Abstract

   Skeletal muscle plays a crucial role in maintaining motor function and
   metabolic homeostasis, with its loss or atrophy leading to significant
   health consequences. Long non‐coding RNAs (lncRNAs) have emerged as key
   regulators in muscle biology; however, their precise roles in muscle
   function and pathology remain to be fully elucidated. This study
   demonstrates that lncRNA maternally expressed gene 3 (MEG3) is
   preferentially expressed in slow‐twitch muscle fibers and dynamically
   regulated during muscle development, aging, and in the context of
   Duchenne muscular dystrophy (DMD). Using both loss‐ and
   gain‐of‐function mice models, this study shows that lncRNA‐MEG3 is
   critical for preserving muscle mass and function. Its depletion leads
   to muscle atrophy, mitochondrial dysfunction, and impaired regenerative
   capacity, while overexpression enhances muscle mass, increases
   oxidative muscle fiber content, and improves endurance. Notably,
   lncRNA‐MEG3 overexpression in MDX mice significantly alleviates muscle
   wasting and adipose tissue infiltration. Mechanistically, this study
   uncovers a novel interaction between lncRNA‐MEG3 and the polycomb
   repressive complex 2 (PRC2), where lncRNA‐MEG3 binds to SUZ12 polycomb
   repressive complex 2 subunit (Suz12), stabilizes PRC2, facilitates
   SUZ12 liquid–liquid phase separation (LLPS), and regulates the
   epigenetic modulation of four and a half lim domains 3 (Fhl3) and ring
   finger protein 128 (Rnf128). These findings not only highlight the
   crucial role of lncRNA‐MEG3 in muscle homeostasis but also provide new
   insights into lncRNA‐based therapeutic strategies for muscle‐related
   diseases.

   Keywords: fat infiltration, lncRNA‐MEG3, muscle atrophy, oxidative
   muscle fibers, SUZ12 LLPS
     __________________________________________________________________

   This study has identified lncRNA‐MEG3 as a key regulator of muscle
   mass, promoting slow‐twitch muscle fibers and preventing muscle
   atrophy. By stabilizing the SUZ12/PRC2 complex through liquid–liquid
   phase separation (LLPS), lncRNA‐MEG3 influences mitochondrial function
   and lipid metabolism. These findings highlight its potential as a
   therapeutic target for muscle‐wasting diseases like Duchenne muscular
   dystrophy (DMD).

   graphic file with name ADVS-12-2417715-g005.jpg

1. Introduction

   Skeletal muscle is a crucial organ for maintaining vital physiological
   functions, including movement, posture maintenance, and energy
   metabolism.^[ [54]^1 ^] As a primary metabolic organ, skeletal muscle
   supports systemic metabolic balance by regulating energy homeostasis.
   Muscle loss and atrophy precipitate functional impairment, degenerative
   remodeling, and the pathogenesis of muscle‐related disorders. Fat
   infiltration and muscle fiber type transition, such as the shift from
   type I to type IIb fibers, are hallmark features of muscle
   degeneration.^[ [55]^2 ^] These changes not only impair muscle function
   but also exacerbate metabolic disorders, contributing to the
   development and progression of related diseases.^[ [56]^3 ^]
   Additionally, mitochondrial dysfunction, closely linked to energy
   metabolism and oxidative stress, serves as a key driver of muscle
   atrophy.^[ [57]^4 ^] Therefore, a deeper understanding of the factors
   regulating skeletal muscle homeostasis and muscle atrophy, along with
   their molecular mechanisms, is of critical importance for preserving
   muscle mass and for the early diagnosis and treatment of muscle
   diseases.

   LncRNA, defined as transcripts longer than 200 nucleotides with no
   protein‐coding potential, have emerged as critical regulators of
   various biological processes, including myogenesis and muscle‐related
   diseases.^[ [58]^5 , [59]^6 ^] For instance, linc‐muscle
   differentiation 1 and lnc‐mitochondrial genome are highly expressed in
   skeletal muscle and regulate muscle cell differentiation, with their
   altered expression being linked to muscle atrophy and hypertrophy.^[
   [60]^7 , [61]^8 ^] LncRNA exerts their regulatory functions through
   various mechanisms. Unlike mRNAs, certain lncRNAs exert their
   regulatory functions by interacting with multiple proteins to form
   intricate regulatory networks that modulate gene expression.^[ [62]^9 ,
   [63]^10 ^] For example, X inactive specific transcript binds to spen
   family transcriptional repressor through its repetitive ring structure
   to silence gene expression.^[ [64]^11 ^] Additionally, some lncRNA
   serve as scaffolds for membrane less subcellular structures or nuclear
   bodies (NBs).^[ [65]^9 ^] These specialized structures often contain
   proteins with low‐complexity domains or intrinsically disordered region
   (IDR), which drive droplet formation via LLPS. Notably, lncRNA can
   accelerate LLPS in vitro by capturing IDR‐containing proteins or
   altering their conformations.^[ [66]^12 ^] However, there have been no
   reports on lncRNA regulating skeletal muscle size, growth, metabolism,
   and diseases through LLPS.

   LncRNA‐MEG3, encoded by the imprinted Dlk1‐Dio3 locus, is widely
   recognized for its tumor‐suppressive functions and is frequently
   downregulated in diverse malignancies, including breast, liver,
   colorectal, cervical, gastric, and ovarian cancers.^[ [67]^13 ^] Beyond
   its roles in oncogenesis, emerging studies have highlighted its
   biological significance in various tissues, including skeletal muscle,
   as well as its involvement in developmental processes.^[ [68]^14 ,
   [69]^15 , [70]^16 ^] Nearly a decade ago, initial investigations
   identified lncRNA‐MEG3 as highly expressed in adult skeletal muscle,
   with functions predominantly associated with the regulation of muscle
   cell proliferation, differentiation, and regeneration.^[ [71]^17 ^] Our
   previous work demonstrated that lncRNA‐MEG3 is conserved across
   species, including mice, pigs, and humans, serves as a marker for
   porcine meat production traits, and regulates myogenesis through the
   miR‐133a‐3p/proline rich transmembrane protein 2 axis.^[ [72]^18 ,
   [73]^19 ^] Furthermore, recent findings revealed that lncRNA‐MEG3
   knockdown in obese mice significantly reduced Akt phosphorylation
   levels in skeletal muscle, leading to impaired insulin sensitivity.^[
   [74]^20 , [75]^21 ^] These findings suggest that lncRNA‐MEG3 plays
   pivotal roles in muscle physiology, extending beyond myogenesis to
   encompass metabolic programming and potential involvement in
   muscle‐related diseases. However, its broader regulatory functions and
   underlying mechanisms remain largely unexplored, warranting further
   investigation.

   In this study, we demonstrated that lncRNA‐MEG3 is crucial for
   regulating muscle size and mass. Using loss‐of‐function mice models in
   skeletal muscle, we showed that lncRNA‐MEG3 depletion led to muscle
   atrophy, increased fat deposition in muscle tissue, impaired skeletal
   muscle regeneration, and reduced mitochondrial ATP production.
   Transgenic overexpression of lncRNA‐MEG3 in skeletal muscle promoted
   the conversion of muscle fibers to an oxidative phenotype, enhanced
   muscle mass and strength, and served as a novel regulator of skeletal
   muscle fat infiltration. By integrating RNA sequencing (RNA‐seq),
   chromatin immunoprecipitation sequencing (CUT&tag), and in vitro/in
   vivo LLPS experiments, we revealed that lncRNA‐MEG3 directly interacts
   with SUZ12 to regulate its LLPS, maintaining histone h3 lysine 27
   tri‐methylation (H3K27me3) modifications of key muscle‐specific genes,
   such as Fhl3 and Rnf128. These findings not only highlight lncRNA‐MEG3
   as a key regulator of muscle size, quality, and metabolic remodeling
   but also suggest its potential as a therapeutic target for muscle
   atrophy and fat infiltration‐related diseases.

2. Results

2.1. Temporal and Tissue‐Specific Expression of LncRNA‐MEG3 in Skeletal
Muscle Development and Disease

   To elucidate the role of lncRNA‐MEG3 in skeletal muscle, we first
   analyzed the publicly available single‐cell RNA sequencing dataset
   ([76]GSE147457),^[ [77]^22 ^] ([78]GSM4732632)^[ [79]^23 ^] and Human
   Muscle Ageing Cell Atlas database
   ([80]https://db.cngb.org/cdcp/hlma/rnaseq/).^[ [81]^24 ^] This dataset
   includes data from human skeletal muscle development, aging, and the
   mouse DMD model (MDX). Our analysis revealed that lncRNA‐MEG3
   expression gradually increased during embryonic development and
   juvenile stages, but declined with aging from puberty to old age in
   muscle stem cells (MuSCs) (Figure [82]1A,B; Figure [83]S1A,B and Table
   [84]S1, Supporting Information). In mice muscle at various
   developmental stages (embryonic day 15, birth, 5 weeks, 10 months, and
   18 months), we observed a similar trend with lncRNA‐MEG3 expression
   first rising and then decreasing over time (Figure [85]1C).
   Additionally, we found that lncRNA‐MEG3 expression in MuSCs from the
   MDX mouse model was lower, compared with wild‐type (WT) controls
   (Figure [86]1D; Figure [87]S2A, Supporting Information). Importantly,
   lncRNA‐MEG3 was highly expressed in slow‐twitch muscle fibers
   (Figure [88]1E). In human tissues from the Genotype‐Tissue Expression
   (GTEx) project, we identified a potential enhancer within the
   lncRNA‐MEG3 locus in muscle tissue (Figure [89]1F). Collectively, these
   findings establish lncRNA‐MEG3 as a highly enriched non coding RNA
   regulator, with potentially significant roles in skeletal muscle
   identity and disease.

Figure 1.

   Figure 1
   [90]Open in a new tab

   LncRNA‐MEG3 controls the development of skeletal muscle mass and
   regeneration. A) Violin plots depicting the expression levels of
   lncRNA‐MEG3 across the MuSCs cluster in human skeletal muscle at
   different developmental stages (embryonic, fetal, juvenile, and adult)
   (n = 3). B) Violin plots showing the expression levels of lncRNA‐MEG3
   in the differentiation cluster of MuSCs from young and aged mice (n =
   3). C) qRT‐PCR analysis of lncRNA‐MEG3 expression at different skeletal
   muscle developmental stages in mice (n = 6). (A, B, C, D, and E
   indicate a highly significant difference). D) Heat map illustrating
   differentially expressed genes in skeletal muscle tissue from wild‐type
   (WT) and MDX mice. E) qRT‐PCR showing lncRNA‐MEG3 expression across
   various tissues (Quadriceps femoris (QUA), Gastrocnemius (GAS), Soleus
   (SOL), Extensor digitorum longus (EDL), Brain, Stomach, Heart, Brown
   adipose tissue (BAT), White adipose tissue (WAT), Ovary, Kidney, Lung
   and Liver) from 6‐week‐old mice (n = 3). F) GTEx database analysis
   shows H3K27ac marks in muscle tissue along the lncRNA‐MEG3 locus,
   indicating the presence of an enhancer. G) Schematic representation of
   knockdown strategy using AAV9‐NC or AAV9‐lncRNA‐MEG3 shRNA (AAV9‐KD)
   injections into neonatal WT mice. Experimental timeline created in
   Adobe Illustrator. H) qRT‐PCR analysis of lncRNA‐MEG3 expression in
   muscle tissue from AAV9‐NC or AAV9‐KD injected mice (n = 6). I) QUA
   muscle weight in 8 weeks post‐AAV9‐NC (n = 5) or AAV9‐KD (n = 6)
   injections mice. J) QUA myofiber size in 8 weeks post‐AAV9‐NC (n = 4)
   or AAV9‐KD (n = 5) injections mice. Scale bar = 100 µm. K) Schematic
   representation of the experimental design for AAV9 injections and
   analysis timeline. Adult mice received AAV9 knockdown lncRNA‐MEG3
   (AAV9‐KD) or a control vector (AAV9‐NC) followed by cardiotoxin
   (CTX)‐induced injury on mice TA muscle. Analysis was performed at
   various time points post‐injury (3.5, 5.5, 10‐, and 25‐days post‐injury
   (dpi)). L) Representative images of injured and uninjured TA muscles
   from AAV9‐NC and AAV9‐KD mice (n = 6). Scale bar = 0.5 cm.
   Representative H&E staining M) and immunofluorescence (IF) staining N)
   of TA muscle cross‐sections (n = 3). Scale bar = 50 µm. O)
   Quantification of eMyHC^+ myofibers in TA muscle cross‐sections at 5.5
   dpi (n = 6). P) Quantification of laminin^+ myofibers in TA muscle
   cross‐sections at 10 dpi (n = 6). Q) Distribution of fiber sizes at 10
   dpi from AAV9‐NC and AAV9‐KD mice TA muscle after CTX injury (n = 4).
   R) qRT‐PCR showing the lncRNA‐MEG3, Myhc and Myog expression at 10 dpi
   after knockdown lncRNA‐MEG3 in TA muscle (n = 6). Data are mean ± SD;
   p‐values were calculated using Student's t‐test. ^* p < 0.05, ^** p
   < 0.01 and ^*** p < 0.001.

2.2. LncRNA‐MEG3 is Essential for Muscle Mass Maintenance and Regeneration

   To investigate the functional role of lncRNA‐MEG3, we examined its
   influence on skeletal muscle growth during postnatal development by
   extensively inhibiting lncRNA‐MEG3 expression. This was achieved
   through the intraperitoneal injection of adenoviral‐associated viral
   (AAV) vectors carrying shRNA targeting lncRNA‐MEG3 (AAV9‐KD) or a
   control vector (AAV9‐NC) into postnatal day 3 mice (Figure [91]1G).
   After 8 weeks of AAV delivery, we confirmed efficient knockdown of
   lncRNA‐MEG3 in skeletal muscles (Figure [92]1H; Figure [93]S3A,B,
   Supporting Information). Notably, the reduction of lncRNA‐MEG3 levels
   resulted in a significant decrease in muscle weight (Figure [94]1I;
   Figure [95]S3C,D, Supporting Information), with further validation of
   atrophic changes at the myofiber level (Figure [96]1J). While muscle
   atrophy is known to exhibit fiber‐type‐specific responses, systemic
   delivery of AAV‐shRNA targeting lncRNA‐MEG3 led to a decrease in the
   cross‐sectional area (CSA) of muscle fibers across all tested muscle
   groups, suggesting that lncRNA‐MEG3 knockdown has a broad, detrimental
   effect on muscle mass regulation (Figure [97]S3E,F, Supporting
   Information). Additionally, lncRNA‐MEG3 knockdown upregulated the
   expression of well‐established muscle atrophy markers, Atrogin‐1 and
   Murf1, at both the mRNA and protein levels (Figure [98]S3G,H,
   Supporting Information). Next, we explored whether lncRNA‐MEG3 could
   induce spontaneous muscle mass reduction in the absence of external
   atrophic stimuli. For this, AAV9‐NC and AAV9‐KD viruses were injected
   into the tibialis anterior (TA) muscle of 5‐month‐old mice (Figure
   [99]S3I, Supporting Information). After 8 weeks of AAV‐mediated
   delivery, efficient knockdown of lncRNA‐MEG3 was confirmed in the TA
   muscle (Figure [100]S3J, Supporting Information). Lower expression of
   lncRNA‐MEG3 in these muscles was associated with a trend toward reduced
   TA muscle weight (Figure [101]S3K, Supporting Information) and a
   significant decrease in muscle fiber size (Figure [102]S3L,M,
   Supporting Information). Furthermore, we investigated the impact of
   lncRNA‐MEG3 knockdown on muscle mass using a mice model of cardiotoxin
   (CTX) induced muscle injury (Figure [103]1K). At 5.5 days post‐injury
   (dpi), lncRNA‐MEG3 knockdown significantly impaired muscle recovery
   compared to WT controls (Figure [104]1L). Immunohistochemical analysis
   further confirmed a substantial accumulation of inflammatory myofibers
   in the muscles of lncRNA‐MEG3 knockdown mice (Figure [105]S3N,O,
   Supporting Information). Notably, AAV9 injection for 2 weeks did not
   significantly affect muscle size or weight, ensuring that the observed
   effects were specifically attributed to lncRNA‐MEG3 depletion (Figure
   [106]S3P, Supporting Information). Histological analysis using
   hematoxylin and eosin (H&E) staining further revealed a smaller
   regenerative area in lncRNA‐MEG3 knockdown muscles at 3.5, 5.5‐, and
   10‐ dpi, with impaired muscle repair (Figure [107]1M). IF staining
   further confirmed that lncRNA‐MEG3 knockdown compromised muscle
   regeneration, as evidenced by a decrease in the number of newly
   regenerated (eMyHC^+) and mature (laminin^+) muscle fibers at both
   early (5.5 dpi) and late (10 dpi) stages of recovery
   (Figure [108]1N–P). At 25 dpi, the distribution of the CSA in muscle
   fibers in the lncRNA‐MEG3 knockdown group significantly decreased,
   accompanied by a reduction in the expression of differentiation marker
   genes (Myog and Myhc) (Figure [109]1Q,R). Together, these findings
   establish that lncRNA‐MEG3 is a crucial regulator of muscle mass.

2.3. LncRNA‐MEG3 Drives Skeletal Muscle Mass and Function Through Modulation
of Muscle Fiber Composition and Lipid Deposition

   To assess whether lncRNA‐MEG3 is sufficient to drive muscle mass, we
   utilized the Cre‐LoxP system for the in vivo overexpression of
   lncRNA‐MEG3 (Figure [110]2A). While there was no significant impact on
   body or heart weight (Figure [111]2B,C), chronic overexpression of
   lncRNA‐MEG3 resulted in an increase in the mass of all skeletal muscles
   analyzed in 8‐month‐old mice (Figure [112]2D,E; Figure [113]S3Q,
   Supporting Information). Quantification of muscle fibers CSA in the TA,
   GAS, and QUA confirmed the observed increase in muscle mass
   (Figure [114]2F). Moreover, we found that compared with WT mice, the
   mass of both white and brown adipose tissue was reduced in lncRNA‐MEG3
   transgenic (TG) mice (Figure [115]2G,H). The muscle phenotype driven by
   lncRNA‐MEG3 was progressive, as a mild phenotype was observed in
   6‐month‐old mice (Figure [116]S4A–H, Supporting Information), with no
   significant atrophy detected following 5 weeks of overexpression
   (Figure [117]S4I–P, Supporting Information). Notably, in aging mice,
   overexpression of lncRNA‐MEG3 not only increased skeletal muscle mass
   but also led to an increase in overall body weight (Figure [118]S4Q–W,
   Supporting Information). However, the effect on adipose tissue was no
   longer evident (Figure [119]S4X, Supporting Information). We
   hypothesize that in younger mice, lncRNA‐MEG3 may influence systemic
   metabolism, while the age‐related decline in metabolic capacity may
   account for the diminished impact on fat deposition in older animals.
   So, we further investigated the impact of lncRNA‐MEG3 on different
   muscle fiber types and muscle lipid deposition. Upon visual inspection,
   each of the different types of muscles from the lncRNA‐MEG3 mice was
   redder in color compared with WT mice (Figure [120]2I). To assess
   muscle fiber types, we performed immunofluorescent staining for myosin
   heavy chain (Myhc) isoforms in the SOL muscles. The results showed that
   the overexpression of lncRNA‐MEG3 led to an increased density of
   oxidative fibers and a reduced number of glycolytic fibers, in contrast
   to that observed in the WT mice (Figure [121]2J). When muscle sections
   were immune‐stained for succinate dehydrogenase (SDH), a marker of
   mitochondrial activity, less than 38% of fibers in the muscle of WT
   mice were SDH‐positive, whereas over 60% of myofibers in lncRNA‐MEG3
   mice exhibited SDH positivity (Figure [122]2K). Similar changes were
   observed in the GAS and TA muscles (Figure [123]S5A,B, Supporting
   Information). Slow fiber‐specific genes were significantly upregulated
   in the TG mice at both mRNA and protein levels, while fast
   fiber‐specific genes were downregulated (Figure [124]2L–N). Oil Red O
   (ORO) staining of the GAS muscle cross‐sections indicated a trend
   toward reduced lipid droplet numbers (Figure [125]2O). Additionally,
   the expression of adipogenesis marker genes (Adipoq and Fabp4) was
   reduced at both the mRNA and protein levels in lncRNA‐MEG3 TG mice
   (Figure [126]2P,Q). Importantly, lncRNA‐MEG3 TG mice exhibited
   accelerated muscle regeneration and an increased proportion of slow
   muscle fibers following CTX‐induced damage to the TA muscle (Figure
   [127]S6A–O, Supporting Information). Additionally, lncRNA‐MEG3 was
   positively correlated with Myh7 (R = 0.52, P = 0.00071), but had no
   correlation with Myh4 (R = 0.17, P = 0.53) during human skeletal muscle
   development (Figure [128]S6P,Q, Supporting Information). Further
   analysis of primary myoblasts isolated from lncRNA‐MEG3 TG mice
   revealed that lncRNA‐MEG3 overexpression resulted in significantly
   increased type I fibers, enlarged myotube sizes, and reduced
   adipogenesis in myotubes (Figure [129]2R–T). To determine the
   functional consequences of the observed muscle atrophy after 8 months
   of lncRNA‐MEG3 deficiency, we performed grip strength measurements,
   which demonstrated that lncRNA‐MEG3 overexpression enhanced muscle
   strength (Figure [130]2U). Additionally, lncRNA‐MEG3 TG mice showed
   improved endurance in treadmill tests, as evidenced by increased
   running time and running distance (Figure [131]2V,W). Furthermore, we
   observed an increase in muscle contraction force in lncRNA‐MEG3 TG
   mice, compared with the WT control (Figure [132]2X). In summary, these
   data suggest that lncRNA‐MEG3 is essential for promoting the
   accumulation of slow muscle fibers, maintaining muscle mass, and
   preserving muscle function over time.

Figure 2.

   Figure 2
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   LncRNA‐MEG3 TG mice maintain muscle mass and promote an increase in
   slow‐twitch muscle fibers. A) Schematic illustrating the experimental
   design for TG mice with lncRNA‐MEG3 overexpression, generated through
   Cre‐LoxP‐mediated homologous recombination at the Rosa26 locus to
   induce lncRNA‐MEG3 expression. B) Body weight, C) Heart weight, D) GAS
   weight, TA weight, EDL weight, and E) SOL weight in wild‐type (WT) and
   lncRNA‐MEG3 TG mice (n = 7). All weights were normalized to tibia
   length (TL) to account for differences in body size WT and TG. F) Fiber
   size of GAS, TA, and QUA muscles in WT and lncRNA‐MEG3 TG mice (n = 4).
   G,H) WAT (G) and BAT weight (H) in WT and lncRNA‐MEG3 TG mice (n = 5).
   I) Representative images of skeletal muscle (SOL, GAS, EDL, QUA, and
   TA) from 6‐month‐old WT and TG mice (n = 3). Scale bar = 0.5 cm. J)
   Representative Myh7, Myh4, and Myh2 immunostaining in SOL muscles from
   WT and lncRNA‐MEG3 TG mice, with quantification of staining intensity
   using ImageJ (n = 3). Quantification is shown (right panel). Scale bar
   = 200 µm. K) Representative SDH staining of SOL muscle sections from WT
   and lncRNA‐MEG3 TG mice, with quantification of SDH‐positive fibers
   using ImageJ (n = 4). Scale bar = 200 µm. L–N) qRT‐PCR (n = 6) and
   Western blot (n = 3) analyses showing the expression of oxidative and
   glycolytic genes relative to marker genes in GAS, TA, and SOL muscles
   from WT and lncRNA‐MEG3 TG mice. O) Representative Oil Red O (ORO)
   staining of lipid droplets in GAS muscle tissue from WT and lncRNA‐MEG3
   TG mice (n = 6). Scale bar = 50 µm. P,Q) qRT‐PCR (n = 6) and Western
   blot (n = 3) analysis of adipogenesis‐related gene expression in GAS
   muscle tissue from WT and lncRNA‐MEG3 TG mice. R) Representative Myh7
   and Myh4 immunostaining in primary myotubes derived from WT and
   lncRNA‐MEG3 TG mice (n = 3). Scale bar = 100 µm. S) Representative Myhc
   staining and quantification of myotube width in primary myotubes from
   WT and lncRNA‐MEG3 TG mice (n = 6). Scale bar = 50 µm. T)
   Representative ORO staining of lipid droplets in primary myotubes from
   WT and lncRNA‐MEG3 TG mice (n = 6). Scale bar = 50 µm. U–W) Forelimb
   grip strength measurements (n = 3) (U) and treadmill endurance
   performance, including running time (V) and distance (W) in WT and
   lncRNA‐MEG3 TG mice (n = 6). Forelimb grip strength was measured at
   15 min intervals and standardized to lean body weight. (X) The
   contraction force was determined by triggering contraction using
   incremental stimulation frequencies (1 ms pulses at 0–300 Hz for 500 ms
   at 100 V) (n = 3). Data are mean ± SD; p‐values were calculated using
   Student's t‐test. ^* p < 0.05, ^** p < 0.01 and ^*** p < 0.001.

2.4. Elevated Expression of LncRNA‐MEG3 Alleviates Muscle Atrophy Induced by
Fat Infiltration

   Given that the overexpression of lncRNA‐MEG3 in adult mice resulted in
   increased muscle mass, we sought to investigate whether modulation of
   lncRNA‐MEG3 levels could influence the pathophysiology of
   muscle‐wasting diseases. In a model of DMD, we observed significant
   muscle fiber atrophy and intramuscular fat infiltration (Figure
   [134]S7A–D, Supporting Information). Grip strength and endurance were
   significantly reduced in MDX mice compared with WT (Figure [135]S7E–G,
   Supporting Information). Moreover, lncRNA‐MEG3 expression was markedly
   decreased, while the expression of muscle atrophy markers Atrogin‐1 and
   Murf‐1 was significantly elevated in the TA, SOL, and GAS muscles of
   MDX mice (Figure [136]S7H, Supporting Information). Similarly, after
   48 h of fasting in C57BL/6 mice, which resulted in reduced muscle mass
   and fiber size, lncRNA‐MEG3 expression decreased in atrophied muscle
   tissue (Figure [137]S7I–K, Supporting Information). These data
   underscore a close relationship between lncRNA‐MEG3 expression and
   muscle atrophy. To explore the therapeutic potential of lncRNA‐MEG3 in
   muscle wasting, we administered an intraperitoneal injection of the
   AAV9 virus to induce lncRNA‐MEG3 overexpression in MDX mice
   (Figure [138] 3A). The overexpression of lncRNA‐MEG3 significantly
   reduced the number of centrally nucleated fibers, and muscle fibrosis
   and increased the CSA of muscle fibers (Figure [139]3B–D).
   Additionally, lncRNA‐MEG3 overexpression notably improved grip strength
   and endurance (Figure [140]3E–G). In primary myoblasts derived from
   lncRNA‐MEG3‐overexpressing MDX mice, lncRNA‐MEG3 maintains calcium
   homeostasis by upregulating the expression of the sarco/endoplasmic
   reticulum Ca^2⁺‐ATPase (SERCA) gene, further demonstrating its critical
   role in regulating muscle strength and function in MDX mice (Figure
   [141]S8A,B, Supporting Information). Importantly, lncRNA‐MEG3
   overexpression also reduced lipid accumulation in the muscles of MDX
   mice (Figure [142]3H), suggesting its potential therapeutic application
   in mitigating muscle atrophy associated with intramuscular fat
   deposition. To confirm the underlying mechanism, we isolated stromal
   vascular fraction (SVF) cells from the GAS muscles of WT and
   lncRNA‐MEG3 TG mice and differentiated them into mature adipocytes
   (Figure [143]3I). Conditioned media (CM) from these cells was then
   applied to dexamethasone (DEX)‐induced C2C12 myotubes, a model of
   muscle atrophy, for morphological and molecular analysis
   (Figure [144]3I). As shown in Figure [145]3J–M, CM from lncRNA‐MEG3 TG
   mice significantly enhanced myotube size, differentiation capacity, and
   reduced the expression of muscle atrophy‐related mRNA and protein
   markers compared with CM from WT cells. These findings suggest that
   lncRNA‐MEG3 mitigates muscle wasting by preventing fat infiltration.
   Additionally, it is well‐established that fat infiltration can trigger
   inflammatory responses and oxidative stress.^[ [146]^25 ^] In primary
   myoblasts isolated from lncRNA‐MEG3‐overexpressing MDX mice, the
   expression of antioxidant enzymes Sod1 and Cat was significantly
   upregulated, while the levels of pro‐inflammatory cytokines IL‐6 and
   IL‐1β were notably decreased (Figure [147]S8B, Supporting Information).
   Moreover, the production of reactive oxygen species and IL‐6 was
   significantly reduced after lncRNA‐MEG3 overexpression in MDX mice
   (Figure [148]S8C,D, Supporting Information). Collectively, these
   results indicate that lncRNA‐MEG3 attenuates fat infiltration‐induced
   oxidative stress and inflammatory responses, highlighting its
   therapeutic potential in treating muscle degenerative diseases such as
   DMD.

Figure 3.

   Figure 3
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   LncRNA‐MEG3 regulates muscle atrophy induced by fat infiltration. A)
   Schematic of the experimental design using AAV9‐based overexpression
   (AAV9‐OV) in MDX mice. B) Expression of lncRNA‐MEG3 in QUA, GAS, TA,
   EDL, and SOL muscles after AAV9‐OV injection in MDX mice (n = 6). C)
   Representative H&E, Masson, and Laminin staining of muscle
   cross‐sections showing structural changes following AAV9‐OV in MDX mice
   (n = 6). scale bar = 50 µm. D) Distribution of myofiber sizes in
   AAV9‐EV and AAV9‐OV MDX mice (n = 6). E–G) Forelimb grip strength
   treadmill endurance performance in MDX mice treated with AAV9‐EV or
   AAV9‐OV (n = 5). (H) Representative BODIPY staining of lipid droplets
   in muscle tissues and quantification of lipid number (n = 3). scale bar
   = 50 µm. I) Schematic illustration of the co‐culture system of beige
   adipocytes and myotubes. J) Representative images of Myhc staining and
   myotube width in DEX‐treated C2C12 myotubes cultured with or without a
   conditioned medium (CM) from differentiated primary beige adipocytes of
   WT and lncRNA‐MEG3 TG mice (n = 6). Scale bar = 100 µm. K–M) qRT‐PCR
   and Western blot showing the expression of adipogenesis, muscle
   atrophy, and differentiation marker genes in DEX‐treated C2C12 myotubes
   cultured with or without CM from differentiated primary beige
   adipocytes of WT and lncRNA‐MEG3 TG mice (n = 6). CM: Conditioned
   medium, DEX: Dexamethasone. Data are mean ± SD; p‐values were
   calculated using Student's t‐test. ^* p < 0.05, ^** p < 0.01 and ^*** p
   < 0.001.

2.5. LncRNA‐MEG3 Regulates Mitochondrial Content and Lipid Metabolism in
Skeletal Muscle

   Electron microscopy analysis of GAS muscle from AAV9‐KD mice revealed a
   significant reduction in mitochondrial density, accompanied by notable
   disruption of mitochondrial cristae, compared with WT controls (Figure
   [150]4A,B). These findings correlate well with the decreased
   mitochondrial DNA content and suppressed expression of genes related to
   mitochondrial biogenesis and oxidative phosphorylation in GAS muscle
   from lncRNA‐MEG3 KD mice (Figure [151]S9A–C, Supporting Information).
   To determine whether the decrease in mitochondrial numbers in
   lncRNA‐MEG3 KD mice was associated with impaired respiratory function,
   we performed an O2K system analysis. The results showed that
   lncRNA‐MEG3 KD suppressed ATP production and mitochondrial oxygen
   consumption rate (OCR) in GAS muscle fibers (Figure [152]S9D–F,
   Supporting Information). Mitochondrial ATP synthesis relies on the
   electron transport chain and oxidative phosphorylation.^[ [153]^26 ^]
   After stimulation with carbonyl cyanide 4‐(trifluoromethoxy)
   phenylhydrazone (FCCP), both basal respirations, following oligomycin
   inhibition of ATP synthesis, and maximal respiration, were
   significantly lower in GAS muscle fibers from lncRNA‐MEG3 KD mice
   compared with control (Figure [154]S9E,F, Supporting Information).
   Similarly, total ATP content was reduced in the GAS muscle of
   lncRNA‐MEG3 KD mice (Figure [155]4C), whereas TG mice exhibited the
   opposite trend (Figure [156]4D–F; Figure [157]S9G–L, Supporting
   Information), indicating that lncRNA‐MEG3 plays a crucial role in
   maintaining mitochondrial function. Furthermore, in isolated
   intramuscular adipocytes, we observed that the knockdown of lncRNA‐MEG3
   suppressed the expression of key lipolysis markers (Hsl and Atgl),
   while overexpression promoted their expression (Figure [158]4G).
   Similarly, in C2C12 myoblasts, lncRNA‐MEG3 knockdown enhanced
   adipogenesis, whereas its overexpression inhibited this process
   (Figure [159]4H,I). These findings indicate that lncRNA‐MEG3 modulates
   lipid metabolism, leading to altered fat deposition in skeletal muscle.
   To explore whether the effects of lncRNA‐MEG3 knockdown on lipid
   accumulation were linked to mitochondrial dysfunction, we employed a
   glucocorticoid‐induced fat infiltration model in both lncRNA‐MEG3 KD
   and TG mice. After 14 days of treatment, lncRNA‐MEG3 KD mice showed
   significant fat infiltration and impaired muscle regeneration in the TA
   muscle (Figure [160]4J). Correspondingly, lncRNA‐MEG3 knockdown led to
   elevated levels of Perilipin‐1 protein (Figure [161]4K), along with the
   upregulation of mature adipocyte markers (Adipoq, Fabp4), and muscle
   atrophy markers, while muscle differentiation and mitochondrial
   biogenesis markers were suppressed (Figure [162]4L,M). In contrast,
   lncRNA‐MEG3 TG mice exhibited reduced lipid accumulation, muscle
   atrophy markers, and enhanced expression of relevant mitochondrial and
   muscle differentiation genes (Figure [163]4N–Q). These results suggest
   that the loss of lncRNA‐MEG3 promotes muscle atrophy induced by lipid
   infiltration, likely through the disruption of mitochondrial biogenesis
   and lipid metabolism.

Figure 4.

   Figure 4
   [164]Open in a new tab

   LncRNA‐MEG3 regulates mitochondrial biogenesis impairment induced by
   fat infiltration. A) Representative transmission electron microscopy
   (TEM) images of mitochondrial ultrastructure in skeletal muscles from
   AAV9‐NC and AAV9‐KD mice (n = 4). B) Quantification of mitochondrial
   density per µm^2 in skeletal muscles in AAV9‐NC and AAV9‐KD mice (n =
   4). C) ATP production levels in AAV9‐NC and AAV9‐KD mice GAS muscle
   tissue (n = 4). D) Representative transmission electron microscopy
   (TEM) images of mitochondrial ultrastructure in GAS muscle tissue from
   WT and lncRNA‐MEG3 TG mice (n = 4). E) Quantification of mitochondrial
   density per µm^2 in skeletal muscles in WT and lncRNA‐MEG3 TG mice (n =
   4). F) ATP production levels in skeletal muscles in WT and TG mice (n =
   4). G) qRT‐PCR showing the expression of lipid metabolism marker genes
   in muscles with lncRNA‐MEG3 knockdown or overexpression (n = 6). H,I)
   Representative BODIPY staining in C2C12 myotubes with lncRNA‐MEG3
   knockdown or overexpression (n = 3). Scale bar = 100 µm. J) H&E
   staining of TA muscle at the 14 dpi after GLY injury with lncRNA‐MEG3
   knockdown (n = 3). Scale bar = 100 µm. K) IF analysis of Perilipin‐1
   (green) and DAPI (blue) at the 14 dpi after GLY injury in muscles with
   lncRNA‐MEG3 knockdown (n = 3). Scale bar = 100 µm. L,M) qRT‐PCR and
   Western blot analysis of muscle atrophy, lipid deposition, and
   mitochondrial biogenesis marker genes in TA muscles at the 14 dpi after
   GLY injury with lncRNA‐MEG3 knockdown (n = 6). N) H&E staining of TA
   muscle at the 14 dpi after GLY injury in WT and lncRNA‐MEG3 TG mice (n
   = 3). Scale bar = 100 µm. O) IF analysis of Perilipin‐1 (green) and
   DAPI (blue) at the 14 dpi after GLY injury in lncRNA‐MEG3 TG mice (n =
   3). Scale bar = 100 µm. P,Q) qRT‐PCR and Western blot analysis of
   muscle atrophy, lipid deposition, and mitochondrial biogenesis marker
   genes in TA muscle at the 14 dpi after GLY injury in WT and lncRNA‐MEG3
   TG mice (n = 6). Data are mean ± SD; p‐values were calculated using
   Student's t‐test. ^* p < 0.05, ^** p < 0.01 and ^*** p < 0.001.

2.6. LncRNA‐MEG3 Stabilizes the PRC2 Complex by Interacting with SUZ12 to
Regulate Muscle Phenotype and Mitochondrial Function

   To investigate the molecular mechanism underlying the role of
   lncRNA‐MEG3, we employed the catRAPID
   ([165]http://s.tartaglialab.com/page/catrapid_group) algorithm to
   predict potential proteins interacting with lncRNA‐MEG3. SUZ12 emerged
   as the highest‐ranking predicted protein, and we confirmed that
   lncRNA‐MEG3 is predominantly localized in the nucleus (Figure
   [166]5A,B). RNA pull‐down assays in mice primary myoblasts lysates
   revealed a direct interaction between lncRNA‐MEG3 and SUZ12
   (Figure [167]5C). Notably, similar interactions were observed with
   lncRNA‐MEG3 from both pigs and humans, suggesting a conserved mechanism
   across species (Figure [168]5D,E). RNA immunoprecipitation (RIP)
   experiments in C2C12 myoblasts further validated the endogenous
   interaction between lncRNA‐MEG3 and SUZ12 (Figure [169]5F). To
   understand the structural basis for this interaction, we predicted the
   secondary structure of lncRNA‐MEG3, which revealed the presence of
   three distinct loops (Figure [170]5G). To identify the specific RNA
   domains involved in the interaction with SUZ12, we generated a series
   of lncRNA‐MEG3 mutants by deleting different loop regions. In vitro,
   RNA‐protein binding assays demonstrated that only the full‐length
   lncRNA‐MEG3 was able to bind SUZ12, while all mutant versions failed
   (Figure [171]5H). Next, we investigated the protein domains of SUZ12
   required for its binding to lncRNA‐MEG3 by generating three truncated
   SUZ12 mutants (B1, B2, and B3) (Figure [172]5I). The B3 fragment (amino
   acids 550–739) was identified as the core region essential for binding
   to lncRNA‐MEG3 (Figure [173]5J). Further truncation studies narrowed
   the critical binding domain to the 651–700 amino acid region
   (Figure [174]5K,L), which is known to interact with other PRC2
   components such as EZH2 and EED (Figure [175]5M).^[ [176]^27 ^] These
   findings suggest that lncRNA‐MEG3 may modulate the stability of the
   PRC2 complex, thereby influencing gene regulation. To test this
   hypothesis, we examined the effects of RNase treatment and lncRNA‐MEG3
   knockdown on the interaction between SUZ12, EED, and EZH2. Both
   treatments led to a loss of these interactions, which could be restored
   by reintroducing lncRNA‐MEG3 (Figure [177]5N–Q). Furthermore, the
   overexpression of lncRNA‐MEG3 promoted the nuclear localization of
   SUZ12 and facilitated the formation of more PRC2 complexes within the
   cell (Figure [178]5R,S). Notably, we found that the knockdown of Suz12
   resulted in a marked reduction in type I muscle fibers, increased lipid
   droplet accumulation, and impaired mitochondrial function (Figure
   [179]S10A–D, Supporting Information). In addition, Suz12 knockdown
   rescued the skeletal muscle phenotype induced by lncRNA‐MEG3
   overexpression (Figure [180]S10E–H, Supporting Information). Moreover,
   neither the knockdown nor the overexpression of lncRNA‐MEG3 affected
   the expression of Suz12 at the total mRNA and protein levels, and Suz12
   did not alter lncRNA‐MEG3 expression (Figure [181]5S; Figure
   [182]S10I–L, Supporting Information). Together, these results
   demonstrate that lncRNA‐MEG3 regulates muscle mass, lipid metabolism,
   and mitochondrial function by stabilizing the PRC2 complex through its
   interaction with SUZ12.

Figure 5.

   Figure 5
   [183]Open in a new tab

   Characterization of the interaction between lncRNA‐MEG3 and SUZ12. A)
   RNA‐binding motif enrichment analysis showing the most frequently
   detected RNA‐binding proteins associated with lncRNA‐MEG3 by catRAPID
   ([184]http://s.tartaglialab.com/page/catrapid_group). B) Fluorescent in
   situ hybridization (FISH) showing lncRNA‐MEG3 localization (green) in
   primary mouse myoblasts and C2C12 myoblasts. Nuclei are stained with
   DAPI (blue) (n = 3). Scale bar = 20 µm. C–E) RNA pull‐down assay of
   SUZ12, β‐actin (negative control), and GAPDH (negative control) using
   antisense and sense probes for lncRNA‐MEG3 in C2C12 myoblast lysates (n
   = 3). Input represents total lysates. F) Quantification of the
   lncRNA‐MEG3 interaction with SUZ12 normalized to input level (n = 3).
   G) Predicted secondary structure of lncRNA‐MEG3 highlighting structural
   domains (D1, D2, and D3). H) RNA pull‐down assay was performed to
   analyze the binding of lncRNA‐MEG3 WT, deletion expression vectors (D1,
   D2, and D3), and SUZ12 protein (n = 3). I,J) Domain mapping of SUZ12
   using truncated fragments (B1, B2, and B3) in pull‐down assays,
   identifying the lncRNA‐MEG3 interaction domain in SUZ12 (n = 3). K)
   Schematic of the deletion constructs in the B3 domain, with the deleted
   regions located at the following amino acid positions: c1 (500‐550 aa
   deletion), c2 (551–600 aa deletion), c3 (601–650 aa deletion), c4
   (651–700 aa deletion), and c5 (701–739 aa deletion). L) RNA pull‐down
   assay showing the interaction between biotinylated lncRNA‐MEG3 and the
   B3 domain deletion variants (n = 3). M) Schematic representation of
   SUZ12 domains, indicating the lncRNA‐MEG3 binding region and the
   EZH2/EED interaction region. N) Co‐IP was performed to analyze the
   interaction between SUZ12 and EZH2/EED after C2C12 myoblasts were
   treated with RNase T1, DNase I, and lncRNA‐MEG3 shRNA (n = 3). O) Co‐IP
   was performed to analyze the interaction between SUZ12 and EZH2/EED
   after overexpression of lncRNA‐MEG3 in C2C12 myoblasts (n = 3). P,Q)
   Co‐IP was performed to analyze the interaction between SUZ12 and
   EZH2/EED in C2C12 myoblasts with knockdown of lncRNA‐MEG3 (n = 3). R)
   IF staining of SUZ12 (red) in control and lncRNA‐MEG3‐overexpressing
   C2C12 cells showing enhanced nuclear localization. DAPI stains nuclei
   (blue) (n = 3). Scale bar = 20 µm. S) Western blot showing the
   expression of Suz12 in the nucleus, cytoplasm, and whole cell lysates
   of C2C12 myoblasts after lncRNA‐MEG3 overexpression (n = 3). Lamin B1
   was used as a marker for the nuclear protein, α‐Tubulin as a marker for
   the cytoplasmic protein, and Gapdh as a marker for the total protein.
   Data are mean ± SD; p‐values were calculated using Student's t‐test.
   ^** p < 0.01.

2.7. LncRNA‐MEG3/SUZ12 Maintains H3K27me3 Modification and Regulates Fhl3 and
Rnf128 Expression

   To investigate the impact of lncRNA‐MEG3 on downstream target genes
   through SUZ12, we performed RNA sequencing and H3K27me3 CUT&Tag
   combined analysis. RNA‐seq analysis revealed that SUZ12 knockdown in
   myotubes resulted in the differential expression of 980 genes,
   including the downregulation of Myh7 and upregulation of Myh4 (Figure
   [185]6A; Table [186]S2, Supporting Information). Gene Ontology (GO) and
   Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment analyses
   identified significant pathways related to muscle fiber type conversion
   and energy metabolism, such as calcium signaling and MAPK signaling
   pathways (Figure [187]6B,C; Tables [188]S3 and [189]S4, Supporting
   Information). CUT&Tag analysis following SUZ12 knockdown showed
   alterations in H3K27me3 modifications at the 2163 genes promoter (Table
   [190]S5, Supporting Information). Overlap analysis revealed that the
   genes with altered expression levels also overlapped with those
   regulated by H3K27me3 (Figure 6D; Table [191]S6, Supporting
   Information). These overlapping genes were enriched in signaling
   pathways associated with myogenesis, muscle contraction, and skeletal
   muscle development, including MAPK, PI3K‐Akt, Hippo, and calcium
   signaling pathways (Figure 6D; Tables [192]S7 and [193]S8, Supporting
   Information). Notably, we identified Fhl3 and Rnf128 within the list of
   differentially enriched genes, and the H3K27me3 modification in their
   promoter regions is influenced by SUZ12 (Figure [194]6E). Previous
   studies have shown that Fhl3 promotes fast‐twitch fiber formation,
   while Rnf128 promotes fat accumulation.^[ [195]^28 , [196]^29 ^]
   ChIP‐qPCR, qRT‐PCR, and Western blot results indicated that knockdown
   of Suz12 and lncRNA‐MEG3 reduced H3K27me3 enrichment at the promoter
   regions of Fhl3 and Rnf128, leading to their upregulation, while Suz12
   and lncRNA‐MEG3 overexpression had the opposite effect
   (Figure [197]6F–K). Moreover, Suz12 knockdown was able to rescue the
   effects of lncRNA‐MEG3 on the expression of both Fhl3 and Rnf128
   (Figure [198]S10M, N, Supporting Information). These results indicate
   that lncRNA‐MEG3 interacts with SUZ12 to regulate the expression of
   Fhl3 and Rnf128 by modulating H3K27me3 deposition at their promoter
   regions.

Figure 6.

   Figure 6
   [199]Open in a new tab

   LncRNA‐MEG3 interacts with SUZ12 to regulate the H3K27me3 modification
   of the Fhl3 and Rnf128 promoter. A) Volcano plot showing differentially
   expressed genes upon SUZ12 knockdown. B) Gene Ontology (GO) enrichment
   analysis of DEGs, highlighting biological processes significantly
   affected by SUZ12 modulation. C) KEGG pathway enrichment analysis of
   DEGs, showing pathways related to muscle development and immune
   responses. D) Venn diagrams showing the overlap between DEGs
   (downregulated and upregulated) and H3K27me3 target genes, with bar
   plots displaying enriched pathways for overlapping genes. E) Schematic
   diagrams of Fhl3 and Rnf128 loci, illustrating H3K27me3 modification
   sites at their promoters. F) ChIP‐qPCR showing the enrichment of
   H3K27me3 at the Fhl3 promoter in Suz12 knockdown (KD) and
   overexpression (OV) myoblasts compared to controls (n = 3). G,H)
   qRT‐PCR (n = 6) (G) and Western blot (n = 3) (H) showing the expression
   levels of Fhl3 in Suz12 OV and KD myoblast. I) ChIP‐qPCR showing the
   enrichment of H3K27me3 at the Rnf128 promoter in lncRNA‐MEG3 knockdown
   (KD) and overexpression (OV) myoblasts compared to controls (n = 3).
   J,K) qRT‐PCR (n = 6) (J) and Western blot (n = 3) (K) showing the
   expression levels of Rnf128 in lncRNA‐MEG3 OV and KD myoblast. Data are
   mean ± SD; P‐values were calculated using Student's t‐test. ^* p
   < 0.05, ^** p < 0.01 and ^*** p < 0.001.

2.8. LLPS Determines the Activity of SUZ12

   To explore the molecular mechanism underlying the influence of SUZ12 on
   Fhl3 and Rnf128, we analyzed the protein structure of SUZ12. Structural
   analysis revealed a prominent IDR at the N‐terminus of SUZ12 (Figure
   [200]7A), suggesting that SUZ12 may function through LLPS.^[ [201]^30
   ^] Both IF staining and GFP‐tagged fluorescence imaging confirmed that
   SUZ12 forms dynamic puncta, a hallmark feature of LLPS
   (Figure [202]7B,C). Treatment with 1,6‐ethylene glycol, a known
   inhibitor of phase separation, prevented the formation of these puncta
   (Figure [203]7D), further supporting the LLPS behavior of SUZ12.
   Photobleaching recovery experiments revealed that SUZ12‐GFP puncta
   exhibited fluorescence recovery, indicating the dynamic and reversible
   nature of the phase‐separated structures (Figure [204]7E). We further
   examined the effect of various factors on SUZ12 LLPS by manipulating
   protein concentration, salt ion concentration, and pH. Increasing the
   concentration of purified SUZ12‐GFP proteins or adjusting ionic
   conditions significantly enhanced both the size and number of puncta
   (Figure [205]7F–H). Moreover, following photobleaching, the purified
   SUZ12‐GFP proteins exhibited recovery and fusion into larger puncta,
   reinforcing the dynamic properties of SUZ12 phase separation
   (Figure [206]7I,J). These findings collectively demonstrate that SUZ12
   exhibits LLPS properties both in vivo and in vitro. To assess the role
   of the IDR in SUZ12‐mediated LLPS, we constructed IDR‐replacement
   vectors using IDRs from FUS and TIA1 proteins to substitute for the IDR
   region in SUZ12‐GFP proteins (Figure [207]7K), resulting in
   FUSIDR‐SUZ12 and TIA1IDR‐SUZ12. The deletion of the IDR from SUZ12‐GFP
   (ΔIDR‐SUZ12) markedly reduced puncta formation, underscoring its
   essential role in LLPS (Figure [208]7L). Consistently, purified
   SUZ12‐GFP proteins lacking the IDR failed to form condensates, whereas
   the IDR‐replacement variants retained LLPS functionality
   (Figure [209]7M). Furthermore, the phase‐separation enhancer PEG8000
   reinforced the necessity of the IDR in facilitating SUZ12 LLPS
   (Figure [210]7N). Collectively, these findings indicate that LLPS
   serves as a critical regulatory mechanism governing SUZ12 activity,
   with the IDR playing a pivotal role in this process.

Figure 7.

   Figure 7
   [211]Open in a new tab

   SUZ12 undergoes LLPS. A) Identification of IDR within the SUZ12 protein
   using bioinformatics analysis. The IDR region (red box) and zinc finger
   (ZnF) domains are highlighted. B) IF showing SUZ12 (green) localization
   in C2C12 myoblasts. Nuclei are stained with DAPI (blue) (n = 3). Scale
   bar = 20 µm. C) GFP‐tagged SUZ12 exhibits nuclear localization in C2C12
   myoblasts. DAPI stains nuclei (n = 3). Scale bar = 20 µm. D)
   Immunostaining of SUZ12 in C2C12 myoblasts treated with 1,6‐ethylene
   glycol, showing disrupted puncta formation. Quantification of
   SUZ12‐positive nuclear puncta per cell is shown (right panel) (n = 6).
   Scale bar = 10 µm. E) Fluorescence recovery after photobleaching (FRAP)
   of SUZ12 puncta in C2C12 myoblasts, demonstrating its liquid‐like
   properties. Representative images before and after bleaching are shown
   with recovery kinetics (right panel). Scale bar = 10 µm. F)
   Concentration‐dependent phase separation of purified SUZ12 protein in
   vitro. Representative images and quantification of puncta formation at
   increasing protein concentrations are shown (n = 6). Scale bar = 10 µm.
   G) Salt sensitivity assay showing SUZ12 puncta disruption with
   increasing NaCl concentrations (300 to 500 mm) (n = 6). Scale bar =
   10 µm. H) pH‐dependent phase separation of SUZ12, showing optimal
   droplet formation at pH 6.0–6.5. Quantification of puncta is shown
   (right panel) (n = 6). Scale bar = 10 µm. I) FRAP analysis of SUZ12
   droplets in vitro, demonstrating rapid fluorescence recovery after
   photobleaching, indicative of liquid‐like dynamics. Scale bar = 10 µm.
   J) Time‐lapse imaging of SUZ12 droplets coalescence in vitro, showing
   fusion events over time. K) Schematic of SUZ12 mutants (ΔIDR‐SUZ12,
   FUSIDR, and TIA1IDR) used to investigate the role of IDR in LLPS. L) IF
   of full‐length (FL) SUZ12 and ΔIDR‐SUZ12 in C2C12 myoblasts. ΔIDR‐SUZ12
   lacks nuclear puncta formation. Quantification is shown (right panel)
   (n = 6). Scale bar = 10 µm. M) Representative images of GFP‐tagged
   SUZ12 different vectors in C2C12 myoblasts. Quantification of the
   puncta number is shown in the right panel (n = 6). Scale bar = 10 µm.
   N) Turbidity assay of purified SUZ12 protein in the presence of
   PEG8000, confirming its ability to undergo phase separation (n = 6).
   Data are mean ± SD; p‐values were calculated using Student's t‐test. ^*
   p < 0.05, ^** p < 0.01 and ^*** p < 0.001.

2.9. LncRNA‐MEG3 Modulates SUZ12 LLPS and Epigenetic Regulation of Fhl3 and
Rnf128

   Subcellular fractionation through differential ultracentrifugation was
   employed to identify proteins involved in LLPS.^[ [212]^31 ^] Our
   analysis revealed that SUZ12 is present in the P20 fraction (pellet
   collected at 2000 × g), indicating that SUZ12 is part of a protein
   subset involved in heavy granule formation. Notably, SUZ12 levels in
   the P20 fraction decreased upon RNase A treatment or following the
   knockdown of lncRNA‐MEG3 in cells (Figure [213]S11A,B, Supporting
   Information). In contrast, the overexpression of lncRNA‐MEG3 led to an
   increase in SUZ12 levels in the same fraction (Figure [214]S11C,
   Supporting Information). Orthogonal experiments confirmed that SUZ12
   underwent LLPS in a dose‐dependent manner in the presence of
   lncRNA‐MEG3 (Figure [215]S11D, Supporting Information). Further
   co‐localization studies demonstrated that both lncRNA‐MEG3 and SUZ12
   are predominantly localized in the nucleus (Figure [216]S11E,
   Supporting Information), suggesting that lncRNA‐MEG3 may promote the
   formation of SUZ12‐containing heavy granules or liquid droplets via
   direct interaction. To further test this hypothesis, we performed gain
   and loss of the function experiment on lncRNA‐MEG3 in C2C12 myoblasts
   to assess its impact on the formation of SUZ12 puncta. The results
   showed that the overexpression of lncRNA‐MEG3 in C2C12 myoblasts
   significantly enhanced the formation of SUZ12 puncta, whereas the
   knockdown of lncRNA‐MEG3 reduced their formation (Figure [217]S11F,G,
   Supporting Information). Functional assays revealed that the SUZ12
   construct lacking the IDR failed to induce H3K27me3 modification at the
   Fhl3 and Rnf128 promoter and could not regulate their expression
   (Figure [218]S11H,I, Supporting Information). Conversely, the
   overexpression of SUZ12 rescued the ability of lncRNA‐MEG3 knockdown to
   inhibit H3K27me3 modification and up‐regulation of FHL3 and Rnf128, an
   effect that was abolished when the SUZ12 IDR was deleted (Figure
   [219]S11J,K, Supporting Information). Together, these results highlight
   that lncRNA‐MEG3 plays a pivotal role in promoting SUZ12 LLPS and
   regulating the epigenetic landscape of Fhl3 and Rnf128.

2.10. LncRNA‐MEG3 Regulates Muscle Mass and Atrophy by Promoting SUZ12 LLPS
to Modulate the Expression of Fhl3 and Rnf128

   To further elucidate the role of the lncRNA‐MEG3/SUZ12/Fhl3/Rnf128
   signaling axis in regulating skeletal muscle mass and preventing
   atrophy, we investigated the functional roles of Fhl3 and Rnf128 in
   skeletal muscle. The overexpression of Fhl3 in myotubes resulted in the
   upregulation of Myh4 and Murf1 expression, but in the downregulation of
   Myh7 and Nrf1 expression, compared with control (Figure [220]S12 A,B,
   Supporting Information). Additionally, Fhl3 overexpression suppressed
   ATP synthesis and cAMP production, with the opposite effects observed
   upon Fhl3 knockdown (Figure [221]S12C–H, Supporting Information). In
   contrast, overexpression of Rnf128 led to an increase in the levels of
   muscle atrophy and adipogenesis markers, while simultaneously reducing
   the levels of mitochondrial biogenesis markers in myotubes (Figure
   [222]S13A,B, Supporting Information). This was accompanied by a
   decrease in myotube size, ATP synthesis, and cAMP generation (Figure
   [223]S13C–E, Supporting Information). Conversely, Rnf128 knockdown
   inhibited this muscle‐wasting phenotypes (Figure [224]S13F–J,
   Supporting Information), highlighting the collaborative roles of Fhl3
   and Rnf128 in regulating muscle mass, fiber‐type conversion, fat
   deposition, and muscle atrophy.

   Furthermore, we explored the potential of rescuing the effects of
   lncRNA‐MEG3 on muscle phenotypes by overexpressing Fhl3 and Rnf128. We
   found that overexpression of these factors could partially restore
   muscle atrophy, muscle fiber type transition, mitochondrial biogenesis,
   fat deposition, and the expression of marker genes under
   lncRNA‐MEG3‐sufficient conditions (Figure [225] 8A–H). However, when
   Suz12 was overexpressed in the wild‐type vector, this rescuing effect
   of Fhl3 and Rnf128 on lncRNA‐MEG3 function was completely abolished
   (Figure [226]8A–H). Importantly, transfection with Suz12 lacking its
   IDR limited the rescue effect, suggesting that the IDR domain of SUZ12
   is essential for the regulatory activity of this pathway
   (Figure [227]8A–H). These findings underscore the critical role of the
   lncRNA‐MEG3/SUZ12/Fhl3/Rnf128 axis in maintaining skeletal muscle mass,
   preventing muscle atrophy, regulating fiber‐type plasticity, and
   modulating fat deposition. Moreover, SUZ12‐mediated LLPS serves as a
   key regulatory switch for this signaling cascade.

Figure 8.

   Figure 8
   [228]Open in a new tab

   LncRNA‐MEG3 enhances muscle function via SUZ12 LLPS‐mediated regulation
   of Fhl3 and Suz12. A) Representative immunostaining for Myh7, Myh4 in
   C2C12 myoblasts after co‐transfected with NC, lncRNA‐MEG3
   overexpression vector (lncRNA‐MEG3 OV), lncRNA‐MEG3 OV + Fhl3
   overexpression vector (Fhl3 OV), lncRNA‐MEG3 OV + Fhl3 OV + Suz12
   overexpression vector (Suz12 OV), lncRNA‐MEG3 OV + Fhl3 OV + Suz12
   deleted IDR region overexpression vector (ΔIDR‐Suz12 OV) (n = 3). Scale
   bar = 100 µm. B) Representative ORO staining and immunostaining for
   Myhc in C2C12 myoblasts after co‐transfected with NC, lncRNA‐MEG3 OV,
   lncRNA‐MEG3 OV + Rnf128 overexpression vector (Rnf128 OV), lncRNA‐MEG3
   OV + Rnf128 OV + Suz12 OV, lncRNA‐MEG3 OV + Rnf128 OV +ΔIDR‐Suz12 OV (n
   = 3). Scale bar = 100 µm. C,D) ATP synthesis and cAMP biogenesis
   detection after co‐transfected with NC, lncRNA‐MEG3 OV, lncRNA‐MEG3 OV
   + Fhl3 OV, lncRNA‐MEG3 OV + Fhl3 OV + Suz12 OV, lncRNA‐MEG3 OV + Fhl3
   OV +ΔIDR‐Suz12 OV in C2C12 myoblasts (n = 3). E,F) ATP synthesis and
   cAMP biogenesis detection after co‐transfected with NC, lncRNA‐MEG3 OV,
   lncRNA‐MEG3 OV + Rnf128 OV, lncRNA‐MEG3 OV + Rnf128 OV + Suz12 OV,
   lncRNA‐MEG3 OV + Rnf128 OV +ΔIDR‐Suz12 OV in C2C12 myoblasts (n = 3).
   G) qRT‐PCR analysis of the expression of marker genes related to muscle
   atrophy, fat deposition, muscle fiber type conversion, and
   mitochondrial biogenesis in C2C12 myoblasts after co‐transfected with
   NC, lncRNA‐MEG3 OV, lncRNA‐MEG3 OV + Fhl3 OV, lncRNA‐MEG3 OV + Fhl3 OV
   + Suz12 OV, lncRNA‐MEG3 OV + Fhl3 OV +ΔIDR‐Suz12 OV (n = 6). (H)
   qRT‐PCR analysis of marker gene expression related to muscle atrophy,
   fat deposition, muscle differentiation, and mitochondrial biogenesis in
   C2C12 myoblasts after co‐transfected with NC, lncRNA‐MEG3 OV,
   lncRNA‐MEG3 OV + Rnf128 OV, lncRNA‐MEG3 OV + Rnf128 OV + Suz12 OV,
   lncRNA‐MEG3 OV + Rnf128 OV +ΔIDR‐Suz12 OV (n = 6). I) qRT‐PCR showing
   lncRNA‐MEG3 expression after C2C12 myoblasts treated with culture media
   for different cancer cell lines (n = 3). J) qRT‐PCR showing lncRNA‐MEG3
   expression after C2C12 myoblasts treated with different cytokines (n =
   3). Data are mean ± SD; p‐values were calculated using Student's
   t‐test. ^* p < 0.05, ^** p < 0.01 and ^*** p < 0.001.

   To investigate the potential impact of cancer‐associated cachexia on
   muscle atrophy, we exposed differentiated C2C12 myotubes to acellular
   conditioned media derived from various cancer cell lines known to
   induce muscle wasting, including 4T1 (breast cancer), C26 (colon
   cancer), and HEPG2 (liver cancer). Notably, we observed a significant
   reduction in lncRNA‐MEG3 levels in conditioned media from HepG2 cells
   (Figure [229]8I). To further identify the signaling molecules
   regulating lncRNA‐MEG3 expression in muscle cells, we treated
   differentiated C2C12 myotubes with cytokines commonly associated with
   cancer cachexia. We found that pro‐inflammatory cytokines, such as
   interleukin IL‐6, IL‐1β, and tumor necrosis factor (TNF), significantly
   downregulated lncRNA‐MEG3 expression (Figure [230]8J). These results
   suggest that lncRNA‐MEG3 expression is tightly regulated by circulating
   cytokines and growth factors, positioning lncRNA‐MEG3 as a crucial
   mediator in muscle loss during cachexia and cancer‐related wasting.

3. Discussion

   The proper development and maintenance of skeletal muscle are pivotal
   for overall human health, as disruptions in muscle homeostasis are
   linked to various diseases, including cachexia, sarcopenia, and
   metabolic disorders. Many factors involved in skeletal muscle
   regulation also play crucial roles in the pathogenesis and treatment of
   these muscle‐related diseases.^[ [231]^32 , [232]^33 ^] The long
   non‐coding RNA (lncRNA) MEG3, first implicated in developmental defects
   following chromosome duplications in both mice and humans,^[ [233]^34 ,
   [234]^35 ^] has since emerged as a critical modulator of tumors and
   cancers.^[ [235]^36 ^] Over the past decade, studies have demonstrated
   that the loss of lncRNA‐MEG3 disrupts myocyte differentiation across
   diverse animal models, including mice, cattle, and pigs.^[ [236]^18 ,
   [237]^19 , [238]^35 ^] Our study extends these findings, demonstrating
   that lncRNA‐MEG3 not only regulates muscle mass and strength but also
   presents itself as a potential therapeutic target for combating muscle
   atrophy. To explore the underlying mechanisms of lncRNA‐MEG3, we
   employed a variety of in vivo models, including MDX mice and CTX‐ and
   GLY‐induced muscle injury models, alongside transgenic technologies and
   gene delivery systems to manipulate the expression of lncRNA‐MEG3 in
   skeletal muscle. Through integrated transcriptomic and CUT&Tag
   analyses, we identified a key role for lncRNA‐MEG3 in regulating
   mitochondrial function and lipid metabolism. Specifically, we found
   that lncRNA‐MEG3 promotes the LLPS of SUZ12, a core component of the
   Polycomb Repressive Complex 2 (PRC2). This interaction stabilizes the
   PRC2 complex, allowing it to catalyze H3K27me3 histone modifications on
   muscle‐related genes such as Fhl3 and Rnf128. In turn, these epigenetic
   changes govern the expression of downstream targets critical for muscle
   function, including genes (Atrogin‐1, Murf‐1, Adipoq, Fabp4,
   Perilipin‐1, Nrf1, Sdhb, Myhc, Myh4, and Myh7) involved in
   mitochondrial biogenesis, lipid metabolism, fiber‐type conversion and
   muscle atrophy (Figure [239]9 ). Skeletal muscle atrophy is a common
   pathological feature in a wide range of diseases, including aging,
   cancer‐associated cachexia, and diabetes.^[ [240]^37 , [241]^38 ,
   [242]^39 ^] In our study, we identified cytokines (IL1β, TNFα, and IL6)
   that suppress lncRNA‐MEG3 expression in cancer cell culture media,
   suggesting that dysregulated lncRNA‐MEG3 expression could contribute to
   muscle loss in these pathological conditions. The molecular pathways
   through which lncRNA‐MEG3 maintains skeletal muscle growth and quality
   provide important insights into its potential as a target for
   therapeutic intervention.

Figure 9.

   Figure 9
   [243]Open in a new tab

   Schematic illustration of the role and mechanism of energy metabolism
   reprogramming. Triggered by lncRNA‐MEG3 overexpression in skeletal
   muscle. LncRNA‐MEG3 is specifically induced in skeletal muscle in
   response to normal muscle development, stimulating the translocation
   and accumulation of SUZ12 protein in the nucleus, where it condenses
   into liquid droplets through the mechanism of LLPS. lncRNA‐MEG3 acts as
   a scaffold, recruiting SUZ12 and increasing its local concentration,
   promoting its interaction with EED and EZH2. This complex is
   sequestered together in the droplets, forming membrane‐less subcellular
   compartments, which catalyze the enhancement of H3K27me3, thereby
   inhibiting the transcriptional activity of downstream target genes.
   Through this mechanism, lncRNA‐MEG3 promotes Suz12 phase separation,
   suppressing the expression of Fhl3 and Rnf128, and further affecting
   the expression of genes involved in lipid metabolism, muscle atrophy,
   muscle fiber type conversion, and mitochondrial biogenesis, such as
   Nrf1, Myh7, Myh4, Adipoq, and Fabp4. These drive enhanced skeletal
   muscle mass and endurance, inhibit fat infiltration, and prevent muscle
   atrophy.

   One of the most intriguing findings of our study is the role of
   lncRNA‐MEG3 in regulating adipogenesis within skeletal muscle. Using
   single‐cell RNA sequencing, we previously observed that lncRNA‐MEG3
   knockdown leads to an increase in fibro‐adipogenic progenitor (FAP)
   cells, underscoring its involvement in adipogenesis and pathological
   fat deposition.^[ [244]^19 ^] Muscle injury and disease often lead to
   the differentiation of mesenchymal stem cells into adipogenic
   precursors, contributing to pathological intramuscular fat
   accumulation.^[ [245]^40 ^] In this study, we demonstrate that
   lncRNA‐MEG3 suppresses this adipogenic shift, preserving skeletal
   muscle mass and function. Overexpression of lncRNA‐MEG3 in both primary
   muscle‐derived adipocytes and injury models partially restored muscle
   fiber morphology, reduced fat infiltration, and improved myotube
   diameter and endurance. This suggests that lncRNA‐MEG3 plays a key role
   in mitigating fat deposition, a hallmark of muscle atrophy in various
   pathological conditions. Muscle atrophy is frequently associated with a
   shift in muscle fiber composition. In diseases such as cancer, sepsis,
   and aging, fast glycolytic fibers (type II) are particularly
   susceptible to atrophy, whereas slow oxidative fibers (type I) are more
   resistant.^[ [246]^41 , [247]^42 ^] Our study revealed that lncRNA‐MEG3
   preferentially induces atrophy in glycolytic muscle fibers and promotes
   a transition towards slow myosin heavy chain (MHC) isoforms. This shift
   is accompanied by enhanced mitochondrial content and improved fatigue
   resistance in mice, which aligns with existing evidence that slow
   fibers contain higher mitochondrial densities and are more
   endurance‐oriented.^[ [248]^43 ^] These findings further underscore the
   role of lncRNA‐MEG3 in regulating both muscle fiber‐type plasticity and
   mitochondrial function.

   A major insight from our work is the identification of SUZ12 as a key
   regulatory protein in the lncRNA‐MEG3 signaling axis. Our data show
   that the full‐length lncRNA‐MEG3 is essential for the LLPS of SUZ12,
   facilitating its interaction with other PRC2 components, including EZH2
   and EED.^[ [249]^27 , [250]^44 ^] This RNA‐dependent interaction plays
   a critical role in stabilizing the PRC2 complex, ensuring the proper
   modification of H3K27me3 at target gene promoters.^[ [251]^45 ^] By
   modulating these epigenetic changes, lncRNA‐MEG3 governs the expression
   of Fhl3 and Rnf128. Fhl3 is a gene that promotes the formation of
   fast‐twitch muscle fibers, while Rnf128 facilitates fat accumulation.^[
   [252]^28 , [253]^29 ^] Herein, we also found that Fhl3 and Rnf128
   jointly regulate the expression of mitochondrial biogenesis genes, ATP
   and cAMP formation, and muscle atrophy. These results establish the
   lncRNA‐MEG3/SUZ12/Fhl3/Rnf128 axis as a key regulatory pathway in
   muscle mass maintenance and atrophy.

   LLPS is essential for the dynamic assembly of biomolecules and the
   formation of membrane‐less organelles that regulate molecular
   interactions and organelle function.^[ [254]^46 ^] Proteins with IDRs
   drive LLPS because of their flexible, chain‐like structures, which
   allow multivalent, low‐affinity interactions that promote biomolecular
   condensate formation.^[ [255]^46 , [256]^47 , [257]^48 ^] Some proteins
   and transcription factors enriched in IDR and low‐complexity domains,
   such as PSPC1, TRA, and Rho, influence gene expression and biological
   processes through LLPS.^[ [258]^49 , [259]^50 , [260]^51 ^] Our study
   provides the first evidence of LLPS in regulating muscle mass, lipid
   deposition, and atrophy. We reveal that SUZ12, through its proline‐rich
   IDR, drives LLPS, a process that we show is essential for its function
   in skeletal muscle. The deletion of the IDR region of SUZ12 impairs its
   ability to undergo LLPS, disrupting the H3K27me3 modifications at the
   promoters of Fhl3 and Rnf128 and, consequently, reducing their
   expression. Notably, LLPS in the skeletal muscle has rarely been
   reported, with factors such as FUS and TDP‐43 being linked to
   amyotrophic lateral sclerosis (ALS), a neurodegenerative disease
   affecting muscle function.^[ [261]^52 , [262]^53 ^] Our findings
   suggest that similar LLPS mechanisms may also be at play in muscle
   cells, with a directly impact on muscle health and disease. These novel
   insights into the molecular mechanisms of phase separation in skeletal
   muscle provide a compelling avenue for future research into
   LLPS‐related therapeutic strategies.

   The role of RNA in LLPS has gained increasing attention, particularly
   in the context of lncRNAs, which have been shown to modulate the
   formation and dynamics of phase‐separated compartments.^[ [263]^54 ^]
   For instance, the lncRNA GIRGL drives CAPRIN1‐mediated LLPS, inhibiting
   the translation of glutamyl‐tRNA synthetase‐1.^[ [264]^55 ^] LncRNA
   MNX1‐AS1 enhances IGF2BP1 phase separation, promoting the
   MNX1‐AS1/IGF2BP1/c‐Myc signaling axis and facilitating cell cycle
   progression and proliferation in lung cancer cells.^[ [265]^56 ^] SNHG9
   regulates the phase separation of the Hippo signaling node kinase
   LATS1, modulating YAP oncogenic signaling and tumorigenesis.^[ [266]^57
   ^] In our study, we demonstrate that lncRNA‐MEG3 promotes the LLPS of
   SUZ12, facilitating the formation of stable RNA‐protein condensates,
   which are essential for maintaining the stability of the PRC2 complex
   and its epigenetic regulatory functions. This work not only highlights
   the importance of RNA‐mediated LLPS in muscle biology but also suggests
   that targeting lncRNA‐MEG3 or its interactions with SUZ12 may offer
   novel therapeutic strategies for muscle‐related diseases.

4. Conclusion

   This study identified lncRNA‐MEG3 as a pivotal regulator of skeletal
   muscle homeostasis, driving the expansion of slow/oxidative muscle
   fibers and influencing muscle mass regulation, regeneration, and
   disease pathogenesis. Our findings establish that lncRNA‐MEG3 is
   essential for maintaining muscle function, with its overexpression
   enhancing muscle strength, endurance, and regeneration, while its
   depletion leads to widespread muscle atrophy and mitochondrial
   dysfunction. Mechanistically, we revealed that lncRNA‐MEG3 exerts its
   effects by stabilizing the SUZ12‐containing PRC2 complex, thereby
   modulating key signaling pathways involved in muscle fiber type
   specification, mitochondrial biogenesis, and lipid metabolism. Notably,
   lncRNA‐MEG3 regulates SUZ12 LLPS, influencing the expression of
   downstream muscle‐specific genes such as Fhl3 and Rnf128. These
   findings not only shed new light on the molecular mechanisms governing
   muscle fiber remodeling but also highlight the therapeutic potential of
   lncRNA‐MEG3 for combating muscle‐wasting conditions, such as Duchenne
   muscular dystrophy. In conclusion, lncRNA‐MEG3 emerges as a powerful
   and novel target, offering promising avenues for the development of
   targeted therapies to address muscle degenerative diseases.

5. Experimental Section

Ethical Declaration

   All animals used in the present study were handled in compliance with
   the guidelines (AGIS2020.04.17) provided by the Biomedical Research
   Ethics Committee of the Agricultural Genomics Institute at Shenzhen,
   Chinese Academy of Agricultural Sciences.

Animal Models and Experimental Procedures

   Female C57BL/6J mice aged 6 weeks, 8 weeks, 5 months, 6 months, 8
   months, and 18 months were used in this study. Mice were housed at 72°
   Fahrenheit under a 12‐h light/12‐h dark cycle and maintained on a
   standard chow diet. Mice had ad libitum access to food and water.
   Transgenic lncRNA‐MEG3 (TG) mice were obtained from the Beijing
   Institute of Animal Science and Veterinary Medicine (2 Yuanmingyuan
   West Road, Beijing, China). Non‐transgenic mice used in the study were
   the non‐transgenic offspring of transgenic parental mice.

   The Adeno‐associated viral (AAV‐9) vectors produced by HanBio
   Biotechnology were used to package the transcript (ENSG00000235116) of
   lncRNA‐MEG3 and shRNA for overexpression and knockdown experiments.
   Neonatal mice were intraperitoneally injected with 1 × 10^12 viral
   genome particles to induce systemic interference, followed by analysis
   after 8 weeks. At 5 weeks of age, the mice were injected with 1 × 10^12
   viral genome particles into the TA muscle for local muscle‐specific
   interference and overexpression, and analysis was performed 8 weeks
   later. shRNA sequence was listed in Table [267]S9 (Supporting
   Information).

   MDX model (Duchenne muscular dystrophy (MDX, hE8‐30) mice were procured
   from CYAGEN Biosciences (Guangzhou, China). For lncRNA‐MEG3
   overexpression in MDX, 8‐week‐old DMD male mice were intraperitoneally
   injected with AAV9 vectors carrying either a scrambled control or a
   lncRNA‐MEG3 overexpression vector (viral titer: 1 × 10¹¹
   particles/mouse; HanBio Biotechnology, Beijing, China)

Regeneration Model

   For the muscle regeneration experiment, 6‐week‐old mice with
   AAV9‐mediated skeletal muscle‐specific knockdown of lncRNA‐MEG3 and
   lncRNA‐MEG3 TG mice were injected with 100 µL of cardiotoxin (CTX;
   10 mm solution, Sigma–Aldrich) into the TA muscle to induce muscle
   injury. TA muscles were collected at 3.5, 5.5, 10, and 25 days post‐CTX
   treatment for histological (H&E staining), IF, and statistical
   analyses. All mice were euthanized by cervical dislocation, with
   measures taken to minimize animal discomfort throughout the
   experimental procedures.

GLY Injection Model

   Fifty microliters of 50% v/v GLY (Sigma, USA) were injected into the
   left TA muscle of six 8‐week‐old lncRNA‐MEG3 TG mice and an equal
   number of age‐matched WT mice. Mice were allowed to recover for 14 days
   post‐injection (dpi) before being euthanized, and muscles were
   collected for analysis.

Muscle Strength and Treadmill Performance Testing

   Limb Grip Strength: Limb grip strength was measured using the grip‐test
   meter system (XR501, XinRuan, Shanghai, China). Mice were allowed to
   grip a metal grid using their front paws while being gently pulled
   backward until their hind limbs released the grid. Each mouse performed
   four trials, and the average grip strength was calculated for each
   individual. The investigator conducting the test was blinded to the
   treatment groups.

   Treadmill Endurance Testing: Treadmill testing was performed using an
   eight‐lane treadmill system (ZH‐PT/5S, Zhenghua, Anhui, China). Mice
   were acclimated to treadmill running at a speed of 6 m min^−1 for 5
   consecutive days before undergoing a 4‐week chronic treadmill exercise
   protocol. On the final day, an exhaustive exercise test was conducted
   on a treadmill inclined at a fixed 10° slope. The protocol involved
   progressive running, starting at 10 m min^−1 for 15 min, followed by
   sequential increases to 13, 16, 19 , and 22 m min^−1, each for 3 min.
   The mice then ran at 25 m min^−1 until exhaustion. Exhaustion was
   defined as the inability to move off an electric grid (35 V) located at
   the treadmill's end for a continuous duration of 5 s. Running time and
   total distance covered by each mouse were recorded. To minimize diurnal
   variability, all tests were conducted at the same time of the day.
   Throughout the experiments, animals were consistently handled and
   tested by the same investigators, who were blinded to group assignments
   to ensure unbiased data collection.

ATP Quantification

   High‐resolution respirometry was performed using an Oxygraph‐2k unit
   (Oroboros Instruments, Innsbruck). The instrument was set to 37 °C with
   a stirring speed of 750 rpm and a data averaging interval of 2 s
   (Datlab 8.1, Oroboros Instruments). Skeletal muscle (5 mg) was isolated
   from mice, cut into small pieces, and homogenized with a glass tissue
   grinder in 400 µL of mitochondrial respiration solution on ice. Tissue
   homogenate was used for the analysis of mitochondrial respiratory
   function. After 5 min equilibration period, 2 µm Magnesium Green (MgG)
   was first titrated to examine the mitochondrial ATP production.

Mitochondrial Oxygen Consumption Analysis in Skeletal Muscle Fibers

   Mitochondrial oxygen consumption in GAS muscle fibers isolated from
   lncRNA‐MEG3 knockdown and overexpression mice was measured using an
   Oxygraph‐2k units (Oroboros Instruments, Innsbruck) at 25 °C. Muscle
   fibers were treated with either 0.05% dimethylformamide (vehicle) or
   300 µm menadione for 15 min while rotating on a MACSMIX rotator
   (Miltenyi Biotec). After centrifugation (100 × g), cells were washed
   with Krebs‐Henseleit buffer, incubated for the designated period, and
   then resuspended in MiR05 medium (2.5 × 10⁵ cells mL^−1) for oxygen
   consumption analysis. The assessment included: 1) basal oxygen
   consumption; 2) proton leak (Leak) after oligomycin (2.5 µm) addition;
   3) maximal mitochondrial capacity after FCCP titration (final
   concentration 0.1 µm); 4) Complex I (CI)‐linked respiration following
   glutamate, malate, and ADP addition, and CI/CII‐dependent respiration
   after succinate. To distinguish CI‐ and CII‐mediated respiration,
   rotenone (0.5 µm) was used. Coupling efficiency and proton leak were
   analyzed post‐oligomycin treatment. Finally, non‐mitochondrial oxygen
   consumption was measured after antimycin A (2.5 µm) addition for
   correction.

In Vivo Muscle Contraction Force Measurement

   The contractile force of the TA muscle was assessed using the Aurora
   Scientific 1300A 3‐in‐1 whole animal system, in accordance with
   previously described protocols.^[ [268]^58 , [269]^59 ^] Mice were
   anesthetized through intraperitoneal injection of
   3‐bromo‐2‐fluorobenzyl alcohol (31.2 g kg^−1 body weight). After
   anesthesia, the TA muscle was carefully isolated from tendon to tendon.
   The distal tendon was secured with surgical suture, looped, and
   attached to the lever arm hook for force measurements. To determine the
   maximum isometric tetanic force, the TA muscle was electrically
   stimulated at varying frequencies (pulse width: 2 ms; frequency range:
   1–300 Hz; duration: 500 ms; voltage: 100 V; inter‐stimulation interval:
   1 min). During the experiment, the muscle was continuously perfused
   with pre‐warmed (37 °C) phosphate‐buffered saline (PBS) to maintain
   physiological conditions.

Sectioning and Staining

   Skeletal muscle tissues were immediately immersed in a 30% sucrose
   solution and embedded in isopentane pre‐cooled with liquid nitrogen.
   The embedded tissues were sectioned into 10 µm thick slices using
   cryosections (Leica CM1950, Leica, Germany) for subsequent staining
   procedures. Hematoxylin and Eosin (H&E) Staining: tissue sections were
   stained with hematoxylin for 20 min to visualize nuclei, followed by
   differentiation and bluing. The cytoplasm was then counterstained with
   eosin for 5 min. To conduct SDH Staining: cryosections (10 µm) were
   incubated at 37 °C for 30 min in a solution containing 0.2 m sodium
   phosphate buffer (pH 7.6), 0.6 mm nitro blue tetrazolium, and 50 mm
   sodium succinate (Sigma–Aldrich, St. Louis, MO, USA). After incubation,
   sections were rinsed with distilled water and mounted using an aqueous
   mounting medium. For sections IF: muscle sections were incubated
   overnight at 4 °C with primary antibodies specific to embryonic Myhc
   (MF‐20, 1:50 dilution, DSHB), Laminin (ab11575, 1:500 dilution, Abcam),
   MyHC1 (Myh7) (BA‐D5, 1:100 dilution, DSH), MyHC2b (Myh4) (BF‐F3, 1:50
   dilution, DSHB), MyHC2a (Myh2) (SC‐71, 1:100 dilution, DSHB) and
   Perilipin‐1 (ab317260, 1:1000 dilution, Abcam). After washing,
   secondary antibodies (Goat anti‐Mouse IgG2b Cross‐Adsorbed Secondary
   Antibody, Alexa Fluor 488, A‐21141, 1:100 dilution, Goat anti‐Mouse
   IgG1 Cross‐Adsorbed Secondary Antibody, Alexa Fluor 594, A‐21125, 1:100
   dilution, and Goat anti‐Mouse IgM (Heavy chain) Secondary Antibody,
   Alexa Fluor 647, A‐21238, 1:100 dilution, Thermo Fisher Scientific,
   Waltham, MA, USA) were incubated for 1 h at 37 °C. Images were acquired
   using a Nikon A1HD25 microscope (Nikon, Japan). To assess lipid
   content, muscle sections were stained with ORO for 30 min at room
   temperature. For fluorescent visualization of lipid droplets, sections
   were stained with BODIPY dye (BODIPY 493/503, MCE, USA), which emits
   green fluorescence. Imaging: All stained sections were imaged using a
   Nikon A1HD25 Confocal Microscope (Nikon, Japan). The setup enabled
   high‐resolution visualization for detailed morphometric and
   histological analyses.

Transmission Electron Microscopy (TEM) and Image Analysis

   For TEM analysis, GAS muscle samples were immediately fixed overnight
   in 2.5% glutaraldehyde after collection. The sample was infiltrated
   with a mixture of epichlorohydrin and Epon 812 resin (EMS, Hatfield,
   PA, USA) for observation and imaging under the Hitachi Bio TEM (Tokyo,
   Japan) electron microscope facility of Wuhan Sevier Biotechnology Co.,
   Ltd. Each slice captured at least five views of mitochondrial
   populations between muscle fibers and beneath the muscle membrane.
   ImageJ was used to calculate the number of mitochondria.

Single‐Cell RNA Sequencing (scRNA‐seq) Analysis

   We obtained publicly available single‐cell RNA sequencing (scRNA‐seq)
   datasets from human skeletal muscle atlas ([270]GSE147457),^[ [271]^22
   ^] spatiotemporal transcriptomic maps of whole mouse embryos
   ([272]https://cells.ucsc.edu/mouse‐limb/10x/200120_10x.h5ad),^[
   [273]^60 ^] human muscle aging cell atlas database
   ([274]https://db.cngb.org/cdcp/hlma/rnaseq/)^[ [275]^24 ^] and DMD
   model ([276]GSM4732632).^[ [277]^23 ^] ScRNA‐seq data were processed
   and analyzed following standard workflows. Low‐quality cells with
   mitochondrial gene content >15% or gene counts <200 or >8000 were
   filtered out, and doublet removal was performed using DoubletFinder to
   ensure data purity. Gene expression data were normalized using the
   log‐normalization method implemented in Seurat (v5.1.0), where UMI
   counts were log‐transformed and scaled to 10 000 to mitigate sequencing
   depth and batch effects. The top 2000 highly variable genes were
   identified and standardized using z‐score normalization to ensure
   comparability across different cell types. Principal component analysis
   (PCA) was performed to capture major biological variations, retaining
   the top 15 principal components (PCs) for downstream analysis. Batch
   effect correction was conducted using Harmony (v1.2.1) based on the PCA
   results. The corrected embeddings were then subjected to dimensionality
   reduction using uniform manifold approximation and projection (UMAP)
   for visualization. Louvain clustering was performed using the
   FindClusters function in Seurat (resolution = 1) to define distinct
   cell populations. Cell populations were then annotated based on
   established marker genes to ensure biological relevance and accuracy.^[
   [278]^22 , [279]^23 , [280]^60 ^] Finally, key marker gene expression
   patterns were visualized using UMAP and violin plots, providing an
   intuitive representation of the transcriptional profiles across
   distinct cell types.

Primers and Oligonucleotides

   Primers were designed using Primer Premier5. All primer sequences are
   provided in Table [281]S10 (Supporting Information). The
   Biotin‐lncRNA‐MEG3, siRNAs of lncRNA‐MEG3 and Suz12, and siRNA NC were
   obtained from GenePharma (Shanghai, China). All siRNA sequences are
   provided in Table [282]S11 (Supporting Information).

Plasmid Construction

   The full‐length lncRNA‐MEG3, the loop structure mutants of lncRNA‐MEG3
   (designated as lncRNA‐MEG3 D1, D2, and D3, collectively referred to as
   lncRNA‐MEG3 ΔSuz12, which lack Suz12 binding sites), the Suz12 CDS
   region and Suz12 CDS mutants for analyzing LLPS (ΔIDR‐Suz12,
   FUSIDR‐Suz12, TIA1IDR‐Suz12, and ΔlncRNA‐MEG3‐Suz12), Fhl3 CDS and
   Rnf128 CDS were cloned into the pcDNA3.1‐GFP/RFP vector for functional
   analyses. Expression vector with a deleted CDS region of Suz12 (B1, B2,
   B3, c1, c2, c3, c4, and c5) were cloned into the pcDNA3.1‐GST‐tag
   vector for binding analyses. All plasmids were purchased from
   GeneCreate (Wuhan, China).

Isolating Primary Myoblasts

   Primary myoblasts were isolated from the hindlimb skeletal muscle of
   male mice. Briefly, the skeletal muscle was minced and digested in type
   II collagenase (Gibco, Gaithersburg, MD, USA) for 30 min, then filtered
   through a mesh to remove debris. The cells were subsequently cultured
   in RPMI‐1640 medium. When the primary myoblasts reached 90% confluence,
   differentiation was induced with Dulbecco's Modified Eagle Medium
   (DMEM) containing 5% horse serum (Gibco, Gaithersburg, MD, USA).

Cell Culture and Transfection

   HEK293T cell (iCell‐h237), C2C12 myoblasts (iCell‐m013), porcine
   myoblasts (iCell‐0017a) and HEPG2 (iCell‐h092), 4T1 (iCell‐m067) and
   C26 (iCell‐m092) were obtained from iCell Bioscience Inc (Shanghai,
   China). HEK293T cell, C2C12, and HEPG2,4T1 and C26 were cultured in
   DMEM (Gibco) supplemented with 10% fetal bovine serum (Gibco) and 1%
   penicillin/streptomycin (PS, Gibco, Grand Island, NY, USA). Porcine and
   mice primary myoblasts were cultured in RPMI‐1640 medium supplemented
   with 10% fetal bovine serum (Gibco, Grand Island, NY, USA) and 1%
   penicillin/streptomycin (Gibco, Grand Island, NY, USA). All cells were
   cultured in an incubator at 37 °C with 2% CO[2]. To induce adipogenic
   differentiation, C2C12 myoblasts were treated with adipogenic DM
   containing 10% FBS, 1 µM dexamethasone (50‐02‐2, Sigma, USA), 0.5 mm
   1‐methyl‐3‐isobutylmethyl‐xanthine (28822‐58‐4, Sigma, USA), and
   10 µg ml^−1 insulin (11061‐68‐0, Sigma, USA). Typically, when the cells
   reached 50–60% confluence, they were transfected using Lipofectamine
   3000 according to the instructions of the manufacturer, and the medium
   was replaced with a complete medium 6 h after transfection.

   When C2C12 myoblasts reached 80–85% confluence in a 6‐well plate, they
   were treated with cytokines (IL‐1β (PA1000‐4827, IPODIX, China), TNFα
   (PA1000‐4468, IPODIX, China), and IL‐6 (PA1000‐4864, IPODIX, China))
   for 48 h, followed by the analysis of lncRNA‐MEG3 expression. After 3
   days of induction in C2C12 myotube, the cells were treated with
   dexamethasone (Sigma, D‐085, USA) and then switched to adipocyte
   induction medium containing SVF extracted from the corresponding WT and
   lncRNA‐MEG3 mice muscles. Analyses were performed after 3 days of
   induction.

Adipogenic Differentiation

   For adipogenic differentiation, intramuscular preadipocytes were
   cultured with an induction medium (IM) containing DMEM, 10% fetal
   bovine serum (FBS), 2.85 mm insulin, 0.3 mm dexamethasone (DEXA), and
   0.63 mm 3‐isobutyl‐methylxanthine (IBMX) for 4 days when the cells
   reached confluence. Subsequently, the cells were switched to a
   differentiation medium (DM) consisting of DMEM, 10% FBS, and 200 nm
   insulin for 2 more days until the adipocytes matured. To avoid the
   impact of cell density on adipogenic differentiation, differentiation
   was induced when the cells reached 90% confluence.

ATP Assay

   The intracellular ATP concentration was measured using an ATP assay
   (MAK473, Sigma, USA) kit following the manufacturer's protocol.
   Briefly, muscle tissues or C2C12 myoblasts were lysed with lysis
   buffer, then centrifuged at 14 400 g for 5 min at 4 °C. The supernatant
   was collected for ATP measurement, and protein concentration was
   determined using a BCA protein assay kit. Next, 20 µl of the
   supernatant was mixed with 100 µl of ATP assay solution, incubated for
   5 min at room temperature, and mixed immediately, and luminescence was
   measured using a dual‐luciferase reporter detection system (Promega,
   Madison, WI). ATP concentration was calculated from the standard curve
   and expressed as nanomoles per microgram of protein.

cAMP Quantification

   The total cellular cAMP content was quantified using the cAMP Direct
   Enzyme Immunoassay Kit (Enzo Life Sciences). Briefly, the supernatants
   were removed, and muscle tissues or C2C12 myoblasts were treated with
   0.1 m HCl for 10 min at room temperature. The cells were scraped off
   the plate surface using a cell scraper and centrifuged at ≥600 × g to
   pelletize cellular debris. The supernatant was then assayed immediately
   according to the manufacturer's instructions.

Bioinformatic Analysis

   The lncRNA‐MEG3 binding proteins were predicted using catRAPID
   ([283]http://s.tartaglialab.com/page/catrapid_group). The SUZ12 IDR
   region was predicted by PLAAC ([284]http://plaac.wi.mit.edu/) and
   PhaSePred ([285]http://lab.phasep.pro/).

RNA‐Seq Analysis

   The RNA‐seq libraries were sequenced on an Illumina NovaSeq 6000
   platform (Novogene, Tianjin, China) using a paired‐end read length of
   150 bp. Raw sequencing reads were subjected to quality control using
   Fastp to remove low‐quality bases and adapter sequences using default
   settings. Clean reads were then aligned to the reference genome GRCm38
   using HISAT2, generating SAM files. The alignment quality score (MAPQ)
   was set to ≥ 20 to ensure reliable mapping. Subsequently, SAM files
   were converted to BAM format, sorted, and indexed using SAMtools
   (v1.21), with the alignment rate exceeding 80% to ensure data quality.
   Transcript assembly was performed using StringTie (v2.2.3) on the
   aligned reads, and gene and transcript expression levels were estimated
   using the gencode.vM25.annotation.gtf annotation file. Expression
   levels were normalized using TPM (Transcripts Per Million). Data
   normalization was performed using the DESeq2 package (v1.42.0), where
   raw counts were normalized by size factors, followed by log2
   transformation to stabilize the variance across different expression
   levels. Principal component analysis (PCA) was used to assess the
   quality of the data, and the top 2000 most variable genes were selected
   for PCA to identify potential outliers. Differential expression
   analysis was conducted using DESeq2 (v1.42.0), with genes having an
   adjusted p‐value < 0.05 and a log2 fold change > 0.58 considered as
   significantly differentially expressed (DEGs). To gain insights into
   the biological processes and pathways associated with the DEGs, KEGG
   (Kyoto Encyclopedia of Genes and Genomes) pathway enrichment analysis
   was performed using ClusterProfiler (v4.10.0) in R (v4.4.2).

Extraction of Total RNA and qRT‐PCR

   Total RNA was extracted from cells and muscle tissues using TRIzol
   Reagent (Invitrogen, Shanghai, China). cDNA was synthesized using the
   HiScript IV All‐in‐One Ultra RT SuperMix for qPCR (R433‐01, Vazyme,
   Nanjing, China) according to the manufacturer's instructions. qRT‐PCR
   was performed on a QuantStudio 3 Real‐Time PCR System (Thermo Fisher
   Scientific) using Taq Pro Universal SYBR qPCR Master Mix (Q712‐02,
   Vazyme, Nanjing, China) following the manufacturer's protocol. The
   reaction conditions were 95 °C for 10 s, followed by 40 cycles of 5 s
   at 95 °C and 30 s at 60 °C. Relative gene expression levels were
   analyzed using the 2^−△△CT method. Mouse GAPDH and U6 were used as
   endogenous reference genes for normalization. The primers are listed in
   Table [286]S10 (Supporting Information).

Western Blot Analysis

   Protein samples from muscle tissues and cells were extracted using
   protein lysis buffer (Thermo Fisher, Waltham, MA, USA), which contained
   50 mm Tris (pH 7.5), 150 mm NaCl, 0.5% Nonidet P40, and protease and
   phosphatase inhibitors. The protein lysates were separated by SDS‐PAGE,
   transferred to a polyvinylidene fluoride (PVDF) membrane, and
   immunoblotted with primary antibodies against MyH7 (1:1000, Sigma
   M8421, Germany), MyH4 (1:1000, Sigma MABT847, Germany), SDHB (1:1000,
   CST #92649, USA), NRF1 (1:1000, CST #46743, USA), Perilipin‐1 (1:1000,
   abcam ab317260 USA), FABP4 (1:1000, abcam ab308110, USA), Atrogin‐1
   (1:1000, abcam ab168372, USA), Murf‐1 (1:1000, abcam ab183094, USA),
   Adipoq (1:1000, abcam ab181281, USA), EED (1:1000, CST E4L6E, USA),
   EZH2 (1:1000, CST D2C9, USA), SUZ12 (1:1000, CST D39F6, USA), LAMP1
   (1:1000, Sigma D2D11, Germany), Lamin B1 (1:1000, Sigma SAB2101352,
   Germany), Fhl3 (1:1000, Affinity DF8826, China), H3K27me3 (1:1000,
   abcam ab6002, USA), β‐actin (1:1000, CST A5441, USA), Rnf128 (1:1000,
   abcam ab137088, USA), GST‐Tag (1:1000, Affinity T0007, China),
   α‐tubulin (1:1000, Affinity AF4651, China) and GAPDH (1:1000, CST
   14C10, USA). Membranes were washed for 30 min, incubated with
   horseradish peroxidase‐conjugated secondary antibodies (GB23303,
   GB23301, Servicebio, China) for 1 h at room temperature, and washed
   again for 30 min. The membranes were then incubated in Detection
   Solution (SuperSignal West Pico PLUS, ThermoFisher, USA) for 1 min at
   room temperature and exposed to a fluorescent chemiluminescence imaging
   system (ChemiScope 6100Touch, Clinx, China).

Immunofluorescence Staining

   Treated cells were fixed with 4% paraformaldehyde, then permeabilized
   with 0.5% Triton for 20 min, blocked with goat serum for 1 h, and then
   incubated with anti‐MyH7 (1:1000, Sigma M8421, Germany) and anti‐MyH4
   (1:1000, Sigma M1570, Germany) for 2 h. Subsequently, cells were
   incubated with a goat anti‐mice secondary antibody (GB25303, GB21301,
   Servicebio, China). Finally, DAPI (1:1000, Invitrogen D3571, Shanghai,
   China) was added, and the cells were observed using a Nikon A1HD25
   microscope (Nikon).

Fluorescence In Situ Hybridization

   RNA‐FISH experiments were performed following the manufacturer's
   protocol for the FISH Kit (G‐fan, Shanghai, China). FISH experiments
   according were performed to the manufacturer's instructions included
   with the RNA FISH Kit, which was obtained from GEFAN (Shanghai, China).
   Briefly, the cells were incubated with the RNA probes in a
   hybridization buffer for 48 h at 68 °C. The cells were washed three
   times with SSC buffer, stained with DAPI (1:1000, Invitrogen D3571,
   Shanghai, China) for 10 min at room temperature, and then examined
   using a confocal microscope (Nikon A1HD25). The probes were listed in
   Table [287]S12 (Supporting Information)

Oil Red O and BODIPY Staining

   Oil red O (ORO) staining was performed as described.^[ [288]^61 ^]
   Briefly, fixed cells were incubated in ORO dye solution for 30 min.
   Staining was observed under a phase‐contrast microscope or in the
   far‐red spectrum under a confocal microscope (Nikon A1HD25). For BODIPY
   staining, intracellular lipid droplets were stained with 0.5 nm BODIPY
   FL (Invitrogen) for 10 min. Following staining, the cells were fixed
   with 4% paraformaldehyde and examined under a fluorescence microscope
   for analysis.

Measurement of Intracellular Free Calcium

   Intracellular Ca^2⁺ levels were quantified by flow cytometry using the
   calcium‐sensitive fluorescent dye Fluo‐3 AM (S1056, Beyotime, China).
   Primary myoblasts from treated MDX and WT mice were seeded in 24‐well
   plates and incubated with 4 µm Fluo‐3 AM at 37 °C for 30 min in the
   absence of light. Following incubation, the cells were collected,
   washed twice with Hank's Balanced Salt Solution (HBSS), and analyzed
   with a FACSCalibur flow cytometer (BD Biosciences, San Jose, CA, USA).
   The mean fluorescence intensity (MFI) was recorded at 488 nm excitation
   to assess intracellular calcium levels.

Detection of Reactive Oxygen Species

   Intracellular reactive oxygen species (ROS) were measured using a ROS
   detection kit (S0035S, Beyotime, China). Treated MDX and WT primary
   myoblasts were seeded in 12‐well plates and incubated with 10 µm
   2′,7′‐dichlorofluorescein diacetate (DCFH‐DA) at 37 °C for 30 min.
   After incubation, the cells were washed three times with PBS containing
   2% FBS to remove excess dye. ROS levels were then assessed by measuring
   the fluorescence intensity using a FACSCalibur flow cytometer (BD
   Biosciences, San Jose, CA, USA).

Enzyme‐Linked Immunosorbent Assay (ELISA)

   To measure IL‐6 secretion, primary myoblasts isolated from treated MDX
   and WT mice were seeded in 24‐well plates and cultured for 24 h. After
   the incubation period, culture supernatants were collected, and IL‐6
   concentrations were determined using an ELISA kit (PI326, Beyotime,
   China) according to the manufacturer's instructions. The optical
   density (OD) at 450 nm was measured using a microplate reader, and IL‐6
   levels were calculated by comparing the OD values with a standard curve
   generated using recombinant IL‐6 of known concentrations.

RNA‐Chromatin Immunoprecipitation (RIP)

   RNA immunoprecipitation (RIP) assays were conducted using the RNA
   Immunoprecipitation Kit (BersinBi, Guangzhou, China) according to the
   manufacturer's instructions. Briefly, C2C12 myoblasts at 80–90%
   confluency in 15 cm dish were scraped off, then lysed in complete RIP
   lysis buffer, after which 100 µL of whole‐cell extract was incubated
   with RIP buffer containing protein A/G magnetic beads (PB101, Vazyme,
   China) conjugated with anti‐ SUZ12 (1:1000, CST D39F6, USA). Samples
   were incubated with Proteinase K (20mg/ml) with shaking to digest the
   protein, followed by the isolation of immunoprecipitated RNA.
   Furthermore, purified RNA was subjected to qRT‐PCR analysis to
   demonstrate the presence of the binding targets using respective
   primers. The primer sequences are listed in Table [289]S10 (Supporting
   Information).

RNA Pull‐Down Assay

   Whole‐cell lysates were prepared from treated cells using RNA pull‐down
   lysis buffer (25 mm Tris‐HCl, pH 7.4; 150 mm KCl; 5 mm EDTA; 5 mm
   MgCl₂; 1% NP‐40; 0.5 mm DTT) supplemented with protease and RNase
   inhibitors. The lysates were clarified by centrifugation at 12000 × g
   for 10 min at 4 °C. Biotin‐labeled lncRNA‐MEG3 probes (0.1 µg µL^−1)
   were added to the lysates, and the mixture was incubated sequentially
   at 37 °C for 15 min, room temperature for 15 min, and then at 4 °C for
   6 h, and finally to overnight with gentle rotation. Streptavidin‐coated
   magnetic beads (Thermo Fisher Scientific) were pre‐washed and added to
   the reaction mix, followed by incubation at 4 °C for 1 h with
   continuous mixing. The beads were washed five times using RNA pull‐down
   wash buffer (25 mm Tris‐HCl, pH 7.4; 150 mm KCl; 5 mm EDTA; 5 mm MgCl₂;
   0.5% NP‐40), with each wash lasting 10 min. After the final wash, bound
   proteins were eluted by boiling the beads in SDS sample buffer at 95 °C
   for 5 min. The eluates were analyzed by western blotting or mass
   spectrometry to identify proteins interacting with lncRNA‐MEG3. All
   steps were performed under RNase‐free conditions, and experimental
   replicates were included to ensure the reliability of the results.

CUT&Tag Analysis

   The CUT&Tag libraries were sequenced on an Illumina NovaSeq 6000
   platform (Novogene, Tianjin, China) using a paired‐end read length of
   150 bp. Raw sequencing reads were processed with Fastp (default
   settings) to remove adapter sequences and low‐quality bases, and any
   reads shorter than 50 bp were excluded. The cleaned reads were then
   aligned to the GRCm38 reference genome using Bowtie2 (v2.5.4), ensuring
   a minimum mapping quality score of MAPQ ≥ 30, while retaining only
   uniquely mapped reads for further analysis. Peak calling was performed
   using MACS2 (v2.2.9.1), with a q‐value threshold of 0.01 to identify
   significant peaks. Differential peak analysis was performed using the
   DiffBind (v3.10.0) R package, incorporating DESeq2 for robust
   statistical analysis of differential binding events across conditions.
   For data visualization, aligned reads were converted to bigWig format
   using deepTools (v3.5.5) for IGV (Integrative Genomics Viewer)
   visualization. Additionally, heatmaps and metaplots of signal intensity
   across peaks were generated using deepTools to compare signal
   enrichment between different conditions. The identified peaks were
   annotated using ChIPseeker (v1.40.0) with the TxDb. Mmusculus.
   UCSC.mm10. known Gene annotation package to determine their genomic
   features and associated genes.

Chromatin Immunoprecipitation (ChIP) qRT‐PCR Assay

   Cells or tissues were fixed with 1% formaldehyde for 10 min at room
   temperature to crosslink protein‐DNA interactions, and the reaction was
   quenched by adding 125 mm glycine for 5 min. After washing with cold
   PBS, the cells or tissues were lysed in a lysis buffer containing
   protease inhibitors, and the nuclear fraction was collected. Chromatin
   was sheared to 200–1000 bp fragments using sonication, and the fragment
   size was verified by agarose gel electrophoresis. Equal amounts of
   chromatin were incubated overnight with H3K27me3 (ab6002, Abcam, USA)
   at 4 °C, with normal IgG (ab190475, Abcam, USA) as a negative control
   and input chromatin as a positive control. Pre‐washed protein A/G
   magnetic beads (PB101, Vazyme, China) were added, and the mixture was
   incubated at 4 °C for 1–2 h. The beads were washed sequentially with
   low‐salt and high‐salt wash buffers to remove nonspecific binding, and
   chromatin‐protein complexes were eluted using an elution buffer.
   Crosslinking was reversed by incubating the eluted samples with
   Proteinase K (20mg/ml) at 65 °C for 4–6 h or overnight. DNA was
   purified using a DNA extraction kit (G1N70, Sigma, Germany) and
   subjected to quantitative PCR (qPCR) analysis using primers specific to
   the target gene promoter region. Enrichment levels were calculated
   relative to input chromatin and normalized to the IgG control. The
   primers are listed in Table [290]S13 (Supporting Information).

Co‐Immunoprecipitation (Co‐IP)

   Co‐IP was performed according to the instructions of the Magnetic
   Co‐Immunoprecipitation (Co‐IP) Kit (Bes3011(S), BersinBio, China). In
   brief, C2C12 myoblasts were harvested by washing with PBS, and then
   lysed in lysis buffer containing protease inhibitors, incubating on ice
   for 10–20 min. The lysate was centrifuged to remove cell debris, and
   the supernatant was collected and incubated with Protein A/G agarose
   beads for 1–2 h. Specific antibodies were then added and incubated at
   4 °C for 2–4 h or overnight, followed by further incubation with
   Protein A/G agarose beads for an additional 2 h. The beads were washed
   3–5 times, and proteins were eluted using SDS‐PAGE loading buffer,
   followed by Western blot analysis.

Protein Expression and Purification

   SUZ12 protein expression was induced by the BL21 receptor state. Next,
   1 µL SUZ12 protein expression plasmid was transformed into 10 µL BL21
   receptive state. The transformed cells were plated on a
   kanamycin‐resistant culture plate, and monoclonal colonies were
   selected and inoculated into 5 mL of LB liquid medium containing
   kanamycin. The culture was incubated overnight at 37 °C in a shaking
   incubator. Next, 5 mL bacterial solution was inoculated into 400 mL LB
   liquid medium containing kanamycin and placed in a 37 °C constant
   temperature shaker at 200 rpm for 3 h. Next, 100 µL of 1 m IPTG was
   added to induce protein expression, followed by incubation in a shaker
   at 16 °C for 20 h. The protein of SUZ12 was then purified using the
   Tianren Protein purification kit HisPur Ni NTA kit (no.SA004K).

In Vitro Phase Separation Assay

   In vitro phase separation assay was performed in a storage buffer with
   indicated protein concentrations, and PEG8000 was also added to a final
   concentration of 10% (w/v). Phase separation assay was carried out on
   glass‐bottomed dishes (NEST), sealed with optically clear adhesive film
   to prevent evaporation and observed under a Nikon A1HD25 confocal
   microscope equipped with 60 × oil immersion objectives. The phase
   separation assay was performed in a physiological LLPS buffer (20 mm
   Tris‐HCl, pH 7.5, 15 mm NaCl, 130 mm KCl, 5 mm KH2PO4, 1.5 mm MgCl2,
   and 1 mg mL^−1 BSA).

SUZ12 Protein LLPS Detection

   In the salt concentration gradient assay, the protein was incubated
   with buffers containing 37.5, 300, and 500 mm NaCl at 20 °C for 30 min,
   and droplet formation was observed using Nikon A1HD25 confocal
   microscopy. The relative droplet area was quantified using ImageJ
   software. In the pH gradient assay, the protein was diluted in buffers
   with pH values ranging from 6.0 to 4.5 and incubated at 20 °C for
   30 min, followed by Nikon A1HD25 confocal imaging and quantitative
   analysis. Finally, in the temperature gradient assay, the protein was
   incubated at different temperatures (4, 20, and 37 °C) in a buffer
   containing 150 mm NaCl and pH 7.4, with phase separation monitored by
   Nikon A1HD25 confocal microscopy.

Fluorescence Redistribution After Photobleaching (FRAP)

   FRAP experiments were performed on a Nikon A1HD25 confocal microscope
   with a 60 × oil immersion objective. For the in vitro experiments,
   droplets were photobleached with 50% laser power for 0.5 s using 488 nm
   lasers. Time‐series images were acquired every 3 s after bleaching for
   5 min. For the in vivo experiments, FRAP assays were carried out on a
   Nikon A1HD25 confocal microscope at 37 °C in a live‐cell imaging
   chamber. Droplets were bleached with a 488‐nm laser pulse (50%
   intensity, 0.5 s). The recovery from photobleaching was recorded for
   the indicated time. Analysis of the recovery curves was carried out
   using ImageJ software.

Statistical Analysis

   For the wet‐lab experiments, statistical analysis was performed using
   GraphPad Prism 8 (GraphPad Software, San Diego, CA, USA). Data are
   presented as mean ± standard deviation (SD), and each statistical
   analysis included at least three biological replicates (n). A
   two‐tailed Student's t‐test was used to assess the statistical
   significance between the two groups, with tests showing p < 0.05
   considered statistically significant. Significance levels are indicated
   as follows: ns (not significant), ^* p < 0.05, ^** p < 0.01, ^*** p
   < 0.001.

Conflict of Interest

   The authors declare no conflict of interest.

Author Contributions

   Y.Y., C.Y., H.H., and S.W. contributed equally to this work. Z.L.T and
   Y.L.Y. conceived of and oversaw the project. Y.L.Y., S.LW., Y.S.Z.,
   D.Y.F., X.H., and Y.W.L performed the experiments, and H.B.H., C.Y.,
   and Y.L. analyzed the data. X.H. generated the mice. Y.L.Y. and C.Y.
   wrote the manuscript. J.Y.L., Y.C., and Y.Y.Z. conducted a muscle
   plasmid electroporation experiment. Z.L.T. and Y.L.Y. provided
   resources and funding for the investigation.

Supporting information

   Supporting Information
   [291]ADVS-12-2417715-s002.docx^ (10.3MB, docx)

   Supporting Information
   [292]ADVS-12-2417715-s001.xlsx^ (57.4MB, xlsx)

Acknowledgements