction [95, 96]. However, there are no approved drugs for the treatment of skeletal muscle atrophy, except for therapies generally targeting systemic inflammation, energy intake, and physical activity [97]. Therefore, elucidating the molecular mechanisms underlying skeletal muscle atrophy is particularly important. Our results provide evidence that the integrity of chromocenter condensates is closely associated with muscle health. Indeed, the relation of chromocenter-enriched proteins to muscle diseases has been reported by several groups [98–102]. For example, the loss of HP1γ and cohesin at D4Z4 repeats is related to facioscapulohumeral muscular dystrophy [98]. The connection between heterochromatin-associated factors and muscle integrity is further supported by cohesinopathy and ATRX syndrome, both characterized by intellectual disability and muscle weakness, caused by the loss of cohesin or ATRX, respectively [101, 103]. In addition, DAXX is downregulated in sarcopenia, an aging-related disease characterized by a decrease in muscle mass and quality [102]. Loss of Suv420h, an H4K20 methyltransferase, or its mislocalization, is coupled to defective muscle differentiation capacity [99, 100]. These findings suggest an interdependence between chromocenter condensate organization and muscle integrity. Maintenance of chromocenter structure appears both critical for and responsive to the physiological status of muscle cells.
In summary, our study uncovers the critical roles of RNA and DNA with similar low-complexity repeat content in orchestrating chromatin organization and driving condensation-mediated compartmentalization. We demonstrate that ChRO1 not only facilitates chromocenter condensate formation but also ensures heterochromatinization of non-muscle compartments. These findings provide a molecular framework for understanding how repetitive DNA elements achieve cell-type specificity by establishing distinct chromatin compartments and maintaining nuclear integrity. Furthermore, the tight coupling between chromocenter integrity and muscle health underscores the therapeutic potential of targeting dynamic heterochromatin condensation for skeletal muscle atrophy and related disorders.
Supplementary Material
gkag168_Supplemental_Files
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