Ablation of satellite cell‐specific clock gene, Bmal1, alters force production, muscle damage, and repair following contractile‐induced injury

Abstract

Following injury, skeletal muscle undergoes repair via satellite cell (SC)‐mediated myogenic progression. In SCs, the circadian molecular clock gene, Bmal1, is necessary for appropriate myogenic progression and repair with evidence that muscle molecular clocks can also affect force production. Utilizing a mouse model allowing for inducible depletion of Bmal1 within SCs, we determined contractile function, SC myogenic progression and muscle damage and repair following eccentric contractile‐induced injury. At baseline, SC‐Bmal1 ^iKO animals exhibited a ~20–25% reduction in normalized force production (ex vivo and in vivo) versus control SC‐Bmal1 ^Cntrl and SC‐Bmal1 ^iKO untreated littermates (p < .05). Following contractile injury, SC‐Bmal1 ^iKO animals displayed reduced muscle damage and subsequent repair post‐injury (Dystrophin^negative fibers 24 h: SC‐Bmal1 ^Cntrl 199 ± 41; SC‐Bmal1 ^iKO 36 ± 13, p < .05) (eMHC⁺ fibers 7 day: SC‐Bmal1 ^Cntrl 217.8 ± 115.5; SC‐Bmal1 ^iKO 27.8 ± 17.3; Centralized nuclei 7 day: SC‐Bmal1 ^Cntrl 160.7 ± 70.5; SC‐Bmal1 ^iKO 46.2 ± 15.7). SC‐Bmal1 ^iKO animals also showed reduced neutrophil infiltration, consistent with less injury (Neutrophil content 24 h: SC‐Bmal1 ^Cntrl 2.4 ± 0.4; SC‐Bmal1 ^iKO 0.4 ± 0.2, % area fraction, p < .05). SC‐Bmal1 ^iKO animals had greater SC activation/proliferation at an earlier timepoint (p < .05) and an unexplained increase in activation 7 days post injury. Collectively, these data suggest SC‐Bmal1 plays a regulatory role in force production, influencing the magnitude of muscle damage/repair, with an altered SC myogenic progression following contractile‐induced muscle injury.

Ablation of SC‐Bmal1 lowered force production and altered the magnitude of muscle damage/repair and SC myogenic progression following eccentric contractile injury.

Abbreviations

½ RT

half relaxation time

ANOVA

analysis of variance

Bmal1

brain and muscle like arnt‐1

BSA

bovine serum albumin

BW

body weight

Ca⁺⁺

calcium

CE

core‐enhancer

Clock

circadian locomotor output cycle kaput

Cry1

cryptochrome circadian regulator 1

Cry2

cryptochrome circadian regulator 2

CSA

cross‐sectional area

DI

distilled

EDL

extensor digitorum longus

eMHC

embryonic myosin heavy chain

ES

electrically stimulated

g

grams

h

hour

H&E

hematoxylin and eosin

Hz

hertz

IgG

immunoglobular domain

IHC

immunohistochemistry

J

joules

Kg

kilograms

KO

knockout

L

liter

L_f

fiber length

L_f/s

fiber lengths per second

L_o

muscle length

M

mass

min

minutes

mL

milliliter

mM

millimolar

MyoD

myoblast determination protein 1

N

newtons

N/cm²

newton/centimeters squared

Nm

newton meters

Pax7

paired box 7

PBS

phosphate‐buffered saline

PCSA

physiological cross‐sectional area

Per1

period 1

Per2

period 2

PFA

paraformaldehyde

P_o

maximal isometric force

s

seconds

SC

satellite cell

SEM

standard error of mean

TA

tibialis anterior

TPT

time‐to‐peak tension

uL

microliter

V

volts

ZT

zeitgeber hour

ρ

fiber density

1. INTRODUCTION

Molecular clocks reside in all cells and regulate numerous physiological processes in a circadian fashion. ¹ , ² Specifically, a transcriptional/translational feedback loop consisting of rhythmic expression of Per/Cry and Clock/ Bmal1 collectively underpin circadian rhythmicity. ¹ , ² Recent evidence demonstrates molecular clocks play a regulatory role in skeletal muscle repair following injury. ³ , ⁴ , ⁵ , ⁶ , ⁷ In skeletal muscles, satellite cells (SCs) are the resident muscle stem cell population that lie quiescent sandwiched between the basal lamina and muscle membrane. Following injury, SCs proceed down a myogenic lineage via activation, proliferation, differentiation, eventually fusing to the host myofiber to facilitate muscle repair. ⁸ , ⁹ SCs express molecular clock genes as well as several contractile and myogenesis‐related genes that oscillate over a 24 h cycle. ¹⁰

Molecular clocks play a critical role in muscle repair with myogenic progression and the magnitude of repair significantly impaired in Per1/2, Cry1/2, Bmal1 muscle‐specific and whole‐body knockout (KO) animals following injury. ¹¹ , ¹² , ¹³ However, these studies have used either muscle or whole‐body clock‐gene ablation/KO methods making it difficult to establish a precise role for SC‐molecular clocks during muscle repair. A previous study from our group examined the time‐of‐day capacity of SC‐mediated repair following cardiotoxin‐induced injury in wildtype mice and reported blunted repair in the morning versus the evening. ¹⁴ Such observations are possibly underpinned by divergent time‐of‐day capabilities of the SC‐molecular clock. To directly assess the role of SC‐molecular clocks in the repair process, the same study then utilized an inducible depletion model capable of SC‐specific Bmal1 ablation to explore injury/repair at a single timepoint. SCs lacking Bmal1 did not proliferate sufficiently following cardiotoxin injury, leading to blunted muscle repair. ¹⁴ SC activation and proliferation hinges on the myogenic regulatory factor, MyoD, ¹⁵ a clock‐controlled gene. ¹⁶ , ¹⁷ , ¹⁸ Bmal1 binds to the core‐enhancer (CE) region of MyoD and ablation of the CE disrupts its diurnal expression and amplitude/timing during myogenic progression, ultimately leading to blunted myogenesis, ¹⁹ , ²⁰ at least in embryonic muscle. In this regard, muscle‐specific Bmal1 KO animals have recently displayed altered diurnal expression amplitudes of MyoD over the course of 24 h. ²¹ While whole‐body Bmal1 KO models alter MyoD's response during SC proliferation ¹² , ²² evidence that SC‐Bmal1 affects MyoD is lacking.

Prior investigations have reported that Bmal1 regulates contractile‐force production in animal models. ¹⁷ , ²³ , ²⁴ , ²⁵ As higher forces typically lead to greater magnitudes of muscle damage, ²⁶ , ²⁷ Bmal1's regulation on force production may lead to varying magnitudes of damage and subsequent repair. However, it is unknown if SC‐specific, Bmal1, affects force production and if there are any subsequent effects on contractile‐induced injury and repair. Recent work from our lab has shown that SC ablation alters time‐of‐day muscle contractility, with the overall extent of contractile injury being associated with the magnitude of eccentric force. ²⁸ These results suggest that SC‐molecular clocks may regulate force production, which may subsequently determine the extent of contractile injury. Accordingly in the present study, we hypothesized that SC‐Bmal1 would alter force production characteristics and the extent of contractile injury, muscle damage, and subsequent muscle repair. Given the role of Bmal1 in myogenic progression, we also postulated that in response to physiological contractile injury, SC‐Bmal1 ablation would dysregulate myogenic progression. Utilizing a mouse model capable of SC‐specific inducible depletion of molecular clock gene, Bmal1, we assessed baseline contractile function, muscle damage, SC myogenic progression, and muscle repair following in vivo eccentric contractile injury.

2. METHODS

2.1. Animals

All animal experiments were undertaken after approval by the Northwestern University Institutional Animal Care and Use Committee. A previously established mouse model was utilized in this study for inducible depletion of SC‐specific clock gene, Bmal1. ¹⁴ To generate these mice, Pax7CreER mice were crossed with a flox‐stop‐flox‐tdTomato mouse line. Second‐generation Pax7CreER/tdTomato mice were then crossed with either SC‐Bmal1 ^fx/fx or SC‐Bmal1 ^+/+ mouse lines to generate Pax7CreER/tdTomato/Bmal1 ^fx/fx (SC‐Bmal1 ^iKO) capable of inducible depletion or Pax7CreER/tdTomato/Bmal1 ^+/+ control mice (SC‐Bmal1 ^Cntrl). Additionally, since the tdTomato is only expressed in the presence of Pax7CreER expression, this induces fluorescent labeling of SCs in this mouse model after treatment by tamoxifen. ¹⁴ Genotyping (TransnetYX, Cordova, TN) was performed on all mice to ensure appropriate gene construct. In experimental mice (SC‐Bmal1 ^iKO), Bmal1 within SCs is floxed to allow for inducible depletion utilizing the Cre‐Lox system, resulting in a >80% loss of Bmal1 in SCs after treatment by tamoxifen. ¹⁴ Both groups were treated with tamoxifen (2 mg/100 μL per day) via oral gavage for 5 consecutive days (5 × 100 μL gavage) with a ten‐day washout period. Both male and female animals between 12 and 16 months of age were used for all experiments (n = 38, male = 19, female = 19) and tamoxifen treatment was only administered when mice were 12–16 months of age. This leads to Bmal1 ablation and tdTomato expression in SCs of SC‐Bmal1 ^iKO animals while only tdTomato expression in SCs of SC‐Bmal1 ^Cntrl mice. Animals were housed in a 14:10 light–dark cycle and consumed food ad libitum. Previous work from our group shows rhythmic oscillation of molecular clock gene expression under a 14:10 light–dark photoperiod. ²⁸ To assess if SC‐Bmal1 played a role in muscle repair following physiological (contractile) injury, all in vivo/ex vivo experiments were performed at a single timepoint between 1000 and 1200 h (ZT4‐ZT6). This study design was in line with past Bmal1 ablation models that utilized a single timepoint to study the effects of Bmal1 absence on either physiology or injury/repair. ¹⁴ , ¹⁷ , ²³

2.2. Baseline in vivo contractile characteristics

In vivo maximal tetanic and eccentric torque of the dorsiflexors were assessed in SC‐Bmal1 ^Cntrl, SC‐Bmal1 ^iKO, and SC‐Bmal1 ^{iKO Cntrl} (untreated) animals for baseline contractile function measures. In vivo muscle contractility of the dorsiflexors was undertaken according to previous methods (Aurora Scientific, 3‐in‐1 contractile apparatus, 1300A Whole Animal Muscle Test System, Ontario, CA). ²⁹ In brief, after general anesthesia, the left hindlimb was shaved, ankle fixed at 90⁰ and inserted onto a foot‐plate attached to a torque motor, capable of bi‐direction torque measurement. Two needle‐electrodes were placed percutaneously in the region of the peroneal nerve to selectively stimulate the dorsiflexors and twitch contractions (3–6 mA, 100 Hz) were administered thereafter to optimize voltage and electrode placement. ³⁰ Dorsiflexor and plantar flexor torque were simultaneously evaluated throughout all contractions to ensure plantar flexors were not co‐contracting. 500 ms maximal tetanic contractions separated by 3 min rest were administered to determine maximal isometric torque. One eccentric contraction was administered to determine maximal eccentric torque by a 300 ms maximal tetanus followed by a lengthening ramp of 50 ms that rotated the foot into 38° of plantarflexion approximately at 200°/s (starting foot position, −19°; ending foot position, +19°). ²⁹ Optimal stimulation frequency was determined during pilot torque‐frequency curves and all subsequent experimental tetanic and eccentric contractions were administered at 120 Hz. All contractile torque data was normalized to mouse body mass (BW) and expressed as Nm/kg BW. ³⁰ , ³¹ , ³²

2.3. In vivo eccentric contractile injury

In vivo eccentric contractile injury in the dorsiflexors was induced via 200 electrically stimulated eccentric contractions, with each contraction separated by 10 s rest. ³³ Maximal tetanic torque was assessed 3 min before and after the injury bout as an indirect measure of mechanical injury (torque‐deficit in maximal tetanic torque). ²⁶ , ²⁷ , ³³ , ³⁴ , ³⁵ , ³⁶ , ³⁷ , ³⁸ Total work was calculated by the sum of work per contraction ((torque × angular displacement)/BW) across all 200 eccentric contractions per mouse. Following the injury protocol, mice were returned to their housing, allowed to recover and euthanized 24, 72 h, or 7 d post‐injury (n = 3‐5/group). TA muscles were harvested to evaluate the injury, cellular responses, and repair processes at these the timepoints.

2.4. Ex vivo baseline contractile characteristics

Ex vivo tetanic and eccentric specific forces were assessed using the EDL of SC‐Bmal1 ^Cntrl and SC‐Bmal1 ^iKO animals. ²⁸ In brief, the left hindlimb was stabilized with pins, 5–0 silk sutures were tied to the proximal and distal tendons of the EDL and transferred to a bath containing Ringer's solution at 37°C (137 mM NaCl, 5 mM KCl, 1 mM NaH₂PO₄, 24 mM NaHCO₃, 2 mM CaCl₂, 1 mM MgSO₄, and 11 mM glucose containing 10 mg/L curare, pH 7.5). The proximal end of the muscle was sutured to a force‐transducer (Aurora 300C, Aurora Scientific, Ontario, Canada) and the distal tendon sutured to a length‐motor with platinum electrodes straddling the muscle end‐to‐end. ³⁹ A custom LabVIEW program recorded force, length, and time data from each contraction. Twitch contractions were used to optimize muscle length/voltage and fiber length was subsequently calculated using a standard fiber‐length‐to‐muscle‐length ratio of 0.51 for the EDL. ⁴⁰ Three 300 ms maximal tetanic contractions were used to determine maximal tetanic force (the highest of the three used as the tetanic force value). A single eccentric contraction was administered to determine maximal eccentric force and consisted of a 200 ms tetanus followed by a 100 ms lengthening ramp. The eccentric portion of the contraction consisted of muscle lengthening by 15% of L_f at a strain‐rate of 2 L_f/s. All contractile data were measured in volts (V), converted to Newtons (N) based on a calibration curve and normalized to calculated physiological cross‐sectional area (PCSA), and expressed as specific force (N/cm²).

Time‐to‐peak tension (TPT) was determined from the maximal tetanic contractions to evaluate differences in contractile kinetics. TPT was calculated as the time from baseline to the first data point of the maximal tetanic force plateau on the ascending limb of the force trace for each muscle. Half relaxation time (HRT) was calculated as the time from the last data point on the tetanic force plateau to the time that reflected half maximum tetanic force on the descending limb of the force trace.

2.5. Histology and Immunohistochemistry

TA muscle was flash frozen in liquid nitrogen‐cooled isopentane immediately after euthanasia and stored at −80°C. Muscles were transferred to a −25°C cryostat and allowed to equilibrate for 1 h prior to sectioning. In brief, the muscle mid‐belly was cut perpendicular to the orientation of the fibers, embedded in a cryomold and flash frozen (10 s) in liquid nitrogen‐cooled isopentane. ⁴¹ Molds were then allowed to equilibrate in the cryostat for 30 min before sectioning. Sections were cut at 10 μm thickness. Slides were air‐dried for 1 h following sectioning and stored at −80°C until experimental procedures.

Prior to all staining, slides were thawed at room temperature for 1 h. Hematoxylin & eosin (H&E) histological staining consisted of sequentially dipping slides into baths of the following: 1% glutaraldehyde, PBS, hematoxylin, tap water, alcoholic acid, tap water, ammonia water, DI water, alcoholic eosin, 70% ETOH, 80% ETOH, 95% ETOH, 100% ETOH (2x), Xylene (2x). Following staining, slides were coverslipped and stored at −20°C until imaging.

For immunohistochemistry (IHC), slides were thawed for 1 h prior to staining and a hydrophobic barrier was drawn around sections and allowed to dry for 20 min. Sections were washed in PBS, fixed in 4% paraformaldehyde, ⁴² rehydrated with PBS washes and blocked in 1% BSA mouse‐on‐mouse blocking buffer for 1 h. Following blocking, primary antibody cocktails were made in 1%BSA and incubated overnight at −4°C. The next day, sections were washed with PBS and incubated in a secondary antibody cocktail made in PBS. Sections were then washed with PBS, mounted using Vectashield with (Vectashield, H‐1200, Vector Laboratories) or without DAPI (Vectashield, H‐1000, Vector Laboratories), as appropriate, coverslipped, and stored at −20°C until imaging. ⁴¹ , ⁴³ Primary and secondary antibody details were as follows: Primary antibodies: anti‐laminin (rabbit, IgG, 1:500, L9393, Sigma, St. Louis, MO, USA), anti‐dystrophin (rabbit, IgG, 1:100, ab15277, Abcam, Cambridge, UK), anti‐MYH3 (eMHC) (mouse, IgG1, 1:50, F1‐652, DSHB, Iowa City, IA, USA), anti‐Ly‐6G/C (rat, 1:30, lot # 553123, BD Biosciences, Franklin Lakes, NJ, USA), anti‐Ki67 (rabbit, IgG, 1:200, ab15580, Abcam, Cambridge, UK), anti‐MyoD (mouse, IgG2b, 1:50, sc‐377 460, Santa Cruz Biotechnologies, Dallas, TX, USA), anti‐SC71 (mouse, IgG1, 1:50, lot # 2147165, DSHB, Iowa City, IA, USA), anti‐BF‐F3 (mouse, IgM, 1:100, lot # 2266724, DSHB, Iowa City, IA, USA). Secondary antibodies: Alexa Flour 488 goat anti‐rabbit IgG (H + L) (1:250, A‐11034, Invitrogen, Waltham, MA, USA), Alexa Flour 488 goat anti‐mouse IgG1 (1:250, A21121, Invitrogen, Waltham, MA, USA), Alexa Flour 488 goat anti‐mouse IgG2b (1:250, 115–545‐207, Jackson ImmunoResearch Laboratories, West Grove, PA, USA), Alex Flour 594 goat anti‐rat IgG (H + L) (1:250, A‐11007, Invitrogen, Waltham, MA, USA), goat anti‐mouse IgM (1:250, A‐21426, Invitrogen, Waltham, MA, USA).

2.6. Image acquisition and quantification

Images were taken at 10 x and 20 x magnification depending on the IHC protocol and whole tile‐scan images (entire cross‐section) were acquired. For quantification of myofiber necrosis H&E images were used. “Necrosing fibers” were defined as fibers that had profuse nuclei covering the entire fiber. Based on Dystrophin IHC, fibers that were missing myofiber borders (dystrophin) but had nuclei surrounding the remaining outline of the border (and inside the fiber in many cases) were classified as Dystrophin^negative fibers. In a subset of images, locations of these fibers were validated by finding the specific Dystrophin^negative fiber using fiber‐borders (i.e., laminin‐stained sections from a different slide of the same muscle). Ly6+ content was calculated as area fraction of the total cross‐sectional area using ImageJ. The number of eMHC+ positive fibers and fibers with centralized nuclei were manually quantified using ImageJ. As noted, tdTomato is expressed in the presence of Pax7Cre in this mouse model ¹⁴ and therefore all identification of SCs were accomplished via use of the tdTomato/Pax7/DAPI construct. Total tdTomato+ cell (SC) abundance in uninjured sections, tdTomato+/Ki67+/DAPI, and tdTomato+/MyoD+/DAPI cells were manually quantified in ImageJ and expressed as cells/100 fibers. Fiber‐type quantification was performed using an automated muscle‐analysis software MuscleJ. ⁴⁴

2.7. Satellite cell isolation and qPCR

As previously described, ¹⁴ hindlimb muscles (TA and gastrocnemius) were dissected and cleaned by carefully removing non‐muscle tissues before being digested in Ham's F‐10 medium (Corning 10‐070‐CV) containing 5 mg/mL collagenase D (Roche 11 088 866 001) and 5 mg/mL dispase II (Gibco 17 105 041). Digestion was performed at 37°C with shaking at 500 rpm for 30 min. The enzymatic process was halted by adding ice‐cold cell suspension buffer (Ham's F‐10 with 10% FBS and 3 mM EDTA) in a fivefold excess. The muscle tissue was then mechanically dissociated by repeated pipetting (20 times) with a 10‐mL pipette to release SCs from muscle fibers. The cell suspension was sequentially filtered through 70‐μm and 40‐μm nylon mesh strainers (BD Biosciences), with the filtrate collected in a 15‐mL conical tube and centrifuged at 500 × g for 5 min. The resulting cell pellet was resuspended in 1 mL of Pre‐Sort Buffer (BD Biosciences 563 503), and Live‐or‐Dye 640/662‐negative, tdTomato‐positive mononuclear SCs were isolated via sorting on a BD FACSAria II cell sorter.

For qPCR analysis, total RNA was extracted from SCs using TRIzol™ Reagent, following the manufacturer's protocol. The quality and concentration of the RNA were verified with a NanoDrop™ 2000 Spectrophotometer. Only RNA samples with an OD260/280 ratio above 1.80 proceeded to first‐strand cDNA synthesis, which was performed using the High‐Capacity cDNA Reverse Transcription Kit (Applied Biosystems™) as per the kit's protocol. Real‐Time quantitative PCR was subsequently conducted using iTaq Universal SYBR Green Supermix (Bio‐Rad) on a CFX Opus 384 Real‐Time PCR System. Primer sequences used for qPCR were as follows: Bmal1‐F (AGGCCCACAGTCAGATTGAA), Bmal1‐R (TGGTACCAAAGAAGCCAATTCAT), b‐Actin‐F (GGCTGTATTCCCCTCCATCG), b‐Actin‐R (CAGTTGGTAACAATGCCATG).

2.8. Statistics

Individual groups were compared using unpaired t‐tests for all contractile data. All IHC group data were analyzed using two‐way ANOVAs with main effects of time and treatment, with post‐hoc Sidak's multiple comparison tests. Simple linear regressions were also used to analyze relationships between selected variables. Statistical analyses were performed using Prism 9.0 (GraphPad, San Diego, CA) and reported throughout as mean ± standard error of mean (SEM). Statistical significance was set as p < .05.

3. RESULTS

3.1. Baseline in vivo and ex vivo contractile characteristics are lower after Bmal1 ablation in satellite cells

Tamoxifen treatment induced a ~90% depletion of SC‐specific Bmal1 in SC‐Bmal1 ^iKO mice compared to SC‐Bmal1 ^Cntrl counterparts (Figure 1D). Baseline in vivo eccentric dorsiflexor torque was significantly lower (~20%) in SC‐Bmal1 ^iKO compared to both SC‐Bmal1 ^Cntrl and littermate‐untreated control SC‐Bmal1 ^{iKO Cntrl} animals (SC‐Bmal1 ^iKO 84 ± 5 Nm/kg BW; SC‐Bmal1 ^Cntrl 105 ± 8 Nm/kg BW; SC‐Bmal1 ^{iKO Cntrl} 110 ± 4 Nm/kg BW; p < .05) (Figure 2A). No differences were observed in tetanic torque. BW and TA weights were significantly greater in SC‐Bmal1 ^iKO animals (BW: SC‐Bmal1 ^iKO 27 ± 1 g; SC‐Bmal1 ^Cntrl 25 ± 1 g; p < .05) (Figure S2A) (TA weight: SC‐Bmal1 ^iKO 41 ± 2 mg; SC‐Bmal1 ^Cntrl 35 ± 2 mg; p < .01) (Figure S2B). No differences in myofiber area were observed between groups (Figure S2C).

FIGURE 1.

Satellite cell‐specific ablation of Bmal1. (A) Mouse model construct for SC‐Bmal1 ^iKO mice. (B) Representative image displaying genotype of SC‐Bmal1 ^Cntrl and SC‐Bmal1 ^iKO (SC‐ Bmal1 ^fx/fx). (C) Plots showing FACS sorting gating strategy. Upper panel shows total cells, single cells, and live cells. Bottom panel shows tdTomato negative control and tdTomato‐positive SCs. (D) qPCR expression of Bmal1 within SCs of SC‐Bmal1 ^iKO and SC‐Bmal1 ^Cntrl mice (n = 3–4). All data shown as mean ± SEM. Groups were compared using unpaired t‐tests (**p < .01).

FIGURE 2.

Force production is reduced in SC‐Bmal1 ^iKO animals. (A) In vivo tetanic and eccentric torque of the dorsiflexors of all groups (Nm/kg BW) (SC‐ Bmal1 ^Cntrl and SC‐Bmal1 ^iKO n = 14–15, SC‐Bmal1 ^{iKO Cntrl} n = 4–5). (B) Ex vivo tetanic and eccentric specific forces of the EDL for Cntrl and iKO animals (N/cm²) (n = 4–5). (C) Representative image of fiber‐type distribution in the TA (blue: Dystrophin; green: IIa; Magenta: IIb; Green/Magenta: IIa‐IIb; Unlabeled: IIx) (scale bar set to 100 μm). (D) Fiber‐type distribution of TA muscle from Cntrl and iKO animals (n = 3). All data shown as mean ± SEM. Contractile data groups were compared using unpaired t‐tests. Fiber‐type data groups compared using two‐way ANOVA (*p < .05).

Ex vivo tetanic, eccentric specific forces were also reduced ~20%, ~26% respectively in EDL of SC‐Bmal1 ^iKO versus control animals (tetanic specific force: SC‐Bmal1 ^Cntrl 21 ± 1 N/cm²; SC‐Bmal1 ^iKO 16 ± 1 N/cm²; eccentric specific force: SC‐Bmal1 ^Cntrl 31 ± 2 N/cm²; SC‐Bmal1 ^iKO 23 ± 1 N/cm²; p < .05) (Figure 2B). There were no differences in ex vivo tetanic TPT, HRT, fiber‐type proportions (Figure 2C,D) or myofiber area (TA).

3.2. In vivo contractile injury

Figure 3A shows a representative trace of eccentric dorsiflexor torque over 200 eccentric contractions. Immediately following contractile injury, torque was decreased by a similar magnitude from maximal tetanic torque values between groups (SC‐Bmal1 ^Cntrl 21 ± 2 (47% reduction); SC‐Bmal1 ^iKO 19 ± 2.6 (49% reduction), units expressed as Nm/kg BW) (Figure 3B). Total work performed during the contractions were ~20% lower in SC‐Bmal1 ^iKO compared to control animals (SC‐Bmal1 ^Cntrl 12.5 ± 1.2 J/g; SC‐Bmal1 ^iKO 9.9 ± 0.6 J/g; p = .06, Figure 3C).

FIGURE 3.

In vivo eccentric contractile injury. (A) A representative trace from a single animal of all 200 eccentric contractions (torque in units of Nm/ kg BW). (B) Pre‐injury torque versus post‐injury torque values following 200 eccentric contractions in both groups (n = 14–15). (C) Total work performed in both groups throughout 200 eccentric contractions (n = 14–15). All data shown as mean ± SEM. Groups were compared using paired t‐tests for torque‐deficit data and unpaired t‐tests for total work data (*p < .05).

3.3. SC‐Bmal1^iKO animals exhibit less fiber necrosis and dystrophin^negative fibers following contractile muscle injury

Figure 4A,B,D,E are representative images of H&E and dystrophin‐stained cross‐sections showing uninjured and 24 h post‐injury necrosing (H&E) and dystrophin^negative fibers, respectively (necrosing fibers and dystrophin^negative fibers indicated with arrowheads). There was a main effect for post‐injury time and an interaction effect in necrosing as well as dystrophin^negative fibers and a main effect of treatment for dystrophin^negative fibers. SC‐Bmal1 ^Cntrl mice demonstrated greater necrosing fibers and dystrophin^negative fibers at 24 h versus 72 h and 7 days (Sidak's multiple comparisons, Figure 4C,F). However, there were no differences within SC‐Bmal1 ^iKO groups between 24, 72 h, and 7 days. SC‐Bmal1 ^iKO animals demonstrated ~62% less necrosing fibers and ~82% less dystrophin^negative fibers compared to SC‐Bmal1 ^Cntrl animals 24 h post‐injury (Sidak's multiple comparisons, 24 h: Necrosing fibers: SC‐Bmal1 ^Cntrl 87 ± 18; SC‐Bmal1 ^iKO 33 ± 15, Dystrophin^negative fibers: SC‐Bmal1 ^Cntrl 199 ± 41; SC‐Bmal1 ^iKO 36 ± 13). While these were higher in the SC‐Bmal1 ^iKO at 72 h, they were not significantly different (Necrosing fibers at 72 h: SC‐Bmal1 ^Cntrl 39 ± 14; SC‐Bmal1 ^iKO 46 ± 14, Dystrophin^negative fibers at 72 h: SC‐Bmal1 ^Cntrl 25 ± 11; SC‐Bmal1 ^iKO 68 ± 19) (Figure 4C,F).

FIGURE 4.

SC‐Bmal1 ^iKO animals demonstrate less necrosing fibers and dystrophin^negative fibers compared to control animals. (A, B, D and E) Representative images of H&E and dystrophin‐stained cross‐sections showing uninjured and 24 h post‐injury necrosing and dystrophin^negative fibers, respectively (arrow indicates necrosing fiber and dystrophin^negative fiber, respectively). (C) Number of necrosing fibers in either group across all timepoints (n = 4–5). (F) Number of dystrophin^negative fibers in either group across all timepoints (n = 4–5). Scale bar for all images set to 100 μm. All data shown as mean ± SEM. Groups were compared using two‐way ANOVA for main effects of hours‐post‐injury and treatment‐group with post‐hoc Sidak's multiple comparison tests (*, refer to results for Cntrl versus iKO comparisons and, $, refer to results for intra‐group significance) (*p < .05) (**p < .005) (***p < .001) (****p < .0001) ($ p < 0.05) ($$$ p < 0.001) ($$$$ p < 0.0001).

3.4. SC‐Bmal1^iKO animals exhibit reduced signs of muscle repair following contractile muscle injury

Figure 5A,C are representative images showing a cross‐section stained for laminin (blue)/eMHC+(green) and an H&E cross‐section reflective of centralized nuclei fibers, respectively, 7 d post‐injury. In both eMHC+ fibers and fibers containing centralized nuclei, there was a main effect of time post‐injury and an interaction effect. SC‐Bmal1 ^Cntrl mice revealed a greater number of fibers containing centralized nuclei and eMHC+ fibers at 7 d versus 24 h and 72 h (Figure 5B,D, Sidak's multiple comparisons). However, no differences were noted across timepoints within SC‐Bmal1 ^iKO groups in eMHC+ fibers or centralized nuclei. Muscle repair (measured as eMHC+ fibers and fibers containing centralized nuclei) was lower in SC‐Bmal1 ^iKO compared to SC‐Bmal1 ^Cntrl animals with ~87% fewer eMHC+ fibers and 71% fewer fibers containing centralized nuclei at 7 d post injury (eMHC⁺ fibers 7 day: SC‐Bmal1 ^Cntrl 217.8 ± 115.5; SC‐Bmal1 ^iKO 27.8 ± 17.3; Centralized nuclei 7 day: SC‐Bmal1 ^Cntrl 160.7 ± 70.5; SC‐Bmal1 ^iKO 46.2 ± 15.7, Sidak's post‐hoc multiple comparisons).

FIGURE 5.

SC‐Bmal1 ^iKO animals exhibit lesser degrees of muscle repair on day seven post‐injury. (A and C) Representative images showing an uninjured and a 7 day post‐injury cross‐section stained for laminin (blue)/eMHC+ (green) and a H&E cross‐section reflective of centralized nuclei fibers, respectively (arrows indicate eMHC+ fiber and a fiber with a centralized nuclei, respectively). (B) Total number of fibers expressing eMHC+ across timepoints and groups (n = 3–5). (D) Total number of fibers containing centralized nuclei across timepoints and groups (n = 4–5). Scale bar for all images set to 100 μm. All data shown as mean ± SEM. Groups were compared using two‐way ANOVA for main effects of hours‐post‐injury and treatment‐group with post‐hoc Sidak's multiple comparison tests (*, refer to results for Cntrl versus iKO comparisons and, $, refer to results for intra‐group significance) (*p < .05) (**p < .005) (***p < .001) ($ p < 0.05).

3.5. Reduced neutrophil response following contractile muscle injury in SC‐Bmal1^iKO animals

Figure 6A is a representative image showing a cross‐section stained for dystrophin, Ly6G/C⁺ content (referred to as Ly6+), DAPI and a merged image. Overall, there was a main effect of treatment‐group and time (h) post‐injury as well as an interaction effect. Control SC‐Bmal1 ^Cntrl mice demonstrated greater Ly6+ content at 24 versus 72 h and 7 d (Figure 6B, Sidak's multiple comparisons). However, no such differences were detected within SC‐Bmal1 ^iKO groups. Neutrophil content (Ly6+ content) was ~83% reduced at 24 h in SC‐Bmal1 ^iKO versus control animals (neutrophil content at 24 h: SC‐Bmal1 ^Cntrl 2.4 ± 0.4; SC‐Bmal1 ^iKO 0.4 ± 0.2, % area fraction, Sidak's multiple comparisons, Figure 6B). Furthermore, across all timepoints in both SC‐Bmal1 ^Cntrl and SC‐Bmal1 ^iKO groups, the number of dystrophin^negative and necrosing fibers were positively associated with neutrophil content (dystrophin^negative fibers r ² = 0.85 and 0.41; necrosing fibers r ² = 0.42 and 0.49, respectively).

FIGURE 6.

SC‐Bmal1 ^iKO animals experience less neutrophil infiltration. (A) Representative image showing a cross‐section stained for DAPI, dystrophin, Ly6⁺ content, and a merge image. (B) Total Ly6⁺ content in either group across all timepoints (n = 4–5). Scale bar for all images set to 100 μm. All data shown as mean ± SEM. Groups were compared using two‐way ANOVA for main effects of hours‐post‐injury and treatment‐group with post‐hoc Sidak's multiple comparison tests (*, refer to results for Cntrl versus iKO comparisons and, $, refer to results for intra‐group significance) (***p < .001) (****p < .0001) ($$$$ p < 0.0001).

3.6. Altered satellite cell activation and proliferation following contractile muscle injury in SC‐Bmal1^iKO animals

There were no group differences in baseline tdTomato⁺ SCs in the uninjured limb in a subset of animals (SC‐Bmal1 ^Cntrl: 24.9 ± 2.2; SC‐Bmal1 ^iKO: 25 ± 2.4 SC/100fibers, Figure S1A). Figure 7A,C are representative images showing a cross‐section with DAPI, tdTomato⁺ (Pax7), MyoD, or Ki67 (respectively) and a merged image. In both SC activation and proliferation, there were main effects of treatment‐group and time as well as an interaction effect (Figure 7B,D). SC‐Bmal1 ^Cntrl mice demonstrated greater SC activation and proliferation at 72 h compared to 24 h and 7 days (Figure 7B,D, Sidak's multiple comparisons). In SC‐Bmal1 ^iKO, SC activation and proliferation was greatest at 24 h versus 72 h, 7 days (Figure 7B,D). Additionally, in SC‐Bmal1 ^iKO, SC activation was increased at 7 days compared to 72 h (Figure 7B). There was greater SC activation in SC‐Bmal1 ^iKO versus SC‐Bmal1 ^Cntrl animals at 24 h and 7 days but, lower levels of activation at 72 h (SC activation 24, 72 h, 7 day: SC‐Bmal1 ^Cntrl 2.9 ± 0.4, 8.5 ± 0.9, 0.2 ± 0.1; SC‐Bmal1 ^iKO 13.6 ± 1.3, 0.1 ± 0.04, 5.6 ± 1.6, all units in tdTomato+/MyoD+ cells/100 fibers, Sidak post‐hoc analysis) (Figure 7B).

FIGURE 7.

Altered SC activation and proliferation kinetics in SC‐ Bmal1 ^iKO animals vs. control animals. (A and C) Representative images showing a cross‐section stained for DAPI, tdTomato (Pax7), MyoD or Ki67 (respectively), and a merge image. (B) Comparison of Pax7⁺/MyoD⁺ activated SCs across timepoints and between groups (n = 4–5). (D) Comparison of Pax7⁺/Ki67⁺ proliferating SCs across timepoints and between groups (n = 4–5). Scale bar for all images set to 100 μm. All data shown as mean ± SEM. Groups were compared using two‐way ANOVA for main effects of hours‐post‐injury and treatment‐group with post‐hoc Sidak's multiple comparison tests (*, refer to results for Cntrl versus iKO comparisons and, $, refer to results for intra‐group significance) (*p < .05) (**p < .005) (****p < .0001) ($ p < 0.05) ($$$$ p < 0.0001).

Similarly, compared to SC‐Bmal1 ^Cntrl animals, SC‐Bmal1 ^iKO animals exhibited significantly greater SC proliferation at 24 h and lower proliferation at 72 h post‐injury (SC proliferation 24 h, 72 h, 7 day: SC‐Bmal1 ^Cntrl 0.4 ± 0.1, 1.3 ± 0.3, 0.3 ± 0.1; SC‐Bmal1 ^iKO 2.4 ± 0.5, 0.3 ± 0.1, 0.4 ± 0.1, all units in tdTomato+/Ki67+ cells/100 fibers) (Figure 7D). There were positive associations between SC proliferation and neutrophil content at 24 h post‐injury, when neutrophil content was greatest (SC‐Bmal1 ^Cntrl r ² = 0.75, p = .058; SC‐ Bmal1 ^iKO r ² = 0.86, p < .05). A positive association was also observed for both SC‐Bmal1 ^Cntrl and SC‐Bmal1 ^iKO for SC activation and SC proliferation (r ² = 0.45, r ² = 0.68, respectively).

4. DISCUSSION

Using an inducible depletion model of SC‐specific clock gene, Bmal1 (SC‐Bmal1 ^iKO), we determined contractile function, contractile‐induced injury muscle damage, repair and myogenic progression in SC‐Bmal1 ^iKO and SC‐Bmal1 ^Cntrl animals. The major findings from this study were (1) ablation of SC‐Bmal1 reduced force production (in vivo and ex vivo) versus control animals, (2) SC‐Bmal1 ^iKO animals experienced less muscle damage (determined by muscle fiber necrosis, dystrophin^negative fibers, and neutrophil content) and subsequently displayed fewer regenerating fibers, and (3) the SC myogenic program (activation and proliferation) was altered in SC‐Bmal1 ^iKO animals with SC activation/proliferation occurring earlier and activated to a greater extent compared to control animals. These data suggest Bmal1 in SCs plays a regulatory role on force production and, following contractile injury, may influence the magnitude of damage/repair and SC myogenic progression.

Following in vivo eccentric injury, both SC‐Bmal1 ^iKO and control animals exhibited reductions in maximal torque, with the ~50% decrease being in close agreement with previous models of contractile‐induced injury. ³⁰ , ³³ , ⁴⁵ Yet despite similar reductions in maximal torque, SC‐Bmal1 ^iKO animals displayed reduced damage/necrosis and neutrophil infiltration following injury. An explanation for the reductions in markers of muscle damage is likely the lower forces produced by SC‐Bmal1 ^iKO animals, although the precise mechanisms underpinning such force reductions are not easily explained and were not associated with differences in muscle fiber‐type, myofiber area, or contractile kinetics. In line with this notion, high‐ versus low‐forces have been shown to induce a greater extent of injury, ²⁶ , ²⁷ with one study demonstrating this phenomenon by partially inhibiting myosin which resulted in reduced forces and extents of injury in healthy and diseased muscle. ⁴⁶ Additionally, SC‐Bmal1 ^iKO animals had both higher BW and muscle mass but still produced lower torque in vivo. Analysis of muscle force production ex vivo which allows for assessment of force normalized to PCSA (specific force) confirmed that the reductions in force did not simply stem from differences in animal BW or muscle mass. As such, one explanation may be that SC‐molecular clocks exert regulation on contractility via modulation of sarcoplasmic reticulum (SR) Ca⁺⁺ similar to our previous findings showing that SCs have time‐of‐day specific influence over contractile function via Ca⁺⁺ availability. ²⁸ Supportive of this, evidence elsewhere has shown muscle molecular clocks harbor regulation over Ca⁺⁺ contractile proteins and signaling pathways. ¹⁸ , ⁴⁷ , ⁴⁸ Alternatively, SC and muscle‐specific molecular clocks have been shown to regulate the diurnal expression of contractile and EC coupling related genes involved in force production. ¹⁰ , ¹⁸ Critical to maximal force production is the phosphorylation of myosin during cross‐bridge cycling ⁴⁹ , ⁵⁰ with genes responsible for phosphorylating myosin under molecular clock regulation. ¹⁸ Therefore, one possible explanation for our results may be that ablation of SC Bmal1 altered the rhythmicity and therefore post‐translational capacities of genes responsible for phosphorylating myosin during contraction. While SCs primary role is muscle repair, similar permissive functions of SCs have recently been observed and have influenced endurance exercise capacity, ⁵¹ , ⁵² contractility, ²⁸ , ⁵³ , ⁵⁴ and injury‐sensing functions as well. ⁵⁵ , ⁵⁶ , ⁵⁷ , ⁵⁸

Previously, whole body and muscle‐specific Bmal1 KO animals have exhibited lower force production with the underlying mechanism in these studies stemming from long‐term dysregulations on titin, leading to sarcomere‐length modifications. ¹⁶ , ¹⁷ , ²³ , ⁵⁹ However, these animal models were either bred from birth lacking Bmal1 or assessed 22 and 58 weeks following muscle‐Bmal1 ablation ²³ making it likely the mechanisms underpinning the observed force‐alterations were different to the animals in the current study that were assessed 10 days following SC‐Bmal1 ablation. We chose to utilize a shorter washout period to avoid any potential deleterious effects that SC‐Bmal1 ablation may have had on muscle structure that might subsequently impact muscle functional capacity. ²³ As such, we suggest that the effects on force production following SC‐Bmal1 ablation are not due to alterations in muscle structure. As our experiments were conducted at a single timepoint (Z4), we are limited in our implications for time‐of‐day variance. However, it is interesting to speculate that forces produced by SC‐Bmal1 ^iKO animals may have been further reduced at a later‐timepoint in the day when peak Bmal1 expression in SCs has been demonstrated. ¹⁰

Following muscle fiber damage, a first wave of immune cells and neutrophils infiltrate damaged‐regions to initiate a necrotic microenvironment that subsequently necessitates a reparative response from SCs upon further immune‐myogenic crosstalk signaling. ⁶⁰ , ⁶¹ , ⁶² In this regard, we show that the amount of damaged/necrosing fibers were associated with the neutrophil response across all timepoints in both SC‐Bmal1 ^Cntrl and SC‐Bmal1 ^iKO animals, confirming that neutrophils are a function of damage‐induced necrosis independent of the extent of damage that occurred in the absence of SC‐Bmal1. Indeed, SC‐Bmal1 ^iKO animals experienced significantly less damaged/necrosing fibers compared to their SC‐Bmal1 ^Cntrl counterparts after 24 h and consequently less neutrophil infiltration. One explanation for this observation is that the lower forces generated by SC‐Bmal1 ^iKO animals did not induce the widespread damage required to trigger a more extensive necrotic and immune response (supported by these animals performing less work (J/g); p = .06). Others have reported that when contractile‐injury is induced via low‐ versus high‐forces, the extent of injury scales as force increases. ²⁶ , ⁴⁶ In line with this, SC‐Bmal1 ^iKO animals exhibited less eMHC+ fibers and centralized nuclei 7 days post‐injury suggesting that these animals also experienced lesser degrees of muscle repair. We note that in cases of non‐physiological injury (i.e., cardiotoxin‐induced) when the extent of damage exceeds that of contractile‐induced injury, the relationship between SC‐molecular clocks and the damage‐induced neutrophil response may manifest differently. ⁶³ This is likely because models of extreme injury can expose the maximum capacities of cellular events in the muscle regenerative cascade compared to contractile injuries. ⁶⁴ In this regard, the regulatory mechanisms observed by Zhu et al 2024 ⁶³ between SC‐molecular clocks and neutrophils following cardiotoxin injury might also be at play in the current study but may be masked by the higher/lower forces that induced the contractile injury. Potential evidence for this notion was the dysregulated temporal relationship of necrosis, dystrophin^negative fibers and neutrophil infiltration observed across timepoints post‐injury in SC‐Bmal1 ^iKO animals.

Following contractile‐induced muscle injury, the necrotic and neutrophil response initiates SC myogenic progression to facilitate muscle repair. ⁸ , ⁶⁰ , ⁶¹ , ⁶⁵ , ⁶⁶ We show that across all animals and timepoints, SC activation/proliferation proceeded following contractile injury induced damage. However, the timeline of SC progression differed between groups as SC activation/proliferation in SC‐Bmal1 ^iKO peaked at 24 h compared to a later response (72 h) in SC‐Bmal1 ^Cntrl animals. One explanation for this earlier timeline of SC progression could be that these animals experienced less damage/neutrophil infiltration permitting SCs to become activated sooner. Indirect support for this premise are human models of eccentric damage in which SC activation following voluntary versus electrically stimulated (ES) eccentric contraction occurs at an earlier timepoint, likely due to there being no “delay” in activation as there is less fiber necrosis that occurs after voluntary versus ES eccentric contractions. ⁸ , ⁶⁷ , ⁶⁸

Despite having experienced less damage/repair, SC‐Bmal1 ^iKO animals displayed higher relative peaks in SC activation/proliferation at 24 h and an unexplained increase in activation at 7 days post‐injury versus control animals. Since SC‐Bmal1 ^iKO animals underwent less damage/necrosis, neutrophil infiltration and muscle repair, the greater magnitude of SC activation/proliferation is somewhat surprising. These findings may be related to the critical role Bmal1 plays in MyoD's rhythmicity ¹⁸ , ¹⁹ where, upon our inducible depletion of Bmal1 in SCs, MyoD's trajectory/amplitude during myogenesis may have been perturbed. Bmal1 binds to the core‐enhancer (CE) region of MyoD during homeostasis which in‐turn induces MyoD's rhythmic expression throughout the day. However, the ablation of the CE region alters such rhythmicity and dysregulates the timing and amplitude of MyoD's expression during myogenesis. ¹⁹ , ²⁰ Furthermore, prior work reveals that muscle‐specific Bmal1 KO leads to altered MyoD expression patterns over 24 h with increases in rhythmic expression amplitude. ²¹ Such findings may partially explain our observations of an increased amplitude in SC‐MYOD content in SC‐Bmal1 ^iKO animals at 24 h and altered timing at 7 days post‐injury. Collectively, these data suggest SC‐Bmal1 may be required for the appropriate timing of MyoD's response throughout regenerative myogenesis, in line with previous work demonstrating that animals lacking Bmal1's binding site on MyoD had dysregulated MyoD expression during embryonic myogenesis ultimately blunting muscle growth. ¹⁹ , ²⁰ , ⁶⁹ Additional evidence is that SCs and SC‐derived myogenic progenitors of Bmal1 KO animals (whole‐body and SC‐specific) harbor alterations in activation and proliferation following injury, offering further support that Bmal1 regulates MyoD and thus SC activation/proliferation. ¹² , ¹⁴ , ¹⁵ , ⁷⁰

5. CONCLUSION

Ablation of SC‐Bmal1 resulted in ~20–25% lower force production in vivo and ex vivo compared to control animals. Following in vivo contractile injury, SC‐Bmal1 ^iKO animals underwent less necrosis and muscle repair, suggesting the reduced forces led to a lesser magnitude of injury. In line with this notion, SC‐Bmal1 ^iKO animals exhibited less neutrophil infiltration post‐injury. SC activation and proliferation occurred earlier (24 h versus 72 h) in SC‐Bmal1 ^iKO animals suggesting the reduced neutrophil response may have allowed for an earlier timeline of SC myogenic progression. Finally, although SC‐Bmal1 ^iKO may have experienced less damage, potentially explaining the earlier peaks in SC progression, these mice exhibited a greater extent of SC activation as well as an unexplained increase in activation 7 days post‐injury. These findings indicate SC‐Bmal1 may be required for the appropriate response and timing of MyoD during regenerative myogenesis. Collectively, ablation of SC‐Bmal1 lowered force production and altered the magnitude of muscle damage/repair and SC myogenic progression following eccentric contractile injury.

6. LIMITATIONS

Our study has several limitations. We did not employ a 12:12 light–dark photoperiod as we have previously reported that animals housed in a 14:10 photoperiod, and in the same room of the vivarium, exhibit typical, rhythmic oscillations in muscle molecular clock gene expression. ²⁸ Additionally, to assess if SC‐Bmal1 played a role in muscle repair following physiological (contractile) injury, we elected to carry out experiments at a single time point to determine if any differences were present. This study design was in line with past Bmal1 ablation models that utilized a single timepoint to study the effects of Bmal1 absence on either physiology or injury/repair. ¹⁴ , ¹⁷ , ²³ Given this study design, we were able to show that SC‐Bmal1 ablation influenced muscle repair following contractile injury. However, we are limited in our interpretations regarding SC‐Bmal1's circadian regulation on force production and the implications that may have on subsequent injury/repair. Future studies exploring the connection between SC‐Bmal1 and force production will require multiple time‐course measurements of force aligned to varying levels of SC‐Bmal1 expression. The impact of SC‐Bmal1 on the regulation of the neutrophil response and the assessment of the time‐of‐day regulation of SC‐Bmal1 on neutrophils is also an area that warrants further investigation.

AUTHOR CONTRIBUTIONS

RK, SD, JAH contributed to conceptualization and preparation of this manuscript. RK performed all experimentation. RK drafted manuscript. CP and PZ provided the animals used in these experiments and were involved in data interpretation. IR provided the in vivo contractile apparatus setup used on all animals. RK, JAH, SD, CP, PZ, IR contributed to manuscript editing and revising.

DISCLOSURES

The authors declare no competing interests.

Supporting information

Figure S1.

Figure S2.

Text S1.

Kahn RE, Zhu P, Roy I, Peek C, Hawley JA, Dayanidhi S. Ablation of satellite cell‐specific clock gene, Bmal1, alters force production, muscle damage, and repair following contractile‐induced injury. The FASEB Journal. 2025;39:e70325. doi: 10.1096/fj.202402145RR

DATA AVAILABILITY STATEMENT

All data used within the results and to create figures are included in the material of this manuscript. Additional analysis and files can be provided upon request.