Functional characterization of orbicularis oculi and extraocular muscles

Excitation–contraction coupling (ECC) is the process whereby an electrical signal, depolarization of the plasma membrane, is converted into a chemical signal, Ca²⁺ release from the SR, leading to muscle contraction (Caputo, 2011). ECC relies on the function of two main Ca²⁺ channels, the voltage-sensing dihydropyridine receptor (DHPR), an L-type Ca²⁺ channel present on the transverse tubules, and the ryanodine receptor (RyR) Ca²⁺ channel, present on the SR terminal cisternae. In mammalian skeletal muscle, ECC is mechanically coupled, that is membrane depolarization causes the α1 subunit (Ca_v1.1) of the DHPR to undergo a conformational change, causing it to come in direct contact with the RyR1 and leading to release of Ca²⁺ from the SR (Ríos and Pizarro, 1991). In mammalian cardiac muscles, in contrast, ECC does not rely on mechanical coupling, but rather influx of Ca²⁺ through the cardiac α1 subunit of the DHPR (Ca_v1.2) activates the cardiac RyR2, leading to release of Ca²⁺ from the SR (Bers, 2002). Thus, the functional requirements of heart and skeletal muscle are assured by the exquisite specificity of protein isoform expression. This general configuration of skeletal and cardiac ECC protein expression was thought to underlie the function of most striated muscles; however, in a recent study on the protein composition of human extraocular muscles (EOMs), we found that hybrid skeletal/cardiac muscle configurations of the ECC machinery can exist. Indeed EOMs express both the skeletal and cardiac isoforms of the α1 subunit of the DHPR, rely on influx of Ca²⁺ from the extracellular environment, and express not only RyR1 but also RyR3 (Sekulic-Jablanovic et al., 2015).

EOMs, the fastest muscles in the body, derive from somitomeres (preotic mesodermal segments; Wright et al., 2006). There are six EOMs distributed in three antagonistic pairs of muscles that finely control eye movements (Spencer and Porter, 1988); they are innervated by three cranial nerves (III, IV, and VI; Sadeh and Stern, 1984). EOMs are highly specialized and can be either singly innervated or multiply innervated. These characteristics together with the unique expression of myosin heavy chain–EO (MyHC13; Schiaffino and Reggiani, 2011) set them apart from all other skeletal muscles, as a distinctive group of highly specialized muscles (Spencer and Porter, 1988). In contrast, the orbicularis oculi muscles comprise the sphincter muscles of the eyelids; they are classified as facial muscles and functionally antagonize the levator palpebrae superioris muscles, which are accessory EOMs (Porter et al., 1998). The orbicularis oculi muscles can be subdivided into orbital, palpebral, and lacrimal portions. The orbital portion firmly closes the eyelids and is controlled by voluntary action; the palpebral portion closes the eyelids gently in involuntary or reflex blinking; the palpebral portion can be further divided into pretarsal, preseptal portion, and ciliary. The lacrimal portion compresses the lacrimal sac, which receives tears from the lacrimal ducts and conveys them into the nasolacrimal duct (Gray and Lewis, 1918). The orbicularis oculi muscles derive from the mesenchyme in the second pharyngeal arch (Moore et al., 2015) and are innervated by the VII cranial nerve (Ouattara et al., 2004).

Weakness of the orbicularis oculi muscles is often apparent in neuromuscular disorders affecting the neuromuscular junction (Miller et al., 2008). Similar to EOMs, they are also one of the first targets of mitochondrial myopathies and myasthenia gravis, resulting in weakness of eyelid closure and ptosis (Miller et al., 2008). Facial weakness and ptosis have also been described in patients with recessive RYR1 mutations (the gene encoding the RyR1; Taylor et al., 2012), affected by multiminicore disease, congenital fiber type disproportion, and centronuclear disease (Jungbluth et al., 2005; Treves et al., 2008; Clarke et al., 2010; Wilmshurst et al., 2010). In contrast, EOMs are spared in aging, Duchenne muscular dystrophy, and congenital muscular dystrophy when all other skeletal muscles are affected (Kaminski et al., 1992; Khurana et al., 1995; Kallestad et al., 2011). Weakness of the facial musculature caused by any disease is almost certainly accompanied by weakness of the orbicularis oculi muscles as well (Miller et al., 2008). The present investigation was undertaken to identify similarities and differences in ECC and calcium homeostasis in human EOMs, orbicularis oculi, and quadriceps, to identify factors that might contribute to the selective involvement in different neuromuscular disorders. The results of the present investigation show that the ECC machinery and calcium regulation of human orbicularis oculi are more similar to those of quadriceps than of EOMs; in contrast, orbicularis oculi and EOMs are similar in that they both express high levels of utrophin, whereas quadriceps expresses dystrophin. This finding explains the sparing of facial and EOMs in Duchenne patients; because the ECC machinery between orbicularis oculi and quadriceps appears to be similar, our study points to a potential use of autologous facial muscle–derived satellite cells for muscle reimplantation in patients with muscular dystrophy.

Primary muscle cell cultures were established as previously described (Censier et al., 1998; Ducreux et al., 2004) from fragments of muscle biopsies from patients aged between 18 and 65 yr. Quadriceps muscle biopsies were obtained from healthy males or females (six donors) undergoing in vitro diagnostic procedures, orbicularis oculi muscle biopsies were from preseptal muscles of patients (≈75% females) undergoing eyelid cosmetic procedure or blepharoplasty (five donors), and EOM biopsies were from lateral or medial rector muscles of patients undergoing squint corrective surgery (four donors). Cells were plated on laminin-coated glass coverslips and allowed to grow and differentiate as previously described (Treves et al., 2011; Sekulic-Jablanovic et al., 2015). This research was performed in accordance with the Declaration of Helsinki (2013) of the World Medical Association and was approved by the Ethikkommission beider Basel (permit N° EK64/12); all subjects gave written informed consent to carry out this work.

Fura-2/AM (EMD Millipore) or fluo-4/AM (Thermo Fisher Scientific) at a final concentration of 5 µM was used to load myotubes for 30 min at 37°C, after which the coverslips were mounted onto a 37°C thermostatically controlled chamber that was continuously perfused with Krebs-Ringer medium; individual cells were stimulated by means of a 12- or 8-way 100-mm-diameter quartz micromanifold computer-controlled microperfuser (ALA Scientific Instruments), as described previously (Ducreux et al., 2004). The ratiometric Ca²⁺ indicator fura-2 was used for monitoring the global changes in the cytoplasmic Ca²⁺ concentration as previously described (Censier et al., 1998; Ducreux et al., 2004). Ratiometric images (340/380 nm) were acquired every 1 s, and the camera was set at a fixed exposure time of 100 ms. Fura-2 fluorescence ratio signals were converted into [Ca²⁺] using the curve generated using the fura-2 Ca²⁺ imaging calibration kit from Molecular Probes as previously described (Vukcevic et al., 2013). The total amount of rapidly releasable Ca²⁺ in the stores was determined as previously described (Rokach et al., 2013) by calculating the area under the curve of the transient induced by the application of 1 µM ionomycin, plus 1 µM thapsigargin in Krebs-Ringer containing 0.5 mM EGTA.

The dynamics of depolarization-induced Ca²⁺ influx were investigated by total internal reflection fluorescence (TIRF) microscopy using the single wavelength Ca²⁺ indicator fluo-4 as detailed previously (Treves et al., 2011; Ullrich et al., 2011). Myotubes were pretreated with 50 µM ryanodine (EMD Millipore) to block Ca²⁺ release through the RyR1 receptor (Treves et al., 2011), and depolarization-induced Ca²⁺ entry was stimulated by the application of 60 mM KCl. Online fluorescence images were acquired every 100 ms using an inverted Nikon TE2000 TIRF microscope equipped with an oil immersion CFI Plan Apochromat 60× TIRF objective (1.49 NA) and an electron multiplier Hamamatsu CCD camera (C9100-13). The MetaMorph imaging software from Molecular Devices was used for analysis of the fluorescence changes.

Total RNA was extracted from muscle biopsies using TRIzol (Invitrogen) according to the manufacturer’s instructions. RNA was first treated with deoxyribonuclease I (Invitrogen), and then 1,000 ng was reverse transcribed using the high-capacity cDNA reverse transcription kit (Applied Biosystems). cDNA was amplified by qPCR using SYBR Green technology (Fast SYBR Green Master Mix; Applied Biosystems) as previously described (Rokach et al., 2013). The sequences of the primers used for gene amplification and quantification are given in Table S1. qPCR was performed on a 7500 Fast Real-Time PCR machine (Applied Biosystems) using the 7500 software v2.3 as previously described (Sekulic-Jablanovic et al., 2015). Gene expression was normalized to expression of ACTN2, which is present in all muscle fiber types. Results are expressed as fold change compared with expression of the gene in quadriceps muscles.

Differentiated human myotubes grown on a glass coverslip coated with laminin were fixed with 4% paraformaldehyde (made in PBS), permeabilized with 1% Triton in PBS for 20 min, and processed as previously described (Treves et al., 2011). The following antibodies were used: mouse anti-RyR1 (MA3-925; Thermo Fisher Scientific), goat anti-Ca_v1.1 (sc-8160; Santa Cruz Biotechnology, Inc.), rabbit anti-Ca_v1.2 (sc-25686; Santa Cruz Biotechnology, Inc.), mouse anti-dystrophin (ab-7164; Abcam), rabbit anti-utrophin (sc-15377; Santa Cruz Biotechnology, Inc.), Alexa Fluor 488–conjugated chicken anti–rabbit, Alexa Fluor 555–conjugated donkey anti–goat IgG (Thermo Fisher Scientific), and Alexa Fluor 647–conjugated goat anti–mouse IgG (Thermo Fisher Scientific). Cells were stained with 4′, 6-diamidino-2-phenylindole (DAPI; Thermo Fisher Scientific) to visualize nuclei and observed using an A1R confocal microscope (Nikon) with a CFI Apo TIRF 100× (1.49 NA) objective.

The total SR fraction was isolated from flash-frozen muscle samples (human orbicularis oculi, EOM, quadriceps muscles, and mouse heart) as previously described (Anderson et al., 2003) and stored in liquid nitrogen. The protein concentration was determined using the Protein Assay kit II (Bio-Rad Laboratories) and BSA as a standard. SDS-PAGE, protein transfer on to nitrocellulose membranes, and immunostaining were performed as previously described (Anderson et al., 2003). The following primary antibodies were used: mouse anti-RyR1 (MA3-925; Thermo Fisher Scientific), goat anti-Ca_v1.1 (sc-8160; Santa Cruz Biotechnology, Inc.), rabbit anti-Ca_v1.2 (sc-25686; Santa Cruz Biotechnology, Inc.), rabbit anti–calsequestrin 1 (C-0743; Sigma-Aldrich), anti–calsequestrin 2 (3516; Abcam), goat anti-SERCA1 (sc-8093; Santa Cruz Biotechnology, Inc.), mouse anti-sarcalumenin (SRL; MA3-932; Thermo Fisher Scientific), anti-dystrophin (ab-7164; Abcam), anti-utrophin (sc-15377; Santa Cruz Biotechnology, Inc.), and mouse anti–myosin heavy chain (05-716; EMD Millipore). Secondary peroxidase conjugates were protein G–peroxidase (P8170; Sigma-Aldrich) and peroxidase-conjugated goat anti–mouse IgG (A2304; Sigma-Aldrich). The immunopositive bands were visualized by chemiluminescence using the Super Signal West Dura kit (Thermo Fisher Scientific) or the Chemiluminescence kit (Roche).

For myosin heavy chain, gels were performed as described by Talmadge and Roy (1993) except that 30 µg total protein extracts were separated and the final acrylamide concentration was 6.5%. Gels were stained with Coomassie brilliant blue.

Statistical analysis was performed using the Student’s t test for two populations. Values were considered significant when P < 0.05. When more than two groups were compared, analysis was performed by the ANOVA test followed by the Bonferroni post hoc test, using Prism 4.0 software (GraphPad Software). OriginPro 8.6 software (OriginLab) was used for generating dose–response curves.

Orbicularis oculi–derived myotubes were patch clamped in the whole cell configuration and stimulated to different depolarizing potentials to elicit SR Ca²⁺ release as previously reported using a EPC10 amplifier (HEKA) controlled by the data acquisition software PatchMaster (Rokach et al., 2013). External recording solution contained (mM) 130 TEA-CH₃SO₃, 2 CaCl₂, 1 MgCl₂, 10 HEPES, 1 µM tetrodotoxin, and 1 4-aminopyridine, pH 7.4 (CsOH). Internal pipette solution contained (mM) 100 Cs-CH₃SO₃, 10 HEPES, 6 MgCl₂, 11.5 CaCl₂, 4 Na₂ATP, 10 EGTA, and 0.1 K₅-fluo-3, pH 7.2 (CsOH). Solution exchange was controlled by an electrically controlled rapid switch microperfuser (VC8; Ala Scientific Instruments). Starting from a holding potential of −80 mV, the voltage clamp protocol consisted of consecutive depolarizing 500-ms step pulses to increasing potentials ranging from −50 to 50 mV with an increment of 10 mV. In parallel, changes in cytosolic Ca²⁺ in clamped cells were recorded by imaging of the fluorescent Ca²⁺-sensitive indicator fluo-3 using a confocal microscope in the line scan mode (IX81, FluoView FV1000, 60× water objective, NA 1.2; Olympus). Excitation of fluo-3 was achieved at 473 nm, and emission was collected between 510 and 560 nm. Line scans were acquired at a rate of 5.6 ms per line. To acquire multiple line scans from each cell, the duration of the single line scan image was limited to 1,000 lines to avoid photobleaching and cell damage. Experiments were performed at constant room temperature. Images were analyzed using ImageJ (National Institutes of Health) and further processed together with current analysis in OriginPro 2015 software.

Sparks were measured in fluo-4–loaded myotubes using an A1R laser-scanning confocal microscope (Nikon) with a 60× oil immersion Plan Apochromat VC objective (Nikon; NA 1.4) as previously described (Lopez et al., 2015). 5-s duration line scan images (x,t) were acquired in resonant mode at 7,680 lps with 512 pixels (0.05 µm/pixel) in the x and 39,936 pixels (0.126 ms/pixel) in the t direction using a pinhole size of 72.27 µm. Four to five images were taken at different positions across each cell. The Ca²⁺ indicator was excited with a laser at 487 nm, and the fluorescence emitted at 525 ± 25 nm was recorded. To minimize photo damage, the laser intensity was set at 3∼4% of the maximal power. Cells were perfused with Krebs-Ringer containing 2 mM Ca²⁺.

Fig. S1 shows the voltage dependence of Ca²⁺ release in orbicularis oculi–derived myotubes. Fig. S2 shows that human myotubes do not show spontaneous Ca²⁺ release events (SPARKS). Table S1 lists the sequence of primers used for qPCR.

The expression levels of the major gene products involved in skeletal ECC are shown in Fig. 1 A; the values obtained from orbicularis oculi were compared with those obtained from quadriceps of healthy donors. The latter muscles were used as reference, and the expression level of different genes therein was set to 1. The results show the mean (±SEM) expression level of genes from five pooled biopsy samples normalized to ACTN2. The expression of RYR1 and SERCA1 transcripts was slightly elevated (approximately twofold; P < 0.05, Student’s t test), as was that of CASQ1, RYR3, and CACNA1C (4-, 5-, and 10-fold, respectively; P < 0.0001, Student’s t test), whereas CACNA1S expression was similar in orbicularis oculi and quadriceps muscles. Interestingly, the expression levels of UTRN and DMD showed an ∼13- and 16-fold increase, respectively, compared with quadriceps muscles (P < 0.0001, Student’s t test). Although it was reported that the MYH13 isoform is exclusively expressed in EOMs (Wieczorek et al., 1985), we also found that it is expressed in orbicularis oculi (Fig. 1 B). JP45 levels remained unchanged.

Total SR fractions were prepared and probed with antibodies against the major skeletal ECC proteins as well as Ca_v1.2 and CASQ2; the intensities of the immunoreactive bands were normalized to SRL content and are expressed as percent expression of that of quadriceps muscles. Surprisingly, at the protein level, the content of RyR1, Ca_v1.1, SERCA1, and CASQ1 were not changed between orbicularis oculi and quadriceps (Fig. 1 C), whereas CASQ2 was significantly increased. We chose to normalize to SRL content and not to SERCA1 or CASQ1 because by qPCR both SERCA1 and CASQ1 transcripts were significantly increased in orbicularis oculi. Ca_v1.2, the cardiac isoform of the DHPR, could not be detected in Western blots of orbicularis oculi and quadriceps, but it was clearly present in human EOM and mouse heart, which served as positive control (Fig. 1 D). Utrophin was highly expressed in both EOM and orbicularis oculi but less so in quadriceps (fourfold in orbicularis oculi vs. quadriceps; P < 0.05), whereas the opposite was true for dystrophin (orbicularis oculi expressed ∼50% compared with quadriceps; P < 0.05; Fig. 1 E).

Because in a previous study we showed that primary cultures of myotubes explanted from human EOM biopsies had different Ca²⁺ handling properties (Sekulic-Jablanovic et al., 2015), in the next series of experiments we compared Ca²⁺ homeostasis in myotubes derived from the three types of muscles. In this context, it should also be mentioned that human myotubes are different from mouse myotubes or fibers in that they do not contract even after exposure to high concentrations of KCl, 4-chloro-m-cresol (4-cmc) or ionomycin plus thapsigargin. In the first set of experiments, cells were perfused with Krebs-Ringer containing 100 µM La³⁺ to prevent influx of extracellular Ca²⁺ and to ensure that the measurements represent skeletal ECC. Fig. 2 A shows representative fura-2 traces of a quadriceps-derived myotube (continuous line), orbicularis oculi–derived myotube (dashed line), and EOM-derived myotube (dotted line) in response to 60 mM KCl. Fig. 2 (B and C) shows the KCl- and 4-cmc–induced Ca²⁺ release curves, respectively. There were no significant differences between the three types of myotubes as far as peak Ca²⁺ release elicited by either KCl or 4-cmc is concerned. Furthermore, the EC₅₀ for KCl (depolarization)-induced Ca²⁺ release was similar in the three types of myotubes, whereas the EC₅₀ for 4-cmc–induced Ca²⁺ release was significantly different only between orbicularis oculi– and EOM-derived myotubes (Table 1). We also verified the voltage dependence of Ca²⁺ release of orbicularis oculi–derived myotubes; the half-maximal release activation V_1/2 was −25 ± 5 mV (Fig. S1), a value similar to that previously reported in quadriceps-derived myotubes (Rokach et al., 2013), confirming that the ECC is very similar in both kinds of myotubes. Interestingly, however, there were differences in the mean global resting [Ca²⁺]_i in the three populations of myotubes, though we could not ascertain whether this was caused by large differences within cytoplasmic micro/nanodomains or by a uniform difference resulting from the presence, for example, of different cytoplasmic calcium buffering proteins. The size of the intracellular Ca²⁺ stores of orbicularis oculi–derived myotubes was similar to those of quadriceps-derived myotubes but significantly lower than those of EOM-derived myotubes (Fig. 2, C and D; P < 0.0001, Student’s t test). We also examined the different types of myotubes for the presence of Ca²⁺ sparks by high-speed confocal microscopy, but none of the myotubes exhibited sparks (Fig. S2).

We next investigated the process by which membrane depolarization activates Ca²⁺ influx through the DHPR- or depolarization-induced Ca²⁺ influx (Bannister et al., 2009) by TIRF microscopy as previously described (Treves et al., 2011). Fig. 3 A shows pseudocolored fluo-4 fluorescent changes in a representative myotube pretreated with 50 µM ryanodine and stimulated with 60 mM KCl; the pretreatment with high concentrations of ryanodine is necessary to block Ca²⁺ release through RyR, ensuring that the change in fluo-4 fluorescence is not caused by Ca²⁺ release from the SR, but rather from Ca²⁺ influx. Fig. 3 B shows a representative Ca²⁺ influx trace initiated by the application of 60 mM KCl in orbicularis oculi–derived myotubes (solid trace) quadriceps-derived myotubes (dashed trace), and EOM-derived myotubes (dotted trace). A control experiment performed on EOM-derived myotubes showing that no Ca²⁺ influx occurs when the same experiment is performed in the presence of 100 µM La³⁺ is also shown (dash-dotted trace). Fig. 3 C summarizes the results confirming that depolarization-induced Ca²⁺ influx in orbicularis oculi myotubes is not significantly different from quadriceps myotubes, but almost threefold lower compared with EOM-derived myotubes.

The difference in Ca²⁺ influx between EOM and orbicularis oculi muscles most likely reflects the differential expression of Ca_v isoforms in these groups of muscles. This is confirmed by results depicted in Fig. 4 showing the subcellular distribution of Ca_v1.1, Ca_v1.2, and RyR1 in orbicularis oculi– and EOM-derived myotubes; the distribution of Ca_v1.1 (Fig. 4 A, green) and RyR1 (Fig. 4 B, red) was punctate and unstructured within an intracellular membrane compartment. Although they lack a mature organization, RyR1 and Ca_v1.1 codistributed within the myotube (Fig. 4 C). However, Ca_v1.2 could not be detected in orbicularis oculi–derived myotubes (Fig. 4 D). A similar subcellular punctate distribution of Ca_v1.1 (Fig. 4 E) and RyR1 (Fig. 4 F) was found in EOM-derived myotubes; however, in the latter cells, Ca_v1.2 was exclusively targeted to the plasma membrane (Fig. 4 H).

Because qPCR and Western blot analysis revealed that orbicularis oculi and EOM but not quadriceps muscle biopsies express high levels of utrophin, we investigated by confocal microscopy the expression and distribution of dystrophin and utrophin in the different kinds of myotubes. Quadriceps-derived myotubes (Fig. 5, A–D) showed punctate distribution of dystrophin, with some patches close to the plasma membrane (Fig. 5 B), but were negative for utrophin staining (Fig. 5 C). Orbicularis oculi–derived myotubes (Fig. 5, E–H) also showed a punctate distribution of dystrophin (Fig. 5 F); these myotubes were also positive for utrophin whose distribution was less homogeneous, had a more filamentous appearance, and was concentrated in areas around the cell periphery (Fig. 5 G). EOM-derived myotubes (Fig. 5, I–L) were similar to orbicularis oculi–derived myotubes in that dystrophin had a punctate distribution (Fig. 5 J), whereas utrophin was distributed less homogeneously, had a filamentous appearance, and was concentrated in areas around the cell periphery (Fig. 5 K). Interestingly, utrophin and dystrophin appeared to have a separate subcellular distribution that does not overlap within the myotubes (Fig. 5, H and L).

In the present study, we investigated the biochemical and physiological characteristics of orbicularis oculi muscles, a group of facial muscles that are selectively spared or involved in different neuromuscular disorders. As far as the expression of proteins involved in ECC is concerned, it appears that orbicularis oculi muscles are closer to quadriceps than to EOMs. Indeed, the content of skeletal muscle SR proteins was similar between quadriceps and orbicularis oculi muscles and differed from that of EOMs, as only the latter express proteins characteristic of both cardiac and skeletal muscle ECC (Sekulic-Jablanovic et al., 2015). Surprisingly, however, the expression of transcripts did not always match the actual protein content, highlighting the importance of validating arrays or qPCR experiments whenever possible. This was particularly relevant for the difference between the expression of Ca_v1.2 and dystrophin whose transcripts were increased >8–10-fold in orbicularis oculi, but at the protein level they were barely detectable. An intriguing result of this study is that although gene expression in orbicularis oculi is quite different from that in quadriceps, the expression of proteins involved in the ECC process (RyR1, Ca_v1.1, SERCA1, CASQ1, and Ca_v1.2) is similar in both cell types. In contrast, in our previous study, the differences in gene expression between EOM and quadriceps were positively correlated with differences in the expression of ECC proteins (Sekulic-Jablanovic et al., 2015). The reason for this discrepancy is at present unclear, but agreement between mRNA and protein content occurs only ∼40% of the time and is influenced by different factors, including mRNA and protein stability, the presence of microRNAs, and posttranscriptional modifications, with the key role in determining protein abundance being played at the level of translation (Tian et al., 2004; Vogel et al., 2010; Schwanhäusser et al., 2011). A limited overlap in genomic and proteomic data were reported by Khanna et al. (2003) in EOM and leg muscles and by Zhou et al. (2010) in mouse EOM myosin heavy chain isoform expression.

From a point of view of calcium homeostasis, it appears that orbicularis oculi muscle–derived myotubes are closer to those derived from quadriceps than EOM because the Ca²⁺ release dose–response curves, the voltage dependence of Ca²⁺ release, the resting [Ca²⁺], and ionomycin/thapsigargin-sensitive intracellular stores from the two muscle types were functionally indistinguishable and were clearly different from EOM-derived myotubes. The latter have a significantly higher EC₅₀ for 4-cmc–induced Ca²⁺ release, larger ionomycin/thapsigargin-sensitive intracellular stores, express Ca_v1.2, and exhibit a threefold larger depolarization-induced Ca²⁺ influx. The difference in sensitivity to 4-cmc–induced Ca²⁺ release but not KCl-induced Ca²⁺ release may reflect the different amounts of RyR1 and RyR3 expressed in EOM-derived myotubes. Such a hypothesis is compatible with previous findings (Fessenden et al., 2000; Matyash et al., 2002; Sekulic-Jablanovic et al., 2015) who showed that (a) EOM biopsies contain ∼100-times more RYR3 transcript than triceps biopsies, whereas orbicularis oculi express only ∼4-times more; (b) RyR3 have a significantly lower sensitivity to 4-cmc; and (c) in reconstituted 1B5 myotubes, RyR3 cannot form functional contacts with the DHPR and therefore would not contribute directly to the KCl-induced Ca²⁺ release.

One of the physiological characteristics of EOMs is that they are fatigue resistant (Fuchs and Binder, 1983), and this may be functionally related to the high levels of expression of Ca_v1.2 and consequent large Ca²⁺ influx. The Ca²⁺ influx may be used as a means (a) to activate ECC more rapidly by enhancing Ca²⁺ induced Ca²⁺ release or (b) to rapidly replenish intracellular stores. Depolarization-induced Ca²⁺ influx is less pronounced in myotubes derived from other skeletal muscles, most likely because they lack the cardiac isoform of the α1 DHPR subunit.

Although orbicularis oculi express a typical skeletal muscle ECC, they also show similarities with EOMs in that they express MyHC13 and high levels of RYR3. In humans, orbicularis oculi are composed of 90% type II fibers and 10% type I fibers (Wirtschafter et al., 1994; Campbell et al., 1999), but our results show that they also express MyH13. In a study on levator palpebrae and retractor bulbi muscles from different species, it was shown that the latter muscles also express MyHC13 and that this is probably responsible for their rapid contracture times (<10 ms); in fact, limb muscles do not express the superfast myosin isoforms, and their contracture time is approximately five times longer (Lucas and Hoh, 1997). As far as RYR3 expression is concerned, our results are enigmatic because, in general, RYR3 transcripts are expressed at low levels, if at all, in adult skeletal muscle (Martin et al., 1998), and the role of RyR3 is unclear. RYR3KO mice are viable, and their muscles show no obvious physiological differences compared with their wild-type littermates, including no changes in electrically induced Ca²⁺ release and contractile properties of adult muscle fibers, although skeletal muscles from neonatal RYR3 KO mice show decreased tension development (Takeshima et al., 1996). Although for the time being the function of RyR3s is speculative, it is possible that these channels act as amplifiers of the Ca²⁺ signals; compatibly, RYR3KO mice were reported to have a mild cognitive impairment when tested on a water maize (Balschun et al., 1999), but retrospectively, this may have also been related to a visual impairment. Studies aimed at understanding the role of RyR3 in EOMs are currently under way.

Our results concerning the expression of utrophin are interesting and most likely explain why in patients with Duchenne muscular dystrophy ocular and facial muscles are spared. Furthermore, the observation that EOMs are spared in Duchenne muscular dystrophy even though they exhibit a large Ca²⁺ influx provides strong evidence that controlled Ca²⁺ influx, per se, is not deleterious to skeletal muscles as recently proposed (Millay et al., 2009), but is actually a physiological mechanism used by some skeletal muscles. Utrophin and dystrophin share considerable sequence and structural similarity (Tinsley et al., 1992; Winder et al., 1995), and utrophin can associate with the dystrophin-associated complex, serving as a link between actin and the extracellular matrix (Matsumura et al., 1992). Interestingly, in cultured myotubes, the subcellular distribution of utrophin differs from that of dystrophin. Indeed, and as previously reported (Trimarchi et al., 2006) and confirmed in the present study, after 7–10 d of differentiation, dystrophin has a punctate membrane distribution, whereas utrophin has a patchy more filamentous distribution concentrated within specific subcellular domains. The reason for this difference is unclear but may reflect distinct binding partners of the two proteins. In mdx mice, it is believed that utrophin compensates for the lack of dystrophin (Wakefield et al., 2000). Based on our results and on the fact that mdx knocked out also for utrophin show EOM involvement (Baker et al., 2006), it appears that utrophin can functionally compensate in vivo for the lack of dystrophin, supporting the current strategies aimed at modulating utrophin expression in the therapy for Duchenne muscular dystrophy (Perkins and Davies, 2002; Chakkalakal et al., 2005; Guiraud et al., 2015).

Collectively, these studies show that subspecialization of skeletal muscles occurs through multiple factors; although it is true that muscle innervation plays a prominent role, the distribution of a specific muscle within a niche devoted to a precise physiological function is also important. Indeed, the term muscle allotype was proposed to describe the different capacities of myogenic cells of different lineages to express a different subset of myofibrillar genes. Because EOMs, orbicularis oculi, and levator palpebrae (and the retractor bulbi which is present in some mammals but not in humans) are derived from cells from a different lineage than those giving rise to limb muscles, their myogenic precursors must be programmed to express different subsets of proteins. The present study substantiates the validity of the muscle allotype hypothesis because we show that satellite cells derived from different muscles are primed and will follow the developmental characteristics of their muscle of origin, a property that can be exploited in laboratories devoted to tissue engineering.

We gratefully acknowledge the technical support of Anne-Sylvie Monnet.

This work was supported by the Swiss National Science Foundation (SNF; grant number 31003A-146198) and by the Department of Anesthesia Basel University Hospital.

The authors declare no competing financial interests.

Eduardo Ríos served as editor.