Postdevelopmental knockout of Orai1 improves muscle pathology in a mouse model of Duchenne muscular dystrophy

Duchenne muscular dystrophy (DMD), an X-linked recessive muscle disorder that appears early during childhood, is characterized by muscle weakness, reduced contractile function, and progressive muscle degeneration. Despite significant advances in understanding DMD pathogenesis and the development of promising therapeutic strategies, there is currently no cure for DMD, and treatment remains primarily supportive in nature. DMD is linked to loss-of-function mutations in the gene that encodes dystrophin, a 427-kD structural protein on the myoplasmic face of the sarcolemma (Monaco et al., 1986). Dystrophin anchors γ-actin filaments of the subsarcolemmal cytoskeleton to a macromolecular assembly of proteins collectively known as the dystrophin-associated protein complex (Hoffman et al., 1987), which makes further connections via laminin to the extracellular basal lamina. Thus, loss of dystrophin disrupts this structural complex that is critical for maintaining sarcolemmal integrity during mechanical stress. However, the precise mechanisms that underlie progressive muscle fiber degeneration and necrosis in DMD continue to be debated. A long-standing hypothesis for muscle fiber degeneration in DMD is that a sustained elevation in myoplasmic Ca²⁺ triggers multiple intracellular downstream pathways/mechanisms, including mitochondrial Ca²⁺ overload/damage, enhanced oxidative stress, and activation of Ca²⁺-dependent proteases that together promote myofiber degeneration and death (Durbeej and Campbell, 2002).

Currently, two main pathomechanisms are hypothesized to underlie abnormalities in myoplasmic Ca²⁺ homeostasis in DMD: (1) enhanced Ca²⁺ leak from the sarcoplasmic reticulum (SR) through oxidized type 1 ryanodine receptor (RyR1) Ca²⁺ release channels and (2) excessive extracellular Ca²⁺ influx. The first pathomechanism is supported by several studies showing that loss of dystrophin is associated with hypernitrosylation of specific cysteine residues in RyR1 leading to FKBP12 dissociation from RyR1, destabilization of the RyR1 channel closed state, and increased RyR1-dependent SR Ca²⁺ leak (Bellinger et al., 2009; Andersson et al., 2012). Abnormal SR Ca²⁺ leak subsequently leads to mitochondrial Ca²⁺ overload and uncontrolled production of reactive species of oxygen and nitrogen (ROS and RNS, respectively) that further promotes RyR1 oxidization/nitrosylation in a destructive feed-forward cycle of increased RyR1 Ca²⁺ leak and ROS/RNS production. Besides enhancing RyR1 opening probability, ROS/RNS-dependent oxidative stress can also reduce the activity of the SR/ER Ca²⁺ ATPase (SERCA), the main Ca²⁺ pump in the SR membrane responsible for Ca²⁺ reuptake (Viner et al., 1996; Sharov et al., 2006; Dremina et al., 2007; Qaisar et al., 2021). Indeed, a reduction of SERCA activity due to increased oxidative stress was reported previously in dystrophic muscles (Divet and Huchet-Cadiou, 2002; Kargacin and Kargacin, 1996). The importance of increased oxidative stress as a key pathomechanism of muscle degeneration in DMD is further supported by improvement of the dystrophic phenotype following treatment with N-acetylcysteine, a potent antioxidant (Whitehead et al., 2008).

The second pathomechanism for altered Ca²⁺ homeostasis in DMD involves a net increase in trans-sarcolemmal Ca²⁺ influx through “microtears” and/or Ca²⁺-permeable ion channels (e.g., leak, stretch-activated, receptor-activated, and store-operated channels; Boittin et al., 2006; Fong et al., 1990; Franco and Lansman, 1990; Vandebrouck et al., 2002; Millay et al., 2009). A growing number of studies provide evidence for a modulatory role of enhanced store-operated Ca²⁺ entry (SOCE) in the pathogenesis of DMD (Edwards et al., 2010; Zhao et al., 2012; Cully et al., 2012). SOCE is a ubiquitous Ca²⁺ influx mechanism triggered by the depletion of intracellular Ca²⁺ stores, that allows for the recovery of Ca²⁺ ions from the extracellular space needed to both refill ER/SR and activate downstream Ca²⁺ signaling pathways (e.g., NFAT). The two proteins that form the core SOCE machinery are (1) stromal interaction molecule-1 (STIM1), which senses changes in luminal ER/SR Ca²⁺ levels (Roos et al., 2005; Zhang et al., 2005; Liou et al., 2005), and (2) Orai1, the Ca²⁺ release-activated Ca²⁺ channel in the surface membrane (Feske et al., 2006; Yeromin et al., 2006; Vig et al., 2006). STIM1 and Orai1 also coordinate SOCE in skeletal muscle (Stiber et al., 2008; Lyfenko and Dirksen, 2008; Dirksen, 2009). Consistent with the idea that dysregulated activation of STIM1/Orai1-dependent SOCE promotes Ca²⁺-mediated muscle fiber degeneration in muscular dystrophy, STIM1 and Orai1 expression levels and SOCE activity were reported to be increased in dystrophin-deficient mdx mice (Edwards et al., 2010; Zhao et al., 2012; Cully et al., 2012). In addition, early developmental inhibition of SOCE function significantly mitigates the dystrophic phenotype observed in both dystrophin-null (mdx) and γ-sarcoglycan–null mice (Goonasekera et al., 2014).

Because enhanced STIM1/Orai1-mediated SOCE is proposed to be an important mechanism of aberrant Ca²⁺ influx in mdx mice, we evaluated the role of Orai1-dependent Ca²⁺ entry in the DMD phenotype by crossing mdx mice with muscle-specific, tamoxifen-inducible, Orai1-knockout (Orai1 KO) mice. The resulting compound offspring enable tamoxifen-inducible ablation of Orai1 in adult mdx mice (mdx-Orai1 KO mice). We found that ablation of Orai1 in young adult (2–3-mo-old) mdx mice abolished Orai1-dependent Ca²⁺ entry (e.g., SOCE), normalized intracellular Ca²⁺ homeostasis, improved muscle performance, and unexpectedly protected muscles from eccentric contraction–induced damage. Furthermore, postdevelopmental Orai1 KO normalized fiber cross-sectional area (CSA) and reduced muscle fibrosis, a marker of muscle damage and necrosis. These results demonstrate an important role of enhanced Orai1-mediated Ca²⁺ entry in exacerbating the dystrophic phenotype of mdx mice, rendering Orai1 an attractive potential therapeutic target for the treatment of DMD.

Tamoxifen-inducible, muscle-specific Orai1-KO (HSA-MCM Orai1^fl/fl) mice were generated as previously described (Carrell et al., 2016). Briefly, we generated dystrophic mice that permit inducible Orai1 ablation in adult skeletal muscle by crossing male HSA-MCM Orai1^fl/fl mice with homozygous female mdx mice (C57Bl/10ScSn-DMDmdxIJ; Jackson Laboratories). This strategy enabled generation of F1 heterozygous female mdx mice carrying a single floxed Orai1 allele (mdx-Orai1 Het). These mice were then mated with male HSA-MCM Orai1^fl/fl mice to obtain dystrophin-deficient mice carrying two floxed Orai1 alleles (mdx-Orai1 KO). Using this breeding strategy (see Fig. 1 A), we obtained the following four genotypes used in this study: (1) WT mice (HSA-MCM negative Orai1^fl/fl or Orai1^+/fl), (2) mdx mice (mdx-HSA-MCM negative Orai1^fl/fl or Orai1^+/fl), (3) Orai1 KO mice (HSA-MCM-Orai1^fl/fl) mice, and (4) mdx-Orai1 KO mice (mdx-HSA-MCM-Orai1^fl/fl). The mdx allele was detected using the following primers: (forward) 5′-AAC​TCA​TCA​AAT​ATG​CGT​GTT​AGT-3′; (reverse) 5′-GCC​CCT​CAA​TCT​CTT​CAA​ATT​C-3′. The MerCreMer and Orai1 floxed alleles were verified according to protocols described previously (Carrell et al., 2016). To induce Orai1 KO, 2-mo-old mice were treated with a tamoxifen-supplemented diet (Envigo; TD.130860) for 4 wk (while monitoring normal weight gain). Mice were then fed regular mouse chow for ≥1 wk. As DMD is an X-linked disease, all the experiments were restricted to 13–16-wk-old male mice. All animal studies were designed to minimize animal suffering and were approved by the University Committee on Animal Resources at the University of Rochester (UCAR2006-114E).

3–4-mo-old mice were pretrained on a 6-lane treadmill (Columbus Instruments) at a modest treadmill speed of 5 m/min for 5 min at a 15° incline over 3 consecutive days. On the fourth day, mice were subjected to a 1-h endurance run on the treadmill (1 km of total distance covered), starting at 5 m/min for 5 min, followed by increments of 1 m/min until reaching a maximum speed of 20 m/min, and then maintaining this speed for 40 min. Continued running was encouraged by delivering brief (<1 s) spurts of air on the mouse’s backside using a Whoosh Duster. The number of rests during each 5-min window of time was recorded for each mouse. Exhaustion was defined as an inability of the mouse to re-engage the treadmill after three consecutive, <1-s spurts of air as described previously (Carrell et al., 2016). The number of rests and the total distance run were recorded for each mouse. In vivo mouse forelimb grip strength was assessed using a digital force gauge (GTX; Dillon) as described previously (Loy et al., 2011).

Semiquantitative RT-PCR was used to assess relative changes in Orai1 transcript as previously reported (Carrell et al., 2016). Briefly, total RNA was isolated from tibialis anterior (TA) muscles using TRIzol (cat. 15596-018; Thermo Fisher Scientific) and quantified. RNA (1 μg) was then treated with DNase according to the manufacturer’s protocol (EN0525; Thermo Fisher Scientific) and reverse transcribed using a Super Script III First-Strand Synthesis System primed with oligonucleotides (dT; cat. 18080-051; Thermo Fisher Scientific). Semiquantitative PCR was carried out on 10 ng cDNA using 5′-end fluorescein (6-FAM)–labeled forward primers (Integrated DNA Technologies). Reactions were quantified every 2 cycles from 22 to 32 cycles to verify that amplification of both control GAPDH and Orai1 cDNAs were within the linear range. Primers used were mouse GAPDH (921 bp): (forward) 5′-AGG​CCG​GTG​CTG​AGT​ATG​TC-3′; (reverse) 5′-GGG​TGC​AGC​GAA​CTT​TAT​TGA​TGG-3′; and mouse Orai1 (307 bp): (forward) 5′-TTT​AGT​GCC​TGC​ACC​ACA​GTG​CTA-3′; (reverse) 5′-TGT​GGT​TGG​CGA​CGA​TGA​CTG​ATT-3′.

Orai1 transcript level was also assessed using a quantitative RT-PCR approach on RNA isolated from TA muscles as described above. Quantitative PCR was performed with 10 ng cDNA on a StepOnePlus Real-Time PCR machine (Applied Biosystems) using SYBR Green FastMix (Quantabio). Relative mRNA transcript levels from each experiment were performed in triplicate and standardized to their own internal GAPDH gene expression, then to the control condition using the 2^−∆∆CT analysis method, where CT is count threshold, in accordance with Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines (Bustin et al, 2009). Primers used were mouse Orai1 (forward): 5′-GAT​CGG​CCA​GAG​TTA​CTC​CG-3′, (reverse): 5′-TGG​GTA​GTC​ATG​GTC​TGT​GTC-3′; and mouse GAPDH (forward): 5′-AGG​TCG​GTG​TGA​ACG​GAT​TTG-3′, (reverse): 5′-GGG​GTC​GTT​GAT​GGC​AAC​A-3′.

Flexor digitorum brevis (FDB) muscles were dissected from mouse hind paws and placed in a dish containing Ringer’s solution consisting of 145 mM NaCl, 5 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂, and 10 mM HEPES, pH 7.4. Muscles were then incubated in Ringer’s solution supplemented with 1 mg/ml collagenase A for 60 min while rocking gently at 37°C to allow for enzymatic dissociation. Single FDB fibers obtained by mechanical dissociation/trituration were plated on glass-bottom dishes and allowed to settle for ≥20 min before conducting single-fiber experiments (detailed below). Only fibers with a clean morphology, clear striations, and no signs of swelling or damage were selected for experiments.

For Mn²⁺ quench studies, isolated FDB fibers were loaded with 5 μM fura-2 AM for 1 h at 37°C in a Ca²⁺-free Ringer’s solution containing 145 mM NaCl, 5 mM KCl, 3 mM MgCl₂, and 0.2 mM EGTA, pH 7.4. During fura-2 loading, fibers were also incubated with two SERCA pump inhibitors (1 μM thapsigargin [TG] and 15 μM cyclopiazonic acid [CPA]) to fully deplete SR Ca²⁺ stores before measurement of the maximal rate of store-dependent Mn²⁺ quench (store-dependent entry or SOCE). In a second set of experiments, FDB fibers were loaded with fura-2 AM in the absence of SERCA pump inhibitors (store-independent entry or constitutive entry). To prevent movement artifacts during Mn²⁺ quench recordings, 30 μM N-benzyl-p-toluene sulfonamide (BTS) was also included in the extracellular solution (Wei-Lapierre et al., 2013; Michelucci et al., 2019). Fibers were then rinsed with Ca²⁺-free Ringer solution and excited at 362 nm (isosbestic point of fura-2), while emission was detected at 510 nm using a DeltaRam illumination system (Photon Technology International). After obtaining an initial baseline rate of fura-2 decay (R_baseline), fibers were exposed to Ca²⁺-free Ringer’s supplemented with 0.5 mM MnCl₂. The maximum rate of change in fura-2 fluorescence in the presence of Mn²⁺ (R_max) was obtained from the peak time derivative of the fura-2 emission trace during Mn²⁺ application. The maximum rate of entry (R_ENTRY; either with or without prior store depletion) was then calculated as R_ENTRY = R_max − R_baseline and expressed as dF/dt in counts/s as described previously (Wei-Lapierre et al., 2013; Michelucci et al., 2019).

Total releasable Ca²⁺ store content was determined from single FDB fibers loaded with 4 µM fura-FF AM, a low-affinity, ratiometric Ca²⁺ dye, for 30 min at room temperature in control Ringer’s solution followed by 30-min incubation in dye-free Ringer’s solution supplemented with 40 µM BTS, as described previously (Zvaritch et al., 2007; Loy et al., 2011; Wei-Lapierre et al., 2013; Linsley et al., 2017; Michelucci et al., 2019; Michelucci et al., 2020). Fibers were then perfused in Ca²⁺-free Ringer’s solution while alternately excited at 340 and 380 nm (510-nm emission) every 250 ms (30-ms exposure per wavelength and 2 × 2 binning) using a monochromator-based illumination system (TILL Photonics). Fura-FF emission at 535 ± 30 nm was captured using a high-speed, digital QE charge-coupled device camera (TILL Photonics). Total releasable Ca²⁺ store content was assessed from the difference between baseline and peak fura-FF ratios (ΔRatio_340/380) upon application of a Ca²⁺ store release cocktail containing 10 µM ionomycin, 30 µM CPA, and 100 µM EGTA in a Ca²⁺-free Ringer’s solution (ICE) as previously described (Loy et al., 2011; Wei-Lapierre et al., 2013; Michelucci et al., 2019, Michelucci et al., 2020). To confirm that the peak of the fura-FF signal during ICE application was not saturated, maximal fura-FF responsiveness was assessed at the end of each experiment by subsequent application of normal Ca²⁺-containing Ringer’s solution. Analysis of the peak ICE-induced change in fura-FF ratio (ΔRatio_340/380) was calculated using Clampfit 10.0 (Molecular Devices).

Single FDB fibers were loaded with 5 µM fura-2 AM for 30 min at room temperature in control Ringer’s solution followed by a 30-min incubation with dye-free Ringer’s solution. Fura-2–loaded fibers were then placed on the stage of an inverted epifluorescence microscope (Nikon Instruments) and alternately excited at 340 and 380 nm (30-ms exposure per wavelength and 2 × 2 binning) using a monochromator-based illumination system. The fluorescence emission at 510 nm was captured using a high-speed, digital QE charge-coupled device camera (TILL Photonics). Fura-2 340/380 ratios from myoplasmic areas of interest were calculated using TILLvisION software, analyzed offline using ImageJ, and then converted to resting free Ca²⁺ concentrations using an in situ calibration curve for fura-2 generated as described previously (Michelucci et al., 2019).

Ex vivo assessments of muscle-specific force, fatigue during repetitive high-frequency stimulation, and eccentric muscle damage were made in excised extensor digitorum longus (EDL). Muscle-specific force and eccentric muscle damage were also assessed in soleus muscles. Briefly, mice were anesthetized by intraperitoneal injection of an anesthetic cocktail as described previously (Wei-Lapierre et al., 2013). EDL and soleus muscles were isolated, tied using 4-0 surgical suture, carefully excised, attached to a servo motor and force transducer (1200 A, Aurora Scientific), and placed between two platinum electrode plates in a chamber continuously perfused with oxygenated Ringer solution containing 137 mM NaCl, 5 mM KCl, 1.2 mM NaH₂PO₄, 1 mM MgSO₄, 2 mM CaCl₂, 10 mM glucose, and 24 mM NaHCO_3, pH 7.4. Before starting each experiment, optimal stimulation intensity and muscle length (L_o) were determined using a series of 1-Hz twitch stimulation trains to guide stretching the muscle to the length that generated maximal force (F_o). After establishing L_o, muscles were first equilibrated using three 500-ms, 150-Hz tetani delivered at 1-min intervals. EDL and soleus muscles were then subjected to a force–frequency stimulation protocol (from 1 to 250 Hz for EDL muscles; from 1 to 200 Hz for soleus muscles). To assess muscle fatigability, EDL muscles were subjected to a repetitive, high-frequency stimulation protocol (40 stimulus trains of 50 Hz and 500 ms in duration delivered every 2.5 s). To assess susceptibility to damage during eccentric contractions, a different set of EDL and soleus muscles were subjected to a repetitive eccentric contraction protocol optimized for eccentric contraction–induced damage as previously described (Lindsay et al., 2020). This protocol consisted of 10 successive 700-ms, 150-Hz eccentric contractions (500 ms of isometric contraction followed by an additional 200 ms during a 10% of stretch of L_o at a speed of 0.5 L_o/s). Muscle physiological CSA and specific force were calculated as described previously (Hakim et al., 2011).

TA muscles were excised and incubated in 30% sucrose overnight at 4°C (Bachman et al., 2018). Muscles were mounted in optimal cutting temperature medium and snap frozen in liquid nitrogen–cooled 2-methylbutane (Carrell et al., 2016). TA muscle cryosections (10 μm thick) were fixed in 4% paraformaldehyde (3–5 min), washed with PBS, and permeabilized by exposure to 0.1% Triton X-100 in PBS for 5 min. Sections were washed again with PBS, blocked with 3% BSA for 30 min at room temperature, and incubated for 2 h at room temperature with anti-laminin antibody (rat monoclonal diluted 1:100; Sigma-Aldrich). Sections were then washed with PBS and incubated with Alexa Fluor 488 (anti-rat polyclonal; 1:400 diluted; Invitrogen) for 1 h at room temperature. Nuclei were stained with Hoechst dye (0.2 μg/μl for 6 min at room temperature) and washed with PBS before being mounted with VectaShield (Vector Laboratories) for analysis by confocal microscopy. Fiber boundaries were defined by the laminin signal, and CSA was calculated using Fiji software as described previously (Tyagi et al., 2017). To examine gross muscle structure and number of centrally nucleated fibers, cryosections were stained with hematoxylin and eosin as described previously (Brennan et al., 2019).

Diaphragm muscles were excised, carefully cleaned to remove attached tendons, weighed, and snap frozen in liquid nitrogen. Collagen content was measured using a hydroxyproline assay kit (MAK008; Sigma-Aldrich). Briefly, diaphragm muscles were homogenized in water and hydrolyzed overnight in 6 M hydrochloric acid at 110°C. 10 µl of homogenate was then transferred to a 96-well plate and incubated at 60°C to dry the sample. Dried samples were resuspended in chloramine T/oxidation buffer mixture, incubated at room temperature for 5 min, mixed with DMAB-perchloric acid/isopropanol reagent, and then incubated for 90 min at 60°C. Sample absorbance was measured at 560 nm using a FlexStation 3 Multi-Mode Microplate Reader (Molecular Devices). Absorbances (ODs) obtained for each sample were referred against a standard curve to quantify the amount of hydroxyproline, which was then normalized to milligrams of tissue wet-weight.

EDL muscles were dissected, snap frozen in liquid nitrogen, and homogenized in RIPA lysis buffer (20 mM Tris-HCl, 150 mM NaCl, 1 mM Na₂EDTA, 1 mM EGTA, 1% NP-40, 1% sodium deoxycholate, 2.5 mM sodium pyrophosphate, 1 mM β-glycerophosphate, 1 mM Na₃VO₄, and 1 μg/ml leupeptin, pH 7.5) supplemented with a cocktail of protease inhibitors. Protein concentration was determined spectrophotometrically using a Lowry method. Briefly, 10 μg of total protein samples were loaded in 10–12% SDS-PAGE polyacrylamide gel, transferred to nitrocellulose membranes, and blocked with 3% BSA in Tris-buffered saline 0.1% and Tween 20 (TBS-T) overnight. Membranes were probed with the following primary antibodies diluted with 1% BSA in TBS-T for 2 h at room temperature: dystrophin (rabbit polyclonal diluted 1:1,000; Abcam), utrophin (mouse monoclonal diluted 1:1,000; Santa Cruz Biotechnology), STIM1 (rabbit polyclonal diluted 1:5,000; Sigma-Aldrich), SERCA (rabbit polyclonal 1:10,000; Santa Cruz Biotechnology), CASQ1 (mouse monoclonal diluted 1:5,000; Thermo Fisher Scientific), β-tubulin (mouse monoclonal diluted 1:2,000; Invitrogen), and GAPDH (mouse monoclonal diluted 1:50,000; Invitrogen). β-Tubulin was used as a loading control for dystrophin and utrophin, whereas GAPDH was used as a loading control for STIM1, SERCA, and CASQ1. Membranes were then washed three times in TBS-T and incubated with either goat anti-mouse IgG-800 or goat anti-rabbit IgG-800 (both diluted 1:10,000; Invitrogen) secondary antibodies diluted in 1% BSA TBS-T for 1 h at room temperature. Proteins were visualized with an Odyssey Infrared imager from Li-Cor. Densitometry analysis and quantification were performed on exported TIFF images using ImageJ (National Institutes of Health).

All data groups were verified to follow a normal distribution using an Anderson–Darling (A²) test before conducting parametric statistical analyses. The following analyses were performed blinded: (1) quantitative analyses of muscle fiber CSA and fibrosis (Fig. 2), (2) in vivo assessment of grip strength and cumulative rests during treadmill endurance protocol (Fig. S1), and (3) quantitative histologic analysis of centrally nucleated fibers (Fig. S3 B). Statistical significance was determined using one-way ANOVA followed by Dunnett’s test or ANOVA followed by post hoc Tukey test for repeated measures. A non-parametric test was used when data did not follow a normal distribution. Data were analyzed using Prism v8.0.2 for Windows (GraphPad Software), Origin 8.0 (OriginLab Corp.), pCLAMP (Molecular Devices), and ImageJ. In all cases, differences were considered statistically significant at P < 0.05. All data are presented as mean ± SEM.

Fig. S1 summarizes the impact of postdevelopmental, muscle-specific Orai1 ablation in mdx mice on grip strength and treadmill endurance. Fig. S2 compares the magnitude of BTP2-sensitive constitutive Ca²⁺ entry in FDB fibers from WT and mdx mice. Fig. S3 shows that postdevelopmental, muscle-specific Orai1 ablation does not reduce the high level of central nuclei observed in TA muscles from mdx mice. Fig. S4 presents the effect of postdevelopmental, muscle-specific Orai1 ablation in mdx mice peak EDL specific force during sustained tetanic stimulation. Fig. S5 summarizes the impact of postdevelopmental, muscle-specific Orai1 ablation in mdx mice on eccentric contraction–induced damage in soleus muscle. Fig. S6 shows the effect of postdevelopmental, muscle-specific Orai1 ablation in mdx mice on peak EDL specific force during repetitive, high-frequency stimulation. Fig. S7 shows that BTP2 exposure does not alter the magnitude of eccentric contraction–induced damage in EDL muscles from either WT or mdx mice. Fig. S8 summarizes the effect of postdevelopmental, muscle-specific Orai1 ablation in mdx mice on the expression levels of key proteins key involved membrane stability (dystrophin and utrophin) and SR Ca²⁺ levels (SERCA, CASQ1, STIM1L, and STIM1S).

The dysregulation of Ca²⁺ homeostasis in the pathophysiology of DMD has been attributed to multiple mechanisms including microscopic membrane “tears,” as well as increased activity of Ca²⁺-permeable mechanosensitive and store-operated channels (Franco and Lansman, 1990; Fong et al., 1990; Vandebrouck et al., 2002; Millay et al., 2009; Goonasekera et al., 2014; Onopiuk et al., 2015). Previous works also found that muscle fibers from dystrophic mice exhibit increased expression levels of STIM1 and Orai1 that correlate with enhanced SOCE activity (Edwards et al., 2010; Zhao et al., 2012; Cully et al., 2012). To investigate the specific role of Orai1 in DMD, we crossed tamoxifen-inducible, muscle-specific Orai1 KO (Orai1 KO) male mice (Carrell et al., 2016) with mdx female mice homozygous for the dystrophin mutation. The resulting first generation of mdx-Orai1 heterozygous female mice were then bred with male Orai1 KO mice to obtain the four different genotypes used in this study (WT, mdx, mdx-Orai1 KO, and Orai1 KO mice; Fig. 1 A).

To assess the efficacy of postdevelopmental Orai1 ablation, we measured Orai1 transcript levels in TA muscle homogenates using both semiquantitative RT-PCR (Fig. 1 B) and quantitative RT-PCR approaches (Fig. 1 C). Similar to data reported previously (Zhao et al., 2012), Orai1 transcript levels, assessed with both approaches, were significantly increased (∼1.5-fold) in muscle of mdx mice compared with that of WT mice, while virtually absent (∼95% reduction in Orai1 mRNA level) in muscle from both mdx-Orai1 KO and Orai1 KO mice. We used the maximum rate of Mn²⁺ quench of fura-2 fluorescence in single FDB fibers following SR Ca²⁺ store depletion (pretreated for 1 h with 1 μM TG plus 15 μM CPA in a Ca²⁺-free Ringer’s solution) as an index of functional Orai1 expression and maximal SOCE in muscle as described previously (Wei-Lapierre et al., 2013). Consistent with the Orai1 transcript levels reported in Fig. 1, B and C, the maximum rate of Mn²⁺ quench of fura-2 fluorescence was ∼1.75 times greater in fibers from mdx mice (3.2 ± 0.8 counts/s × 10³) compared with that of WT mice (1.8 ± 0.8 counts/s × 10³), while essentially undetectable in fibers from both mdx-Orai1 KO and Orai1 KO mice (Fig. 1, D and E). Interestingly, postdevelopmental Orai1 ablation resulted in a partial rescue of reduced grip strength and increased fatigue during forced treadmill running (Fig. S1).

Prior studies reported increased constitutive Ca²⁺ entry (e.g., Ca²⁺ entry in the absence of active pharmacological store depletion) and resting Ca²⁺ levels in muscle fibers from mdx mice that are both reduced upon removal of extracellular Ca²⁺ (Altamirano et al., 2012). Thus, we addressed whether mdx fibers exhibited significant constitutive Orai1-dependent Ca²⁺ entry by quantifying Mn²⁺ quench of fura-2 fluorescence in WT and mdx fibers in the absence of actively depleting SR Ca²⁺ stores (naive fibers). To do this, Mn²⁺ quench of fura-2 fluorescence was measured in naive fibers first before and then after addition of BTP2, a potent SOCE channel inhibitor (Zitt et al., 2004). Results from these studies demonstrated that mdx fibers exhibited a significant constitutive BTP2-dependent Ca²⁺ entry that is not observed in WT fibers (Fig. S2).

Increases in fiber size (hypertrophy), incidence of central nucleation (nuclei localized at the center of the fiber), inflammation, fibrosis, and necrosis are all hallmarks of muscular dystrophy (Verhaart and Aartsma-Rus, 2019; Deconinck and Dan, 2007). Thus, we assessed muscle fiber CSA in 10-μm-thick slices obtained from frozen TA muscles immunolabeled with anti-laminin antibody and nuclei stained with Hoechst dye (Fig. 2 A). As expected, average CSA of muscle fibers from mdx mice (2,126 ± 61 μm²) was approximately two times greater than that of fibers from WT mice (1,128 ± 21 μm²). Importantly, the CSA of fibers from mdx-Orai1 KO mice (1,214 ± 22 μm²) was significantly reduced compared with that observed for mdx mice (Fig. 2 B). We also measured hydroxyproline content, an index of muscle fibrosis, in diaphragm muscle homogenates from each of the four genotypes. While hydroxyproline content was markedly increased in diaphragm of mdx mice compared with that observed for both WT and Orai1 KO, postdevelopmental Orai1 ablation significantly reduced the increase in hydroxyproline content observed in mdx mice (Fig. 2 C). Finally, we evaluated the percentage of fibers from TA muscle that exhibit central nuclei, an index of muscle regeneration (Folker and Baylies, 2013). While virtually all muscle fibers from WT and Orai1 KO mice lacked centrally located nuclei, >80% of fibers from both mdx and mdx-Orai1 KO mice exhibited the presence of nuclei positioned at the center of the myofiber (Fig. S3). Thus, while postdevelopmental Orai1 ablation normalized muscle hypertrophy and hydroxyproline content in mdx mice, it did not mitigate a widely accepted biomarker of muscle regeneration (central nucleation).

SOCE is a mechanism involved in the refilling of intracellular Ca²⁺ stores during repetitive, high-frequency stimulation that requires the presence of functional Orai1 (Wei-Lapierre et al., 2013; Boncompagni et al., 2017, Boncompagni et al., 2018; Sztretye et al., 2017; Michelucci et al., 2019; Michelucci et al., 2020). Therefore, we measured total releasable Ca²⁺ store content under resting conditions (e.g., without electrical stimulation) in single FDB fibers loaded with a low-affinity ratiometric Ca²⁺ dye, fura-FF, during application of a Ca²⁺ store release cocktail (ICE; 10 μM ionomycin, 30 μM CPA, and 100 μM EGTA). Total releasable Ca²⁺ store content in FDB fibers from mdx mice was significantly reduced (∼40%) compared with that observed in FDB fibers from WT mice, while being partially restored in fibers from mdx-Orai1 KO mice (Fig. 3, A and B). No difference in total releasable Ca²⁺ store content was observed between fibers from WT and Orai1 KO mice. Consistent with enhanced constitutive Ca²⁺ entry (Fig. S2) and SOCE (Fig. 1, D and E) in muscle fibers from mdx mice, we also observed a modest, but statistically significant, increase in resting myoplasmic free Ca²⁺ concentration (133 ± 5.6 nM) in FDB fibers from mdx mice compared with fibers from WT mice (111 ± 2.8 nM), which was normalized following postdevelopmental ablation of Orai1 in mdx-Orai1 KO mice (109 ± 8.2 nM; Fig. 3 C).

Previous studies reported that muscle fibers from mdx mice are characterized by a significant impairment of excitation–contraction coupling (Woods et al., 2004; Woods et al., 2005; Hollingworth et al., 2008; DiFranco et al., 2008; Capote et al., 2010), the process by which an action potential in the surface membrane is used to trigger intracellular Ca²⁺ release from the SR terminal cisternae at the triad junction, ultimately leading to muscle contraction. Therefore, we quantified the amplitude and kinetics of electrically evoked Ca²⁺ transients in FDB fibers during twitch (0.5-Hz) stimulation (Fig. 4). Results from these experiments revealed that peak Ca²⁺ transient amplitude in fibers from mdx mice was significantly reduced compared with that of fibers from WT mice, while Ca²⁺ transient amplitude returned to normal levels in fibers from mdx-Orai1 KO mice (Fig. 4, A and B). We fitted the decay phase of the Ca²⁺ transient to a second-order exponential equation, where the fast component of decay (A_fast and τ_fast) primarily reflects Ca²⁺ binding to fast myoplasmic Ca²⁺ buffers, while the slow component of decay (A_slow and τ_slow) is dominated by SERCA-mediated SR Ca²⁺ reuptake (Baylor and Hollingworth, 2003; Capote et al., 2005; Carroll et al., 1999). Results from these analyses revealed that both τ_fast (Fig. 4 C) and τ_slow (Fig. 4 D) were significantly greater in fibers from mdx mice compared with fibers from WT mice, while τ_fast and τ_slow in fibers from mdx-Orai1 KO and Orai1 KO mice were not significantly different from those of WT mice (Fig. 4, C and D). Thus, postdevelopmental ablation of Orai1 in mdx mice normalized Ca²⁺ transient amplitude and decay kinetics to values similar to those observed in WT mice.

A marked reduction in the ability to generate force is a hallmark feature of DMD (Lynch et al., 2001). Therefore, we determined the effect of postdevelopmental Orai1 ablation on specific force production in intact EDL muscles. Excised EDL muscles of all genotypes were subjected to a force–frequency protocol with frequencies ranging from 1 to 250 Hz (Fig. 5 A). Results from these experiments revealed that absolute force (mN) was greater for EDL muscles from both mdx and mdx-Orai1 KO mice compared with that observed for EDL muscles from either WT or Orai1 KO mice (Fig. 5 B). This finding was consistent with the significantly larger muscle mass (Fig. 5 C) and physiological CSA (Fig. 5 D) of EDL muscles from 13–16-wk-old mdx mice. However, EDL muscles from mdx mice exhibited a reduction of peak specific force (mN/mm²) for all frequencies ≥50 Hz (Fig. 5 E). Specifically, peak EDL specific force at 200 Hz was 193.7 ± 8.6 mN/mm² for WT mice, but only 139.4 ± 3.1 mN/mm² for EDL muscles from mdx mice (Fig. 5 F). EDL muscles from mdx-Orai1 KO mice exhibited peak specific force at 200 Hz that was not significantly different from that of WT mice (174.6 ± 7.8 mN/mm²). These results were confirmed during application of a 2-s tetanus at 150 Hz, which showed that postdevelopmental ablation of Orai1 normalized the marked reduction in peak specific force observed in EDL muscles from mdx mice (Fig. S4). Similar results were observed in soleus muscles (Fig. S5, A and B).

Orai1 ablation leads to increased muscle fatigue during repetitive, high-frequency stimulation (Wei-Lapierre et al., 2013; Boncompagni et al., 2017; Michelucci et al., 2019, Michelucci et al., 2020). To directly assess the impact of postdevelopmental Orai1 ablation on muscle fatigue during prolonged activity, we subjected excised EDL muscles to a standard repetitive, high-frequency stimulation paradigm (40 stimulus trains of 50 Hz and 500 ms in duration delivered every 2.5 s). EDL muscles from WT mice exhibited an initial drop in specific force during the second stimulus train followed by a rebound increase (∼30%) in specific force production (Fig. S6, A and B) similar to that previously reported to be associated with SOCE activity (Michelucci et al., 2019; Michelucci et al., 2020). While EDL muscles from mdx mice exhibited reduced specific force production during the first 50-Hz stimulation (consistent with results in Fig. 5 E), EDL muscles from mdx mice nevertheless also exhibited an ∼30% rebound increase in specific force after the second stimulus (Fig. S6, A and B). Consistent with prior findings (Michelucci et al., 2019), Orai1 ablation abolished this rebound increase in specific force production after the second stimulus in both mdx-Orai1 KO and Orai1 KO mice.

Besides a reduction in isometric force generation, dystrophic muscles are also characterized by an increased susceptibility to eccentric contraction–induced damage (Dellorusso et al., 2001; Liu et al., 2005; Whitehead et al., 2008). To evaluate susceptibility to damage, a different set of EDL muscles were subjected to a previously validated ex vivo eccentric contraction damage protocol (Lindsay et al., 2020) consisting of 10 consecutive, 700-ms stretching tetani delivered at 150 Hz (500 ms of isometric contraction followed by a 200 ms 10% stretch of L_o at a speed of 0.5 L_o/s) applied every 120 s (Fig. 6 A). The ability to develop force following subsequent eccentric contractions declined modestly in EDL muscles from WT mice (61.8 ± 1.9% of initial specific force during the 10th eccentric contraction). In contrast, the eccentric contraction–induced force drop was considerably more pronounced in EDL muscles from mdx mice (35.5 ± 3.8% of initial specific force during the 10th eccentric contraction), consistent with a higher susceptibility to mechanical stress-induced damage (Fig. 6, B–D). Postdevelopmental ablation of Orai1 significantly reduced eccentric contraction–induced force drop in EDL muscles from mdx mice (46.1 ± 2.6% of residual force at the 10th eccentric contraction). Interestingly, postdevelopmental KO of Orai1 also significantly reduced eccentric contraction–induced force drop compared with that observed in EDL muscles from WT mice (72.2 ± 1.2% of initial force during the 10th eccentric contraction). Similar results were also obtained in soleus muscles (Fig. S5, C and D).

To determine if muscle damage is augmented by Orai1-dependent SOCE during each eccentric contraction, we performed experiments on intact EDL muscles from WT and mdx mice subjected to the same eccentric contraction protocol conducted in the presence of either control Ringer’s solution or Ringer’s solution supplemented with 10 μM BTP2. Results from these experiments revealed that eccentric contraction–induced damage was unaltered by acute block of SOCE with BTP2 in EDL muscles from either WT (Fig. S7 A) or mdx (Fig. S7 B) mice. Together, these studies suggest that chronic inhibition of Orai1-dependent SOCE is required to protect skeletal muscle from eccentric contraction–induced injury.

In this study, we report that enhanced Orai1-dependent Ca²⁺ entry contributes to the pathogenesis of DMD. Consistent with prior reports (Edwards et al., 2010; Zhao et al., 2012), skeletal muscle from mdx mice exhibits increased Orai1 transcript levels and enhanced constitutive Ca²⁺ entry and SOCE (Fig. 1 and Fig. S2), which correlated with (1) reduced total releasable Ca²⁺ store content and increased myoplasmic free Ca²⁺ concentration (Fig. 2); (2) increased myofiber CSA, fibrosis, and central nucleation (Fig. 3); (3) reduced peak electrically evoked Ca²⁺ transient amplitude and prolonged Ca²⁺ transient decay (Fig. 4); (4) reduced specific force production (Fig. 5), and (5) increased susceptibility to eccentric contraction–induced damage (Fig. 6). Collectively, this constellation of functional and histopathological alterations represents the dystrophic phenotype of mdx mice.

Importantly, we found that postdevelopmental, muscle-specific KO of Orai1 significantly ameliorates the dystrophic phenotype of young adult (3–4-mo-old) mdx mice. Specifically, ablation of Orai1-dependent Ca²⁺ entry in adult mdx-Orai1 KO mice partially restores total releasable Ca²⁺ store content and normalizes the resting myoplasmic free Ca²⁺ concentration to levels similar to that of WT. Furthermore, both the peak amplitude and slowed decay phase of electrically evoked Ca²⁺ transients in single FDB fibers, as well as peak specific force production in excised EDL and soleus muscles, are significantly improved. However, consistent with that reported previously in WT mice (Michelucci et al., 2019), postdevelopmental ablation of Orai1 in mdx mice also results in a reduced ability to maintain force production during repetitive high-frequency stimulation (Fig. S6).

Finally, postdevelopmental Orai1 KO also reduces myofiber hypertrophy and fibrosis, as well as protects muscles from eccentric contraction–induced muscle damage. However, Orai1 KO does not alter the percentage of centrally nucleated fibers, consistent with ongoing muscle regeneration even in the absence of Orai1 expression. Overall, these results extend prior findings (Goonasekera et al., 2014) by demonstrating that postdevelopmental functional Orai1 deficiency reduces the severity of muscular dystrophy in mdx mice through normalization of both Ca²⁺ homeostasis and sarcolemmal integrity.

A long-standing observation in the field is that Ca²⁺ homeostasis is disrupted in muscular dystrophy. Several interrelated mechanisms have been proposed to contribute to altered Ca²⁺ homeostasis in muscular dystrophy including (1) enhanced trans-sarcolemmal Ca²⁺ entry (Michelucci et al., 2018; Zabłocka et al., 2021; Mareedu et al., 2021), (2) increased RyR1 Ca²⁺ leak (Bellinger et al., 2009; Andersson et al., 2012), (3) impaired SR Ca²⁺ reuptake (Viner et al., 1996; Sharov et al., 2006; Dremina et al., 2007), (4) increased oxidative stress (Whitehead et al., 2008; Petrillo et al., 2017), and (5) mitochondrial dysfunction (Mareedu et al., 2021; Budzinska et al., 2021). Indeed, it is likely that multiple mechanisms work in concert, and possibly at different levels in different muscles, to contribute to altered Ca²⁺ homeostasis in muscular dystrophy.

Although several areas of controversy remain, there is a general consensus that Ca²⁺ homeostatic mechanisms are altered and intracellular free Ca²⁺ concentration is modestly increased in dystrophic muscles. Different mechanisms are proposed to contribute to altered Ca²⁺ homeostasis including excessive Ca²⁺ entry (e.g., microtears, mechanosensitive channels, TrpC channels, and SOCE channels). Prior studies demonstrated a clear relationship between increases in sarcolemmal Ca²⁺ entry and resting intracellular Ca²⁺ concentration in muscle fibers from mdx mice and other genetic models of muscular dystrophy (Yeung et al., 2005; Altamirano et al., 2012). In one of the most comprehensive studies in this regard, Altamirano et al. (2012) used double-barrel Ca²⁺ electrode measurements to quantify subsarcolemmal free Ca²⁺ concentration in fibers from WT and mdx mice under different conditions designed to inhibit extracellular Ca²⁺ entry (e.g., low Ca²⁺, 0 Ca²⁺, 20 μM Gd³⁺, 0 Ca²⁺ + 20 μM Gd³⁺). Consistent with prior findings (Mallouk et al., 2000), this study confirmed that subsarcolemmal Ca²⁺ is elevated in muscle fibers from mdx mice and that this increase is at least partially reduced by removing extracellular Ca²⁺ and/or blocking Ca²⁺ entry channels with Gd³⁺. Our findings further indicate that this increased constitutive Ca²⁺ entry observed in fibers from mdx mice is blocked by BTP2 (Fig. S2).

By altering net trans-sarcolemmal Ca²⁺ influx/efflux, enhanced Ca²⁺ entry via an Orai1-dependent pathway could contribute to the increase in steady-state myoplasmic free Ca²⁺ concentration observed in fibers from mdx mice, consistent with the cell boundary theorem (Friel and Tsien, 1992; Ríos, 2010). The cell boundary theorem states that a change in steady-state resting free Ca²⁺ concentration requires a net change in Ca²⁺ influx/efflux across the surface membrane. We found that muscle fibers from mdx mice exhibit a constitutive BTP2-sensitive Ca²⁺ influx. Assuming that Ca²⁺ efflux mechanisms are unaltered in fibers from mdx mice as suggested previously (Cully et al., 2012), an increase in net Ca²⁺ influx would be expected to result in an increase in steady-state resting free Ca²⁺ concentration. A limitation of our study is that we are not able to provide a more quantitative description, since we did not directly measure net Ca²⁺ influx (only Mn²⁺ influx) and efflux rates in fibers from WT and mdx mice.

In addition to increases in both trans-sarcolemmal Ca²⁺ influx and steady-state resting Ca²⁺ levels, acute or dynamic changes in intracellular Ca²⁺ concentration also depend on (1) myoplasmic Ca²⁺ binding proteins (e.g., parvalbumin), (2) SR Ca²⁺ reuptake (e.g., SERCA), and (3) SR Ca²⁺ leak/release via RyR1. Interestingly, a reduction of SERCA activity (Kargacin and Kargacin, 1996; Divet and Huchet-Cadiou, 2002; Divet et al., 2005; see also Fig. 4) is observed in skeletal muscle of mdx mice. Impaired SR Ca²⁺ reuptake due to reduced SERCA expression and/or activity could at least in part explain the reduction in total releasable Ca²⁺ store content (Fig. 3 B) and slowed Ca²⁺ transient decline (Fig. 4, C and D) observed in FDB fibers from mdx mice. The slower rate of Ca²⁺ transient decline in fibers from mdx mice is in line with results reported previously. For instance, Goonasekera et al. (2014) reported a similar slowed rate of Ca²⁺ transient decay in FDB fibers from δ-sarcoglycan–null (Sgcd^−/−) mice (Goonasekera et al., 2014). In addition, prior studies reported a significant reduction in SERCA function in EDL muscle of mdx mice (Kargacin and Kargacin, 1996; Divet and Huchet-Cadiou, 2002; Divet et al., 2005; Gehrig et al., 2012). In line with this idea, muscle-specific overexpression of SERCA1 restores both peak Ca²⁺ transient amplitude and decay kinetics, as well as dramatically attenuates the dystrophic phenotype in two different muscular dystrophy mouse models (mdx and Sgcd^−/− mice; Goonasekera et al., 2011).

Importantly, we found that postdevelopmental ablation of Orai1 similarly restores key functional measures of SERCA activity in FDB fibers including total releasable Ca²⁺ store content (Fig. 3) and Ca²⁺ transient amplitude/decay (Fig. 4). While the changes in the rate of Ca²⁺ transient decay observed in mdx mice and restoration after Orai1 KO could be explained by deceased SERCA1 expression in muscle of mdx mice that is normalized following Orai1 ablation, we did not find any statistically significant differences in SERCA1 expression across any of the four genotypes (Fig. S8 B). Thus, the precise mechanism by which Orai1 ablation normalizes these functional measures of SERCA activity without altering SERCA expression remains unclear. As one possibility, Orai1 ablation could reduce the inhibitory effects of oxidative/nitrosative modifications on SERCA function reported previously (Viner et al., 1996; Sharov et al., 2006; Dremina et al., 2007; Qaisar et al., 2021). Another possible explanation is that loss of Orai1 could either reduce expression of an endogenous SERCA inhibitor (e.g., sarcolipin or myoregulin; Schneider et al., 2013; Chambers et al., 2022) or increase expression of an endogenous SERCA activator (e.g., DWORF; Nelson et al., 2016). Further studies will be required to evaluate the role of these potential mechanisms.

The reduction in total releasable Ca²⁺ store content likely contributes to the reduction in the amplitude of electrically evoked Ca²⁺ release observed in our study and also reported by others (Head, 2010; Hollingworth et al., 2008; Woods et al., 2004). In addition to the contribution of SERCA pumps, total releasable Ca²⁺ store content also depends on the Ca²⁺ buffering capacity of the SR, which in fast-twitch muscle is largely dependent on the expression of CSQ1 (Michelucci et al., 2020). While some studies reported a significant decrease in CSQ content in young (∼8-wk-old) mdx muscles (Doran et al., 2004; Pertille et al., 2010), this is not typically observed at the older age (13–16 wk old) used in this study (Culligan et al., 2002; Ferretti et al., 2009; Matsumura et al., 2013; Barros Maranhão et al., 2015). Consistent with this, we did not find significant differences in CSQ1 expression in Western blot analyses of EDL muscle homogenates from 3–4-mo-old mice across all four genotypes (Fig. S8). However, the potential impact of postdevelopmental Orai1 ablation on the expression level of other lower-abundance SR Ca²⁺ binding proteins in muscle (e.g., histidine-rich Ca²⁺ binding protein, junctate, sarcolumenin, and CSQ-like binding proteins; Rossi and Dirksen, 2006; Culligan et al., 2002) remains to be determined.

In addition to decreased SERCA activity, the observed reduction in releasable Ca²⁺ store content could be due in part to increased RyR1 Ca²⁺ leak. Bellinger et al. (2009) demonstrated that RyR1 isolated from mdx skeletal muscle exhibits posttranslational modifications (e.g., RyR1-nitrosylation) that result in reduced binding of the regulatory protein FKBP12, with a consequent increase of RyR1 opening and Ca²⁺ leak. The leak of Ca²⁺ through oxidized RyR1 channels (together with reduced SERCA activity and potentially reduced SR Ca²⁺ buffering) would be expected to lead to a reduction in releasable SR Ca²⁺ store content. Consistent with this, treatment for 6 wk with an antioxidant (N-acetylcysteine) significantly reduced the dystrophic phenotype of mdx mice (Whitehead et al., 2008). A similar pathogenic role for altered Ca²⁺ handling and excessive ROS production was demonstrated for other muscle disorders (malignant hyperthermia and central core disease) characterized by enhanced nitrosative/oxidative stress, RyR1 nitrosylation, SR Ca²⁺ leak, and altered Ca²⁺ homeostasis (Durham et al., 2008; Lanner et al., 2012; Michelucci et al., 2015; Michelucci et al., 2017a; Michelucci et al., 2017b; Michelucci et al., 2021). However, whether functionally relevant ROS/RNS-dependent posttranslational modifications to proteins of the SOCE machinery (STIM1 and Orai1) occur in dystrophic muscle is unknown and certainly warrants future investigation.

An unexpected finding of this study was the protective effect of postdevelopmental Orai1 ablation on the susceptibility of muscle to eccentric contraction–induced damage. Specifically, Orai1 KO in EDL and soleus muscles in both WT (Orai1-KO) and mdx (mdx-Orai1 KO) mice exhibit a significantly reduced force drop following a series of eccentric contractions. Interestingly, this reduction in muscle damage was independent of acute Orai1-dependent Ca²⁺ entry, since the eccentric contraction–induced force drop was unaffected by exposure to 10 μM BTP2, a validated inhibitor of SOCE in muscle (Wei-LaPierre et al., 2022). These results suggest that the enhanced protection from eccentric contraction–induced damage does not result from acute Orai1-dependent Ca²⁺ entry, but rather from long-term effects of the loss of Orai1-dependent Ca²⁺ entry (e.g., reduced activation of calpains or other Ca²⁺ dependent processes). Indeed, uncontrolled protein degradation promoted by Ca²⁺-activated calpains is recognized as a key pathophysiological aspect of muscular dystrophy (Spencer et al., 1995; Tidball and Spencer, 2002; Spencer and Mellgren, 2002). Alternatively, a potential scaffolding role of the Orai1 complex in membrane integrity cannot be excluded.

Zhao et al. (2012) reported that chronic (2 wk) in vivo inhibition of SOCE with BTP2 significantly reduces both the rate of proteolytic events mediated by calpains and the progression of the dystrophic phenotype of mdx mice. In an analogous manner, early developmental (or constitutive) inhibition of SOCE in muscle following muscle-specific expression of dominant-negative Orai1 (dnOrai1) also mitigated the dystrophic phenotype in both mdx mice and Sgcd^−/− mice (Goonasekera et al., 2014). However, BTP2 and dnOrai1 inhibit both Orai1 and TrpC channels, and constitutive muscle-specific expression of dnOrai1 results in several additional important developmental effects in muscle (e.g., reduced type 1 fiber content and CSA; Wei-Lapierre et al., 2013). Our results show that specifically targeting Orai1 in fully developed muscle of adult mice is sufficient to mitigate the dystrophic phenotype of mdx mice. These results provide evidence that inhibition of Orai1 in DMD patients may provide some level of protection or slow disease progression even after disease onset. However, the important role of Orai1 in mitigating muscle fatigue during repetitive stimulation (Michelucci et al., 2019; Fig. S6) may limit the therapeutic benefit of inhibiting Orai1 function in individuals with DMD.

We also investigated if the protective effect of postdevelopmental Orai1 ablation could be explained by compensatory changes in the expression of utrophin, a structural and functional paralogue of dystrophin (Ibraghimov-Beskrovnaya et al., 1992). Utrophin localizes at the neuromuscular junction of normal adult fibers (Love et al., 1989; Tinsley et al., 1992), while also being abundantly present throughout the sarcolemma during muscle development and damage-induced muscle regeneration (Ohlendieck et al., 1991; Amenta et al., 2011). Utrophin is also upregulated and localized along the sarcolemma in regenerating fibers of patients with muscular dystrophy (Helliwell et al., 1992), and transgenic overexpression of utrophin suppresses the dystrophic phenotype of mdx mice (Tinsley et al., 1998; Fisher et al., 2001). However, our results indicate that the reduced eccentric contraction–induced force drop observed following postdevelopmental Orai1 ablation occurs in the absence of a detectable change in utrophin expression (Fig. S8).

The findings reported in this study provide new insights regarding the role of Orai1 in DMD pathogenesis. Collectively, our results support the hypothesis that increased Orai1-dependent SOCE potentiates a cascade of downstream pathways that ultimately culminate in increased myofiber fragility and death. An important implication of these studies is that mitigation of the dystrophic phenotype can be achieved by targeting these Orai1-dependent pathways even after muscle development is complete. Nevertheless, several unresolved issues remain to be addressed. For instance, whether other Orai isoforms (Orai2 and Orai3) are involved in the DMD phenotype remains an open and unsolved issue. While we did not directly assess potential compensatory changes in Orai2 and/or Orai3 expression in muscle of Orai1 KO and mdx-Orai1 KO mice, the lack of detectable store-dependent Mn²⁺ quench in fibers from Orai1 KO and mdx-Orai1 KO mice (Fig. 1 E) argues against a compensatory upregulation of alternative Orai1-independent Ca²⁺ influx mechanisms. However, a comprehensive assessment of this possibility will require further study. Finally, future studies are needed to elucidate the precise downstream mechanisms responsible for how enhanced Orai1-mediated SOCE potentiates the dystrophic phenotype, as well as to determine if therapeutic strategies designed to target these mechanisms can be effectively translated to the treatment of children with DMD.

Eduardo Ríos served as editor.

We thank Dr. Victor Yarotskyy for initially handling the mouse colony and developing the breeding scheme. We thank Linda Groom for support with mouse genotyping.

This work was supported by grants from National Institutes of Health (AR059646 and NS097868 to R.T.Dirksen).

The authors declare no competing financial interests.

Author contributions: M. García-Castañeda designed and performed experiments, analyzed and interpreted data, and wrote, edited, and revised the manuscript. A. Michelucci designed and performed experiments, analyzed and interpreted data, and wrote, edited, and revised the manuscript. N. Zhao designed and performed experiments and analyzed and interpreted data. S. Malik designed and performed experiments and analyzed and interpreted data. R.T. Dirksen conceived and directed the study, interpreted data, provided resources, and wrote, edited, and revised the manuscript.