{"paper_id":"45f280a7-a509-4b05-8ca3-0fa4e867e161","body_text":"Mll4 in Skeletal Muscle Fiber Maintains Muscle Stem Cells by Regulating Notch Ligands | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article Mll4 in Skeletal Muscle Fiber Maintains Muscle Stem Cells by Regulating Notch Ligands Yea-Eun Kim, Sang-Hyeon Hann, Young-Woo Jo, Kyusang Yoo, Ji-Hoon Kim, and 2 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-5413133/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 23 Dec, 2024 Read the published version in Skeletal Muscle → Version 1 posted 9 You are reading this latest preprint version Abstract Background Muscle stem cells (MuSCs) undergo numerous state transitions throughout life, which are critical for supporting normal muscle growth and regeneration. Therefore, it is crucial to investigate the regulatory mechanisms governing the transition of MuSC states across different postnatal developmental stages. Methods To assess if myofiber-expressed Mll4 contributes to the maintenance of MuSCs, we crossed MCK Cre/+ or HSA MerCreMer/+ mice to Mll4 f/f mice to generate myofiber-specific Mll4 -deleted mice. Investigations were conducted using 8-week-old and 4-week-old MCK Cre/+ ; Mll4 f/f mice Investigations were conducted using 8-week-old and 4-week-old HSA Cre/+ ; Mll4 f/f mice were utilized. Results During postnatal myogenesis, Mll4 deleted muscles were observed with increased number of cycling MuSCs that proceeded to a differentiation state, leading to MuSC deprivation. This phenomenon occurred independently of gender. When Mll4 was ablated in adult muscles using the inducible method, adult MuSCs lost their quiescence and differentiated into myoblasts, also causing the depletion of MuSCs. Such roles of Mll4 in myofibers coincided with decreased expression levels of distinct Notch ligands: Jag1 and Dll1 in pubertal and Jag2 and Dll4 in adult muscles. Conclusions Our study suggests that Mll4 is crucial for maintaining MuSCs in both pubertal and adult muscles, which may be accomplished through the modulation of distinct Notch ligand expressions in myofibers. These findings offer new insights into the role of myofiber-expressed Mll4 as a master regulator of MuSCs, highlighting its significance not only in developmental myogenesis but also in adult muscle, irrespective of sex. Skeletal muscle Myofiber Muscle stem cells Myeloid/lymphoid or mixed-lineage leukemia 4 (Mll4) Exercise Notch signaling Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Figure 8 Background Muscle stem cells (MuSCs) are resident stem cells of skeletal muscle that contribute to muscle development, growth, and regeneration. They actively proliferate and differentiate into myocytes to contribute to myonuclei accretion and muscle growth [ 1 , 2 ]. In the adult stage, MuSCs enter a quiescent state and remain as reserve stem cells [ 3 , 4 ]. Upon injury, MuSCs become activated, providing myogenic cells to repair the muscle tissue. During muscle development and regenerative myogenesis, the MuSC niche regulates the dynamic transitions of MuSCs, including their activation, proliferation, differentiation, and self-renewal. Myofiber is an important cellular component of the MuSC niche. Unlike other cells that compose the MuSC niche, myofibers are in direct contact with MuSCs [ 5 ]. This enables them to regulate the MuSC state through both paracrine and contact-dependent juxtacrine signaling [ 6 ]. As paracrine signaling, myofibers secrete Wnt-4 to repress aberrant activation and maintain MuSC quiescence in adult homeostatic muscle [ 7 ]. FGF6 is another paracrine factor produced in myofibers to promote MuSC expansion during both developmental and regenerative myogenesis [8.9]. Myofibers also provide juxtacrine signals such as N-cadherin and M-cadherin to maintain MuSCs. These cell adhesion molecules are expressed at myofiber sites that are in direct contact with MuSCs to repress stem cell activation and maintain MuSC quiescence in adult muscles [ 10 ]. Among the various molecular signals derived from myofibers, Notch signaling plays a particularly crucial role in the maintenance of the stem cell pool and the regulation of cell fate decisions in MuSCs. In mammals, Notch signal-sending cells express Notch ligands (Dll1, 4 and Jag1, 2) and signal-receiving cells express Notch receptors (Notch1-4) [ 11 , 12 ]. Myofibers activate Notch signaling at different developmental stages primarily to generate a quiescent population of myogenic progenitor cells [ 2 , 13 ]. Several Notch ligands are expressed in myofibers to facilitate diverse Notch signaling to regulate the MuSC niche effectively at different developmental stages. Dll1, a Notch ligand mainly expressed in pubertal myofibers, interacts with the activated MuSCs to promote self-renewal, which is crucial for maintaining MuSCs [ 14 , 15 , 16 ]. In adult myofibers, Dll4 represses MuSC cell cycle entry, thus retaining the quiescent state of MuSCs [ 17 , 18 ]. While various studies have indicated that Notch signaling originating from myofibers has a role in regulating the state of MuSCs, there is limited understanding of whether there is a regulator present in myofibers that coordinates the expression of Notch ligands during the postnatal period. Mixed-lineage leukemia 4 (MLL4; also known as Kmt2d), a major H3K4 mono- and di-methyltransferase, is an essential histone modification enzyme for enhancer activation [ 19 , 20 , 21 ]. H3K4me1 marking by MLL4 is required for H3K27 acetylation and the recruitment of cell type-specific transcription factors [ 19 , 20 , 21 , 22 ]. Deletion of Mll4 results in the disturbance of H3K4me1 and H3K27ac on active enhancers, leading to defects in transcribing both newly activated genes, as well as genes that were already being expressed [ 19 , 20 , 21 , 22 ]. Recent study addressed that Mll4 depletion in myofibers turns off the slow type I fiber-specific genes, leading to fiber-type transition [ 23 ]. Myofibers are classified into two primary types: Type 1 (slow-twitch) and Type 2 (fast-twitch). Type 1 fibers rely on oxidative metabolism, which endows them with endurance capabilities. In contrast, Type 2 fibers utilize glycolytic pathways to generate rapid force [ 24 ]. This diversity in myofiber types contributes to muscle performance and adaptability to various stimuli [ 25 ]. Liu [ 23 ] reported that endurance exercise capacity was reduced in the Mll4-KO mice due to a slow-to-fast fiber-type shift. However, this study exclusively utilized male mice and focused solely on the function of Mll4 in developing myofibers. Building on these findings, our research aims to further investigate the role of Mll4 in myofibers across both sexes and various developmental stages. In this paper, we report that MLL4 in myofiber is required to maintain MuSCs during both the pubertal and adult stages, regardless of gender. Mouse models with myofiber-specific deletion of Mll4 exhibited reduced myofiber length, fewer myonuclei, and MuSC depletion. When Mll4 was ablated at the adult stage using an inducible method, MuSCs underwent differentiation into myoblasts, either with or without entering the cell cycle, leading to a depletion of adult MuSCs. Furthermore, the expression of specific Notch ligands at both pubertal and adult stages was significantly reduced in Mll4 -knockout myofibers. Together, our data suggest the importance of MLL4 in skeletal muscle fibers for maintaining the MuSC number, which might be achieved by affecting Notch ligand expression, in different postnatal periods. Methods Animals MCK Cre/+ (stock 006475), HSA MerCreMer/+ (stock 031934), and Mll4 f/f (stock 032152) mice were acquired from The Jackson Laboratory (Bar Harbor, ME, USA). The mice were backcrossed to C57BL/6 mice at least 6 times. To generate mice with myofiber-specific deletion of Mll4 , Mll4 f/f mice were crossed with MCK Cre/+ ( MCK Cre/+ ; Mll4 f/f – Mll4 ΔMCK – ). To develop myofiber-specific Mll4 conditional knockout mice using tamoxifen-inducible Cre, Mll4 f/f mice were crossed with HSA MerCreMer/+ ( HSA MerCreMer/+ ; Mll4 f/f – Mll4 ΔHSA – ). Following a breeding strategy from The Jackson Laboratory (Bar Harbor, ME, USA), we bred heterozygous Mll4 f/+ females with Cre-recombinase to homozygous Mll4 f/f males. Both male and female mice were used in the experiments. Mll4 ΔHSA mice used for experiments were adults, between 3–6 months of age. Control littermates lacking Cre-recombinase (Mll4 WT ) were utilized for analysis. All mouse lines were housed under controlled conditions with specific pathogen-free and handled according to the guidelines of the Seoul National University Institutional Animal Care and Use Committee (Protocol number: SNU-240103-3). Animal procedures Tamoxifen (Sigma-Aldrich) was dissolved in corn oil at a concentration of 20mg/ml. For tamoxifen-induced Cre recombination in Mll4 ΔHSA mice, both control and experimental mice were administered tamoxifen at a concentration of 150 mg/kg of mouse per day for five continuous days by intraperitoneal injection. For detection of cell cycle entry, 5-ethynyl-2’-deoxyuridine (EdU; Thermo Fisher Scientific) dissolved in sterile phosphate-buffered saline (PBS) was injected at a concentration of 40 mg/kg of mouse intraperitoneally daily. Muscle injury For BaCl 2 muscle injury, mice were anesthetized with 2.4% 2, 2, 2-Tribromoethanol (Avertin; Sigma-Aldrich) in PBS (240 mg/kg of mouse) and injected with 50 µl 1.2% BaCl 2 in saline (Sigma-Aldrich) to the tibialis anterior (TA) muscles. At 10 days after the injury, TA muscles were dissected, frozen in optimal cutting temperature compound (O.C.T.; Sakura Finetek) with liquid nitrogen, and stored at -80°C until analysis. Measurement of CSA distribution Laminin-stained section was imaged with EVOS FL Auto 2 (Thermo Fisher Scientific) with the same laser setting, exposure, and magnification. To measure CSA, semiautomatic muscle analysis using segmentation of histology (SMASH) was used with a segmentation filter (CSA between 200 µm2 and 6000 µm2; eccentricity ≤ 0.95; convexity ≥ 0.80). For injured muscles, CSA of regenerating myofibers with centralized nuclei was analyzed. The segmentation filter was adjusted as follows: CSA between 150 µm2 and 6000 µm2; eccentricity ≤ 0.95; convexity ≥ 0.80. Single myofiber isolation Single myofiber isolation was performed according to a previously reported protocol with modifications [ 47 ]. Dissected hindlimb extensor digitorum longus (EDL) muscles were enzymatically digested in Dulbecco’s modified Eagle’s medium (DMEM; Hyclone) containing 2.5% HEPES (Sigma-Aldrich) and collagenase II (800 units/mL, Worthington) at 37℃ for 60 min. Digested muscles were blocked in Dulbecco’s modified Eagle’s medium and 10% horse serum (Hyclone). The single myofibers were released by gentle trituration. Undamaged and noncontracted single myofibers were then washed with PBS several times and collected for immunocytochemistry and RNA extraction. Muscle stem cell (MuSC) isolation Isolation of MuSC was performed according to a previously reported protocol with modifications [ 48 ]. Limb muscles were dissected and mechanically dissociated in DMEM containing 10% horse serum, collagenase II (800 units/mL), and dispase (1.1 units/mL, Thermo Fisher Scientific) at 37℃ for 40 min. Digested suspensions were subsequently triturated by sterilized syringes with 20G 1/2 needle (BD Biosciences) and washed with DMEM to harvest mononuclear cells. Mononuclear cells were stained with anti-Sca-1-Pacific blue (Biolegend), anti-CD31-APC (Biolegend), anti-CD45–APC (Biolegend), and anti-Vcam1-Biotin (BD Biosciences). PE-Cy7- Streptavidin (Biolegend) was used as a secondary reagent. To exclude dead cells, 7-AAD (Sigma-Aldrich) was used. Stained cells were analyzed and Vcam1 + Sca1 − 7-AAD – CD31– CD45– MuSCs were isolated using FACS Aria III cell sorter (BD Biosciences) with 4-way-purity precision. FACS gating strategy was referred to the previously reported protocol [ 48 ]. Isotype control density plots were used as a reference for positive gating. Freshly isolated MuSCs were attached to a slide glass by cytospin for immunocytochemistry, or collected for RNA and protein extraction. Immunohistochemistry Freshly dissected TA or Soleus muscles were embedded in O.C.T., snap-frozen in liquid nitrogen, and stored at -80°C prior to sectioning. Cross-sectional 7 µm-thick sections were obtained from the embedded muscles using a cryostat. For myosin heavy chain (MyHC) staining, unfixed muscle sections were incubated overnight at 4°C with mouse anti-MyHC type 1 (DSHB, BA-D5, 1:10) or mouse anti-MyHC type 2x (DSHB, 6H-1, 1:5) in addition to rat anti-laminin (Abcam, ab11576, 1:1000 dilution) in 3% BSA blocking buffer. After washes in PBS, sections were incubated for 1 h with 1:500 dilution of Alexa Fluor 488-goat anti-mouse MIgG2b (Invitrogen), or Alexa Fluor 488-conjugated anti-mouse IgM (Invitrogen), and Alexa Fluor 594-conjugated anti-rat IgG (Invitrogen). For staining myoblasts, sections were fixed in 4% paraformaldehyde (PFA) for 10 min, and washed in PBS. Antigen retrieval was then performed in citrate buffer (10 mM citric acid, pH 6) at 95°C 15 min. The sections were blocked by mouse Ig blocking reagent and blocking buffer from M.O.M. Kit (Vector Laboratories), according to the manufacturer’s protocol. Then, the sections were incubated with primary antibodies in the blocking buffer at 4°C overnight. The primary antibodies used include mouse anti-Pax7 (1:100, DSHB), rabbit anti-Ki67 (1:500, Sigma-Aldrich), mouse anti-MyoG (1:100, DSHB), rabbit anti-Dystrophin (1:500, Abcam), rabbit anti-Laminin (1:500, Sigma-Aldrich), and rabbit anti-cleaved Caspase-3 (1:200, Cell Signaling Technologies). After washing the sections with PBS, the sections were stained with secondary antibodies for 1 hr at RT, washed, and mounted. The secondary antibodies were used at a concentration of 1:400 and include goat anti-Rabbit IgG-Alexa Fluor 488 (Thermo Fisher Scientific), goat anti-Rabbit IgG-Alexa Fluor 594 (Thermo Fisher Scientific), and goat anti-Mouse IgG-Alexa Fluor 594 (Thermo Fisher Scientific). Hoechst 33342 (1:2,000, Thermo Fisher Scientific) was used to visualize nuclei. For EdU staining, we used the Click-iT EdU Alexa Fluor 488 Imaging Kit (Thermo Fisher Scientific) following the manufacturer’s protocol before the blocking step. The number of each cell type and myofibers was counted in the total TA or Soleus area, and representative images were selected in the same region of the section used in the cell counting. Imaging was conducted with EVOS FL Auto 2 (Thermo Fisher Scientific). Immunocytochemistry For isolated myofibers staining, freshly isolated myofibers were fixed in 4% PFA for 10 min at room temperature (RT), quenched in 0.1M glycine in PBS for 10 min at RT, and blocked for 1 hour at RT by blocking buffer (5% goat serum and 5% bovine serum albumin in PBS/0.4% Triton X-100). Then, the myofibers were incubated with mouse anti-Pax7 (1:100, DSHB) in the blocking buffer at 4°C overnight. After washing the myofibers three times with PBS/0.1% Triton X-100, the myofibers were stained with goat anti-Mouse IgG-Alexa Fluor 594 (1:400, Thermo Fisher Scientific) and Hoechst 33342 (1:5,000, Thermo Fisher Scientific) for 1 hour at RT, washed and mounted on slide glass. Imaging was conducted with EVOS FL Auto 2 (Thermo Fisher Scientific). For isolated MuSCs staining, freshly isolated MuSCs were attached to a slide glass by cytospin and fixed by 4% PFA for 10 min at RT. The fixed MuSCs were washed with PBS/0.4% Triton X-100 several times and blocked with blocking buffer (5% goat serum and 5% bovine serum albumin in PBS/0.4% Triton X-100) for 1 hour at RT and incubated with mouse anti-Pax7 (1:100, DSHB) and rabbit anti-MyoD (1:200, Santa Cruz) overnight at 4°C. The slides were washed with PBS/0.1% Triton X-100 several times and incubated with goat anti-Mouse IgG-Alexa Fluor 594 (1:400, Thermo Fisher Scientific) and goat anti-Rabbit IgG-Alexa Fluor 488 (1:400, Thermo Fisher Scientific). For EdU staining, we used the Click-iT EdU Alexa Fluor 647 Imaging Kit (Thermo Fisher Scientific) following the manufacturer’s protocol before the blocking step. The slides were counterstained with Hoechst 33342 (Thermo Fisher Scientific) and mounted. Imaging was conducted with EVOS FL Auto 2 (Thermo Fisher Scientific). Four limb grip strength measurement Grip strength was assessed by using a grip strength test meter (grip strength test BIO-GS3, Bioseb). Mice were allowed to grasp a grid attached to the tester with 4 limbs and were manually pulled in a horizontal direction by the tip of the tail. The test was performed 5 times with 10 min of resting between each measurement. The average of the top 3 result value (N, Newton) was normalized to body weight (g) (N/g). All experiments were performed in a blinded fashion. Chronic exercise training and endurance running test Randomized mice were pre-acclimated to the treadmill (DJ2-242, Dual Treadmill, Daejeon, Korea) before training. The scheme consists of exploration (0m/min for 5 min), and subsequent running (5m/min for 5 min, 10m/min for 5 min, 15m/min for 5 min). After 3 days of acclimation, mice were subjected to chronic exercise training for 5 weeks, 5 days per week with a protocol of 5m/min for 5min, 10m/min for 5min, 15m/min for 30 min. To test endurance running capacity, mice were allowed to run until exhaustion with speed set to 10m/min for 30 min and incremented by 2m/min every 20 min with no inclination. Exhaustion was defined as the condition where mice remained stationary at the end of the treadmill for more than 10 seconds despite mechanical stimulation. All experiments were performed in a blinded fashion. RNA extraction and quantitative real-time polymerase chain (qRT-PCR) Total RNA was extracted from freshly isolated myofibers and MuSCs using a TRIzol Reagent (Life Technologies) and analyzed by qRT-PCR. First-strand complementary DNA was synthesized from 1 µg of RNA using ReverTra Ace (Toyobo) containing random oligomer according to the manufacturer’s instructions. qRT-PCR (Qiagen) was performed with SYBR Green technology (SYBR Premix Ex Taq, Qiagen) using specific primers against indicated genes. Relative mRNA levels were determined using the 2 −ΔΔCt method and normalized to Gapdh . Primers are listed in Supplementary Table 1. Western blot Dissected TA muscles were homogenized in RIPA buffer (50 mM Tris-HCl, pH 7.5, 0.5% SDS, 20 µg/ mL aprotinin, 20 µg/ mL leupeptin, 10 µg/ mL phenylmethylsulfonyl fluoride, 1 mM sodium orthovanadate, 10 mM sodium pyrophosphate, 10 mM sodium fluoride, and 1 mM dithiothreitol). Cell lysates were centrifuged at 13,000 rpm for 15 min. Supernatants were collected and subjected to immunoblot. BCA protein assay (Thermo Fisher Scientific) was used for estimating total protein concentrations. Normalized total proteins were analyzed by electrophoresis in 12% polyacrylamide gels and transferred to PVDF membranes (Millipore). Membranes were blocked in 5% skim milk (BD Biosciences) in TBS with 0.1% Tween-20 and incubated with rabbit anti-Histone H3 (Cell Signaling Technology) and rabbit anti-H3K4me1 (Cell Signaling Technology) overnight at 4°C. After incubation with goat anti-Rabbit IgG-HRP (Promega), the membranes were developed using a Fusion solo chemiluminescence imaging system (Vilber). Histone H3 was used as a loading control. Primary and secondary antibodies were diluted 1:1000 and 1:10000 with PBS containing 0.1% Tween-20 and 3% BSA, respectively. Statistical analysis Sample size determination was based on anticipated variability and effect size that was observed in the investigator’s lab for similar experiments. For quantification, individuals performing the counts were blinded to sample identity and randomized. All statistical analyses were performed using GraphPad Prism 9 (GraphPad Software). For comparison of significant differences in multiple groups for normally distributed data, statistical analysis was performed by one-way or two-way ANOVA followed by Tukey’s pairwise comparison post hoc test. For non-normally distributed data, Brown–Forsythe and Welch ANOVA followed by the Games-Howell multiple comparisons test was used. For the comparison of the two groups, Student’s unpaired t-test assuming a two-tailed distribution with Welch’s correction was used. Unless otherwise noted, all error bars represent s.e.m. The number of biological replicates and statistical analyses for each experiment were indicated in the figure legends. Independent experiments were performed at least in triplicates. Results Mll4 deletion in myofiber alters CSA and slow fiber composition in males, but not in female mice Previously, Liu and colleagues [ 23 ] reported that the deletion of Mll4 in developing myofibers did not impact overall muscle mass but resulted in changes to muscle characteristics, including an increased cross-sectional area (CSA) and a shift from slow to fast fiber types. However, the results were derived exclusively from male mice. Given that Mll4 is expressed in both sexes (Supplementary Fig. 1A), we investigated whether female mice would exhibit a similar phenotype to males following Mll4 ablation. We crossed Mll4 f/f mice that carry loxP sites of the Mll4 gene [ 19 ], with MCK Cre/+ mice [ 26 ], which produced Mll4 ΔMCK mice ( MCK Cre/+ ; Mll4 f/f ) [ 23 ]. By performing quantitative real-time PCR (qRT-PCR), the deletion of Mll4 was confirmed in 8-week-old Mll4 ΔMCK myofibers (Supplementary Fig. 1B-C). Muscle weight remained unchanged in 8-week-old Mll4 ΔMCK mice, regardless of sex (Supplementary Fig. 1D-E). As reported [ 23 ], male Mll4 ΔMCK mice displayed increased CSA (Fig. 1 A) and fiber type shifts toward a decreased ratio of slow-twitch fiber in soleus muscle (Fig. 1 C-E). In contrast, no significant differences in these characteristics were observed in Mll4 ΔMCK female mice compared to Mll4 WT female mice (Fig. 1 B, 1 C, and 1 F-G), suggesting that the alterations in CSA and fiber type composition following the deletion of Mll4 in myofibers may represent a phenotype selectively characteristic of male musculature, potentially attributable to male-specific factors or microenvironments. MuSC depletion in adult Mll4 ΔMCK mice Unexpectedly, when isolating single myofibers to prove the ablation of Mll4 in Mll4 ΔMCK mice (Fig. 2 A), we observed that myofiber length was significantly shorter in both male and female Mll4 ΔMCK mice compared to those of controls (Fig. 2 B and 2 E). Since shortened myofibers may be due to a reduced number of myonuclei [ 27 ], we quantified myonuclear number, which was markedly decreased in Mll4 ΔMCK EDL myofibers compared to the controls (Fig. 2 C and 2 F). To assess whether myonuclear density was also reduced, we calculated the ratio of myonuclear number to myofiber length. This analysis revealed a decrease in myonuclear density in Mll4 ΔMCK myofibers (Fig. 2 D and 2 G). During postnatal development in mice, myofiber length and myonuclear number increase rapidly through myoblast fusion until puberty and then become relatively stable at the adult stage when postnatal myogenesis ceases [ 1 ]. Thus, reduced myonuclear density in adult Mll4 ΔMCK mice might be due to impaired myoblast fusion, defective MuSC differentiation, or even MuSC depletion. To address this issue, we first assessed the number of MuSCs in Mll4 -deleted myofibers (Fig. 2 H). Intriguingly, Pax7 + MuSCs greatly decreased in the myofibers of both male and female 8-week-old Mll4 ΔMCK mice (Fig. 2 I-J). Our data suggest that MuSC depletion in Mll4 -lacking myofibers during postnatal myogenesis resulted in decreased myonuclei accretion and myofiber growth, and that Mll4 in myofiber may play an important role in maintaining the MuSC population, irrespective of gender. MuSC depletion in Mll4 deleted muscle during postnatal muscle growth MuSCs that have actively proliferated during the juvenile stage enter a quiescent state at puberty to establish a reserve stem cell pool in adult muscles [ 2 ]. To investigate if the deletion of Mll4 in myofibers affects MuSC number during postnatal myogenesis, we conducted a histological analysis to quantify Pax7-positive cells in TA muscles during and after postnatal myogenesis. Considering that MCK-Cre mediated ablation of Mll4 occurs after 7 days of birth [ 23 ], 0-week-old perinatal muscles were expected to have comparable MuSC numbers between Mll4 WT and Mll4 ΔMCK muscles. The number of MuSCs remained consistent between the control and Mll4 ΔMCK muscles until 2 weeks of age (Fig. 3 D). However, a decrease of Pax7 + MuSCs was prominent in the pubertal 4-week-old Mll4 ΔMCK TA muscle, with a further decline noted in the 8-week-old muscle (Fig. 3 A and 3 D). This indicates that the deletion of myofiber-specific Mll4 disrupts the MuSC number during postnatal myogenesis, resulting in a depletion of the population in adult muscles. TA and EDL muscles primarily consist of fast-twitch type 2 fibers [ 28 ]. To examine if MuSC depletion occurs in slow-twitch type 1 fiber-rich muscles, the soleus muscle was analyzed. Undoubtedly, the Pax7-positive MuSC number was reduced in the pubertal 4-week-old muscles and further diminished in the adult 8-week-old soleus muscles of Mll4 ΔMCK mice (Fig. 3 B and 3 E). During puberty, cycling MuSCs exit the cell cycle and contribute to quiescent MuSC populations [ 2 ]. To test if the reduced MuSC in Mll4 ΔMCK mice was due to impaired cell cycle exit in cycling pubertal MuSCs, we quantified Ki67-positive MuSCs in the 4-week-old Mll4 ΔMCK mice. Compared to the control, Mll4 ΔMCK mice showed a twofold increase in proliferating Ki67 + Pax7 + MuSCs (Fig. 3 C and 3 F). To investigate if these proliferating MuSCs enter the differentiation state, we isolated MuSCs via cytometry (Supplementary Fig. 2A-B) and quantified MyoD-positive MuSCs. Compared to the Mll4 WT mice, Mll4 ΔMCK mice showed an increase in MyoD-positive MuSCs (Fig. 3 H-I). Furthermore, to label cycling MuSCs, EdU was treated for 2 consecutive days before isolating MuSCs (Fig. 3 G). Notably, the number of MyoD-positive cells also increased among cycling MuSCs (Edu + Pax7 + ) in Mll4 ΔMCK mice (Fig. 3 J). This indicates that while normal MuSCs exit the cell cycle and enter a quiescent state during puberty, MuSCs in Mll4 ΔMCK mice maintain their cell cycle and differentiate into committed myoblasts. Altogether, the deletion of Mll4 in myofibers leads to the loss of MuSCs during postnatal muscle growth. Deletion of Mll4 in adult myofibers does not alter muscle CSA and fiber type composition Following postnatal myogenesis, the adult muscle tissue reaches a steady state characterized by the cessation of myofiber growth and the entry of MuSCs into a quiescent phase. To investigate if induced ablation of Mll4 in muscles after postnatal myogenesis would affect myofiber maintenance, we examined muscle features such as CSA distribution and fiber type composition of Mll4 ablated adult muscles. Adult HSA MerCreMer/+ ; Mll4 f/f mice (Mll4 ΔHSA ) were treated with tamoxifen for consecutive 5 days to induce deletion of the Mll4 gene in myofibers (Fig. 4 A). This resulted in the ablation of the Mll4 gene in the myofibers of Mll4 ΔHSA mice after 2 weeks of tamoxifen administration (Fig. 4 B). The histological analysis showed that CSA distribution, fiber number, and fast-twitch (MyHC2x) fiber composition in the TA muscles of Mll4 ΔHSA mice did not change following 2 weeks (Fig. 4 C-E, Supplementary Fig. 3A) and even 4 weeks (Fig. 4 H-J, Supplementary Fig. 3B) of Mll4 ablation, compared to the control mice. Similarly, in the soleus muscle, the composition of slow-twitch (MyHC1) fibers also remained unchanged after both 2 weeks (Fig. 4 F-G, Supplementary Fig. 3A) and 4 weeks (Fig. 4 K-L, Supplementary Fig. 3B) of Mll4 ablation in Mll4 ΔHSA mice. This indicates that the induced deletion of Mll4 in adult muscles does not affect myofiber maintenance and intracellular features such as fiber CSA and fiber type composition regardless of gender. Mll4 deletion in myofibers does not affect exercise capacity Since MLL4 functions as an enhancer activator [ 19 ], its deletion may disrupt various gene transcription processes. Although the deletion of Mll4 in adult muscles does not impact myofiber characteristics such as CSA and fiber type composition, we investigated whether the deletion affects exercise capacity. To test this, tamoxifen-treated mice were accustomed to chronic exercise training for 5 weeks (hereafter, Mll4 WT − EX and Mll4 ΔHSA−EX ) (Fig. 5 A). The protocol of the chronic exercise provides prolonged contractions of muscles, promoting adaptations such as increased muscle mass while minimizing exercise-induced muscle damage [ 29 ]. In TA muscles, CSA distribution and fast fiber composition remained consistent between Mll4 WT − EX and Mll4 ΔHSA−EX mice. In addition, slow fiber composition was unchanged in Mll4 ΔHSA−EX soleus muscle (Fig. 5 B-F, Supplementary Fig. 4A). To assess whether exercise capacity was affected by Mll4 deletion, we measured grip strength and endurance running capability. In line with the observed similarity in CSA and fiber type composition, Mll4 ΔHSA−EX mice showed comparable grip strength (Fig. 5 G) and endurance running capability (Fig. 5 H-I) relative to the control group. To investigate whether the expression of genes related to fiber type and metabolism was altered in Mll4 ΔHSA−EX mice, we conducted qRT-PCR analyses. Transcriptional profiling revealed that before exercise training, the genes were generally downregulated in Mll4 ΔHSA mice (Supplementary Fig. 5A-B). Notably, no particular gene exhibited higher expression levels that could induce a shift in fiber type composition or metabolic activity. After 5 weeks of chronic exercise, the expression of genes related to fiber type and metabolism of Mll4 ΔHSA−EX mice also showed overall downregulation, with the exception that certain slow-twitch muscle genes were marginally upregulated (Supplementary Fig. 5C). Collectively, the results suggest that the induced knockout of Mll4 in myofibers at the adult stage does not influence exercise capacity or muscle characteristics, such as CSA and fiber type composition, even after physiological exercise stimulation. Loss of adult MuSCs in Mll4 ΔHSA mice To investigate whether the deletion of Mll4 in adult myofibers disturbs the quiescence of MuSCs, TA and soleus muscles were analyzed to quantify Pax7-positive MuSCs. Mll4 ΔHSA mice showed depletion of MuSCs in both muscles (Fig. 6 A and 6 C). To assess whether the loss of adult quiescent MuSCs in Mll4 ΔHSA mice is associated with their entry into the cell cycle, we quantified Ki67-positive MuSCs in the TA muscles. In Mll4 WT TA muscles, there was a negligible presence of Ki67 + Pax7 + cells, whereas Mll4 ΔHSA muscles showed an increase of Ki67 + Pax7 + cells (Fig. 6 D and 6 E). This suggests that ablation of Mll4 in myofibers at the adult stage causes MuSCs to exit quiescence and enter the cell cycle, leading to the depletion of MuSCs. To examine if the cell cycle entry of adult Mll4 ΔHSA mice leads to a differentiation state, we performed an immunocytochemistry assay on sorted MuSCs (Supplementary Fig. 2C-D) to quantify MyoD-positive cells. Compared to the Mll4 WT MuSCs, Mll4 ΔHSA MuSCs showed an increased number of MyoD-expressing cells (Fig. 6 G-H). For detecting MuSCs that entered the cell cycle, we treated EdU for 2 weeks in Mll4 ΔHSA mice (Fig. 6 F). This long-term EdU labeling method was applied to identify the scarcely dividing MuSCs in adult muscle tissue [ 30 ]. We found that the population of MyoD-expressing cells among EdU-positive, dividing MuSCs was also increased in Mll4 ΔHSA muscles (Fig. 6 I). On the other hand, given that adult quiescent MuSCs can differentiate without entering the cell cycle [ 30 ], we also quantified MyoD-expressing cells among EdU-negative, non-dividing MuSCs. Interestingly, Mll4 ΔHSA MuSCs showed an increased number of MyoD-positive cells among EdU-negative MuSCs (Fig. 6 J). This indicates that the deletion of Mll4 in adult muscles causes adult MuSCs to lose their quiescence and undergo differentiation, either with or without dividing. To test whether the differentiated myogenic progeny of Mll4 ΔHSA muscles fuse into myofibers, we conducted a histological analysis and quantified EdU-positive nuclei located on the inner side of dystrophin structure [ 30 ] (Fig. 5 K). While the number of fiber-incorporating EdU-positive nuclei was extremely low in control muscles, it was sevenfold higher in Mll4 ΔHSA muscles compared to the control group (Fig. 5 L). This suggests that Mll4 deficiency in adult muscles causes MuSCs to exit quiescence and undergo differentiation, with at least some, or perhaps all, of these differentiated cells subsequently fusing into myofibers. This eventually results in severe MuSC loss in adult muscles. Lack of Mll4 impairs muscle regeneration following injury MuSCs are the primary cell type that contributes to muscle regeneration capacity. To investigate if MuSC depletion in Mll4 ΔHSA mice leads to hindered muscle regeneration, we subjected TA muscles of Mll4 WT and Mll4 ΔHSA mice to injury using BaCl 2 (hereafter, Mll4 WT − inj and Mll4 ΔHSA−inj ) (Fig. 7 A). Muscles were analyzed 10 days post-injury, as the majority of regenerating fibers are restored [ 31 ]. TA muscles of Mll4 ΔHSA−inj mice showed reduced muscle mass compared to that of Mll4 WT − inj mice, while adjacent muscles such as the EDL, GA, and soleus remained unaffected (Fig. 7 B). Histological analysis of muscle sections revealed active muscle regeneration in Mll4 WT − inj muscles, as indicated by the predominance of myofibers with centrally located nuclei and relatively homogenous fiber sizes. Conversely, Mll4 ΔHSA−inj muscle was observed with disorganized tissue architecture with residual damaged fibers that failed to undergo effective regeneration. (Fig. 7 C). Moreover, Mll4 ΔHSA−inj mice showed reduced CSA of regenerating fibers (Fig. 7 D). These results suggest that lack of MuSCs due to the deletion of myofiber-specific Mll4 resulted in severely impinged muscle regeneration capacity. Mll4 in myofibers affects the MuSC niche by regulating Notch ligand expression To explore how Mll4 in myofibers may have affected the MuSC niche, we screened for downstream effector candidate genes by analyzing public datasets. For one, we analyzed data curated by Liu et al. [ 23 ]. This provided a list of downregulated genes in muscle from MLL4-SET-knockout (KO) mice, where the enzymatic SET domain of MLL4 is ablated, compared to control mice. In addition, we analyzed data from Lee et al. [ 19 ], to obtain the list of downregulated genes in cultured, differentiated Mll4-KO myocytes versus control. Sixty-six genes were identified as commonly downregulated from the two datasets. Since myofiber can directly regulate the MuSC niche via signaling through ligand-receptor interactions [ 32 ], we then identified ligands from the 66 candidate genes by comparing them to the mouse ligand database. Consequently, 5 genes were identified as ligand-coding genes that are downregulated by Mll4 KO in both whole muscle and differentiated myocytes. To our surprise, the Notch ligand Jag2 was among the 5 candidate genes. Also, Dll1, another Notch ligand, was identified as downregulated in Mll4 KO myocytes. (Fig. 8 A). For MuSCs, Notch signaling is a major signaling pathway that maintains the stem cell pool. When the Notch downstream effector Rbpj is deleted in adult MuSCs, which are predominantly in a quiescent state, they exit the quiescent state and undergo aberrant differentiation [ 30 , 33 ]. Myofiber-specific deletion of Dll4, a Notch ligand that is mainly expressed in adult myofibers, induces premature differentiation of MuSCs, resulting in a reduced number of stem cells [ 17 , 34 ]. These studies suggest that the maintenance of MuSC quiescence is highly dependent on Notch signaling between MuSCs and myofibers. Considering that the depletion of MuSCs was observed in both Notch signaling-reduced MuSCs and myofiber-specific Mll4- deleted (Mll4 ΔMCK and Mll4 ΔHSA ) MuSCs, we sought to validate the downregulation of Notch ligands in Mll4 ΔMCK and Mll4 ΔHSA mice. A previous study found that Dll4 and Jag2 are dominantly expressed in adult myofibers, compared to other notch ligands [ 17 ]. In addition, we reported that myofibers of 4-week-old mice exhibited robust expression of Dll1 and Jag1 proteins [ 2 ]. Considering that Notch ligands have a fluctuating expression pattern in muscles throughout the developmental stages, we compared mRNA quantity for Notch ligands in the myofibers of wild-type pubertal 4-week-old and adult mice (Fig. 8 B). Interestingly, genes having dominant expression during each time point correlated with genes that were downregulated due to Mll4 depletion in muscle fibers. While expression of major Notch ligands of pubertal 4-week-old myofibers – Jag1 and Dll1 – decreased in 4-week-old Mll4 ΔMCK myofibers (Fig. 8 C), the primary Notch ligands of adult myofibers – Jag2 and Dll4 – decreased in adult Mll4 ΔHSA myofibers (Fig. 8 D). In other words, Mll4 insufficiency in myofibers disturbed Notch ligand expression that is dominant in each pubertal or adult muscle. To test if Notch signaling is indeed reduced in 4-week-old Mll4 ΔMCK and adult Mll4 ΔHSA MuSCs, the mRNA levels of canonical Notch effectors – HeyL, Hey1 , and Hes1 – were quantified via qRT-PCR. As expected, the overall expression of genes mentioned above was downregulated in both Mll4 ΔMCK and adult Mll4 ΔHSA MuSCs (Fig. 8 E-F). Considering the molecular feature of MLL4, we analyzed public ChIP-seq data of MLL4 in myocytes [ 19 ], to examine whether it may modulate the transcription of Notch ligands on the chromosomal level. This revealed the genomic binding of MLL4 on Dll1 and Jag2 gene loci, where Mll4 deletion reduced H3K4me1 and H3K27ac levels on enhancers for both Dll1 and Jag2 genes (Fig. 8 G-H). This implicates the possibility of MLL4 directly regulating the induction of different Notch ligand gene expressions. Taken together, MLL4 can control diverse Notch ligand expression in myofibers, which is necessary for regulating the MuSC niche during and after postnatal myogenesis. Discussion Skeletal muscle has a resilient characteristic due to its resident stem cell populations. Thus, uncovering the mechanism of regulating MuSC fate is crucial for understanding the biological process of developmental and regenerative myogenesis. In this paper, we explored the role of Mll4 in myofibers regarding the MuSC state regulation and discovered that myofiber-expressed Mll4 is important for maintaining MuSCs in both muscles during and after postnatal myogenesis. In the pubertal Mll4 ΔMCK muscle, lack of Mll4 in myofibers resulted in increased population of differentiating myogenic cells, leading to a decrease of MuSCs. Furthermore, induced ablation of Mll4 in adult myofibers resulted in the quiescence exit of MuSCs, which also caused dramatic depletion of MuSCs. During postnatal myogenesis, juvenile MuSCs constantly proliferate for muscle development [ 1 , 2 ]. This proliferating cell population decreases due to cell cycle exit during puberty to establish a reserve pool of quiescent MuSCs in adult muscles [ 3 , 4 ]. Our findings indicate that Mll4 in myofibers are critical for maintaining MuSCs in pubertal muscles, where cycling MuSCs begin to enter quiescence, as well as in adult muscles, where MuSCs remain in a quiescent state. This suggests that Mll4 plays a critical role in myofibers by creating a microenvironment that supports the maintenance of a healthy population of MuSCs in muscle tissue. The ability to maintain an adequate number of MuSCs is crucial regardless of gender and age. In this study, we elucidate two critical properties of Mll4 in preserving the stemness of MuSCs. First, Mll4 regulates the MuSC number in both sexes. Skeletal muscle exhibits sexual dimorphism in terms of mass, fiber type composition, and contractility attributed to variations in gene expression and hormonal profiles between genders [ 35 , 36 ]. These differences may have contributed to the disparate muscle phenotypes after the deletion of myofiber- Mll4 in male and female mice (Fig. 1 C-I). However, gender did not influence the extent of MuSC depletion resulting from Mll4 ablation. Secondly, Mll4 regulates MuSC quiescence across different developmental stages, including both during and after postnatal myogenesis. Previous research indicated that myofiber-specific deletion of Mll4 led to a slow-to-fast fiber type shift [ 23 ]. However, our findings reveal that this phenotype is not present following Mll4 deletion in adult muscle tissue. This suggests that Mll4 may play a role in the development of myofibers, but not in their maintenance during adulthood, at which developmental myogenesis is complete. Indeed, it is reported that during developmental myogenesis, Foxo/Notch signaling regulates fiber type specification, leading to a reduction in slow fibers and an increase in fast fibers when disrupted in muscle [ 37 ]. Considering that (1) this phenotype is in line with that of Mll4 -mKO mice, as reported previously, and (2) Mll4 has the possibility of regulating the gene expression of Notch ligands, the deletion of Mll4 might have disrupted Notch signaling in developing myofibers, leading to aberrant fiber type specification. Altogether, our data provide new insights into the role of Mll4 as a crucial regulator of the MuSC quiescence, highlighting its significance across developmental stages and irrespective of gender. Chromatin modification of enhancers within myofibers can modulate the expression of extracellular matrix (ECM) components or growth factors, thereby indirectly influencing the MuSC niche [ 38 , 39 ]. Our study suggests that MLL4, an enhancer activator, regulates Notch ligand expression in myofibers to directly control MuSC quiescence. By analyzing and validating transcriptome databases from previous studies, we verified that the expression of Notch ligands was downregulated in Mll4-KO myocyte. Moreover, an analysis of ChIP-seq data revealed genomic binding of MLL4 on Notch ligand gene loci. The downregulation of Notch ligands in myofibers led to a reduced expression of canonical Notch target genes in MuSCs. This indicates that MLL4 can regulate the signaling pathway that affects adjacent cells. This finding is particularly intriguing given that MLL4 has primarily been studied as a critical factor for activating intracellular signaling pathways, including those related to cancer and cell fate determination [ 19 , 20 , 40 , 41 , 42 ]. Specifically, in myofibers, Mll4 was reported to activate the transcription of slow-twitch genes [ 23 ]. By inspecting the physiological impact of Mll4 deletion in myofibers on MuSCs, we revealed that Mll4 regulates not only intracellular signaling pathways, as previously reported, but also signaling pathways that affect adjacent cells, by controlling expressions of ligand genes. This underscores the importance of exploring the potential gene-regulating activity of MLL4, which may impact other cellular processes, such as differentiation and tumorigenesis, in neighboring cells. It has been well-established that Notch signaling is a fundamental pathway regulating the MuSC niche in prenatal and postnatal muscles to maintain an appropriate stem cell pool [ 11 , 12 ]. Previous studies reported that the myofiber is an important source of Notch ligands, sending signals to MuSCs to control their niche and hence their cell fate [ 2 , 13 , 43 ]. Notch ligands have distinct expression patterns in myofibers during development, affecting the MuSC niche in different ways. In this study, we investigated the Notch ligands with prevailing expression in different stages; Jag1 and Dll1 in pubertal myofibers, and Jag2 and Dll4 in adult muscle fibers. Interestingly, Mll4 deletion in pubertal and adult myofibers disturbed the expression of Notch ligands that were principally expressed in each stage. Previously, Eliazer and colleagues reported that myofiber-specific deletion of Dll4 resulted in a reduction of MuSCs. However, the decrease in MuSCs was more pronounced when the pan-Notch regulator Mib1 was deleted from myofibers [ 17 ]. This implies the presence of a complementary Notch ligand acting as a signaling factor to maintain MuSC quiescence in adult muscles, together with Dll4. Our findings suggest that along with the well-known factor Dll4, Jag2 may be another Notch ligand in adult myofibers contributing to maintaining the Pax7 + quiescent stem cells, both of which were found to be regulated by Mll4. Taken together, this study suggests that MLL4 functions as a regulator that modulates the expression of various Notch ligands in myofibers during both pubertal and adult stages. This regulation is essential for the precise control of MuSC quiescence throughout developmental stages. Mll4 deletion notably hindered H3K4me1 and H3K27ac levels on enhancers for both the Dll1 gene in pubertal fibers and the Jag2 gene in adult fibers, indicating different gene regulation of MLL4 in the two developmental stages. This may be attributed to the distinct pioneer transcription factors that recruit MLL4 to induce Notch ligand expression at different developmental stages. Several transcription factors – such as CCAAT/enhancer-binding protein family, myocyte enhancer factor 2 family, and Nrf1 – are identified to bind the DNA to recruit the MLL4 complex [ 19 , 21 , 23 , 44 ]. Depending on the cell type and differentiation stage, different transcription factors recruit MLL4 to regulate the expression of various genes. Transcription factors associated with Notch signaling have also been identified. A study on chicken embryos found that a transcription coregulator, Yap, binds to the enhancer of Jag2 [ 45 ]. In mice, Notch1 ICD can act as a transcription activator in muscle fibers to activate the gene expression of Jag2 and Dll4 [ 46 ]. Following these studies, it is plausible that Yap and Notch ICD may recruit MLL4 to regulate the expression of different Notch ligands in myofibers. Further investigation is required to elucidate the molecular mechanism by which MLL4 regulates Notch ligand gene expression in myofibers. This includes identifying the specific pioneer transcription factors that interact with MLL4 across different developmental stages. Conclusions Our results suggest a unique function of Mll4 in myofibers controlling MuSC state, possibly by orchestrating different Notch ligand expressions in various developmental stages. Moreover, despite skeletal muscle being known to exhibit sexual dimorphism, the role of Mll4 regulating MuSCs was valid in both male and female mice. By elucidating an additional mechanism governing MuSC maintenance, this research opens new avenues for the biological manipulation of muscle stem cells. Abbreviations MuSCs Muscle stem cells Mll4 Myeloid/lymphoid or mixed-lineage leukemia 4 Jag1, Jag2 Jagged 1, Jagged 2 Dll1, Dll2 Delta-like protein 1, Delta-like protein 2 TA Tibialis anterior Sol Soleus GA Gastrocnemius EDL Extensor digitorum longus CSA Cross-sectional area MyHC1 Myosin Heavy Chain 1 MyHC2x Myosin Heavy Chain 2x Declarations Ethics approval and consent to participate The care and treatment of animals in this study were approved by the Institutional Animal Care and Use Committee (IACUC) protocols (SNU-240103-3) of Seoul National University. Consent for publication Not applicable Availability of data and materials All data generated or analyzed during this study are included in the published article and are available from the corresponding author upon reasonable request. Competing interests The authors declare that they have no competing interests. Funding This work was supported by NRF-2022R1A2C3007621 (Y.Y.K.), NRF-2020R1A5A1018081 (Y.Y.K.), and R01 NS118748 (S.-K.L. and J.W.L.). Author’s Contributions Conceptualization: S.H.H., Y.E.K., J.H.K., Y.Y.K.; Methodology: Y.E.K. and S.H.H.; Validation: Y.E.K. and S.H.H.; Formal analysis: Y.E.K. and S.H.H.; Investigation: Y.E.K., S.H.H., and Y.W.J.; Writing-original draft preparation: Y.E.K. and S.H.H.; Writing-review and editing: Y.E.K., S.H.H., Y.W.J., K.Y., and Y.Y.K.; Visualization: Y.E.K.; Supervision: Y.Y.K.; Project administration: Y.Y.K.; Funding acquisition: J.W.L. and Y.Y.K. Acknowledgments We express our gratitude to the Kong laboratory members for their valuable feedback during the project. 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Supplementary Files Additionalfile1.SupplementaryMaterial1.docx Additional file 1 (PDF) Supplementary Material 1 Additionalfile2.Supplementarytable1.xlsx Additional file 2 (xls) Supplementary Table 1: Mouse primers used for genotyping and qRT-PCR Cite Share Download PDF Status: Published Journal Publication published 23 Dec, 2024 Read the published version in Skeletal Muscle → Version 1 posted Editorial decision: Revision requested 27 Nov, 2024 Reviews received at journal 26 Nov, 2024 Reviewers agreed at journal 24 Nov, 2024 Reviews received at journal 13 Nov, 2024 Reviewers agreed at journal 11 Nov, 2024 Reviewers invited by journal 11 Nov, 2024 Editor assigned by journal 11 Nov, 2024 Submission checks completed at journal 07 Nov, 2024 First submitted to journal 07 Nov, 2024 You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. 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Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {\"props\":{\"pageProps\":{\"initialData\":{\"identity\":\"rs-5413133\",\"acceptedTermsAndConditions\":true,\"allowDirectSubmit\":false,\"archivedVersions\":[],\"articleType\":\"Research Article\",\"associatedPublications\":[],\"authors\":[{\"id\":381888900,\"identity\":\"401e43f7-65c5-4b47-beca-0295b2e78f28\",\"order_by\":0,\"name\":\"Yea-Eun Kim\",\"email\":\"\",\"orcid\":\"\",\"institution\":\"Seoul National University\",\"correspondingAuthor\":false,\"prefix\":\"\",\"firstName\":\"Yea-Eun\",\"middleName\":\"\",\"lastName\":\"Kim\",\"suffix\":\"\"},{\"id\":381888902,\"identity\":\"cf3e0a78-a321-4497-bb92-f32cfffc967c\",\"order_by\":1,\"name\":\"Sang-Hyeon Hann\",\"email\":\"\",\"orcid\":\"\",\"institution\":\"Seoul National University\",\"correspondingAuthor\":false,\"prefix\":\"\",\"firstName\":\"Sang-Hyeon\",\"middleName\":\"\",\"lastName\":\"Hann\",\"suffix\":\"\"},{\"id\":381888905,\"identity\":\"eda1dba5-5738-4b57-b7e4-c6a32bb13fee\",\"order_by\":2,\"name\":\"Young-Woo Jo\",\"email\":\"\",\"orcid\":\"\",\"institution\":\"Seoul National University\",\"correspondingAuthor\":false,\"prefix\":\"\",\"firstName\":\"Young-Woo\",\"middleName\":\"\",\"lastName\":\"Jo\",\"suffix\":\"\"},{\"id\":381888906,\"identity\":\"f0496b46-e0fb-4581-836e-56d906dcbc55\",\"order_by\":3,\"name\":\"Kyusang Yoo\",\"email\":\"\",\"orcid\":\"\",\"institution\":\"Seoul National University\",\"correspondingAuthor\":false,\"prefix\":\"\",\"firstName\":\"Kyusang\",\"middleName\":\"\",\"lastName\":\"Yoo\",\"suffix\":\"\"},{\"id\":381888908,\"identity\":\"67523d80-08ed-408f-808c-c721c9f7bf82\",\"order_by\":4,\"name\":\"Ji-Hoon Kim\",\"email\":\"\",\"orcid\":\"\",\"institution\":\"Korea Institute of Science and Technology\",\"correspondingAuthor\":false,\"prefix\":\"\",\"firstName\":\"Ji-Hoon\",\"middleName\":\"\",\"lastName\":\"Kim\",\"suffix\":\"\"},{\"id\":381888909,\"identity\":\"78a2b7fd-497d-4e15-8775-95aebd354d78\",\"order_by\":5,\"name\":\"Jae W. Lee\",\"email\":\"\",\"orcid\":\"\",\"institution\":\"University at Buffalo\",\"correspondingAuthor\":false,\"prefix\":\"\",\"firstName\":\"Jae\",\"middleName\":\"W.\",\"lastName\":\"Lee\",\"suffix\":\"\"},{\"id\":381888910,\"identity\":\"42bf6970-8d02-4472-bfe9-97ac80b54ef6\",\"order_by\":6,\"name\":\"Young-Yun Kong\",\"email\":\"data:image/png;base64,iVBORw0KGgoAAAANSUhEUgAAAZAAAAAyAQMAAABI0h/eAAAABlBMVEX///8AAABVwtN+AAAACXBIWXMAAA7EAAAOxAGVKw4bAAAAuUlEQVRIiWNgGAWjYDACHhBRAeMdIFrLGZK1MLaRosXgzOFjD7/Oq8szOMD88APDmXtEaDnblm4su+1wscEBNmMJhhvFRGg5z2MmLbntQOKGAwxmDAwfEojVMqcOqIX9G5FazvaYSX5sYAZq4QHacoMILZJnjqVJMxw7nDjzME+xRMIZIrTwnUk+Jvmjpi6x73j7xg8fjhGhReEAAwMzODqZgZgIDQwM8g3AmPxBjMpRMApGwSgYuQAAJJ49PDgIDakAAAAASUVORK5CYII=\",\"orcid\":\"\",\"institution\":\"Seoul National University\",\"correspondingAuthor\":true,\"prefix\":\"\",\"firstName\":\"Young-Yun\",\"middleName\":\"\",\"lastName\":\"Kong\",\"suffix\":\"\"}],\"badges\":[],\"createdAt\":\"2024-11-08 02:53:32\",\"currentVersionCode\":1,\"declarations\":\"\",\"doi\":\"10.21203/rs.3.rs-5413133/v1\",\"doiUrl\":\"https://doi.org/10.21203/rs.3.rs-5413133/v1\",\"draftVersion\":[],\"editorialEvents\":[{\"content\":\"https://doi.org/10.1186/s13395-024-00369-9\",\"type\":\"published\",\"date\":\"2024-12-23T15:57:48+00:00\"}],\"editorialNote\":\"\",\"failedWorkflow\":false,\"files\":[{\"id\":69862201,\"identity\":\"8234ad26-4096-4b9d-b785-21f5c7a13dea\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:00:51\",\"extension\":\"jpg\",\"order_by\":1,\"title\":\"Figure 1\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":1594304,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eFiber CSA and fiber type composition in adult Mll4\\u003c/strong\\u003e\\u003csup\\u003e\\u003cstrong\\u003eWT \\u003c/strong\\u003e\\u003c/sup\\u003e\\u003cstrong\\u003eand Mll4\\u003c/strong\\u003e\\u003csup\\u003e\\u003cstrong\\u003eΔMCK\\u003c/strong\\u003e\\u003c/sup\\u003e\\u003cstrong\\u003e mice.\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003e(A-B) \\u003c/strong\\u003ePercentage of myofibers within each indicated range of CSA in TA muscle of Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e and Mll4\\u003csup\\u003eWT \\u003c/sup\\u003emice. n=3 mice for each genotype.\\u003cstrong\\u003e (C)\\u003c/strong\\u003e Representative image of soleus muscles immunolabeled with anti-MyHC1 (green) and anti-laminin (red). Scale bar, 500 μm. \\u003cstrong\\u003e(D, F)\\u003c/strong\\u003e The fiber number of whole TA muscles from Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and\\u003csup\\u003e \\u003c/sup\\u003eMll4\\u003csup\\u003eΔMCK \\u003c/sup\\u003emice of both males and females. n=3 mice for each genotype.\\u003cstrong\\u003e (E, G)\\u003c/strong\\u003e The percentage of slow fiber among total fiber of soleus muscles from Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and\\u003csup\\u003e \\u003c/sup\\u003eMll4\\u003csup\\u003eΔMCK \\u003c/sup\\u003emice of both males and females. n=3 mice for each genotype. Data are presented as mean ± SEM of biological replicates. Statistical analyses were performed using unpaired t-test with Welch’s correction.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Figure1.jpg\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/017bdf8c0494a1bb790a92af.jpg\"},{\"id\":69862208,\"identity\":\"0eeee631-0519-426a-a282-6cbe6b974662\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:00:51\",\"extension\":\"jpg\",\"order_by\":2,\"title\":\"Figure 2\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":1253718,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eAltered myofiber phenotype of 8-week-old Mll4\\u003c/strong\\u003e\\u003csup\\u003e\\u003cstrong\\u003eΔMCK \\u003c/strong\\u003e\\u003c/sup\\u003e\\u003cstrong\\u003emice.\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003e(A)\\u003c/strong\\u003e Representative image of the isolated single myofiber. DAPI staining was applied to visualize nuclei. Scale bar, 500 μm.\\u003cstrong\\u003e (B, E)\\u003c/strong\\u003e Myofiber length, \\u003cstrong\\u003e(C, F)\\u003c/strong\\u003e myonuclei accretion, and \\u003cstrong\\u003e(D, G)\\u003c/strong\\u003e myonuclear density were quantified. \\u003cstrong\\u003e(H) \\u003c/strong\\u003eImmunocytochemistry of isolated myofibers of 4-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e and Mll4\\u003csup\\u003eWT \\u003c/sup\\u003emice with DAPI (blue) and anti-Pax7 (red). MuSCs are marked with arrowheads. Scale bars, 100 μm.\\u003cstrong\\u003e (I, J) \\u003c/strong\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e MuSC number per fiber of Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and\\u003csup\\u003e \\u003c/sup\\u003eMll4\\u003csup\\u003eΔMCK \\u003c/sup\\u003emice of both genders. (B-G, and I-J) n=3 mice for each genotype; \\u0026gt;20 fibers per mouse was quantified. Data are presented as mean ± SEM of biological replicates. Statistical analyses were performed using unpaired t-test with Welch’s correction.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Figure2.jpg\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/f1c6c82950eeb14170580aa3.jpg\"},{\"id\":69862202,\"identity\":\"75a2a1fd-43dd-4cc9-92d1-3cd2da166374\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:00:51\",\"extension\":\"jpg\",\"order_by\":3,\"title\":\"Figure 3\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":1640821,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eMuSC depletion due to increased population of differentiating myoblasts in 4-week-old Mll4\\u003c/strong\\u003e\\u003csup\\u003e\\u003cstrong\\u003eΔMCK\\u003c/strong\\u003e\\u003c/sup\\u003e\\u003cstrong\\u003e muscles.\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eImmunohistochemistry on TA \\u003cstrong\\u003e(A)\\u003c/strong\\u003e and soleus \\u003cstrong\\u003e(B)\\u003c/strong\\u003e muscle section with DAPI (blue), anti-laminin (green), and anti-Pax7 (red). Scale bars, 20 μm.\\u003cstrong\\u003e (D) \\u003c/strong\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e MuSC number per 100 fibers was quantified in TA muscles of 0, 2, 4, and 8-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e and littermate control mice. \\u003cstrong\\u003e(E) \\u003c/strong\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e MuSC number per 100 fibers was quantified in soleus muscles of 4 and 8-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e and littermate control mice. \\u003cstrong\\u003e(C) \\u003c/strong\\u003eImmunohistochemistry on 4W TA muscle section with DAPI (blue), anti-Ki67 (green), and anti-Pax7 (red). Scale bars, 20 μm.\\u003cstrong\\u003e (F) \\u003c/strong\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003eKi67\\u003csup\\u003e+\\u003c/sup\\u003e cell number per total Pax7\\u003csup\\u003e+\\u003c/sup\\u003e cells of 4W TA muscle. \\u003cstrong\\u003e(G) \\u003c/strong\\u003eSchematic diagram of EdU treatment. \\u003cstrong\\u003e(H) \\u003c/strong\\u003eImmunocytochemistry of sorted MuSCs with DAPI (blue), anti-Pax7 (red), anti-MyoD (green), and EdU (white – pseudo color for Alexa Fluor 647). MyoD\\u003csup\\u003e+ \\u003c/sup\\u003ecells and EdU\\u003csup\\u003e+\\u003c/sup\\u003e cells are marked with arrowheads and sharps, respectively. Scale bars, 20 μm. For enlarged images, Scale bars represent 10 μm.\\u003cstrong\\u003e (I) \\u003c/strong\\u003eMyoD\\u003csup\\u003e+\\u003c/sup\\u003e cell number per total Pax7\\u003csup\\u003e+\\u003c/sup\\u003e cells, and \\u003cstrong\\u003e(J)\\u003c/strong\\u003e MyoD\\u003csup\\u003e+\\u003c/sup\\u003e cell number per EdU\\u003csup\\u003e+\\u003c/sup\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e cells were quantified. (D-F) n=3-4 mice for each genotype. (I-K)\\u003cstrong\\u003e \\u003c/strong\\u003en=3-4 mice for each genotype; \\u0026gt;200 sorted MuSCs per mouse were quantified. (D-F, and I-K) Data are presented as mean ± SEM of biological replicates. Statistical analyses were performed using unpaired t-test with Welch’s correction.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Figure3.tif.jpg\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/689214220bf421544d5521bd.jpg\"},{\"id\":69862999,\"identity\":\"998a67fd-474d-49a3-b478-9e01d4c3ea8d\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:16:51\",\"extension\":\"jpg\",\"order_by\":4,\"title\":\"Figure 4\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":1453932,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eInduced deletion of Mll4 in the adult stage does not impact myofiber maintenance.\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003e(A)\\u003c/strong\\u003e Schematic diagram of mouse preparation.\\u003cstrong\\u003e (B) \\u003c/strong\\u003eqRT-PCR analysis of myofibers to confirm the downregulation of the \\u003cem\\u003eMll4\\u003c/em\\u003e gene in +2W Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice. \\u003cstrong\\u003e(C) \\u003c/strong\\u003ePercentage of myofibers within each indicated range of CSA, \\u003cstrong\\u003e(D)\\u003c/strong\\u003e gross fiber number, and \\u003cstrong\\u003e(E) \\u003c/strong\\u003epercentage of MyHC2x fibers in TA muscle of Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and +2W Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice. \\u003cstrong\\u003e(F) \\u003c/strong\\u003eGross fiber number and \\u003cstrong\\u003e(G)\\u003c/strong\\u003e percentage of MyHC1 fibers in soleus muscle of Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice. \\u003cstrong\\u003e(H) \\u003c/strong\\u003ePercentage of myofibers within each indicated range of CSA, \\u003cstrong\\u003e(I) \\u003c/strong\\u003egross fiber number, and \\u003cstrong\\u003e(J)\\u003c/strong\\u003e percentage of MyHC2x fibers in TA muscle of Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and +4W Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice. \\u003cstrong\\u003e(K) \\u003c/strong\\u003eGross fiber number \\u003cstrong\\u003e(L)\\u003c/strong\\u003e and percentage of MyHC1 fibers in soleus muscle of Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and +4W Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice. (B-L) n=3 mice for each genotype.\\u003cstrong\\u003e \\u003c/strong\\u003eData are presented as mean ± SEM of biological replicates. Statistical analyses were performed using unpaired t-test with Welch’s correction.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Figure4.jpg\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/444b6d278d375b0ee3126d5e.jpg\"},{\"id\":69862800,\"identity\":\"1cdaa25d-999a-45d8-9e8c-bb055819b6a0\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:08:51\",\"extension\":\"jpg\",\"order_by\":5,\"title\":\"Figure 5\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":1176387,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eNormal exercise capacity and muscle features of myofiber-\\u003c/strong\\u003e\\u003cem\\u003e\\u003cstrong\\u003eMll4\\u003c/strong\\u003e\\u003c/em\\u003e\\u003cstrong\\u003e deleted muscles.\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003e(A)\\u003c/strong\\u003e Schematic diagram of mouse training and preparation. \\u003cstrong\\u003e(B)\\u003c/strong\\u003e The measurement values of grip strength (N/g) of Mll4\\u003csup\\u003eΔWT-EX\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA-EX\\u003c/sup\\u003e mice. Grip strength (N) was normalized to body weight (g). \\u003cstrong\\u003e(C, D) \\u003c/strong\\u003eThe measurement values of endurance running test. Total running time (min) (C) and total running distance (m) (D) of Mll4\\u003csup\\u003eΔWT-EX\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA-EX\\u003c/sup\\u003e mice. \\u003cstrong\\u003e(E) \\u003c/strong\\u003ePercentage of myofibers within each indicated range of CSA, \\u003cstrong\\u003e(F)\\u003c/strong\\u003e gross fiber number, and \\u003cstrong\\u003e(G) \\u003c/strong\\u003epercentage of MyHC2x fibers in TA muscle of Mll4\\u003csup\\u003eΔWT-EX\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA-EX\\u003c/sup\\u003e mice. \\u003cstrong\\u003e(H) \\u003c/strong\\u003eGross fiber number and \\u003cstrong\\u003e(I)\\u003c/strong\\u003e percentage of MyHC1 fibers in soleus muscle of Mll4\\u003csup\\u003eΔWT-EX\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA-EX\\u003c/sup\\u003e mice. (B-I) n=6 mice for each genotype.\\u003cstrong\\u003e \\u003c/strong\\u003eData are presented as mean ± SEM of biological replicates. Statistical analyses were performed using unpaired t-test with Welch’s correction.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Figure5.jpg\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/1ebb53317f0664af1b86530a.jpg\"},{\"id\":69863000,\"identity\":\"07f590d3-1ce0-44d1-a6b2-20314390c0fb\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:16:51\",\"extension\":\"jpg\",\"order_by\":6,\"title\":\"Figure 6\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":1823474,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eSevere MuSC deprivation in \\u003c/strong\\u003e\\u003cem\\u003e\\u003cstrong\\u003eMll4\\u003c/strong\\u003e\\u003c/em\\u003e\\u003cstrong\\u003e deleted adult myofibers.\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003e(A)\\u003c/strong\\u003e Immunohistochemistry on TA muscle section with DAPI (blue), anti-laminin (green), and anti-Pax7 (red). \\u003cstrong\\u003e(B) \\u003c/strong\\u003ePax\\u003csup\\u003e+\\u003c/sup\\u003e MuSC number per 100 fibers of Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e TA and \\u003cstrong\\u003e(C)\\u003c/strong\\u003e soleus muscle. Scale bars, 20 μm.\\u003cstrong\\u003e (D)\\u003c/strong\\u003e Immunohistochemistry on TA muscle section with DAPI (blue), anti-Ki67 (green), and anti-Pax7 (red). Scale bars, 20 μm.\\u003cstrong\\u003e (E)\\u003c/strong\\u003e Pax7\\u003csup\\u003e+\\u003c/sup\\u003eKi67\\u003csup\\u003e+\\u003c/sup\\u003e cell number per total Pax7\\u003csup\\u003e+\\u003c/sup\\u003e cells. \\u003cstrong\\u003e(F) \\u003c/strong\\u003eSchematic diagram of EdU treatment.\\u003cstrong\\u003e (G)\\u003c/strong\\u003e Immunocytochemistry of sorted MuSCs with DAPI (blue), anti-Pax7 (red), anti-MyoD (green), and EdU (white – pseudo color for Alexa Fluor 647). MyoD\\u003csup\\u003e+\\u003c/sup\\u003e cells and EdU\\u003csup\\u003e+\\u003c/sup\\u003e cells are marked with arrowheads and sharps, respectively. Scale bars, 20 μm. For enlarged images, Scale bars represent 10 μm.\\u003cstrong\\u003e (H)\\u003c/strong\\u003e MyoD\\u003csup\\u003e+\\u003c/sup\\u003e cell number per total Pax7\\u003csup\\u003e+\\u003c/sup\\u003e cells, \\u003cstrong\\u003e(I)\\u003c/strong\\u003e MyoD\\u003csup\\u003e+\\u003c/sup\\u003e cell number per EdU\\u003csup\\u003e+\\u003c/sup\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e cells, and \\u003cstrong\\u003e(J)\\u003c/strong\\u003e MyoD\\u003csup\\u003e+\\u003c/sup\\u003e cell number per EdU\\u003csup\\u003e-\\u003c/sup\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e cells were quantified. \\u003cstrong\\u003e(K) \\u003c/strong\\u003eImmunohistochemistry on TA muscle section with DAPI (blue), EdU (green), and anti-dystrophin (red). Scale bars, 20 μm. For enlarged images, Scale bars represent 10 μm.\\u003cstrong\\u003e (L) \\u003c/strong\\u003eThe number of fiber incorporated EdU\\u003csup\\u003e+\\u003c/sup\\u003e cells per 100 fibers. (B, C, E, and K) n=3-4 mice for each genotype. (G-I)\\u003cstrong\\u003e \\u003c/strong\\u003en=3-4 mice for each genotype; \\u0026gt;200 sorted MuSCs per mouse were quantified. (B, C, E, K, G-I, and K) Data are presented as mean ± SEM of biological replicates. Statistical analyses were performed using unpaired t-test with Welch’s correction.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Figure6.jpg\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/953c8746d60d1efb2be87c4b.jpg\"},{\"id\":69862206,\"identity\":\"1fa25b1a-7e59-48a7-9885-f7d45738d148\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:00:51\",\"extension\":\"jpg\",\"order_by\":7,\"title\":\"Figure 7\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":1213860,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eDefective muscle regeneration capacity after injury in \\u003c/strong\\u003e\\u003cem\\u003e\\u003cstrong\\u003eMll4\\u003c/strong\\u003e\\u003c/em\\u003e\\u003cstrong\\u003e deleted TA muscles.\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003e(A)\\u003c/strong\\u003e Schematic diagram of muscle injury and mouse preparation. \\u003cstrong\\u003e(B)\\u003c/strong\\u003e Muscle mass of TA, EDL, GA, and soleus muscles of Mll4\\u003csup\\u003eWT-inj\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA-inj\\u003c/sup\\u003e mice. \\u003cstrong\\u003e(C)\\u003c/strong\\u003e Representative image of TA muscle labeled with anti-laminin (green) and Hoechst 33342. Scale bars, 100 μm. \\u003cstrong\\u003e(D)\\u003c/strong\\u003e Percentage of myofibers within each indicated range of CSA in TA muscle of Mll4\\u003csup\\u003eWT-inj\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA-inj\\u003c/sup\\u003e mice. (B and D) n=3 mice for each genotype. Data are presented as mean ± SEM of biological replicates. Statistical analyses were performed using unpaired t-test with Welch’s correction.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Figure7.jpg\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/bd0245fb5aaaf4c008fbb847.jpg\"},{\"id\":69862210,\"identity\":\"ddd3aa0a-6647-4138-b2ab-31e1a39fc0a2\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:00:51\",\"extension\":\"jpg\",\"order_by\":8,\"title\":\"Figure 8\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":1263086,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eAltered Notch ligand expression in \\u003c/strong\\u003e\\u003cem\\u003e\\u003cstrong\\u003eMll4\\u003c/strong\\u003e\\u003c/em\\u003e\\u003cstrong\\u003e deleted myofibers, leading to downregulated notch signaling in MuSCs.\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003e(A)\\u003c/strong\\u003e Evaluation of three public databases proved downregulation of Notch ligands in \\u003cem\\u003eMll4\\u003c/em\\u003e-deleted muscle and myocyte. \\u003cstrong\\u003e(B) \\u003c/strong\\u003eqRT-PCR analysis to compare Notch ligand expression in pubertal (4W) and adult myofibers.\\u003cstrong\\u003e (C-D)\\u003c/strong\\u003e qRT-PCR analysis to quantify mRNA expression of Notch ligands in myofibers of 4-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice and adult Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice.\\u003cstrong\\u003e (E-F) \\u003c/strong\\u003eqRT-PCR of canonical Notch effectors confirmed general downregulation of Notch signaling in Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e MuSCs. \\u003cstrong\\u003e(G-H) \\u003c/strong\\u003eChIP-seq data of MLL4 in myocytes revealing the binding of MLL4 to \\u003cem\\u003eDll1\\u003c/em\\u003e and \\u003cem\\u003eJag2\\u003c/em\\u003e gene loci. (B-F) n=3 mice for each genotype.\\u003cstrong\\u003e \\u003c/strong\\u003eData are presented as mean ± SEM of biological replicates. Statistical analyses were performed using unpaired t-test with Welch’s correction.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Figure8.jpg\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/52676b2de2e483cf13ce9c94.jpg\"},{\"id\":72640848,\"identity\":\"a5191416-859a-4426-85a4-2c4ad5475240\",\"added_by\":\"auto\",\"created_at\":\"2024-12-30 16:10:24\",\"extension\":\"pdf\",\"order_by\":0,\"title\":\"\",\"display\":\"\",\"copyAsset\":false,\"role\":\"manuscript-pdf\",\"size\":12487702,\"visible\":true,\"origin\":\"\",\"legend\":\"\",\"description\":\"\",\"filename\":\"manuscript.pdf\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/705f6d95-20b4-4e1d-965e-72e31317ce73.pdf\"},{\"id\":69862211,\"identity\":\"d039cc0d-b288-4ee8-823d-9222183188fd\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:00:52\",\"extension\":\"docx\",\"order_by\":1,\"title\":\"\",\"display\":\"\",\"copyAsset\":false,\"role\":\"supplement\",\"size\":8721588,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eAdditional file 1 (PDF)\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eSupplementary Material 1\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Additionalfile1.SupplementaryMaterial1.docx\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/dd86f618b979a98675b1e1a1.docx\"},{\"id\":69862797,\"identity\":\"39853416-d7cd-4585-a9ad-b12d49a2de2b\",\"added_by\":\"auto\",\"created_at\":\"2024-11-26 06:08:51\",\"extension\":\"xlsx\",\"order_by\":2,\"title\":\"\",\"display\":\"\",\"copyAsset\":false,\"role\":\"supplement\",\"size\":11303,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eAdditional file 2 (xls)\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eSupplementary Table 1: Mouse primers used for genotyping and qRT-PCR\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"Additionalfile2.Supplementarytable1.xlsx\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-5413133/v1/11dae8348829f3bf5cd24871.xlsx\"}],\"financialInterests\":\"No competing interests reported.\",\"formattedTitle\":\"Mll4 in Skeletal Muscle Fiber Maintains Muscle Stem Cells by Regulating Notch Ligands\",\"fulltext\":[{\"header\":\"Background\",\"content\":\"\\u003cp\\u003eMuscle stem cells (MuSCs) are resident stem cells of skeletal muscle that contribute to muscle development, growth, and regeneration. They actively proliferate and differentiate into myocytes to contribute to myonuclei accretion and muscle growth [\\u003cspan citationid=\\\"CR1\\\" class=\\\"CitationRef\\\"\\u003e1\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR2\\\" class=\\\"CitationRef\\\"\\u003e2\\u003c/span\\u003e]. In the adult stage, MuSCs enter a quiescent state and remain as reserve stem cells [\\u003cspan citationid=\\\"CR3\\\" class=\\\"CitationRef\\\"\\u003e3\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR4\\\" class=\\\"CitationRef\\\"\\u003e4\\u003c/span\\u003e]. Upon injury, MuSCs become activated, providing myogenic cells to repair the muscle tissue. During muscle development and regenerative myogenesis, the MuSC niche regulates the dynamic transitions of MuSCs, including their activation, proliferation, differentiation, and self-renewal.\\u003c/p\\u003e \\u003cp\\u003eMyofiber is an important cellular component of the MuSC niche. Unlike other cells that compose the MuSC niche, myofibers are in direct contact with MuSCs [\\u003cspan citationid=\\\"CR5\\\" class=\\\"CitationRef\\\"\\u003e5\\u003c/span\\u003e]. This enables them to regulate the MuSC state through both paracrine and contact-dependent juxtacrine signaling [\\u003cspan citationid=\\\"CR6\\\" class=\\\"CitationRef\\\"\\u003e6\\u003c/span\\u003e]. As paracrine signaling, myofibers secrete Wnt-4 to repress aberrant activation and maintain MuSC quiescence in adult homeostatic muscle [\\u003cspan citationid=\\\"CR7\\\" class=\\\"CitationRef\\\"\\u003e7\\u003c/span\\u003e]. FGF6 is another paracrine factor produced in myofibers to promote MuSC expansion during both developmental and regenerative myogenesis [8.9]. Myofibers also provide juxtacrine signals such as N-cadherin and M-cadherin to maintain MuSCs. These cell adhesion molecules are expressed at myofiber sites that are in direct contact with MuSCs to repress stem cell activation and maintain MuSC quiescence in adult muscles [\\u003cspan citationid=\\\"CR10\\\" class=\\\"CitationRef\\\"\\u003e10\\u003c/span\\u003e].\\u003c/p\\u003e \\u003cp\\u003eAmong the various molecular signals derived from myofibers, Notch signaling plays a particularly crucial role in the maintenance of the stem cell pool and the regulation of cell fate decisions in MuSCs. In mammals, Notch signal-sending cells express Notch ligands (Dll1, 4 and Jag1, 2) and signal-receiving cells express Notch receptors (Notch1-4) [\\u003cspan citationid=\\\"CR11\\\" class=\\\"CitationRef\\\"\\u003e11\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR12\\\" class=\\\"CitationRef\\\"\\u003e12\\u003c/span\\u003e]. Myofibers activate Notch signaling at different developmental stages primarily to generate a quiescent population of myogenic progenitor cells [\\u003cspan citationid=\\\"CR2\\\" class=\\\"CitationRef\\\"\\u003e2\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR13\\\" class=\\\"CitationRef\\\"\\u003e13\\u003c/span\\u003e]. Several Notch ligands are expressed in myofibers to facilitate diverse Notch signaling to regulate the MuSC niche effectively at different developmental stages. Dll1, a Notch ligand mainly expressed in pubertal myofibers, interacts with the activated MuSCs to promote self-renewal, which is crucial for maintaining MuSCs [\\u003cspan citationid=\\\"CR14\\\" class=\\\"CitationRef\\\"\\u003e14\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR15\\\" class=\\\"CitationRef\\\"\\u003e15\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR16\\\" class=\\\"CitationRef\\\"\\u003e16\\u003c/span\\u003e]. In adult myofibers, Dll4 represses MuSC cell cycle entry, thus retaining the quiescent state of MuSCs [\\u003cspan citationid=\\\"CR17\\\" class=\\\"CitationRef\\\"\\u003e17\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR18\\\" class=\\\"CitationRef\\\"\\u003e18\\u003c/span\\u003e]. While various studies have indicated that Notch signaling originating from myofibers has a role in regulating the state of MuSCs, there is limited understanding of whether there is a regulator present in myofibers that coordinates the expression of Notch ligands during the postnatal period.\\u003c/p\\u003e \\u003cp\\u003eMixed-lineage leukemia 4 (MLL4; also known as Kmt2d), a major H3K4 mono- and di-methyltransferase, is an essential histone modification enzyme for enhancer activation [\\u003cspan citationid=\\\"CR19\\\" class=\\\"CitationRef\\\"\\u003e19\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR20\\\" class=\\\"CitationRef\\\"\\u003e20\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR21\\\" class=\\\"CitationRef\\\"\\u003e21\\u003c/span\\u003e]. H3K4me1 marking by MLL4 is required for H3K27 acetylation and the recruitment of cell type-specific transcription factors [\\u003cspan citationid=\\\"CR19\\\" class=\\\"CitationRef\\\"\\u003e19\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR20\\\" class=\\\"CitationRef\\\"\\u003e20\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR21\\\" class=\\\"CitationRef\\\"\\u003e21\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR22\\\" class=\\\"CitationRef\\\"\\u003e22\\u003c/span\\u003e]. Deletion of \\u003cem\\u003eMll4\\u003c/em\\u003e results in the disturbance of H3K4me1 and H3K27ac on active enhancers, leading to defects in transcribing both newly activated genes, as well as genes that were already being expressed [\\u003cspan citationid=\\\"CR19\\\" class=\\\"CitationRef\\\"\\u003e19\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR20\\\" class=\\\"CitationRef\\\"\\u003e20\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR21\\\" class=\\\"CitationRef\\\"\\u003e21\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR22\\\" class=\\\"CitationRef\\\"\\u003e22\\u003c/span\\u003e].\\u003c/p\\u003e \\u003cp\\u003eRecent study addressed that \\u003cem\\u003eMll4\\u003c/em\\u003e depletion in myofibers turns off the slow type I fiber-specific genes, leading to fiber-type transition [\\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e]. Myofibers are classified into two primary types: Type 1 (slow-twitch) and Type 2 (fast-twitch). Type 1 fibers rely on oxidative metabolism, which endows them with endurance capabilities. In contrast, Type 2 fibers utilize glycolytic pathways to generate rapid force [\\u003cspan citationid=\\\"CR24\\\" class=\\\"CitationRef\\\"\\u003e24\\u003c/span\\u003e]. This diversity in myofiber types contributes to muscle performance and adaptability to various stimuli [\\u003cspan citationid=\\\"CR25\\\" class=\\\"CitationRef\\\"\\u003e25\\u003c/span\\u003e]. Liu [\\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e] reported that endurance exercise capacity was reduced in the Mll4-KO mice due to a slow-to-fast fiber-type shift. However, this study exclusively utilized male mice and focused solely on the function of Mll4 in developing myofibers. Building on these findings, our research aims to further investigate the role of Mll4 in myofibers across both sexes and various developmental stages.\\u003c/p\\u003e \\u003cp\\u003eIn this paper, we report that MLL4 in myofiber is required to maintain MuSCs during both the pubertal and adult stages, regardless of gender. Mouse models with myofiber-specific deletion of \\u003cem\\u003eMll4\\u003c/em\\u003e exhibited reduced myofiber length, fewer myonuclei, and MuSC depletion. When \\u003cem\\u003eMll4\\u003c/em\\u003e was ablated at the adult stage using an inducible method, MuSCs underwent differentiation into myoblasts, either with or without entering the cell cycle, leading to a depletion of adult MuSCs. Furthermore, the expression of specific Notch ligands at both pubertal and adult stages was significantly reduced in \\u003cem\\u003eMll4\\u003c/em\\u003e-knockout myofibers. Together, our data suggest the importance of MLL4 in skeletal muscle fibers for maintaining the MuSC number, which might be achieved by affecting Notch ligand expression, in different postnatal periods.\\u003c/p\\u003e\"},{\"header\":\"Methods\",\"content\":\"\\u003cdiv id=\\\"Sec3\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eAnimals\\u003c/h2\\u003e \\u003cp\\u003e\\u003cem\\u003eMCK\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eCre/+\\u003c/em\\u003e\\u003c/sup\\u003e (stock 006475), \\u003cem\\u003eHSA\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eMerCreMer/+\\u003c/em\\u003e\\u003c/sup\\u003e (stock 031934), and \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e (stock 032152) mice were acquired from The Jackson Laboratory (Bar Harbor, ME, USA). The mice were backcrossed to C57BL/6 mice at least 6 times. To generate mice with myofiber-specific deletion of \\u003cem\\u003eMll4\\u003c/em\\u003e, \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e mice were crossed with \\u003cem\\u003eMCK\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eCre/+\\u003c/em\\u003e\\u003c/sup\\u003e (\\u003cem\\u003eMCK\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eCre/+\\u003c/em\\u003e\\u003c/sup\\u003e; \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e\\u003cem\\u003e\\u0026ndash; Mll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eΔMCK\\u003c/em\\u003e\\u003c/sup\\u003e\\u003cem\\u003e\\u0026ndash;\\u003c/em\\u003e). To develop myofiber-specific \\u003cem\\u003eMll4\\u003c/em\\u003e conditional knockout mice using tamoxifen-inducible Cre, \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e mice were crossed with \\u003cem\\u003eHSA\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eMerCreMer/+\\u003c/em\\u003e\\u003c/sup\\u003e (\\u003cem\\u003eHSA\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eMerCreMer/+\\u003c/em\\u003e\\u003c/sup\\u003e; \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e\\u003cem\\u003e\\u0026ndash; Mll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eΔHSA\\u003c/em\\u003e\\u003c/sup\\u003e\\u003cem\\u003e\\u0026ndash;\\u003c/em\\u003e). Following a breeding strategy from The Jackson Laboratory (Bar Harbor, ME, USA), we bred heterozygous \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/+\\u003c/em\\u003e\\u003c/sup\\u003e females with Cre-recombinase to homozygous \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e males. Both male and female mice were used in the experiments. Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice used for experiments were adults, between 3\\u0026ndash;6 months of age. Control littermates lacking Cre-recombinase (Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e) were utilized for analysis. All mouse lines were housed under controlled conditions with specific pathogen-free and handled according to the guidelines of the Seoul National University Institutional Animal Care and Use Committee (Protocol number: SNU-240103-3).\\u003c/p\\u003e \\u003c/div\\u003e\\n\\u003ch3\\u003eAnimal procedures\\u003c/h3\\u003e\\n\\u003cp\\u003eTamoxifen (Sigma-Aldrich) was dissolved in corn oil at a concentration of 20mg/ml. For tamoxifen-induced Cre recombination in Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice, both control and experimental mice were administered tamoxifen at a concentration of 150 mg/kg of mouse per day for five continuous days by intraperitoneal injection. For detection of cell cycle entry, 5-ethynyl-2\\u0026rsquo;-deoxyuridine (EdU; Thermo Fisher Scientific) dissolved in sterile phosphate-buffered saline (PBS) was injected at a concentration of 40 mg/kg of mouse intraperitoneally daily.\\u003c/p\\u003e\\n\\u003ch3\\u003eMuscle injury\\u003c/h3\\u003e\\n\\u003cp\\u003eFor BaCl\\u003csub\\u003e2\\u003c/sub\\u003e muscle injury, mice were anesthetized with 2.4% 2, 2, 2-Tribromoethanol (Avertin; Sigma-Aldrich) in PBS (240 mg/kg of mouse) and injected with 50 \\u0026micro;l 1.2% BaCl\\u003csub\\u003e2\\u003c/sub\\u003e in saline (Sigma-Aldrich) to the tibialis anterior (TA) muscles. At 10 days after the injury, TA muscles were dissected, frozen in optimal cutting temperature compound (O.C.T.; Sakura Finetek) with liquid nitrogen, and stored at -80\\u0026deg;C until analysis.\\u003c/p\\u003e\\n\\u003ch3\\u003eMeasurement of CSA distribution\\u003c/h3\\u003e\\n\\u003cp\\u003eLaminin-stained section was imaged with EVOS FL Auto 2 (Thermo Fisher Scientific) with the same laser setting, exposure, and magnification. To measure CSA, semiautomatic muscle analysis using segmentation of histology (SMASH) was used with a segmentation filter (CSA between 200 \\u0026micro;m2 and 6000 \\u0026micro;m2; eccentricity\\u0026thinsp;\\u0026le;\\u0026thinsp;0.95; convexity\\u0026thinsp;\\u0026ge;\\u0026thinsp;0.80). For injured muscles, CSA of regenerating myofibers with centralized nuclei was analyzed. The segmentation filter was adjusted as follows: CSA between 150 \\u0026micro;m2 and 6000 \\u0026micro;m2; eccentricity\\u0026thinsp;\\u0026le;\\u0026thinsp;0.95; convexity\\u0026thinsp;\\u0026ge;\\u0026thinsp;0.80.\\u003c/p\\u003e\\n\\u003ch3\\u003eSingle myofiber isolation\\u003c/h3\\u003e\\n\\u003cp\\u003eSingle myofiber isolation was performed according to a previously reported protocol with modifications [\\u003cspan citationid=\\\"CR47\\\" class=\\\"CitationRef\\\"\\u003e47\\u003c/span\\u003e]. Dissected hindlimb extensor digitorum longus (EDL) muscles were enzymatically digested in Dulbecco\\u0026rsquo;s modified Eagle\\u0026rsquo;s medium (DMEM; Hyclone) containing 2.5% HEPES (Sigma-Aldrich) and collagenase II (800 units/mL, Worthington) at 37℃ for 60 min. Digested muscles were blocked in Dulbecco\\u0026rsquo;s modified Eagle\\u0026rsquo;s medium and 10% horse serum (Hyclone). The single myofibers were released by gentle trituration. Undamaged and noncontracted single myofibers were then washed with PBS several times and collected for immunocytochemistry and RNA extraction.\\u003c/p\\u003e \\u003cdiv id=\\\"Sec8\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eMuscle stem cell (MuSC) isolation\\u003c/h2\\u003e \\u003cp\\u003eIsolation of MuSC was performed according to a previously reported protocol with modifications [\\u003cspan citationid=\\\"CR48\\\" class=\\\"CitationRef\\\"\\u003e48\\u003c/span\\u003e]. Limb muscles were dissected and mechanically dissociated in DMEM containing 10% horse serum, collagenase II (800 units/mL), and dispase (1.1 units/mL, Thermo Fisher Scientific) at 37℃ for 40 min. Digested suspensions were subsequently triturated by sterilized syringes with 20G 1/2 needle (BD Biosciences) and washed with DMEM to harvest mononuclear cells. Mononuclear cells were stained with anti-Sca-1-Pacific blue (Biolegend), anti-CD31-APC (Biolegend), anti-CD45\\u0026ndash;APC (Biolegend), and anti-Vcam1-Biotin (BD Biosciences). PE-Cy7- Streptavidin (Biolegend) was used as a secondary reagent. To exclude dead cells, 7-AAD (Sigma-Aldrich) was used. Stained cells were analyzed and Vcam1\\u003csup\\u003e+\\u003c/sup\\u003eSca1\\u003csup\\u003e\\u0026minus;\\u003c/sup\\u003e 7-AAD\\u003csup\\u003e\\u0026ndash;\\u003c/sup\\u003eCD31\\u0026ndash; CD45\\u0026ndash; MuSCs were isolated using FACS Aria III cell sorter (BD Biosciences) with 4-way-purity precision. FACS gating strategy was referred to the previously reported protocol [\\u003cspan citationid=\\\"CR48\\\" class=\\\"CitationRef\\\"\\u003e48\\u003c/span\\u003e]. Isotype control density plots were used as a reference for positive gating. Freshly isolated MuSCs were attached to a slide glass by cytospin for immunocytochemistry, or collected for RNA and protein extraction.\\u003c/p\\u003e \\u003c/div\\u003e\\n\\u003ch3\\u003eImmunohistochemistry\\u003c/h3\\u003e\\n\\u003cp\\u003eFreshly dissected TA or Soleus muscles were embedded in O.C.T., snap-frozen in liquid nitrogen, and stored at -80\\u0026deg;C prior to sectioning. Cross-sectional 7 \\u0026micro;m-thick sections were obtained from the embedded muscles using a cryostat. For myosin heavy chain (MyHC) staining, unfixed muscle sections were incubated overnight at 4\\u0026deg;C with mouse anti-MyHC type 1 (DSHB, BA-D5, 1:10) or mouse anti-MyHC type 2x (DSHB, 6H-1, 1:5) in addition to rat anti-laminin (Abcam, ab11576, 1:1000 dilution) in 3% BSA blocking buffer. After washes in PBS, sections were incubated for 1 h with 1:500 dilution of Alexa Fluor 488-goat anti-mouse MIgG2b (Invitrogen), or Alexa Fluor 488-conjugated anti-mouse IgM (Invitrogen), and Alexa Fluor 594-conjugated anti-rat IgG (Invitrogen). For staining myoblasts, sections were fixed in 4% paraformaldehyde (PFA) for 10 min, and washed in PBS. Antigen retrieval was then performed in citrate buffer (10 mM citric acid, pH 6) at 95\\u0026deg;C 15 min. The sections were blocked by mouse Ig blocking reagent and blocking buffer from M.O.M. Kit (Vector Laboratories), according to the manufacturer\\u0026rsquo;s protocol. Then, the sections were incubated with primary antibodies in the blocking buffer at 4\\u0026deg;C overnight. The primary antibodies used include mouse anti-Pax7 (1:100, DSHB), rabbit anti-Ki67 (1:500, Sigma-Aldrich), mouse anti-MyoG (1:100, DSHB), rabbit anti-Dystrophin (1:500, Abcam), rabbit anti-Laminin (1:500, Sigma-Aldrich), and rabbit anti-cleaved Caspase-3 (1:200, Cell Signaling Technologies). After washing the sections with PBS, the sections were stained with secondary antibodies for 1 hr at RT, washed, and mounted. The secondary antibodies were used at a concentration of 1:400 and include goat anti-Rabbit IgG-Alexa Fluor 488 (Thermo Fisher Scientific), goat anti-Rabbit IgG-Alexa Fluor 594 (Thermo Fisher Scientific), and goat anti-Mouse IgG-Alexa Fluor 594 (Thermo Fisher Scientific). Hoechst 33342 (1:2,000, Thermo Fisher Scientific) was used to visualize nuclei. For EdU staining, we used the Click-iT EdU Alexa Fluor 488 Imaging Kit (Thermo Fisher Scientific) following the manufacturer\\u0026rsquo;s protocol before the blocking step. The number of each cell type and myofibers was counted in the total TA or Soleus area, and representative images were selected in the same region of the section used in the cell counting. Imaging was conducted with EVOS FL Auto 2 (Thermo Fisher Scientific).\\u003c/p\\u003e\\n\\u003ch3\\u003eImmunocytochemistry\\u003c/h3\\u003e\\n\\u003cp\\u003eFor isolated myofibers staining, freshly isolated myofibers were fixed in 4% PFA for 10 min at room temperature (RT), quenched in 0.1M glycine in PBS for 10 min at RT, and blocked for 1 hour at RT by blocking buffer (5% goat serum and 5% bovine serum albumin in PBS/0.4% Triton X-100). Then, the myofibers were incubated with mouse anti-Pax7 (1:100, DSHB) in the blocking buffer at 4\\u0026deg;C overnight. After washing the myofibers three times with PBS/0.1% Triton X-100, the myofibers were stained with goat anti-Mouse IgG-Alexa Fluor 594 (1:400, Thermo Fisher Scientific) and Hoechst 33342 (1:5,000, Thermo Fisher Scientific) for 1 hour at RT, washed and mounted on slide glass. Imaging was conducted with EVOS FL Auto 2 (Thermo Fisher Scientific). For isolated MuSCs staining, freshly isolated MuSCs were attached to a slide glass by cytospin and fixed by 4% PFA for 10 min at RT. The fixed MuSCs were washed with PBS/0.4% Triton X-100 several times and blocked with blocking buffer (5% goat serum and 5% bovine serum albumin in PBS/0.4% Triton X-100) for 1 hour at RT and incubated with mouse anti-Pax7 (1:100, DSHB) and rabbit anti-MyoD (1:200, Santa Cruz) overnight at 4\\u0026deg;C. The slides were washed with PBS/0.1% Triton X-100 several times and incubated with goat anti-Mouse IgG-Alexa Fluor 594 (1:400, Thermo Fisher Scientific) and goat anti-Rabbit IgG-Alexa Fluor 488 (1:400, Thermo Fisher Scientific). For EdU staining, we used the Click-iT EdU Alexa Fluor 647 Imaging Kit (Thermo Fisher Scientific) following the manufacturer\\u0026rsquo;s protocol before the blocking step. The slides were counterstained with Hoechst 33342 (Thermo Fisher Scientific) and mounted. Imaging was conducted with EVOS FL Auto 2 (Thermo Fisher Scientific).\\u003c/p\\u003e \\u003cdiv id=\\\"Sec11\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eFour limb grip strength measurement\\u003c/h2\\u003e \\u003cp\\u003eGrip strength was assessed by using a grip strength test meter (grip strength test BIO-GS3, Bioseb). Mice were allowed to grasp a grid attached to the tester with 4 limbs and were manually pulled in a horizontal direction by the tip of the tail. The test was performed 5 times with 10 min of resting between each measurement. The average of the top 3 result value (N, Newton) was normalized to body weight (g) (N/g). All experiments were performed in a blinded fashion.\\u003c/p\\u003e \\u003c/div\\u003e \\u003cdiv id=\\\"Sec12\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eChronic exercise training and endurance running test\\u003c/h2\\u003e \\u003cp\\u003eRandomized mice were pre-acclimated to the treadmill (DJ2-242, Dual Treadmill, Daejeon, Korea) before training. The scheme consists of exploration (0m/min for 5 min), and subsequent running (5m/min for 5 min, 10m/min for 5 min, 15m/min for 5 min). After 3 days of acclimation, mice were subjected to chronic exercise training for 5 weeks, 5 days per week with a protocol of 5m/min for 5min, 10m/min for 5min, 15m/min for 30 min. To test endurance running capacity, mice were allowed to run until exhaustion with speed set to 10m/min for 30 min and incremented by 2m/min every 20 min with no inclination. Exhaustion was defined as the condition where mice remained stationary at the end of the treadmill for more than 10 seconds despite mechanical stimulation. All experiments were performed in a blinded fashion.\\u003c/p\\u003e \\u003c/div\\u003e \\u003cdiv id=\\\"Sec13\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eRNA extraction and quantitative real-time polymerase chain (qRT-PCR)\\u003c/h2\\u003e \\u003cp\\u003eTotal RNA was extracted from freshly isolated myofibers and MuSCs using a TRIzol Reagent (Life Technologies) and analyzed by qRT-PCR. First-strand complementary DNA was synthesized from 1 \\u0026micro;g of RNA using ReverTra Ace (Toyobo) containing random oligomer according to the manufacturer\\u0026rsquo;s instructions. qRT-PCR (Qiagen) was performed with SYBR Green technology (SYBR Premix Ex Taq, Qiagen) using specific primers against indicated genes. Relative mRNA levels were determined using the 2\\u003csup\\u003e\\u0026minus;ΔΔCt\\u003c/sup\\u003e method and normalized to \\u003cem\\u003eGapdh\\u003c/em\\u003e. Primers are listed in Supplementary Table\\u0026nbsp;1.\\u003c/p\\u003e \\u003c/div\\u003e \\u003cdiv id=\\\"Sec14\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eWestern blot\\u003c/h2\\u003e \\u003cp\\u003eDissected TA muscles were homogenized in RIPA buffer (50 mM Tris-HCl, pH 7.5, 0.5% SDS, 20 \\u0026micro;g/ mL aprotinin, 20 \\u0026micro;g/ mL leupeptin, 10 \\u0026micro;g/ mL phenylmethylsulfonyl fluoride, 1 mM sodium orthovanadate, 10 mM sodium pyrophosphate, 10 mM sodium fluoride, and 1 mM dithiothreitol). Cell lysates were centrifuged at 13,000 rpm for 15 min. Supernatants were collected and subjected to immunoblot. BCA protein assay (Thermo Fisher Scientific) was used for estimating total protein concentrations. Normalized total proteins were analyzed by electrophoresis in 12% polyacrylamide gels and transferred to PVDF membranes (Millipore). Membranes were blocked in 5% skim milk (BD Biosciences) in TBS with 0.1% Tween-20 and incubated with rabbit anti-Histone H3 (Cell Signaling Technology) and rabbit anti-H3K4me1 (Cell Signaling Technology) overnight at 4\\u0026deg;C. After incubation with goat anti-Rabbit IgG-HRP (Promega), the membranes were developed using a Fusion solo chemiluminescence imaging system (Vilber). Histone H3 was used as a loading control. Primary and secondary antibodies were diluted 1:1000 and 1:10000 with PBS containing 0.1% Tween-20 and 3% BSA, respectively.\\u003c/p\\u003e \\u003c/div\\u003e \\u003cdiv id=\\\"Sec15\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eStatistical analysis\\u003c/h2\\u003e \\u003cp\\u003eSample size determination was based on anticipated variability and effect size that was observed in the investigator\\u0026rsquo;s lab for similar experiments. For quantification, individuals performing the counts were blinded to sample identity and randomized. All statistical analyses were performed using GraphPad Prism 9 (GraphPad Software). For comparison of significant differences in multiple groups for normally distributed data, statistical analysis was performed by one-way or two-way ANOVA followed by Tukey\\u0026rsquo;s pairwise comparison post hoc test. For non-normally distributed data, Brown\\u0026ndash;Forsythe and Welch ANOVA followed by the Games-Howell multiple comparisons test was used. For the comparison of the two groups, Student\\u0026rsquo;s unpaired t-test assuming a two-tailed distribution with Welch\\u0026rsquo;s correction was used. Unless otherwise noted, all error bars represent s.e.m. The number of biological replicates and statistical analyses for each experiment were indicated in the figure legends. Independent experiments were performed at least in triplicates.\\u003c/p\\u003e \\u003c/div\\u003e\"},{\"header\":\"Results\",\"content\":\"\\u003cp\\u003e \\u003cb\\u003eMll4\\u003c/b\\u003e \\u003cb\\u003edeletion in myofiber alters CSA and slow fiber composition in males, but not in female mice\\u003c/b\\u003e\\u003c/p\\u003e \\u003cp\\u003ePreviously, Liu and colleagues [\\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e] reported that the deletion of \\u003cem\\u003eMll4\\u003c/em\\u003e in developing myofibers did not impact overall muscle mass but resulted in changes to muscle characteristics, including an increased cross-sectional area (CSA) and a shift from slow to fast fiber types. However, the results were derived exclusively from male mice. Given that Mll4 is expressed in both sexes (Supplementary Fig.\\u0026nbsp;1A), we investigated whether female mice would exhibit a similar phenotype to males following \\u003cem\\u003eMll4\\u003c/em\\u003e ablation. We crossed \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e mice that carry loxP sites of the \\u003cem\\u003eMll4\\u003c/em\\u003e gene [\\u003cspan citationid=\\\"CR19\\\" class=\\\"CitationRef\\\"\\u003e19\\u003c/span\\u003e], with \\u003cem\\u003eMCK\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eCre/+\\u003c/em\\u003e\\u003c/sup\\u003e mice [\\u003cspan citationid=\\\"CR26\\\" class=\\\"CitationRef\\\"\\u003e26\\u003c/span\\u003e], which produced Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice (\\u003cem\\u003eMCK\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eCre/+\\u003c/em\\u003e\\u003c/sup\\u003e; \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e) [\\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e]. By performing quantitative real-time PCR (qRT-PCR), the deletion of \\u003cem\\u003eMll4\\u003c/em\\u003e was confirmed in 8-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e myofibers (Supplementary Fig.\\u0026nbsp;1B-C). Muscle weight remained unchanged in 8-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice, regardless of sex (Supplementary Fig.\\u0026nbsp;1D-E). As reported [\\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e], male Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice displayed increased CSA (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eA) and fiber type shifts toward a decreased ratio of slow-twitch fiber in soleus muscle (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eC-E). In contrast, no significant differences in these characteristics were observed in Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e female mice compared to Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e female mice (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eB, \\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eC, and \\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eF-G), suggesting that the alterations in CSA and fiber type composition following the deletion of Mll4 in myofibers may represent a phenotype selectively characteristic of male musculature, potentially attributable to male-specific factors or microenvironments.\\u003c/p\\u003e \\u003cp\\u003e \\u003c/p\\u003e \\u003cdiv id=\\\"Sec17\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eMuSC depletion in adult Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice\\u003c/h2\\u003e \\u003cp\\u003eUnexpectedly, when isolating single myofibers to prove the ablation of Mll4 in Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig2\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eA), we observed that myofiber length was significantly shorter in both male and female Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice compared to those of controls (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig2\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eB and \\u003cspan refid=\\\"Fig2\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eE). Since shortened myofibers may be due to a reduced number of myonuclei [\\u003cspan citationid=\\\"CR27\\\" class=\\\"CitationRef\\\"\\u003e27\\u003c/span\\u003e], we quantified myonuclear number, which was markedly decreased in Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e EDL myofibers compared to the controls (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig2\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eC and \\u003cspan refid=\\\"Fig2\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eF). To assess whether myonuclear density was also reduced, we calculated the ratio of myonuclear number to myofiber length. This analysis revealed a decrease in myonuclear density in Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e myofibers (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig2\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eD and \\u003cspan refid=\\\"Fig2\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eG).\\u003c/p\\u003e \\u003cp\\u003e \\u003c/p\\u003e \\u003cp\\u003eDuring postnatal development in mice, myofiber length and myonuclear number increase rapidly through myoblast fusion until puberty and then become relatively stable at the adult stage when postnatal myogenesis ceases [\\u003cspan citationid=\\\"CR1\\\" class=\\\"CitationRef\\\"\\u003e1\\u003c/span\\u003e]. Thus, reduced myonuclear density in adult Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice might be due to impaired myoblast fusion, defective MuSC differentiation, or even MuSC depletion. To address this issue, we first assessed the number of MuSCs in \\u003cem\\u003eMll4\\u003c/em\\u003e-deleted myofibers (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig2\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eH). Intriguingly, Pax7\\u003csup\\u003e+\\u003c/sup\\u003e MuSCs greatly decreased in the myofibers of both male and female 8-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig2\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eI-J). Our data suggest that MuSC depletion in \\u003cem\\u003eMll4\\u003c/em\\u003e-lacking myofibers during postnatal myogenesis resulted in decreased myonuclei accretion and myofiber growth, and that \\u003cem\\u003eMll4\\u003c/em\\u003e in myofiber may play an important role in maintaining the MuSC population, irrespective of gender.\\u003c/p\\u003e \\u003cp\\u003e \\u003cb\\u003eMuSC depletion in\\u003c/b\\u003e \\u003cb\\u003eMll4\\u003c/b\\u003e \\u003cb\\u003edeleted muscle during postnatal muscle growth\\u003c/b\\u003e\\u003c/p\\u003e \\u003cp\\u003eMuSCs that have actively proliferated during the juvenile stage enter a quiescent state at puberty to establish a reserve stem cell pool in adult muscles [\\u003cspan citationid=\\\"CR2\\\" class=\\\"CitationRef\\\"\\u003e2\\u003c/span\\u003e]. To investigate if the deletion of Mll4 in myofibers affects MuSC number during postnatal myogenesis, we conducted a histological analysis to quantify Pax7-positive cells in TA muscles during and after postnatal myogenesis. Considering that MCK-Cre mediated ablation of \\u003cem\\u003eMll4\\u003c/em\\u003e occurs after 7 days of birth [\\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e], 0-week-old perinatal muscles were expected to have comparable MuSC numbers between Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e muscles. The number of MuSCs remained consistent between the control and Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e muscles until 2 weeks of age (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eD). However, a decrease of Pax7\\u003csup\\u003e+\\u003c/sup\\u003e MuSCs was prominent in the pubertal 4-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e TA muscle, with a further decline noted in the 8-week-old muscle (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eA and \\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eD). This indicates that the deletion of myofiber-specific \\u003cem\\u003eMll4\\u003c/em\\u003e disrupts the MuSC number during postnatal myogenesis, resulting in a depletion of the population in adult muscles.\\u003c/p\\u003e \\u003cp\\u003e \\u003c/p\\u003e \\u003cp\\u003eTA and EDL muscles primarily consist of fast-twitch type 2 fibers [\\u003cspan citationid=\\\"CR28\\\" class=\\\"CitationRef\\\"\\u003e28\\u003c/span\\u003e]. To examine if MuSC depletion occurs in slow-twitch type 1 fiber-rich muscles, the soleus muscle was analyzed. Undoubtedly, the Pax7-positive MuSC number was reduced in the pubertal 4-week-old muscles and further diminished in the adult 8-week-old soleus muscles of Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eB and \\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eE).\\u003c/p\\u003e \\u003cp\\u003eDuring puberty, cycling MuSCs exit the cell cycle and contribute to quiescent MuSC populations [\\u003cspan citationid=\\\"CR2\\\" class=\\\"CitationRef\\\"\\u003e2\\u003c/span\\u003e]. To test if the reduced MuSC in Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice was due to impaired cell cycle exit in cycling pubertal MuSCs, we quantified Ki67-positive MuSCs in the 4-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice. Compared to the control, Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice showed a twofold increase in proliferating Ki67\\u003csup\\u003e+\\u003c/sup\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e MuSCs (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eC and \\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eF). To investigate if these proliferating MuSCs enter the differentiation state, we isolated MuSCs via cytometry (Supplementary Fig.\\u0026nbsp;2A-B) and quantified MyoD-positive MuSCs. Compared to the Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e mice, Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice showed an increase in MyoD-positive MuSCs (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eH-I). Furthermore, to label cycling MuSCs, EdU was treated for 2 consecutive days before isolating MuSCs (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eG). Notably, the number of MyoD-positive cells also increased among cycling MuSCs (Edu\\u003csup\\u003e+\\u003c/sup\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e) in Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eJ). This indicates that while normal MuSCs exit the cell cycle and enter a quiescent state during puberty, MuSCs in Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e mice maintain their cell cycle and differentiate into committed myoblasts. Altogether, the deletion of Mll4 in myofibers leads to the loss of MuSCs during postnatal muscle growth.\\u003c/p\\u003e \\u003c/div\\u003e \\u003cdiv id=\\\"Sec18\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eDeletion of Mll4 in adult myofibers does not alter muscle CSA and fiber type composition\\u003c/h2\\u003e \\u003cp\\u003eFollowing postnatal myogenesis, the adult muscle tissue reaches a steady state characterized by the cessation of myofiber growth and the entry of MuSCs into a quiescent phase. To investigate if induced ablation of \\u003cem\\u003eMll4\\u003c/em\\u003e in muscles after postnatal myogenesis would affect myofiber maintenance, we examined muscle features such as CSA distribution and fiber type composition of \\u003cem\\u003eMll4\\u003c/em\\u003e ablated adult muscles. Adult \\u003cem\\u003eHSA\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eMerCreMer/+\\u003c/em\\u003e\\u003c/sup\\u003e; \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e mice (Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e) were treated with tamoxifen for consecutive 5 days to induce deletion of the \\u003cem\\u003eMll4\\u003c/em\\u003e gene in myofibers (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig4\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eA). This resulted in the ablation of the \\u003cem\\u003eMll4\\u003c/em\\u003e gene in the myofibers of Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice after 2 weeks of tamoxifen administration (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig4\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eB). The histological analysis showed that CSA distribution, fiber number, and fast-twitch (MyHC2x) fiber composition in the TA muscles of Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice did not change following 2 weeks (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig4\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eC-E, Supplementary Fig.\\u0026nbsp;3A) and even 4 weeks (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig4\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eH-J, Supplementary Fig.\\u0026nbsp;3B) of \\u003cem\\u003eMll4\\u003c/em\\u003e ablation, compared to the control mice. Similarly, in the soleus muscle, the composition of slow-twitch (MyHC1) fibers also remained unchanged after both 2 weeks (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig4\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eF-G, Supplementary Fig.\\u0026nbsp;3A) and 4 weeks (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig4\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eK-L, Supplementary Fig.\\u0026nbsp;3B) of \\u003cem\\u003eMll4\\u003c/em\\u003e ablation in Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice. This indicates that the induced deletion of \\u003cem\\u003eMll4\\u003c/em\\u003e in adult muscles does not affect myofiber maintenance and intracellular features such as fiber CSA and fiber type composition regardless of gender.\\u003c/p\\u003e \\u003cp\\u003e \\u003c/p\\u003e \\u003c/div\\u003e \\u003cdiv id=\\\"Sec19\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eMll4 deletion in myofibers does not affect exercise capacity\\u003c/h2\\u003e \\u003cp\\u003eSince MLL4 functions as an enhancer activator [\\u003cspan citationid=\\\"CR19\\\" class=\\\"CitationRef\\\"\\u003e19\\u003c/span\\u003e], its deletion may disrupt various gene transcription processes. Although the deletion of Mll4 in adult muscles does not impact myofiber characteristics such as CSA and fiber type composition, we investigated whether the deletion affects exercise capacity. To test this, tamoxifen-treated mice were accustomed to chronic exercise training for 5 weeks (hereafter, Mll4\\u003csup\\u003eWT\\u0026thinsp;\\u0026minus;\\u0026thinsp;EX\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA\\u0026minus;EX\\u003c/sup\\u003e) (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig5\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eA). The protocol of the chronic exercise provides prolonged contractions of muscles, promoting adaptations such as increased muscle mass while minimizing exercise-induced muscle damage [\\u003cspan citationid=\\\"CR29\\\" class=\\\"CitationRef\\\"\\u003e29\\u003c/span\\u003e]. In TA muscles, CSA distribution and fast fiber composition remained consistent between Mll4\\u003csup\\u003eWT\\u0026thinsp;\\u0026minus;\\u0026thinsp;EX\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA\\u0026minus;EX\\u003c/sup\\u003e mice. In addition, slow fiber composition was unchanged in Mll4\\u003csup\\u003eΔHSA\\u0026minus;EX\\u003c/sup\\u003e soleus muscle (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig5\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eB-F, Supplementary Fig.\\u0026nbsp;4A).\\u003c/p\\u003e \\u003cp\\u003e \\u003c/p\\u003e \\u003cp\\u003eTo assess whether exercise capacity was affected by Mll4 deletion, we measured grip strength and endurance running capability. In line with the observed similarity in CSA and fiber type composition, Mll4\\u003csup\\u003eΔHSA\\u0026minus;EX\\u003c/sup\\u003e mice showed comparable grip strength (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig5\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eG) and endurance running capability (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig5\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eH-I) relative to the control group. To investigate whether the expression of genes related to fiber type and metabolism was altered in Mll4\\u003csup\\u003eΔHSA\\u0026minus;EX\\u003c/sup\\u003e mice, we conducted qRT-PCR analyses. Transcriptional profiling revealed that before exercise training, the genes were generally downregulated in Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice (Supplementary Fig.\\u0026nbsp;5A-B). Notably, no particular gene exhibited higher expression levels that could induce a shift in fiber type composition or metabolic activity. After 5 weeks of chronic exercise, the expression of genes related to fiber type and metabolism of Mll4\\u003csup\\u003eΔHSA\\u0026minus;EX\\u003c/sup\\u003e mice also showed overall downregulation, with the exception that certain slow-twitch muscle genes were marginally upregulated (Supplementary Fig.\\u0026nbsp;5C). Collectively, the results suggest that the induced knockout of \\u003cem\\u003eMll4\\u003c/em\\u003e in myofibers at the adult stage does not influence exercise capacity or muscle characteristics, such as CSA and fiber type composition, even after physiological exercise stimulation.\\u003c/p\\u003e \\u003c/div\\u003e \\u003cdiv id=\\\"Sec20\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eLoss of adult MuSCs in Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice\\u003c/h2\\u003e \\u003cp\\u003eTo investigate whether the deletion of \\u003cem\\u003eMll4\\u003c/em\\u003e in adult myofibers disturbs the quiescence of MuSCs, TA and soleus muscles were analyzed to quantify Pax7-positive MuSCs. Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice showed depletion of MuSCs in both muscles (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig6\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eA and \\u003cspan refid=\\\"Fig6\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eC). To assess whether the loss of adult quiescent MuSCs in Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice is associated with their entry into the cell cycle, we quantified Ki67-positive MuSCs in the TA muscles. In Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e TA muscles, there was a negligible presence of Ki67\\u003csup\\u003e+\\u003c/sup\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e cells, whereas Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e muscles showed an increase of Ki67\\u003csup\\u003e+\\u003c/sup\\u003ePax7\\u003csup\\u003e+\\u003c/sup\\u003e cells (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig6\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eD and \\u003cspan refid=\\\"Fig6\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eE). This suggests that ablation of \\u003cem\\u003eMll4\\u003c/em\\u003e in myofibers at the adult stage causes MuSCs to exit quiescence and enter the cell cycle, leading to the depletion of MuSCs. To examine if the cell cycle entry of adult Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice leads to a differentiation state, we performed an immunocytochemistry assay on sorted MuSCs (Supplementary Fig.\\u0026nbsp;2C-D) to quantify MyoD-positive cells. Compared to the Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e MuSCs, Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e MuSCs showed an increased number of MyoD-expressing cells (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig6\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eG-H). For detecting MuSCs that entered the cell cycle, we treated EdU for 2 weeks in Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig6\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eF). This long-term EdU labeling method was applied to identify the scarcely dividing MuSCs in adult muscle tissue [\\u003cspan citationid=\\\"CR30\\\" class=\\\"CitationRef\\\"\\u003e30\\u003c/span\\u003e]. We found that the population of MyoD-expressing cells among EdU-positive, dividing MuSCs was also increased in Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e muscles (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig6\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eI). On the other hand, given that adult quiescent MuSCs can differentiate without entering the cell cycle [\\u003cspan citationid=\\\"CR30\\\" class=\\\"CitationRef\\\"\\u003e30\\u003c/span\\u003e], we also quantified MyoD-expressing cells among EdU-negative, non-dividing MuSCs. Interestingly, Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e MuSCs showed an increased number of MyoD-positive cells among EdU-negative MuSCs (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig6\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eJ). This indicates that the deletion of Mll4 in adult muscles causes adult MuSCs to lose their quiescence and undergo differentiation, either with or without dividing.\\u003c/p\\u003e \\u003cp\\u003e \\u003c/p\\u003e \\u003cp\\u003eTo test whether the differentiated myogenic progeny of Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e muscles fuse into myofibers, we conducted a histological analysis and quantified EdU-positive nuclei located on the inner side of dystrophin structure [\\u003cspan citationid=\\\"CR30\\\" class=\\\"CitationRef\\\"\\u003e30\\u003c/span\\u003e] (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig5\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eK). While the number of fiber-incorporating EdU-positive nuclei was extremely low in control muscles, it was sevenfold higher in Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e muscles compared to the control group (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig5\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eL). This suggests that \\u003cem\\u003eMll4\\u003c/em\\u003e deficiency in adult muscles causes MuSCs to exit quiescence and undergo differentiation, with at least some, or perhaps all, of these differentiated cells subsequently fusing into myofibers. This eventually results in severe MuSC loss in adult muscles.\\u003c/p\\u003e \\u003cp\\u003e \\u003cb\\u003eLack of\\u003c/b\\u003e \\u003cb\\u003eMll4\\u003c/b\\u003e \\u003cb\\u003eimpairs muscle regeneration following injury\\u003c/b\\u003e\\u003c/p\\u003e \\u003cp\\u003eMuSCs are the primary cell type that contributes to muscle regeneration capacity. To investigate if MuSC depletion in Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice leads to hindered muscle regeneration, we subjected TA muscles of Mll4\\u003csup\\u003eWT\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice to injury using BaCl\\u003csub\\u003e2\\u003c/sub\\u003e (hereafter, Mll4\\u003csup\\u003eWT\\u0026thinsp;\\u0026minus;\\u0026thinsp;inj\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA\\u0026minus;inj\\u003c/sup\\u003e) (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig7\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eA). Muscles were analyzed 10 days post-injury, as the majority of regenerating fibers are restored [\\u003cspan citationid=\\\"CR31\\\" class=\\\"CitationRef\\\"\\u003e31\\u003c/span\\u003e]. TA muscles of Mll4\\u003csup\\u003eΔHSA\\u0026minus;inj\\u003c/sup\\u003e mice showed reduced muscle mass compared to that of Mll4\\u003csup\\u003eWT\\u0026thinsp;\\u0026minus;\\u0026thinsp;inj\\u003c/sup\\u003e mice, while adjacent muscles such as the EDL, GA, and soleus remained unaffected (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig7\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eB). Histological analysis of muscle sections revealed active muscle regeneration in Mll4\\u003csup\\u003eWT\\u0026thinsp;\\u0026minus;\\u0026thinsp;inj\\u003c/sup\\u003e muscles, as indicated by the predominance of myofibers with centrally located nuclei and relatively homogenous fiber sizes. Conversely, Mll4\\u003csup\\u003eΔHSA\\u0026minus;inj\\u003c/sup\\u003e muscle was observed with disorganized tissue architecture with residual damaged fibers that failed to undergo effective regeneration. (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig7\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eC). Moreover, Mll4\\u003csup\\u003eΔHSA\\u0026minus;inj\\u003c/sup\\u003e mice showed reduced CSA of regenerating fibers (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig7\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eD). These results suggest that lack of MuSCs due to the deletion of myofiber-specific Mll4 resulted in severely impinged muscle regeneration capacity.\\u003c/p\\u003e \\u003cp\\u003e \\u003c/p\\u003e \\u003c/div\\u003e \\u003cdiv id=\\\"Sec21\\\" class=\\\"Section2\\\"\\u003e \\u003ch2\\u003eMll4 in myofibers affects the MuSC niche by regulating Notch ligand expression\\u003c/h2\\u003e \\u003cp\\u003eTo explore how \\u003cem\\u003eMll4\\u003c/em\\u003e in myofibers may have affected the MuSC niche, we screened for downstream effector candidate genes by analyzing public datasets. For one, we analyzed data curated by Liu et al. [\\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e]. This provided a list of downregulated genes in muscle from MLL4-SET-knockout (KO) mice, where the enzymatic SET domain of MLL4 is ablated, compared to control mice. In addition, we analyzed data from Lee et al. [\\u003cspan citationid=\\\"CR19\\\" class=\\\"CitationRef\\\"\\u003e19\\u003c/span\\u003e], to obtain the list of downregulated genes in cultured, differentiated Mll4-KO myocytes versus control. Sixty-six genes were identified as commonly downregulated from the two datasets. Since myofiber can directly regulate the MuSC niche via signaling through ligand-receptor interactions [\\u003cspan citationid=\\\"CR32\\\" class=\\\"CitationRef\\\"\\u003e32\\u003c/span\\u003e], we then identified ligands from the 66 candidate genes by comparing them to the mouse ligand database. Consequently, 5 genes were identified as ligand-coding genes that are downregulated by Mll4 KO in both whole muscle and differentiated myocytes. To our surprise, the Notch ligand Jag2 was among the 5 candidate genes. Also, Dll1, another Notch ligand, was identified as downregulated in Mll4 KO myocytes. (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eA).\\u003c/p\\u003e \\u003cp\\u003e \\u003c/p\\u003e \\u003cp\\u003eFor MuSCs, Notch signaling is a major signaling pathway that maintains the stem cell pool. When the Notch downstream effector Rbpj is deleted in adult MuSCs, which are predominantly in a quiescent state, they exit the quiescent state and undergo aberrant differentiation [\\u003cspan citationid=\\\"CR30\\\" class=\\\"CitationRef\\\"\\u003e30\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR33\\\" class=\\\"CitationRef\\\"\\u003e33\\u003c/span\\u003e]. Myofiber-specific deletion of Dll4, a Notch ligand that is mainly expressed in adult myofibers, induces premature differentiation of MuSCs, resulting in a reduced number of stem cells [\\u003cspan citationid=\\\"CR17\\\" class=\\\"CitationRef\\\"\\u003e17\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR34\\\" class=\\\"CitationRef\\\"\\u003e34\\u003c/span\\u003e]. These studies suggest that the maintenance of MuSC quiescence is highly dependent on Notch signaling between MuSCs and myofibers. Considering that the depletion of MuSCs was observed in both Notch signaling-reduced MuSCs and myofiber-specific \\u003cem\\u003eMll4-\\u003c/em\\u003edeleted (Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e) MuSCs, we sought to validate the downregulation of Notch ligands in Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e and Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e mice.\\u003c/p\\u003e \\u003cp\\u003eA previous study found that \\u003cem\\u003eDll4\\u003c/em\\u003e and \\u003cem\\u003eJag2\\u003c/em\\u003e are dominantly expressed in adult myofibers, compared to other notch ligands [\\u003cspan citationid=\\\"CR17\\\" class=\\\"CitationRef\\\"\\u003e17\\u003c/span\\u003e]. In addition, we reported that myofibers of 4-week-old mice exhibited robust expression of \\u003cem\\u003eDll1\\u003c/em\\u003e and \\u003cem\\u003eJag1\\u003c/em\\u003e proteins [\\u003cspan citationid=\\\"CR2\\\" class=\\\"CitationRef\\\"\\u003e2\\u003c/span\\u003e]. Considering that Notch ligands have a fluctuating expression pattern in muscles throughout the developmental stages, we compared mRNA quantity for Notch ligands in the myofibers of wild-type pubertal 4-week-old and adult mice (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eB). Interestingly, genes having dominant expression during each time point correlated with genes that were downregulated due to \\u003cem\\u003eMll4\\u003c/em\\u003e depletion in muscle fibers. While expression of major Notch ligands of pubertal 4-week-old myofibers \\u0026ndash; \\u003cem\\u003eJag1\\u003c/em\\u003e and \\u003cem\\u003eDll1\\u003c/em\\u003e \\u0026ndash; decreased in 4-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e myofibers (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eC), the primary Notch ligands of adult myofibers \\u0026ndash; \\u003cem\\u003eJag2\\u003c/em\\u003e and \\u003cem\\u003eDll4\\u003c/em\\u003e \\u0026ndash; decreased in adult Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e myofibers (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eD). In other words, \\u003cem\\u003eMll4\\u003c/em\\u003e insufficiency in myofibers disturbed Notch ligand expression that is dominant in each pubertal or adult muscle.\\u003c/p\\u003e \\u003cp\\u003eTo test if Notch signaling is indeed reduced in 4-week-old Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e and adult Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e MuSCs, the mRNA levels of canonical Notch effectors \\u0026ndash; \\u003cem\\u003eHeyL, Hey1\\u003c/em\\u003e, and \\u003cem\\u003eHes1\\u003c/em\\u003e \\u0026ndash; were quantified via qRT-PCR. As expected, the overall expression of genes mentioned above was downregulated in both Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e and adult Mll4\\u003csup\\u003eΔHSA\\u003c/sup\\u003e MuSCs (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eE-F).\\u003c/p\\u003e \\u003cp\\u003eConsidering the molecular feature of MLL4, we analyzed public ChIP-seq data of MLL4 in myocytes [\\u003cspan citationid=\\\"CR19\\\" class=\\\"CitationRef\\\"\\u003e19\\u003c/span\\u003e], to examine whether it may modulate the transcription of Notch ligands on the chromosomal level. This revealed the genomic binding of MLL4 on \\u003cem\\u003eDll1\\u003c/em\\u003e and \\u003cem\\u003eJag2\\u003c/em\\u003e gene loci, where \\u003cem\\u003eMll4\\u003c/em\\u003e deletion reduced H3K4me1 and H3K27ac levels on enhancers for both \\u003cem\\u003eDll1\\u003c/em\\u003e and \\u003cem\\u003eJag2\\u003c/em\\u003e genes (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eG-H). This implicates the possibility of MLL4 directly regulating the induction of different Notch ligand gene expressions. Taken together, MLL4 can control diverse Notch ligand expression in myofibers, which is necessary for regulating the MuSC niche during and after postnatal myogenesis.\\u003c/p\\u003e \\u003c/div\\u003e\"},{\"header\":\"Discussion\",\"content\":\"\\u003cp\\u003eSkeletal muscle has a resilient characteristic due to its resident stem cell populations. Thus, uncovering the mechanism of regulating MuSC fate is crucial for understanding the biological process of developmental and regenerative myogenesis. In this paper, we explored the role of \\u003cem\\u003eMll4\\u003c/em\\u003e in myofibers regarding the MuSC state regulation and discovered that myofiber-expressed \\u003cem\\u003eMll4\\u003c/em\\u003e is important for maintaining MuSCs in both muscles during and after postnatal myogenesis. In the pubertal Mll4\\u003csup\\u003eΔMCK\\u003c/sup\\u003e muscle, lack of \\u003cem\\u003eMll4\\u003c/em\\u003e in myofibers resulted in increased population of differentiating myogenic cells, leading to a decrease of MuSCs. Furthermore, induced ablation of \\u003cem\\u003eMll4\\u003c/em\\u003e in adult myofibers resulted in the quiescence exit of MuSCs, which also caused dramatic depletion of MuSCs. During postnatal myogenesis, juvenile MuSCs constantly proliferate for muscle development [\\u003cspan citationid=\\\"CR1\\\" class=\\\"CitationRef\\\"\\u003e1\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR2\\\" class=\\\"CitationRef\\\"\\u003e2\\u003c/span\\u003e]. This proliferating cell population decreases due to cell cycle exit during puberty to establish a reserve pool of quiescent MuSCs in adult muscles [\\u003cspan citationid=\\\"CR3\\\" class=\\\"CitationRef\\\"\\u003e3\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR4\\\" class=\\\"CitationRef\\\"\\u003e4\\u003c/span\\u003e]. Our findings indicate that Mll4 in myofibers are critical for maintaining MuSCs in pubertal muscles, where cycling MuSCs begin to enter quiescence, as well as in adult muscles, where MuSCs remain in a quiescent state. This suggests that \\u003cem\\u003eMll4\\u003c/em\\u003e plays a critical role in myofibers by creating a microenvironment that supports the maintenance of a healthy population of MuSCs in muscle tissue.\\u003c/p\\u003e \\u003cp\\u003eThe ability to maintain an adequate number of MuSCs is crucial regardless of gender and age. In this study, we elucidate two critical properties of \\u003cem\\u003eMll4\\u003c/em\\u003e in preserving the stemness of MuSCs. First, \\u003cem\\u003eMll4\\u003c/em\\u003e regulates the MuSC number in both sexes. Skeletal muscle exhibits sexual dimorphism in terms of mass, fiber type composition, and contractility attributed to variations in gene expression and hormonal profiles between genders [\\u003cspan citationid=\\\"CR35\\\" class=\\\"CitationRef\\\"\\u003e35\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR36\\\" class=\\\"CitationRef\\\"\\u003e36\\u003c/span\\u003e]. These differences may have contributed to the disparate muscle phenotypes after the deletion of myofiber-\\u003cem\\u003eMll4\\u003c/em\\u003e in male and female mice (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eC-I). However, gender did not influence the extent of MuSC depletion resulting from \\u003cem\\u003eMll4\\u003c/em\\u003e ablation. Secondly, Mll4 regulates MuSC quiescence across different developmental stages, including both during and after postnatal myogenesis. Previous research indicated that myofiber-specific deletion of \\u003cem\\u003eMll4\\u003c/em\\u003e led to a slow-to-fast fiber type shift [\\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e]. However, our findings reveal that this phenotype is not present following \\u003cem\\u003eMll4\\u003c/em\\u003e deletion in adult muscle tissue. This suggests that Mll4 may play a role in the development of myofibers, but not in their maintenance during adulthood, at which developmental myogenesis is complete. Indeed, it is reported that during developmental myogenesis, Foxo/Notch signaling regulates fiber type specification, leading to a reduction in slow fibers and an increase in fast fibers when disrupted in muscle [\\u003cspan citationid=\\\"CR37\\\" class=\\\"CitationRef\\\"\\u003e37\\u003c/span\\u003e]. Considering that (1) this phenotype is in line with that of \\u003cem\\u003eMll4\\u003c/em\\u003e-mKO mice, as reported previously, and (2) Mll4 has the possibility of regulating the gene expression of Notch ligands, the deletion of \\u003cem\\u003eMll4\\u003c/em\\u003e might have disrupted Notch signaling in developing myofibers, leading to aberrant fiber type specification. Altogether, our data provide new insights into the role of Mll4 as a crucial regulator of the MuSC quiescence, highlighting its significance across developmental stages and irrespective of gender.\\u003c/p\\u003e \\u003cp\\u003eChromatin modification of enhancers within myofibers can modulate the expression of extracellular matrix (ECM) components or growth factors, thereby indirectly influencing the MuSC niche [\\u003cspan citationid=\\\"CR38\\\" class=\\\"CitationRef\\\"\\u003e38\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR39\\\" class=\\\"CitationRef\\\"\\u003e39\\u003c/span\\u003e]. Our study suggests that MLL4, an enhancer activator, regulates Notch ligand expression in myofibers to directly control MuSC quiescence. By analyzing and validating transcriptome databases from previous studies, we verified that the expression of Notch ligands was downregulated in Mll4-KO myocyte. Moreover, an analysis of ChIP-seq data revealed genomic binding of MLL4 on Notch ligand gene loci. The downregulation of Notch ligands in myofibers led to a reduced expression of canonical Notch target genes in MuSCs. This indicates that MLL4 can regulate the signaling pathway that affects adjacent cells. This finding is particularly intriguing given that MLL4 has primarily been studied as a critical factor for activating intracellular signaling pathways, including those related to cancer and cell fate determination [\\u003cspan citationid=\\\"CR19\\\" class=\\\"CitationRef\\\"\\u003e19\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR20\\\" class=\\\"CitationRef\\\"\\u003e20\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR40\\\" class=\\\"CitationRef\\\"\\u003e40\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR41\\\" class=\\\"CitationRef\\\"\\u003e41\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR42\\\" class=\\\"CitationRef\\\"\\u003e42\\u003c/span\\u003e]. Specifically, in myofibers, Mll4 was reported to activate the transcription of slow-twitch genes [\\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e]. By inspecting the physiological impact of \\u003cem\\u003eMll4\\u003c/em\\u003e deletion in myofibers on MuSCs, we revealed that \\u003cem\\u003eMll4\\u003c/em\\u003e regulates not only intracellular signaling pathways, as previously reported, but also signaling pathways that affect adjacent cells, by controlling expressions of ligand genes. This underscores the importance of exploring the potential gene-regulating activity of MLL4, which may impact other cellular processes, such as differentiation and tumorigenesis, in neighboring cells.\\u003c/p\\u003e \\u003cp\\u003eIt has been well-established that Notch signaling is a fundamental pathway regulating the MuSC niche in prenatal and postnatal muscles to maintain an appropriate stem cell pool [\\u003cspan citationid=\\\"CR11\\\" class=\\\"CitationRef\\\"\\u003e11\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR12\\\" class=\\\"CitationRef\\\"\\u003e12\\u003c/span\\u003e]. Previous studies reported that the myofiber is an important source of Notch ligands, sending signals to MuSCs to control their niche and hence their cell fate [\\u003cspan citationid=\\\"CR2\\\" class=\\\"CitationRef\\\"\\u003e2\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR13\\\" class=\\\"CitationRef\\\"\\u003e13\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR43\\\" class=\\\"CitationRef\\\"\\u003e43\\u003c/span\\u003e]. Notch ligands have distinct expression patterns in myofibers during development, affecting the MuSC niche in different ways. In this study, we investigated the Notch ligands with prevailing expression in different stages; \\u003cem\\u003eJag1\\u003c/em\\u003e and \\u003cem\\u003eDll1\\u003c/em\\u003e in pubertal myofibers, and \\u003cem\\u003eJag2\\u003c/em\\u003e and \\u003cem\\u003eDll4\\u003c/em\\u003e in adult muscle fibers. Interestingly, \\u003cem\\u003eMll4\\u003c/em\\u003e deletion in pubertal and adult myofibers disturbed the expression of Notch ligands that were principally expressed in each stage. Previously, Eliazer and colleagues reported that myofiber-specific deletion of Dll4 resulted in a reduction of MuSCs. However, the decrease in MuSCs was more pronounced when the pan-Notch regulator Mib1 was deleted from myofibers [\\u003cspan citationid=\\\"CR17\\\" class=\\\"CitationRef\\\"\\u003e17\\u003c/span\\u003e]. This implies the presence of a complementary Notch ligand acting as a signaling factor to maintain MuSC quiescence in adult muscles, together with Dll4. Our findings suggest that along with the well-known factor Dll4, Jag2 may be another Notch ligand in adult myofibers contributing to maintaining the Pax7\\u003csup\\u003e+\\u003c/sup\\u003e quiescent stem cells, both of which were found to be regulated by Mll4. Taken together, this study suggests that MLL4 functions as a regulator that modulates the expression of various Notch ligands in myofibers during both pubertal and adult stages. This regulation is essential for the precise control of MuSC quiescence throughout developmental stages.\\u003c/p\\u003e \\u003cp\\u003e \\u003cem\\u003eMll4\\u003c/em\\u003e deletion notably hindered H3K4me1 and H3K27ac levels on enhancers for both the \\u003cem\\u003eDll1\\u003c/em\\u003e gene in pubertal fibers and the \\u003cem\\u003eJag2\\u003c/em\\u003e gene in adult fibers, indicating different gene regulation of MLL4 in the two developmental stages. This may be attributed to the distinct pioneer transcription factors that recruit MLL4 to induce Notch ligand expression at different developmental stages. Several transcription factors \\u0026ndash; such as CCAAT/enhancer-binding protein family, myocyte enhancer factor 2 family, and Nrf1 \\u0026ndash; are identified to bind the DNA to recruit the MLL4 complex [\\u003cspan citationid=\\\"CR19\\\" class=\\\"CitationRef\\\"\\u003e19\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR21\\\" class=\\\"CitationRef\\\"\\u003e21\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR23\\\" class=\\\"CitationRef\\\"\\u003e23\\u003c/span\\u003e, \\u003cspan citationid=\\\"CR44\\\" class=\\\"CitationRef\\\"\\u003e44\\u003c/span\\u003e]. Depending on the cell type and differentiation stage, different transcription factors recruit MLL4 to regulate the expression of various genes. Transcription factors associated with Notch signaling have also been identified. A study on chicken embryos found that a transcription coregulator, Yap, binds to the enhancer of \\u003cem\\u003eJag2\\u003c/em\\u003e [\\u003cspan citationid=\\\"CR45\\\" class=\\\"CitationRef\\\"\\u003e45\\u003c/span\\u003e]. In mice, Notch1 ICD can act as a transcription activator in muscle fibers to activate the gene expression of \\u003cem\\u003eJag2\\u003c/em\\u003e and \\u003cem\\u003eDll4\\u003c/em\\u003e [\\u003cspan citationid=\\\"CR46\\\" class=\\\"CitationRef\\\"\\u003e46\\u003c/span\\u003e]. Following these studies, it is plausible that Yap and Notch ICD may recruit MLL4 to regulate the expression of different Notch ligands in myofibers. Further investigation is required to elucidate the molecular mechanism by which MLL4 regulates Notch ligand gene expression in myofibers. This includes identifying the specific pioneer transcription factors that interact with MLL4 across different developmental stages.\\u003c/p\\u003e\"},{\"header\":\"Conclusions\",\"content\":\"\\u003cp\\u003eOur results suggest a unique function of Mll4 in myofibers controlling MuSC state, possibly by orchestrating different Notch ligand expressions in various developmental stages. Moreover, despite skeletal muscle being known to exhibit sexual dimorphism, the role of Mll4 regulating MuSCs was valid in both male and female mice. By elucidating an additional mechanism governing MuSC maintenance, this research opens new avenues for the biological manipulation of muscle stem cells.\\u003c/p\\u003e\"},{\"header\":\"Abbreviations\",\"content\":\"\\u003cdiv class=\\\"DefinitionList\\\"\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eMuSCs\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eMuscle stem cells\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eMll4\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eMyeloid/lymphoid or mixed-lineage leukemia 4\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eJag1, Jag2\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eJagged 1, Jagged 2\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eDll1, Dll2\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eDelta-like protein 1, Delta-like protein 2\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eTA\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eTibialis anterior\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eSol\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eSoleus\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eGA\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eGastrocnemius\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eEDL\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eExtensor digitorum longus\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eCSA\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eCross-sectional area\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eMyHC1\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eMyosin Heavy Chain 1\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003cdiv class=\\\"DefinitionListEntry\\\"\\u003e \\u003cdiv class=\\\"Term\\\"\\u003eMyHC2x\\u003c/div\\u003e \\u003cdiv class=\\\"Description\\\"\\u003e \\u003cp\\u003eMyosin Heavy Chain 2x\\u003c/p\\u003e \\u003c/div\\u003e \\u003c/div\\u003e \\u003c/div\\u003e\"},{\"header\":\"Declarations\",\"content\":\"\\u003cp\\u003eEthics approval and consent to participate\\u003c/p\\u003e\\n\\u003cp\\u003eThe care and treatment of animals in this study were approved by the Institutional Animal Care and Use Committee (IACUC) protocols (SNU-240103-3) of Seoul National University.\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eConsent for publication\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eNot applicable\\u003c/p\\u003e\\n\\u003cp\\u003eAvailability of data and materials\\u003c/p\\u003e\\n\\u003cp\\u003eAll data generated or analyzed during this study are included in the published article and are available from the corresponding author upon reasonable request.\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eCompeting interests\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eThe authors declare that they have no competing interests.\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eFunding\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eThis work was supported by NRF-2022R1A2C3007621 (Y.Y.K.), NRF-2020R1A5A1018081 (Y.Y.K.), and R01 NS118748 (S.-K.L. and J.W.L.).\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eAuthor\\u0026rsquo;s Contributions\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eConceptualization: S.H.H., Y.E.K., J.H.K., Y.Y.K.; Methodology: Y.E.K. and S.H.H.; Validation: Y.E.K. and S.H.H.; Formal analysis: Y.E.K. and S.H.H.; Investigation: Y.E.K., S.H.H., and Y.W.J.; Writing-original draft preparation: Y.E.K. and S.H.H.; Writing-review and editing: Y.E.K., S.H.H., Y.W.J., K.Y., and Y.Y.K.; Visualization: Y.E.K.; Supervision: Y.Y.K.; Project administration: Y.Y.K.; Funding acquisition: J.W.L. and Y.Y.K.\\u003c/p\\u003e\\n\\u003cp\\u003eAcknowledgments\\u003c/p\\u003e\\n\\u003cp\\u003eWe express our gratitude to the Kong laboratory members for their valuable feedback during the project.\\u0026nbsp;\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eAuthor\\u0026rsquo;s information\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eSchool of Biological Sciences, Seoul National University, Seoul 08826, Republic of Korea\\u003c/p\\u003e\\n\\u003cp\\u003eYea-Eun Kim, Sang-Hyeon Hann, Young-Woo Jo, Kyusang Yoo, Young-Yun Kong\\u003c/p\\u003e\\n\\u003cp\\u003eMolecular Recognition Research Center, Korea Institute of Science and Technology, Seoul 02792,\\u0026nbsp;Republic of Korea\\u003c/p\\u003e\\n\\u003cp\\u003eJi-Hoon Kim\\u003c/p\\u003e\\n\\u003cp\\u003eDepartment of Biological Sciences, University at Buffalo, Buffalo, NY 142604, USA\\u003c/p\\u003e\\n\\u003cp\\u003eJae W. Lee\\u003c/p\\u003e\"},{\"header\":\"References\",\"content\":\"\\u003col\\u003e\\u003cli\\u003e\\u003cspan\\u003eWhite RB, Bi\\u0026eacute;rinx AS, Gnocchi VF, Zammit PS. Dynamics of muscle fibre growth during postnatal mouse development. BMC Dev Biol. 2010;10:21.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eKim JH, Han GC, Seo JY, Park I, Park W, Jeong HW, Lee SH, Bae SH, Seong J, Yum MK, Hann SH, Kwon YG, Seo D, Choi MH, Kong YY. Sex hormones establish a reserve pool of adult muscle stem cells. Nat Cell Biol. 2016;18(9):930\\u0026ndash;40.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eMukund K, Subramaniam S. Skeletal muscle: A review of molecular structure and function, in health and disease. Wiley Interdiscip Rev Syst Biol Med. 2020;12(1):1\\u0026ndash;46.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eFrontera WR, Ochala J. Skeletal Muscle: A Brief Review of Structure and Function. 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Physiology. 2015;30(1):30\\u0026ndash;9.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eTakahashi F, Baba T, Christianto A, Yanai S, Lee-Okada HC, Ishiwata K, Nakabayashi K, Hata K, Ishii T, Hasegawa T, Yokomizo T, Choi MH, Morohashi K. Development of sexual dimorphism of skeletal muscles through the adrenal cortex, caused by androgen-induced global gene suppression. Cell Rep 2024, 43(2).\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eKitamura T, Kitamura YI, Funahashi Y, Shawber CJ, Castrillon DH, Kollipara R, DePinho RA, Kitajewski J, Accili D. A Foxo/Notch pathway controls myogenic differentiation and fiber type specification. J Clin Invest. 2007;117(9):2477\\u0026ndash;85.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eZhou J, So KK, Li Y, Yuan J, Ding Y, Chen F, Huang Y, Liu J, Lee W, Li G, Ju Z, Sun H, Wang H. Elevated H3K27ac in aged skeletal muscle leads to increase in extracellular matrix and fibrogenic conversion of muscle satellite cells. Aging Cell. 2019;18(5):1\\u0026ndash;13.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eLazure F, Blackburn DM, Corchado AH, Sahinyan K, Karam N, Sharanek A, Nguyen D, Lepper C, Najafabadi HS, Perkins TJ, Jahani-Asl A, Soleimani VD. Myf6/MRF4 is a myogenic niche regulator required for the maintenance of the muscle stem cell pool. EMBO Rep 2020:1\\u0026ndash;19.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eRao RC, Dou Y. Hijacked in cancer: The KMT2 (MLL) family of methyltransferases. Nat Rev Cancer. 2015;15(6):334\\u0026ndash;46.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eDhar SS, Zhao D, Lin T, Gu B, Pal K, Wu SJ, Alam H, Lv J, Yun K, Gopalakrishnan V, Flores ER, Northcott PA, Rajaram V, Li W, Shilatifard A, Sillitoe RV, Chen K, Lee MG. MLL4 is required to maintain broad H3K4me3 peaks and super-enhancers at tumor suppressor genes. Mol Cell. 2018;70(5):825\\u0026ndash;e8416.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eAng SY, Uebersohn A, Spencer CI, Huang Y, Lee JE, Ge K, Bruneau BG. KMT2D regulates specific programs in heart development via histone H3 lysine 4 di-methylation. Development. 2016;143(5):810\\u0026ndash;21.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eFujimaki S, Seko D, Kitajima Y, Yoshioka K, Tsuchiya Y, Masuda S, Ono Y. Notch1 and Notch2 coordinately regulate stem cell function in the quiescent and activated states of muscle satellite cells. Stem Cells. 2018;36(2):278\\u0026ndash;85.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eHuisman C, Kim YA, Jeon S, Shin B, Choi J, Lim SJ, Youn SM, Park Y, Medha KC, Kim S, Lee SK, Lee S, Lee JW. The histone H3-lysine 4-methyltransferase Mll4 regulates the development of growth hormone-releasing hormone-producing neurons in the mouse hypothalamus. Nat Commun. 2021;12(1):1\\u0026ndash;14.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eEsteves de Lima J, Bonnin MA, Birchmeier C, Duprez D. Muscle contraction is required to maintain the pool of muscle progenitors via YAP and NOTCH during fetal myogenesis. ELife. 2016;5(AUGUST):1\\u0026ndash;25.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eBi P, Yue F, Sato Y, Wirbisky S, Liu W, Shan T, Wen Y, Zhou D, Freeman J, Kuang S. Stage-specific effects of Notch activation during skeletal myogenesis. ELife. 2016;5:1\\u0026ndash;22.\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eGallot Y, Hindi S, Mann A, Kumar A. Isolation, culture, and staining of single myofibers. Bio-protocol 2016, 6(19).\\u003c/span\\u003e\\u003c/li\\u003e \\u003cli\\u003e\\u003cspan\\u003eLiu L, Cheung TH, Charville GW, Rando TA. Isolation of skeletal muscle stem cells by fluorescence-activated cell sorting. Nat Protoc. 2015;10(10):1612\\u0026ndash;24.\\u003c/span\\u003e\\u003c/li\\u003e\\u003c/ol\\u003e\"}],\"fulltextSource\":\"\",\"fullText\":\"\",\"funders\":[],\"hasAdminPriorityOnWorkflow\":false,\"hasManuscriptDocX\":true,\"hasOptedInToPreprint\":true,\"hasPassedJournalQc\":\"\",\"hasAnyPriority\":false,\"hideJournal\":false,\"highlight\":\"\",\"institution\":\"\",\"isAcceptedByJournal\":true,\"isAuthorSuppliedPdf\":false,\"isDeskRejected\":\"\",\"isHiddenFromSearch\":false,\"isInQc\":false,\"isInWorkflow\":false,\"isPdf\":false,\"isPdfUpToDate\":true,\"isWithdrawnOrRetracted\":false,\"journal\":{\"display\":true,\"email\":\"info@researchsquare.com\",\"identity\":\"skeletal-muscle\",\"isNatureJournal\":false,\"hasQc\":true,\"allowDirectSubmit\":false,\"externalIdentity\":\"skem\",\"sideBox\":\"Learn more about [Skeletal Muscle](http://skeletalmusclejournal.biomedcentral.com/)\",\"snPcode\":\"13395\",\"submissionUrl\":\"https://submission.nature.com/new-submission/13395/3\",\"title\":\"Skeletal Muscle\",\"twitterHandle\":\"@BioMedCentral\",\"acdcEnabled\":true,\"dfaEnabled\":true,\"editorialSystem\":\"em\",\"reportingPortfolio\":\"BMC/SO AJ\",\"inReviewEnabled\":true,\"inReviewRevisionsEnabled\":true},\"keywords\":\"Skeletal muscle, Myofiber, Muscle stem cells, Myeloid/lymphoid or mixed-lineage leukemia 4 (Mll4), Exercise, Notch signaling\",\"lastPublishedDoi\":\"10.21203/rs.3.rs-5413133/v1\",\"lastPublishedDoiUrl\":\"https://doi.org/10.21203/rs.3.rs-5413133/v1\",\"license\":{\"name\":\"CC BY 4.0\",\"url\":\"https://creativecommons.org/licenses/by/4.0/\"},\"manuscriptAbstract\":\"\\u003ch2\\u003eBackground\\u003c/h2\\u003e \\u003cp\\u003eMuscle stem cells (MuSCs) undergo numerous state transitions throughout life, which are critical for supporting normal muscle growth and regeneration. Therefore, it is crucial to investigate the regulatory mechanisms governing the transition of MuSC states across different postnatal developmental stages.\\u003c/p\\u003e\\u003ch2\\u003eMethods\\u003c/h2\\u003e \\u003cp\\u003eTo assess if myofiber-expressed Mll4 contributes to the maintenance of MuSCs, we crossed \\u003cem\\u003eMCK\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eCre/+\\u003c/em\\u003e\\u003c/sup\\u003e or \\u003cem\\u003eHSA\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eMerCreMer/+\\u003c/em\\u003e\\u003c/sup\\u003e mice to \\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e mice to generate myofiber-specific \\u003cem\\u003eMll4\\u003c/em\\u003e-deleted mice. Investigations were conducted using 8-week-old and 4-week-old \\u003cem\\u003eMCK\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eCre/+\\u003c/em\\u003e\\u003c/sup\\u003e;\\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e mice Investigations were conducted using 8-week-old and 4-week-old \\u003cem\\u003eHSA\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003eCre/+\\u003c/em\\u003e\\u003c/sup\\u003e;\\u003cem\\u003eMll4\\u003c/em\\u003e\\u003csup\\u003e\\u003cem\\u003ef/f\\u003c/em\\u003e\\u003c/sup\\u003e mice were utilized.\\u003c/p\\u003e\\u003ch2\\u003eResults\\u003c/h2\\u003e \\u003cp\\u003eDuring postnatal myogenesis, \\u003cem\\u003eMll4\\u003c/em\\u003e deleted muscles were observed with increased number of cycling MuSCs that proceeded to a differentiation state, leading to MuSC deprivation. This phenomenon occurred independently of gender. When \\u003cem\\u003eMll4\\u003c/em\\u003e was ablated in adult muscles using the inducible method, adult MuSCs lost their quiescence and differentiated into myoblasts, also causing the depletion of MuSCs. Such roles of \\u003cem\\u003eMll4\\u003c/em\\u003e in myofibers coincided with decreased expression levels of distinct Notch ligands: \\u003cem\\u003eJag1\\u003c/em\\u003e and \\u003cem\\u003eDll1\\u003c/em\\u003e in pubertal and \\u003cem\\u003eJag2\\u003c/em\\u003e and \\u003cem\\u003eDll4\\u003c/em\\u003e in adult muscles.\\u003c/p\\u003e\\u003ch2\\u003eConclusions\\u003c/h2\\u003e \\u003cp\\u003eOur study suggests that \\u003cem\\u003eMll4\\u003c/em\\u003e is crucial for maintaining MuSCs in both pubertal and adult muscles, which may be accomplished through the modulation of distinct Notch ligand expressions in myofibers. These findings offer new insights into the role of myofiber-expressed Mll4 as a master regulator of MuSCs, highlighting its significance not only in developmental myogenesis but also in adult muscle, irrespective of sex.\\u003c/p\\u003e\",\"manuscriptTitle\":\"Mll4 in Skeletal Muscle Fiber Maintains Muscle Stem Cells by Regulating Notch Ligands\",\"msid\":\"\",\"msnumber\":\"\",\"nonDraftVersions\":[{\"code\":1,\"date\":\"2024-11-26 06:00:46\",\"doi\":\"10.21203/rs.3.rs-5413133/v1\",\"editorialEvents\":[{\"type\":\"communityComments\",\"content\":0},{\"type\":\"decision\",\"content\":\"Revision requested\",\"date\":\"2024-11-27T12:40:10+00:00\",\"index\":\"\",\"fulltext\":\"\"},{\"type\":\"editorInvitedReview\",\"content\":\"\",\"date\":\"2024-11-26T11:40:39+00:00\",\"index\":\"hide\",\"fulltext\":\"\"},{\"type\":\"reviewerAgreed\",\"content\":\"250432188838828761072668610511347781393\",\"date\":\"2024-11-24T10:49:14+00:00\",\"index\":\"hide\",\"fulltext\":\"\"},{\"type\":\"editorInvitedReview\",\"content\":\"\",\"date\":\"2024-11-14T02:17:45+00:00\",\"index\":\"hide\",\"fulltext\":\"\"},{\"type\":\"reviewerAgreed\",\"content\":\"266281847310568553599747290677973259401\",\"date\":\"2024-11-11T12:52:11+00:00\",\"index\":\"hide\",\"fulltext\":\"\"},{\"type\":\"reviewersInvited\",\"content\":\"\",\"date\":\"2024-11-11T12:35:30+00:00\",\"index\":\"\",\"fulltext\":\"\"},{\"type\":\"editorAssigned\",\"content\":\"\",\"date\":\"2024-11-11T12:29:00+00:00\",\"index\":\"\",\"fulltext\":\"\"},{\"type\":\"checksComplete\",\"content\":\"\",\"date\":\"2024-11-08T04:20:09+00:00\",\"index\":\"\",\"fulltext\":\"\"},{\"type\":\"submitted\",\"content\":\"Skeletal Muscle\",\"date\":\"2024-11-08T02:39:38+00:00\",\"index\":\"\",\"fulltext\":\"\"}],\"status\":\"published\",\"journal\":{\"display\":true,\"email\":\"info@researchsquare.com\",\"identity\":\"skeletal-muscle\",\"isNatureJournal\":false,\"hasQc\":true,\"allowDirectSubmit\":false,\"externalIdentity\":\"skem\",\"sideBox\":\"Learn more about [Skeletal Muscle](http://skeletalmusclejournal.biomedcentral.com/)\",\"snPcode\":\"13395\",\"submissionUrl\":\"https://submission.nature.com/new-submission/13395/3\",\"title\":\"Skeletal Muscle\",\"twitterHandle\":\"@BioMedCentral\",\"acdcEnabled\":true,\"dfaEnabled\":true,\"editorialSystem\":\"em\",\"reportingPortfolio\":\"BMC/SO AJ\",\"inReviewEnabled\":true,\"inReviewRevisionsEnabled\":true}}],\"origin\":\"\",\"ownerIdentity\":\"9c15434c-e788-44d4-8f77-ed00552405e2\",\"owner\":[],\"postedDate\":\"November 26th, 2024\",\"published\":true,\"recentEditorialEvents\":[],\"rejectedJournal\":[],\"revision\":\"\",\"amendment\":\"\",\"status\":\"published-in-journal\",\"subjectAreas\":[],\"tags\":[],\"updatedAt\":\"2024-12-30T16:05:51+00:00\",\"versionOfRecord\":{\"articleIdentity\":\"rs-5413133\",\"link\":\"https://doi.org/10.1186/s13395-024-00369-9\",\"journal\":{\"identity\":\"skeletal-muscle\",\"isVorOnly\":false,\"title\":\"Skeletal Muscle\"},\"publishedOn\":\"2024-12-23 15:57:48\",\"publishedOnDateReadable\":\"December 23rd, 2024\"},\"versionCreatedAt\":\"2024-11-26 06:00:46\",\"video\":\"\",\"vorDoi\":\"10.1186/s13395-024-00369-9\",\"vorDoiUrl\":\"https://doi.org/10.1186/s13395-024-00369-9\",\"workflowStages\":[]},\"version\":\"v1\",\"identity\":\"rs-5413133\",\"journalConfig\":\"researchsquare\"},\"__N_SSP\":true},\"page\":\"/article/[identity]/[[...version]]\",\"query\":{\"redirect\":\"/article/rs-5413133\",\"identity\":\"rs-5413133\",\"version\":[\"v1\"]},\"buildId\":\"qtupq5eGEP_6zYnWcrvyt\",\"isFallback\":false,\"isExperimentalCompile\":false,\"dynamicIds\":[84888],\"gssp\":true,\"scriptLoader\":[]}","source_license":"CC-BY-4.0","license_restricted":false}