A novel protocol for the direct isolation of a highly pure and regenerative population of satellite stem cells

Significance

Cell-based therapy using satellite stem cells (SSCs) holds tremendous potential for enhancing muscle regeneration. However, conventional isolation methods often yield heterogeneous cell populations contaminated with nonmyogenic cell types, requiring additional purification steps that can compromise the regenerative quality of SSCs. In this study, we present a streamlined and efficient protocol for the direct isolation of a highly pure (~97% purity) population of SSCs, eliminating the need for time-consuming and expensive purification processes. Our method preserves the regenerative capacity of the isolated cells, offering a promising and clinically relevant approach for advancing skeletal muscle regenerative engineering.

Abstract

The utility of a pure population of highly regenerative satellite stem cells (SSCs) is a prerequisite for successful cell-based muscle therapies. Previous works have reported several methods for the SSC isolation. However, the majority of cells isolated using previous methods are fibroblasts and other nonmyogenic cell types, necessitating further expensive and time-consuming purification steps often affecting the regenerative quality of the isolated SSCs. Here, we describe a simple, time-effective, and robust protocol for the isolation of a pure population of SSCs in a single direct step, eliminating the need for further purification steps. By separating the muscle fascicles from the adjacent connective tissues (i.e., epimysium and perimysium) and utilizing a defined dissociation medium, a cell pool enriched in SSCs was successfully obtained. Immunofluorescent staining confirmed the stemness and the myogenic purity of the isolated cells (~97%). Upon myogenic induction, SSCs gave rise to multinucleated myofibers that exhibited spontaneous contraction in the culture dish for up to 21 d. Efforts to optimize the culture conditions revealed that tissue culture plates (TCPs) coated with a tissue-specific extract significantly enhanced SSCs’ attachment, growth, and differentiation compared to collagen I, Matrigel-coated TCPs, or noncoated TCPs. Further studies confirmed the robust myogenic regenerative capacity of the isolated cells, as evidenced by their ability to display key regenerative characteristics, demonstrating the mild effects of our isolation protocol on their regenerative capacity. The isolation protocol presented herein can potentially be used to obtain SSCs with high myogenic purity for skeletal muscle regenerative engineering and clinical indications.

Among all the tissues present in the body, the skeletal muscle tissue constitutes the most, accounting for 40 to 45% of the total body weight (1). The primary function of the skeletal muscle is to facilitate body movement through moving bones and other body parts (2, 3). However, given its superficial anatomical location, the skeletal muscle is susceptible to injury (1).

It is now widely recognized that skeletal muscle has the capacity for in vivo regeneration following an injury (1). However, this regeneration does not always occur, particularly in cases of major muscle injuries (4). For this reason, several strategies have been established in the past several years to promote skeletal muscle regeneration following excessive injuries (5, 6). Among these strategies, cell-based therapies are superior in improving muscle regeneration and functional recovery after muscular trauma, which are based on the delivery of cells that can contribute to de novo muscle regeneration and repair (7, 8). However, to exploit the potential benefits of using such a strategy for skeletal muscle repair, it is vital to identify a proper cell source to ensure the therapy’s imminent success.

Satellite stem cells (SSCs) are native precursors of the skeletal muscle residing on the periphery of myofibers (9). They exhibit a self-renewal capacity and remarkable myogenic differentiation capability, playing a crucial role in the regeneration and maintenance of skeletal muscle tissue (10). In response to tissue injury, a substantial reservoir of quiescent (inactive) satellite cells becomes activated at the injury site due to local signals. These cells transition into proliferating myoblasts, ultimately undergoing differentiation and fusion to form myofibers. This process contributes significantly to muscle repair, making them a potent source for cell-based muscle therapies (6, 11–18).

SSCs can be isolated from skeletal muscle biopsies, and various methods for their isolation and culture have been established in the literature to date (18–25). However, the majority of the isolated cells through these methods are nonmyogenic cells, primarily consisting of fibroblasts. This necessitates further time-consuming, technically challenging, and costly purification steps [e.g., preplating, fluorescence-activated cell sorting (FACS), or magnetic-activated cell sorting (MACS)] which may result in cell loss, heightened risk of bacterial contamination, and frequently impact the viability and regenerative potential of the isolated cells (26–28). Moreover, these protocols require growing the isolated population on tissue culture plates (TCPs) coated with Matrigel (29–31). However, Matrigel is derived from cancer cells, and its suitability for clinical applications is not feasible (32–34). Therefore, as an alternative to Matrigel, collagen type I has been predominantly used (35, 36). Nevertheless, collagen type I lacks major components present in the native SSCs’ niche, including laminin, fibronectin, collagen type IV, sulfate proteoglycans, and various growth factors, which are crucial for stimulating enhanced SSC attachment, growth, and differentiation (37).

To address the constraints of current isolation and culture methods, the proposed isolation protocol should ideally yield a pure population of SSCs without compromising their regenerative capacity. Furthermore, the suggested culture protocol should be designed to effectively support the growth and differentiation of SSCs, utilizing methods that hold the potential for future clinical applications. In line with this, we have established a simple yet robust protocol for the direct isolation of a pure and highly regenerative population of SSCs, eliminating the need for further purification steps. Light microscopy and immunofluorescence analyses confirmed the stemness and the myogenic purity of the isolated cells using our protocol. Upon myogenic induction, these cells gave rise to multinucleated and contracting myofibers. Furthermore, the attachment, growth, and differentiation of SSCs were shown to be significantly enhanced when grown on tissue TCPs coated with a skeletal muscle extracellular matrix (smECM) extract compared to those grown on Matrigel, collagen I–coated TCPs, or noncoated-TCPs. Further studies confirmed the mild effects of our isolation protocol on the functional myogenic regenerative capacity of the isolated cells, as evidenced by their ability to display essential regenerative characteristics.

Results and Discussion

Obtaining a Pure SSC Population.

The utility of a pure population of highly regenerative SSCs is one of the essential requirements for a successful cell-based muscle therapy (38). Despite the many protocols established in the literature for SSC isolation (19–25), they suffer from complexity and time consumption. Moreover, they require additional postisolation purification steps that come with a myriad of additional disadvantage (26–28). Here, we have successfully developed a simple yet robust protocol for the direct isolation of a pure population of SSCs from skeletal muscle tissue biopsies, drawing upon insights from skeletal muscle anatomy (Fig. 1). This direct isolation method eliminated the need for additional costly and time-consuming purification steps, which typically impact the quality and regenerative potential of the isolated cells (26–28). Our isolation technique is based on purifying the muscle biopsy by separating the muscle fascicles from the surrounding connective tissues (i.e., epimysium and perimysium), which predominantly contain nonmyogenic cells such as fibroblasts (SI Appendix, Fig. S1). We anticipate that these purified muscle fascicles will contain a highly pure population of SSCs.

Fig. 1.

A schematic illustrating the various steps involved during the isolation of SSCs.

Notably, the epimysium and perimysium are primarily composed of collagen types I and III (37). Therefore, to further maximize the purity of the isolated population, the purified muscle fascicles underwent digestion in a dispase and collagenase IV-based solution. Dispase, a natural protease, exhibits high specificity in digesting key proteins within the satellite cells' niche, such as fibronectin, laminin, and collagen type IV, without affecting collagen types I and III (39). Recognizing that the sole use of dispase might not efficiently digest collagen-based ECM proteins (40), collagenase type IV was included in the digestion recipe. Subsequently, we hypothesized that through purifying the muscle biopsy from the surrounding connective tissues and employing digestive enzymes targeting exclusively the proteins constituting the SSCs' niche, a highly pure population of SSCs can be obtained.

Morphological and Immunofluorescence Phenotypic Characterizations.

Our hypothesis was proved through a series of in vitro experiments, including morphological and immunofluorescence phenotypic characterizations, differentiation, and functional assessments. To establish a control baseline, a subset of the isolated cells underwent a single round of preplating for comparative analysis. Hereafter, SSCs directly used after isolation are referred to as “initial plate.” In contrast, SSCs used after being subjected to a single round of preplating are referred to as “preplate.”

Morphological evaluations over time revealed that the isolated cells in both groups displayed a similar morphology to that previously reported for in vitro cultured SSCs (41), characterized by a swell nuclei region and extended cytoplasmatic processes from both poles of the cells (Fig. 2A).

Fig. 2.

Morphological phenotypic characterizations of the isolated SSCs. (A) Morphological evaluations of freshly isolated SSCs at days 0, 3, and 7 postseeding (Passage 0) and (B) at passages 1 and 2 in the initial plate and preplate groups.

It is well known that fibroblast-contaminated myogenic populations suffer from fibroblast outgrowth and predominancy during expansion (42). Therefore, we expanded the cells to passages 1 and 2, during which their morphology was evaluated to observe the presence of any cells with a distinct morphology compared to the initial observation at passage 0. Our findings indicated that cells in both groups retained the same SSCs-like morphology observed at passage 0 without any indications of cells displaying a distinct morphology (Fig. 2B). This suggests the purity of the isolated cells and affirms their origin from the same population.

Cells at passage 2 from both groups were then used for immunofluorescence phenotypic characterizations against early SSC makers, including Pax-7 and MyoD, as well as late myogenic markers expressed in differentiated myogenic cells such as Desmin and MHC. Additionally, negative markers such as α-SMA (an endothelial cell marker) and VIM (a fibroblast cell marker) were assessed to confirm the cells' stemness and myogenic purity.

We chose to phenotypically characterize the cells by immunofluorescence at passage 2, as any outgrowth of nonmyogenic cells, if present, would be more evident at this stage compared to passages 0 and 1. This would ensure a profound representation of the overall phenotype of the isolated population. Results from these characterizations revealed that nearly 97% of the isolated cells in both groups stained positive for the early SSC markers, with no obvious expression to the late myogenic markers nor the negative markers. This observation suggests the stemness and myogenic purity of the cells, respectively (Fig. 3 A–C). In addition, the relatively similar expression patterns to these markers between the initial plate and preplate groups indicate the robustness of our isolation protocol and its ability to directly yield a pure myogenic population without additional purification steps, such as preplating.

Fig. 3.

Immunofluorescence phenotypic characterizations of the isolated SSCs. (A and B) Immunofluorescence phenotypic characterizations of freshly isolated SSCs and expanded to passage 2 from the initial and preplate groups against early SSC markers (Pax-7 and MyoD), late myogenic markers (Desmin and MHC), and negative markers (endothelial α-SMA, and fibroblasts Vim markers). (C) Immunofluorescence images were used to calculate the % of positive cells per field (PF), and the resultant values were compared between the groups.

Myogenic Differentiation, Long-Term Fusion, and Functionality Maintenance.

Our growth medium is supplemented with basic fibroblast growth factor (bFGF) and epidermal growth factor (EGF) to enhance the proliferation of SSCs during expansion (43–45). Specifically, bFGF not only elicits a powerful proliferative response but also impedes the differentiation of SSCs into postmitotic myofibers (45, 46). This synergic effect ensures the maintenance of a culture of undifferentiated and actively dividing cells. For this reason, these growth factors are withdrawn from our differentiation medium to facilitate the differentiation and fusion of SSCs into multinucleated myofibers. Nevertheless, given both bFGF and HGF are abundantly secreted by fibroblasts (47), we speculated that fibroblasts-contaminated myogenic cell cultures could hinder the ability of muscle cells to differentiate and subsequently fuse into myofibers. This hindrance is attributed to the persistent secretion of these growth factors, with particular emphasis on bFGF. This speculation is in agreement with prior research findings, which demonstrated a substantial reduction in muscle cell differentiation and fusion when muscle cells were directly or indirectly cocultured with fibroblasts, attributed to the elevated secretion of FGF (48, 49). Therefore, we conducted a long-term differentiation and fusion assessment for up to 21 d. The objective of this long-term culture duration was to provide an opportunity for fibroblasts, if present, to proliferate and outgrow. This in turn allowed us to observe the influence of fibroblast outgrowth on SSC differentiation and fusion, serving as a clear indicator of the presence of an impure myogenic population.

Immunofluorescence staining for Desmin revealed that the isolated SSCs in both groups were able to differentiate, fuse into myofibers, and maintain this fusion for up to 21 d—a characteristic of a pure myogenic population (50) (Fig. 4A). Subsequently, we calculated the fusion index using the Desmin-stained images to quantify and compare the myogenic differentiation between the initial plate and preplate conditions. Both groups showed a relatively similar fusion index, with no statistically significant differences (Fig. 4B). This outcome reinforces the robustness of our isolation protocol and its ability to directly yield a pure myogenic population without necessitating additional purification steps such as preplating.

Fig. 4.

Myogenic differentiation potential of the isolated SSCs and long-term fusion maintenance in both groups. (A) Immunofluorescence staining images for Desmin at day 21 postdifferentiation. (B) Immunofluorescence images were used to calculate the fusion index in %, and the resultant values were compared between the groups.

The same differentiated cultures in both groups were examined under microscopy at day 21 postdifferentiation to detect signs of spontaneous contractions. Our findings revealed that these cells not only sustained long-term fusion but also maintained their functionality and contractility consistently for up to 21 d at a similar magnitude (Movies S3–S6). The sustained functionality of the developed myofibers over the long term strongly indicates the absence of contaminating fibroblasts in the culture since numerous growth factors secreted by fibroblasts have been demonstrated to induce senescence in primary cells, particularly in prolonged culture systems (51).

Of note, the observed long-term maintenance of functionality in the differentiated cultures from passage 2 was comparable in magnitude to that observed in differentiated cultures from passages 0 and 1 within both groups (http://www.pnas.org/lookup/doi/10.1073/pnas.2426081122#supplementary-materialsMovies S2–S5). This further confirms the myogenic purity of the isolated cells, indicated by their ability to maintain their myogenic potential upon expansion.

Based on the above analyses, we concluded that both the initial plate and preplate cultures demonstrate equivalent myogenic purity. Consequently, we decided to proceed with subsequent experiments utilizing cells from the initial plate group. However, the relatively low fusion index observed in cells from both groups implies a need for further optimizations in the culture conditions to enhance the overall behavior and fusion index of SSCs.

Optimization of Primary SSC Culture Conditions.

A critical determinant influencing the differentiation of SSCs, their subsequent fusion into multinucleated myofibers, as well as their attachment and growth is the choice of extracellular matrix (ECM) protein used to coat the culture plates (52, 53). Recognizing the significance of this factor, we aimed to optimize the culture conditions by growing SSCs on TCPs coated with various proteins, including Matrigel, collagen type I, and an smECM-derived extract. Subsequently, we assessed their attachment, growth, and differentiation. Collagen type I and Matrigel are routinely used to coat TCPs to enhance SSCs’ attachment, growth, and myogenic fusion (29–31, 35, 36). However, collagen type I lacks essential ECM components inherently present in the SSCs’ niche, such as laminin, fibronectin, collagen type IV, sulfate proteoglycans, and various growth factors (37). These ECM components play crucial roles in creating a conductive microenvironment for SSCs’ adhesion, growth, differentiation, and self-renewal (37). While Matrigel contains all of these essential components, its origin (i.e., ECM-derived cancerous cells) limits its potential clinical utility (32–34).

Given the limitations of the routinely used coating materials, we included a coating extract directly derived from the smECM in our optimization studies to evaluate its efficacy in comparison to collagen type I and Matrigel coatings. Besides constituting all the essential components needed by muscle cells to function properly and preserve their linage specificity (37), a coating substrate derived from the smECM holds greater clinical relevance. As such, it represents itself as an ideal alternative for clinical indications to the currently used coatings.

For all analyses, we coated the TCPs with each coating material at a fixed concentration of 1 mg/mL to examine the sole effects of each coating material on SSC behavior. Noncoated TCPs served as a control group. When assessing SSC attachment on the different coatings, we observed poor adherence of SSCs to the noncoated TCPs (Fig. 5A). In contrast, SSCs exhibited robust adhesion and a more spread morphology on all coated TCPs, regardless of the coating material. This phenomenon aligns with the well-established observation that cells generally display superior attachment and a well-spread morphology on ECM-coated surfaces compared to noncoated tissue culture plastics, owing to the presence of arginine-glycine-aspartic acid (RGD) peptides in the ECM, which facilitate enhanced cell attachment (54). Therefore, this outcome was anticipated and aligns with a wealth of previously published research (55, 56).

Fig. 5.

Optimization of SSC culture conditions—attachment and growth evaluations. SSCs were seeded on TCPs coated with Matrigel, Collagen type I, and smECM, and their attachment and growth were evaluated while grown on the different coatings. (A) Whole 24-well and (B) 10X mag crystal violet staining images showing the degree of SSC attachment on the different coatings 24 h postseeding, and (C) quantification data of SSC attachment at 0.5, 1, 8, and 24 h postseeding. (D) Growth of SSCs on the different coatings at days 1 and 3 postseeding measured by MTS.

However, in comparison to various coating materials, SSCs demonstrated the most robust overall attachment on smECM-coated TCPs, followed by collagen type I, and then Matrigel-coated TCPs (Fig. 5A–C). At the 0.5 h postseeding, no statistical differences in cell attachment were observed between the groups. Nevertheless, starting at 1 h postseeding until the end of the study, SSCs showed a significantly higher level of attachment on the smECM-coated TCPs compared to the noncoated TCPs. This statistical difference, in contract to the noncoated TCPs, became evident starting at 8-h postseeding and continued when SSCs were cultured on collagen type I and Matrigel-coated TCPs. At 24 h postseeding, SSC attachment was statistically higher in the smECM-coated TCP group compared to the Matrigel-coated TCP and noncoated TCP groups. However, no statistically significant differences were observed between the smECM and collagen type I–coated TCP groups at all time points.

Subsequently, we evaluated the growth of SSCs on TCPs coated with different materials using the MTS assay. The results mirrored the trends observed in the attachment assay, with SSCs exhibiting the most robust growth on smECM-coated TCPs after 1 and 3 d of culture, followed by collagen type I and Matrigel-coated TCPs, then noncoated TCPs (Fig. 5D). At every timepoint, SSC growth on the smECM-coated TCPs was significantly greater than that observed on the TCPs coated with the other materials, including the noncoated TCPs. These observations suggest that growing SSCs on cultures containing ECM from the tissue of origin can accelerate and enhance their attachment and growth capabilities. Despite Matrigel and smECM being ECM derivatives, SSCs exhibited superior attachment and growth when cultured on TCPs coated with smECM.

Indeed, due to the variations in the ECM's components among the different tissues, tissue-specific ECM products can direct better physiologically relevant cellular responses over nonmatched tissue sources (57). Therefore, the enhanced attachment and growth of SSCs on the smECM-coated TCPs compared to the Matrigel-coated indicate a better appreciation of their tissue-specific microenvironment. Although no statistically significant differences were noted between the smECM and collagen type I–coated TCP groups in the attachment study, SSCs exhibited a significantly superior growth rate on smECM-coated TCPs compared to collagen I–coated TCPs. This further emphasizes the positive effects of employing a tissue-specific ECM extract as a coating substrate for primary cell cultures.

In addition to assessing the attachment and growth of SSCs, we also examined their differentiation on the different coatings. For this assessment, we initially planned to evaluate the differentiation among the groups on day 10 postinduction. However, as early as day 4 postinduction, we observed that SSCs could rapidly fuse and fill the entire TCP with multinucleated myofibers during a routine light microscopy inspection. To prevent myofiber detachment due to confluency, we terminated the experiment on day 4 and performed immunofluorescence staining for Desmin (Fig. 6A). Desmin immunofluorescence images showed that SSCs were able to rapidly fuse, forming very long multinucleated myofibers on all coated TCPs. In contrast, SSCs exhibited limited fusion ability on the noncoated TCPs, resulting in the formation of short myotubes, as anticipated. These findings are consistent with numerous previously published studies demonstrating that myogenic cells exhibit significantly improved differentiation and fusion abilities on ECM-coated TCPs compared to noncoated ones (55, 56).

Fig. 6.

Optimization of SSC culture conditions—differentiation evaluations. SSCs were seeded on TCPs coated with Matrigel, Collagen I, and smECM, and their differentiation was evaluated while grown on the different coatings. (A) Whole 24-well and 10X mag immunofluorescence staining images for Desmin 4 d postdifferentiation. (B) Immunofluorescence images were used to calculate the fusion index in %, and the resultant values were compared between the groups.

The Desmin immunofluorescence images were then used to calculate the fusion index among the groups. According to this analysis, all ECM-coated TCP groups exhibited a high fusion index compared to the noncoated TCP group, with statistically significant differences observed (Fig. 6B). SSCs demonstrated the highest fusion index when grown on smECM-coated TCPs compared to TCPs coated with collagen type I or Matrigel, showing a statistically significant difference in comparison to the collagen type I–coated TCP group. However, no statistical differences were observed in the fusion index between the smECM and Matrigel-coated TCP groups. Matrigel, a commercially available cancer ECM derivative, is often supplemented with various growth factors known to stimulate early and enhanced myogenic differentiation (35, 58). Therefore, the relatively similar fusion index observed in the Matrigel-coated TCP group compared to the smECM-coated TCP group is likely attributed to the effect of the supplemented growth factors rather than the effect of ECM itself.

Overall, these data further support and contribute to the widely acknowledged understanding in the literature that the use of ECM-coated TCPs is absolutely necessary for optimally maintaining, expanding, and differentiating primary SSC cultures (55, 56). However, in comparison to the commercially available and routinely used coating materials such as collagen type I and Matrigel, we demonstrate that a coating solution derived from the smECM provides a superior biological and clinical alternative for culturing primary SSCs. While the impact of the smECM coating on muscle cell behavior has been previously demonstrated (59–61), our comprehensive study, comparing it with routinely used coating materials, highlights the robustness of the smECM coating, emphasizing its potential for clinical applications. Further investigations will focus on characterizing the biochemical and biophysical properties of ECM preparations across different batches to ensure consistency and reproducibility of results.

Functional Myogenic Regenerative Capacity.

Highly regenerative SSCs exhibit several key characteristics. For instance, they possess the capability to secrete muscle-specific ECM proteins, such as laminin (Lam) and collagen type IV (COL IV). Additionally, upon fusion into multinucleated myofibers, these cells demonstrate functionality through contractibility. Moreover, they develop acetylcholine receptors (AChRs), facilitating accelerated neural integration upon transplantation in vivo. Furthermore, these SSCs undergo partial self-renewal, transitioning to quiescent SSCs, thereby replenishing the stem cell pool in case any repair is needed in case of injury (3, 62). Therefore, we next sought to evaluate the functional myogenic regenerative capacity of the isolated SSCs to 1) determine the impact of our isolation protocol on their quality and regenerative potential and 2) examine their suitability for cell-based muscle therapies, where highly regenerative SSCs are typically needed for successful outcomes.

The myogenic regenerative capacity of these cells was assessed by growing them on smECM-TCPs. Subsequently, the cells were induced to differentiate for 4 d,

followed by processing them for immunofluorescence and functional assessments. Immunofluorescence-stained samples for Lam, COL type IV, AChRs, and Pax-7 revealed the robust myogenic regenerative capacity of our isolated SSCs, as they showed intense deposition of muscle-specific ECM proteins such as Lam and COL type IV, AChRs formation along the periphery of the developing myofibers, and a large pool of self-renewed Pax-7 positive SSCs that were present within the differentiated culture (Fig. 7).

Fig. 7.

Myogenic regenerative capacity of the isolated SSCs. SSCs were grown on smECM-coated TCPs, and their myogenic regenerative capacity was evaluated via immunofluorescence staining against (A) Lam, (B) COL IV, (C) AChRs, and (D) Pax-7 at day 4 of differentiation.

To further validate the functional myogenic regenerative capacity of these cells, we induced their contractions at day 4 postdifferentiation. This was achieved either through electrical stimulation using various frequencies to induce both twitch (1 and 3 Hz) and tetanus (10 Hz) contractions, or via biochemical stimulation using acetylcholine (ACh) chloride. It is widely recognized that the myofibers’ ability to follow the applied frequency pattern during electrical stimulation is indicative of a highly functional state. This functionality allows them to influx calcium ions in a magnitude corresponding to the applied stimulation frequency, as shown in previous studies (63). Moreover, the capacity of the myofibers to contract in response to biochemical stimulation with ACh chloride serves as an indicator of the functionality of the formed AChRs. AChRs play a crucial role in generating an action potential on the myofibers' surface upon contact with ACh chloride, subsequently triggering the influx of calcium ions and causing the myofiber to contract (63).

Notably, our observations in this study revealed that the formed myofibers exhibited spontaneous contractions and responded effectively to all applied electrical stimulation frequencies and biochemical stimulation with ACh chloride (Movie S7). These findings strongly suggest that the myofibers are in a highly functional state.

Having observed the functionality of myofibers and the formation of functional AChRs on the surface of developing myofibers, our subsequent focus was to evaluate the calcium handling within the myofibers following both electrical and biochemical stimulations. This additional assessment aimed to further confirm their functional myogenic regenerative capacity.

To achieve this, we stimulated the myofibers with the same frequencies employed earlier during electrical stimulation to closely monitor the calcium handling response to each applied frequency. Our observations revealed that the magnitude of calcium influx was directly influenced by the stimulation frequency applied (Movie S8), a pattern that mimics the muscle response observed in vivo (64, 65).

Furthermore, upon biochemical stimulation with ACh chloride, a rapid calcium influx was observed within the myofibers (Movie S8), providing additional evidence of the functional activity of the formed AChRs. These findings further confirm the myofibers’ functional and regenerative capacity.

Collectively, these data confirm the myogenic purity of our isolated SSCs and their robust functional myogenic regenerative capacity. These results suggest the effectiveness of the isolation method established herein, indicating its suitability for application in skeletal muscle regenerative engineering and clinical applications (6, 17, 66–70). Nevertheless, it is crucial to address certain limitations inherent in our isolation protocol.

One notable challenge is the technical difficulty posed by the intramuscular fat, making the efficient separation of muscles from fat challenging. The accumulation of intramuscular fat in skeletal muscle, particularly associated with age and obesity, presents a hurdle in achieving optimal isolation efficiency (71). Therefore, whether similar myogenic purity can be achieved from muscles of varying ages and health conditions remains a subject for further investigation. Future studies will focus on optimizing the enzymatic digestion protocol to account for variations in tissue composition, particularly in samples with high intramuscular fat content, to maximize cell yield and purity.

Additionally, the use of rabbit calf muscles as the source for obtaining SSCs in this study introduces another limitation. Rabbit calf muscles, being relatively large compared to human biopsies (0.5 cm³ or 100 to 200 mg) or muscles from smaller species such as rats and mice, facilitated the identification and purification of muscle from the epimysium and perimysium while still providing a sufficient muscle volume for cell isolation (72). However, replicating this process with human muscle biopsies or muscles from smaller species may be challenging without sacrificing a significant muscle volume. Therefore, a study using human muscle biopsies or muscles from smaller species will be mandatory to validate the efficacy and applicability of the isolation protocol presented in this study to muscles of varying sizes and origins.

Conclusion

In conclusion, we present a simple and robust protocol for the direct isolation of a highly pure and regenerative population of SSCs from skeletal muscles. Our protocol eliminates the need for additional costly and time-consuming purification steps. This efficiency is achieved through the removal of the connective tissues that primarily comprise nonmyogenic cells such as fibroblasts, during the preparation process. By targeting the cells’ native niche during the digestion process, our protocol maximizes the yield of SSCs isolated from the tissues. Moreover, our findings highlight the essential role of growing primary SSCs on ECM-coated TCPs for optimal attachment, growth, and differentiation. Notably, among all the coating materials tested in this study, SSCs cultured on TCPs coated with a tissue-specific ECM extract demonstrated superior properties. This emphasizes the advantages of employing a tissue-specific ECM as a coating substrate for primary cell cultures. Finally, the isolated SSCs in this study exhibited regenerative characteristics and demonstrated high functionality, suggesting minimal impact of our isolation protocol on their functional myogenic regenerative capacity. These results collectively confirm the efficacy and mild effects of our approach, marking it as a promising method for the isolation of SSCs with potential applications in regenerative medicine.

Materials and Methods

Isolation and Culture of Primary SSCs.

Primary SSCs were isolated from the calf muscles of (3.0 to 3.5 kg) New Zealand white rabbits (Envigo, USA) in accordance with the guidelines and regulations approved by the University of Connecticut Health Center Institutional Animal Care and Use Committee (IACUC) under protocol number TE-101976-0122. Briefly, rabbits were killed with an overdose of ketamine. The euthanized rabbits were shaved, disinfected with 70% ethanol and betadine, and placed in a prone position. The calf muscles from both hindlimbs were harvested and rinsed in phosphate-buffered saline (PBS, Gibco, USA) containing 1% Penicillin Streptomycin (P/S, Gibco) to remove excess blood. Next, the muscle tissues were thoroughly purified by removing the fascial tissues, epimysium, fat, and blood vessels. After that, the muscle tissues were longitudinally cut in half using a scalpel to expose the belly region of the muscle. Subsequently, the muscle fascicles were purified from the perimysium under the guidance of a dissecting microscope and collected. The muscle fascicles were then further purified by removing any visible intramuscular fat, blood vessels, or connective tissues, followed by mincing into 1 mm³. Next, the minced muscle tissue was digested in Dulbecco's Modified Eagle Medium-F:12 (DMEM-F:12, Gibco) containing 0.2% (w/v) collagenase IV (Gibco) and 0.4% (w/v) dispase II (Sigma Aldrich, USA) for 2 h at 37 °C with continuous agitation. Next, an equal volume of DMEM-F:12 containing 10% Fetal Bovine Serum (FBS, Gibco) and 1% P/S was added to the digested tissue to terminate the digestion reaction, followed by passing the digestion solution through a 100 µm cell strainer and centrifugation at 400×g for 5-min. Next, the cells were plated in 10 cm polyester dishes at a density of 6 × 10⁵ cells/ dish and maintained in a humidified tissue culture incubator at 37 °C and 5% CO₂ in DMEM-F:12 growth media (GM) containing 18% FBS, 10 ng/ml recombinant human Epidermal Growth Factor (rhEGF), 1 ng/mL recombinant human Basic Fibroblast Growth Factor (rhbFGF) (All from R&D systems), 10 µg/mL human insulin, 0.4 µg/mL dexamethasone (all from Sigma Aldrich), and 1% P/S. For the preplate group, the supernatant containing the nonadherent cells was collected 24 h post–initial plating and transferred to fresh 10 cm polyester dishes. From this point, the medium was changed on days 4 and 7. Once 80% confluency was reached, the cells were detached using 0.25% EDTA-trypsin (Gibco), and further expanded in T-150 flasks and maintained as described above. Cells at passage 2 were used for all experiments. Differentiation medium (DM) composed of DMEM with high glucose (DMEM-HG, Gibco) supplemented with 5% Horse Serum (HS, Gibco) and 1% P/S was used to promote the formation of multinucleated myofibers upon reaching 90 to 100% confluency or at day 4 of culture. GM and DM were changed every 2 d for all experiments.

smECM Extract Preparation.

The smECM extract was prepared from porcine hindlimb muscles. Briefly, skeletal muscles were harvested from the hindlimb of skeletally mature porcine (Animals Technologies Inc.) and purified from the fat tissues and blood vessels under aseptic conditions. Next, the purified muscle tissues were grounded into 1 to 2 mm³ small fragments and subjected to 7 freeze-thaw cycles (12 h freezing at −80 °C, followed by thawing at room temperature (RT) until completely thawed) for decellularization. After that, muscle fragments were placed in glass bottles containing sterile-filtered isopropanol to eliminate any residual lipids in the sample and left under slow stirring for 24 h at RT, followed by three washes with sterile distilled water (DH₂O) and 1% P/S. After that, muscle fragments were incubated with 5 × 10⁷ U/l deoxyribonuclease I (DNase-I) and 1 × 10⁶ U/l ribonuclease (RNase) (all from Sigma Aldrich) solution in sterile DH₂O containing 1% P/S and left under slow stirring for 12 h at RT. Next, the solution was aspirated, and the muscle fragments were extensively washed in sterile DH₂O containing 1% P/S for 72 h to remove any DNase/RNase residue, with replacing the solution every 24 h. The muscle fragments were then frozen at −20 °C for 48 h, followed by freeze-drying for 48 h. Freeze-dried muscle fragments were milled using a cryomiller (6750, SPEX SamplePrep, USA) to obtain smECM in powder form. To prepare the smECM coating solution, the powder was digested at a stock concertation of 40 mg/mL in 0.1 N acetic acid (Bio-Rad) and 1 mg/mL pepsin (Sigma Aldrich) solution in DH₂O and incubated for 48 h at RT under fast stirring. Next, the digested solution was either used immediately for coating or stored at −80 °C for later use.

TCPs Coating.

The stock solutions were diluted to a final concentration of 1 mg/mL using 0.1 N acetic acid (Collagen I, Collagen Solutions, UK, and smECM) or precooled DMEM-HG (Matrigel, Corning, USA). Next, 200 µL of each of the diluted solution was added to 24 wells and incubated at RT for 2 h under gentle agitation (Collagen I and smECM) or at 4 °C for 7 min (Matrigel). The coating solutions were then aspirated, and the coated plates were either used immediately or sealed with parafilm, stored at 4 °C, and used within a week after coating.

Morphological Phenotypic Characterizations.

The gross morphology of the isolated cells was monitored at days 0, 3, and 7 post–initial plating and after preplating and at the first and second passages using normal light microscopy (OLYMPUS, Japan).

Immunofluorescence Staining.

For the immunofluorescence phenotypic characterizations, the cells were plated in 24-well plates at a density of 5.7 K cells/ well, maintained in culture as described above, and subjected to staining after 30% confluency was reached. For the immunofluorescence differentiation characterizations, the cells were plated in 24-well plates at a density of 30 K cells/ well, maintained in culture as described above, and subjected to staining either on days 4 or 21 postinduction. Briefly, cells were fixed with 4% paraformaldehyde (Electron Microscopy Sciences, USA) for 15 to 20 min, permeabilized with 0.1% Triton X-100 (Sigma Aldrich) for 20 min, and blocked with 1% bovine serum albumin (BSA, Sigma Aldrich) solution for 1 h. Next, cells were incubated with primary antibodies for 2 h, washed twice with PBS, and then incubated with secondary antibodies mixed with DAPI (1:3000 dilution, DAPI) for 2 h. Primary antibodies used were mouse anti-Pax7 (1:10 dilution, DSHB, USA), mouse anti-MyoD (1:200 dilution, LSBio, USA), mouse anti-Desmin (1:100 dilution, ThermoFisher), mouse anti-MHC (1:4 dilution, DSHB), mouse anti-αSMA (1:100 dilution, ThermoFisher), mouse anti-VIM (1:100 dilution, Abcam), rabbit anti-Laminin (1:200 dilution, ThermoFisher), rabbit anti-COL IV (1:100 dilution, Abcam), and α-Bungarotoxin Alexa Fluor 594 conjugate (1:100 dilution, ThermoFisher) which targets AChRs. Goat anti-mouse IgG (Alexa Fluor 594) and goat anti-rabbit IgG (Alexa Fluor 488) were used as secondary antibodies (1:500 dilution, all from Abcam). To visualize Filamentous actin (F-actin), Alexa Fluor 488 Phalloidin and Alexa Fluor 594 Phalloidin (1:40 dilution, all from ThermoFisher) were used during the secondary antibody incubation. All primary and secondary antibodies were diluted using 1% BSA. The entire staining process was carried out at RT. Cells were visualized using the inverted fluorescence microscopes (Zeiss LSM 880, Germany) or (Leica DMi8, Germany).

Phenotypic Quantification.

The immunofluorescence images were used to quantify the % of positive cells per field (PF) using the ImageJ software (version 1.4 g, NIH, USA). Four to six different images from every sample/ group (N = 4/ group) were used for the quantification.

Fusion Index.

The fusion index was quantified using the Desmin-stained immunofluorescence images. Briefly, the number of nuclei within the myofibers and the total nuclei number were manually counted using the ImageJ software. To calculate the fusion index, the following equation was used:

$$FusionIndex\left(\%\right)=\frac{\mathit{Total\; number\; of\; nuclei\; within\; myofibers}}{\mathit{Total\; number\; of\; nuclei}}x100,$$

Myofibers containing more than three nuclei were used for the fusion index analysis. Four different images/ sample were used for the analysis (N = 4/ group).

Crystal Violet Staining.

Cells were plated at a density of 20 K cells/ well and maintained in culture as described above. At 0.5, 1, 8, and 24 h, their attachment was qualitatively and quantitatively evaluated (N = 4/ group) using the crystal violet assay kit (Sigma Aldrich) following the manufacturer's instructions. Briefly, the cells were washed once with PBS, fixed with 4% paraformaldehyde, washed twice with PBS, and incubated with 400 µL of the crystal violet reagent for 5 min at RT. Cells were washed with PBS until the solution remained completely clear. At this point, the cells were visualized using an inverted microscope (Leica DMi8). To quantify the attachment, 400 µL of 100% methanol was added to each sample to dissolve the bound reagent, and 100 µL from the supernatant of each sample was transferred to 96-well plates and the absorbance was read in duplicate at 570 nm using a plate reader (BioTek, Synergy H1, USA). To eliminate the interface of crystal violet reagent with the TCPs, the crystal violet reagent was added to acellular TCPs, and their absorbance values were deducted from the absorbance values measured for each sample.

MTS Assay.

Cells were plated at a density of 5.7 K cells/ well and maintained in culture as described above. At 1 and 3 d, their growth rate was quantified (N = 4/ group) using the CellTiter 96® AQueous One Solution Cell Proliferation Assay (MTS, Promega Inc, USA) following the manufacturer's instructions. Briefly, the cells were washed once with PBS, and 500 µL of the MTS reagent in a ratio of 5:1 (media: MTS) was added to each well and then incubated for 2 h at 37 °C. Next, 100 µL from the supernatant of each sample was transferred to 96-well plates, and the absorbance was read in duplicate at 490 nm using a plate reader (BioTek, Synergy H1). To eliminate the interface of MTS reagent with the TCPs, the MTS reagent was added to acellular TCPs, and their absorbance values were deducted from the absorbance values measured for each sample.

Electrical and Biochemical Stimulation.

Myofibers were electrically stimulated with 1, 3 Hz (twitch), and 10 Hz (tetanus) at 30 V and 3.6 ms pulse duration using C-PACE EP Cell Culture Stimulator (ION OPTIX, USA). Biochemical stimulation was performed by directly adding acetylcholine (ACh) chloride (Sigma Aldrich) solution into the culture medium at a final concentration of 2 mM. Spontaneous and electrically and biochemically induced myofiber contractions were imaged and recorded in the form of movies using an inverted microscope (Leica DMi8).

Fluo-4 Calcium Staining.

For the calcium handling assessment, the cells were stained using the Fluo-4 Calcium Imaging Kit (ThermoFisher) following the manufacturer's instructions. Briefly, the cells were washed once with PBS, and then incubated with the Fluo-4 staining solution at 37 °C for 30 min, followed by incubation at RT for another 30 min. Next, the Fluo-4 staining solution was aspirated, and the cells were washed once with PBS. After the staining, the cells were electrically and biochemically stimulated as described above, and the calcium handling was recorded in the form of movies using an Apple® iPhone XR camera mounted on the eyepiece of an inverted microscope (Leica DMi8).

Statistics.

All quantitative data were expressed as mean ± SD. All statistical analyses were performed using the statistical software Prism GraphPad version 8 (GraphPad, USA). Statistical analyses were performed using the unpaired Student's t test and two-way ANOVA with Tukey’s post hoc test. Statistical significance was evaluated at *P <0.05, **P <0.01, ***P < 0.001, and ****P < 0.0001.

Supplementary Material

Appendix 01 (PDF)

Movie S1.

Spontaneous contraction of differentiated SSCs from the initial plate group 21-days post-induction at passage 0.

Movie S2.

Spontaneous contraction of differentiated SSCs from the initial plate group 21-days post-induction at passage 1.

Movie S3.

Spontaneous contraction of differentiated SSCs from the initial plate group 21-days post-induction at passage 2.

Movie S4.

Spontaneous contraction of differentiated SSCs from the pre-plate group 21-days post-induction at passage 0.

Movie S5.

Spontaneous contraction of differentiated SSCs from the pre-plate group 21-days post-induction at passage 1.

Movie S6.

Spontaneous contraction of differentiated SSCs from the pre-plate group 21-days post-induction at passage 2.

Movie S7.

Spontaneous, electrically, and biochemically induced contractions of differentiated SSCs 4-days post-induction. Note that the appearance and disappearance of the red (°) during tetanus (10Hz) contraction highlights the beginning and end of the stimulation train, respectively.

Movie S8.

Calcium handling of differentiated SSCs 4-days post-induction during spontaneous, electrically, and biochemically stimulated contractions.

Data, Materials, and Software Availability

All study data are included in the article and/or SI Appendix.