A Method for Fabricating Long Decellularized Scaffolds from Skeletal Muscle

Introduction

Skeletal muscle tissue engineering (SMTE) is a rapidly evolving field with diverse interdisciplinary applications, spanning from regenerative medicine to biorobotics. ¹ In the biomedical domain, engineered muscle constructs are being developed for tissue grafting,^2,3 disease modeling, and drug screening, ⁴ offering controlled environments to study muscle development and physiology. Simultaneously, SMTE advances inform the development of biohybrid systems, ⁵ where muscle-inspired constructs are integrated into robotic platforms to enable biologically inspired movement. This broad range of desired applications highlights the high potential of SMTE, underscoring the importance of developing physiologically relevant muscle constructs suitable for both biomedical and biohybrid applications. Yet, despite significant recent progress, reproducing the structural scale and organization of native skeletal muscle remains a major challenge.

SMTE strategies generally rely on the interplay of three key components: a suitable cell source, a scaffold that guides tissue organization, and biochemical or mechanical cues that promote maturation.^6,7 Among these, the scaffold plays a crucial role. Its physical and biochemical properties provide structural support for three-dimensional (3D) tissue formation ⁸ and strongly influence cellular behavior. ⁹ To be suitable for SMTE, a scaffold must possess properties tailored to the specificities of skeletal muscle and should therefore: (1) be biocompatible, (2) possess sufficient mechanical strength to withstand the cyclic strains used in muscle maturation protocols, (3) mimic the anisotropic, hierarchical architectural organization of native muscle, with aligned pores or fibrillar structures, (4) support muscle cells' growth, differentiation, and alignment, (5) be sized to allow myofibers to reach physiologically relevant dimensions, and (6) degrade at a rate compatible with tissue development.^1,10

Many natural and synthetic materials have been explored to meet these criteria, ¹¹ but they often fail to capture the biochemical and architectural complexity of the native extracellular matrix (ECM). ¹ Consequently, there is growing interest in decellularized ECM scaffolds, which can more faithfully capture some of the native matrix’s features. ¹² Multiple decellularization techniques have been investigated for skeletal muscle, falling into three broad categories known as chemical, enzymatic, and physical decellularization.^13,14 Each method balances cell removal and ECM preservation differently; however, detergent-based chemical protocols, typically using sodium dodecyl sulfate (SDS), are the most widely employed. ¹⁴ Still, identifying an optimal decellularization strategy remains challenging as it must be adapted to tissue geometry. The process must be efficient enough to remove cellular material while being gentle enough to preserve ECM structure and mechanical integrity. ¹³ Consequently, most existing decellularization methods are applied to small muscle fragments or homogenized tissue, ¹³ which are subsequently reprocessed using electrospinning, 3D bioprinting, ¹⁵ or hydrogel casting. ¹⁶ While these approaches offer customizability, they rarely reproduce the highly aligned and hierarchical native architecture of the native muscle ECM. ¹⁷ Conversely, whole-muscle decellularization better preserves architecture but often suffers from poor reagent penetration into deeper tissue regions, ⁸ leaving the tissue core only partially decellularized.

Beyond these biochemical challenges, a major and often overlooked limitation in SMTE is the small size of engineered constructs. Most engineered muscle tissues reported to date measure only millimeters to a few centimeters in length and a few millimeters in thickness,^18–39 constrained by diffusion limits and mechanical fragility. While such scales may suffice for many in vitro studies, they may not fully reproduce the mechanical loading environment or long-range fiber alignment of native muscle, in which myotubes can extend up to 10 cm. ⁴⁰ As a result, current systems may be limited for applications that require macroscale force generation, macroscale contractions, biomimetic strain distribution, or more physiologically relevant dimensions. Extending construct length while maintaining 3D organization would therefore provide an opportunity to better investigate these length-dependent aspects of muscle function and help bridge the gap between microengineered models and larger-scale systems.

In this context, the development of decellularized scaffolds that preserve long, aligned muscle bundles is of particular interest. Based on this rationale, a method is presented here for preparing decellularized skeletal muscle bundles using an approach that lies between whole-organ decellularization and finely minced tissue processing. Intact muscle fibers are isolated through targeted dissection to retain the native matrix architecture, and decellularization conditions are optimized to achieve removal of cellular material while preserving ECM integrity. The resulting scaffolds exceed 10 cm in length, exhibit physiologically relevant anisotropy, and possess sufficient mechanical strength to support downstream handling and stimulation. This work therefore provides a platform to investigate skeletal muscle behavior at extended length scales and contributes toward addressing current dimensional limitations in SMTE.

Methods

Experiment

Scaffold preparation

For scaffold preparation (Fig. 1A), whole pig psoas major muscles were incubated in glycerol, dissected into fiber bundles, and finally decellularized by freeze-thawing and SDS treatment, yielding decellularized muscle scaffolds suitable for further characterization. Muscle bundles were reproducibly dissected to lengths exceeding 10 cm while maintaining fiber integrity, as shown by scanning electron microscopy (SEM) imaging of glycerinated tissue prior to decellularization (Fig. 1B). After detergent treatment, scaffolds appeared macroscopically translucent, consistent with cellular removal, while retaining visible overall fiber alignment (Fig. 1C). At higher magnification, SEM imaging confirmed preservation of ECM fiber orientation, resulting in scaffolds with a well-defined anisotropic architecture (Fig. 1D). Higher SDS concentrations caused marked fiber disruption, indicating that SDS concentration was the primary contributor to structural damage over longer incubation times. Scaffold diameter decreased after decellularization under all conditions (Fig. 1E), with 24 h treatments yielding submillimeter scaffolds, a thickness compatible with diffusion-mediated cell survival in the absence of vascular support.^41,42

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FIG. 1.

Preparation and preliminary morphological characterization of decellularized skeletal muscle scaffolds. (A) Workflow of scaffold preparation. (B) SEM imaging of glycerinated tissue shows intact aligned fibers (scale bar = 200 μm). (C) Postdetergent treatment photograph showing a translucent scaffold (>10 cm) with preserved macroscopic fiber alignment (highlighted by the red line). (D) SEM imaging confirms overall preservation of extracellular matrix architecture and anisotropic fiber orientation. Excessive SDS concentrations results in increased fiber disruption (scale bar = 200 μm). (E) Scaffold diameter after decellularization. Average diameter before SDS treatments was 2.50 mm (SD = 0.8). Diameter decreases under all conditions with 24 h treatments yielding submillimeter constructs. SEM, scanning electron microscopy; SDS, sodium dodecyl sulfate; SD, standard deviation.

Assessment of decellularization

DNA quantification confirmed effective decellularization under all conditions (Fig. 2A). Relative to native muscle, DNA content was significantly reduced by 87% following 0.5% SDS treatment for 8 h, by 96% following 0.5% SDS for 24 h, by 90% following 1% SDS for 8 h, and by 92% following 1% SDS for 24 h. PicoGreen analysis yielded absolute dsDNA values of 39.7 ± 9.1 ng/mg, 4.9 ± 4.3 ng/mg, 27.1 ± 11.9 ng/mg, and 4.3 ± 1.9 ng/mg dry weight, respectively (Fig. 2B). All values were below the widely accepted 50 ng dsDNA/mg of ECM dry weight threshold, ⁴³ confirming successful cell removal. DAPI staining and histological assessment provided complementary qualitative information on decellularization efficacy. Native muscle controls showed abundant, well-defined, and aligned nuclei by both DAPI (Fig. 2C, left) and H&E staining (Fig. 2C, right). In comparison, DAPI staining of 8-h-treated samples exhibited only a small number of diffuse, poorly defined signals, consistent with residual nuclear material, whereas no discernible discrete nuclear staining was observed in 24 h scaffolds aside from minor background autofluorescence (Fig. 2D). H&E staining of transverse sections (Fig. 2E) corroborated these findings, showing scattered remaining nuclei following short decellularization and no detectable nuclei after longer treatments. These observations are in agreement with the quantitative PicoGreen results, confirming more complete removal of nuclear material with extended decellularization duration.

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FIG. 2.

Assessment of scaffold decellularization. (A) DNA reduction in decellularized scaffolds compared to native muscle as measured by NanoDrop. (B) PicoGreen quantification of residual dsDNA in decellularized scaffolds. All decellularization methods yielded scaffolds containing below the threshold of 50 ng dsDNA/mg ECM dry weight (dashed line) (43). (C) Native predecellularization tissue: DAPI staining (left) and H&E (right) shows numerous, well-defined nuclei aligned along the fiber axis. (D,E) Decellularized scaffolds: DAPI (D) and H&E (E) show consistent trends across conditions, with scattered nuclear remnant after short treatments and no apparent residual nuclei after longer treatments. Structural integrity is largely preserved under milder conditions, with striation retained at 0.5% for 8 h, but is clearly disrupted in the harsher 1% 24 h condition. DAPI, 4′,6-diamidino-2-phenylindole; ECM, extracellular matrix; H&E, hematoxylin and eosin.

Biochemical, structural, and mechanical characterization

Microcomputed tomography (µCT) imaging of glycerinated controls showed a uniformly dense structure (representative image, Fig. 3A, middle), consistent with the presence of intact cellular and extracellular components. Scaffolds treated with 0.5% SDS for 8 h and 24 h (Fig. 3A, left) exhibited longitudinally aligned tubular voids throughout the construct, indicative of effective cell removal while preserving the native ECM architecture. In contrast, scaffolds treated with 1% SDS showed disruption of structural features. In the 1% SDS for 8 h condition (Fig. 3A, top right), some channels appeared misaligned and folded, as visible in the lower-right region of the image. In the 1% SDS for 24 h condition (Fig. 3A, bottom right), scaffold collapse was observed, consistent with the loss of structural integrity observed in SEM imaging. Across noncollapsed conditions, the mean cavity width was 143 µm. Preservation of structure at lower SDS concentrations was further supported by histological analysis. H&E staining of longitudinal sections from 0.5% SDS for 24 h scaffolds (Fig. 3Bi) revealed the same longitudinally aligned tubular architecture, while transverse sections (Fig. 3Bii) displayed the associated characteristic honeycomb pattern, confirming retention of the native ECM structure. Sirius Red (Fig. 3Biii) and Alcian Blue (Fig. 3Biv) staining of longitudinal sections respectively verified the presence of collagen and glycosaminoglycans (GAGs), consistent with preservation of key ECM components following decellularization. Residual striation patterns were occasionally visible in H&E sections under milder conditions, predominantly in the 0.5% SDS 8 h samples, suggesting that minor amounts of contractile proteins may remain.

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FIG. 3.

Biochemical, structural and mechanical characterization of decellularized muscle scaffolds. (A) Representative μCT images of glycerinated controls (middle) and treated scaffolds (scale bars = 500 μm). (B) Histological assessment of scaffold architecture and composition of scaffold treated with 0.5% SDS for 24 h: (i) H&E stain, longitudinal section (scale bar = 500 μm); (ii) H&E stain, cross-section (scale bar = 100 μm); (iii) Sirius Red staining for collagen, longitudinal section (scale bar = 500 μm); (iv) Alcian Blue staining for glycosaminoglycans, longitudinal section (scale bar = 500 μm). (C) Mechanical testing showed (i) retained tensile strength and (ii) strain behavior comparable to controls. Higher (iii) stress values after prolonged SDS exposure reflected reduced cross-sectional area following decellularization. (iv) Schematic representation of stress–strain used to determine toe and linear-region moduli. (v) Toe modulus. (vi) Linear region modulus. μCT, microcomputed tomography.

Mechanical testing demonstrated that decellularized scaffolds retained tensile performance comparable to that of glycerinated controls (Fig. 3Bi–iii), with no significant deterioration across treatments. Maximum force remained stable, indicating preserved load-bearing capacity of the bundles, while increased maximum stress after longer SDS exposure reflected reduced cross-sectional area post-decellularization. Strain at maximum stress was two- to threefold higher in prepared scaffolds compared to glycerinated samples, with higher values observed in more intensively treated scaffolds. However, these differences were not statistically significant. Strain at maximum stress was therefore not considered to differ meaningfully across conditions. Further analysis of the stress–strain curves (Fig. 3Biv) showed that, in the toe region (0–10% strain), decellularized samples exhibited a trend toward lower stiffness compared to glycerinated controls (Fig. 3Bv). In contrast, within the higher-strain apparent linear region, decellularized samples tended to exhibit higher stiffness, with the 24 h samples generally displaying higher modulus values than the other conditions (Fig. 3Bvi). These values fall within the lower range of those usually reported for native skeletal muscle tissue.^44,45

In vitro assessment of biocompatibility

Biocompatibility of decellularized scaffolds was evaluated via PrestoBlue metabolic activity assays and live/dead staining. Initial cell attachment to the scaffolds was around 30% for all conditions (34.5 ± 7.3% for 0.5% SDS 8 h, 38.2 ± 16.3% for 0.5% SDS 24 h, 24.6 ± 7.5% for 1% SDS 8 h, 37.0 ± 16.9% for 1% SDS 24 h), as estimated by normalizing the signal from cells on scaffolds to the combined signal from cells on scaffolds and cells remaining attached to the culture wells after transfer of the scaffolds to fresh wells, as described in the Methods section (Supplementary Fig. S1A). Over the subsequent 7 days of culture, C2C12-seeded scaffolds exhibited a progressive increase in metabolic activity, reaching up to 30-fold higher than day 1 levels by day 7 across all scaffold types, with no significant differences between decellularization conditions at any time point (Fig. 4A). Metabolic activity in scaffold-seeded samples remained comparable to that of well controls up to day 5. Data for well controls at day 7 are not shown due to overconfluence and subsequent cell detachment in 2D culture beyond day 5 of culture. Importantly, metabolic activity in all scaffold types was significantly higher than that in unseeded scaffold controls, confirming a negligible contribution from any potential residual cells (Supplementary Fig. S1B).

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FIG. 4.

Biocompatibility of decellularized muscle scaffolds. (A) PrestoBlue metabolic activity assay shows progressive increase in cell viability over 7 days, in line with cell behavior on well-plate controls. (B) Live/dead fluorescence imaging of C2C12 cells cultured on scaffolds for 1 and 7 days. Live cells appear green (calcein AM) and dead cells red (ethidium homodimer-1) (scale bar = 200 μm). (C) Fluorescent staining and imaging of C2C12 differentiated for 5 days on 0.5% 24 h scaffold, with phalloidin (green) and DAPI (blue). The red arrow indicates scaffold fiber orientation. The bottom image shows zoomed-in view of the region of interest from the image above (scale bars = 200 μm).

Live-cell staining corroborated these results (Fig. 4B), revealing predominantly viable cells on all scaffolds at days 1 and 7, with increased cell density over time. Scaffolds exposed to longer SDS incubation appeared to support greater surface coverage and deeper infiltration, suggesting that extended decellularization may enhance scaffold porosity and cell colonization. Complementary dead-cell staining confirmed minimal cytotoxicity, with faint, uniform red signal primarily attributable to scaffold autofluorescence. Collectively, these findings demonstrate that the rise in metabolic activity reflects cell proliferation and infiltration.

Choice of scaffold preparation conditions

To summarize scaffold performance, Table 1 compiles key evaluation criteria encompassing structural, biochemical, mechanical, and biological properties. These criteria were defined as follows: (1) scaffold diameter, with a target of <1 mm to support oxygen and nutrient diffusion; (2) preservation of fiber anisotropy following decellularization, as evaluated by SEM imaging; (3) effective decellularization, defined by DNA content <50 ng/mg ECM and the absence of visible nuclei or nuclear material (H&E, DAPI); (4) porosity, assessed by the presence of µCT-visible channels and evidence of cell penetration into the scaffold interior; (5) mechanical properties, including retention of load-bearing capacity relative to glycerinated controls, strain at maximum stress >40%, and a Young’s modulus compatible with myogenic differentiation; and (6) biocompatibility, evaluated through cell viability and proliferation assays. This overview provides a rationale for selecting the 0.5% SDS for 24 h condition as the most balanced strategy.

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Table 1.

Experiments - Choice of Scaffold Preparation Conditions

Criteria	0.5%		1%
	8 h	24 h	8 h	24 h
Diameter	0	1	0	1
Anisotropy of dissected fibers	1	1	0.5	0
Decellularization
DNA content	0.5	1	1	1
H&E and DAPI staining	0	1	0	1
Porosity
µCT	1	1	0.5	0
Cell penetration	0	1	0.5	1
Mechanical properties
Load-bearing	0.5	0.5	0.5	0.5
Strain resistance	1	1	1	1
Young’s modulus	0.5	1	0.5	1
Biocompatibility	1	1	1	1
Overall score	5.5/10	9.5/10	5.5/10	7/10
Conclusion	0.5% for 24 h proposed as optimal parameters

DAPI, 4′, 6-diamidino-2-phenylindole; H&E, hematoxylin and eosin; µCT, microcomputed tomography.

Degradation study

Scaffolds generated using the optimal 0.5% SDS 24 h protocol were then evaluated for degradation over 28 days following incubation in culture medium and freeze-drying (Fig. 5A). Dry weight decreased by ∼20% during the first week (21 ± 10%), with a gradual further decrease to 40% by day 28. However, these changes were not statistically significant across time points. Mechanical testing performed at days 1, 14, and 28 (Fig. 5B) showed a general reduction in maximum stress, roughly halving over the study period, although this trend was not statistically significant. Maximum strain remained unchanged, indicating that the scaffolds largely maintained their mechanical integrity despite modest mass loss and a nonsignificant decrease in strength over time.

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FIG. 5.

Degradation of decellularized muscle scaffolds in vitro. (A) Scaffold dry weight over 28 days in culture medium shows gradual mass loss without significant differences between time points. (B) Mechanical testing at days 0, 14, and 28 demonstrates overall stable maximum stress and strain, indicating preserved mechanical integrity throughout incubation.

Discussion

Among scaffold preparation strategies for SMTE, decellularization has emerged as particularly promising. Decellularization can preserve the biochemical complexity of the native ECM ¹² and, when applied on whole tissues, it has the additional benefit of retaining the 3D organization characteristic of the tissue. ⁸ Compared with synthetic or single-component biomaterials, decellularized matrices provide a more physiologically relevant environment for cell adhesion, growth, and differentiation. ¹ However, several challenges limit their wider adoption. Key issues include achieving efficient nuclear material removal, maintaining architectural features that support myoblast maturation, attaining physiologically relevant dimensions, and preserving mechanical robustness. Importantly, these requirements are closely interconnected. Harsher decellularization protocols may improve DNA clearance but often compromise ECM structure and mechanics. ¹³ Reducing scaffold size can facilitate decellularization but requires complex reconstruction to restore functional architecture.^13,15,16 Similarly, increasing scaffold dimensions enhances physiological relevance but impedes detergent penetration, heightening the risk of residual DNA being present. ¹³ Therefore, identifying an appropriate balance between DNA removal, structural preservation, mechanical integrity, and physiologically relevant dimensions remains a major challenge. The work discussed here presents a strategy designed to meet these requirements.

The scaffold preparation strategy developed here was designed to be simple, reproducible, and broadly accessible, relying only on standard laboratory tools and reagents. Although this study was limited by the relatively small number of biological replicates (n = 3–4), which may limit the statistical power to detect subtle differences, consistent trends were observed across independent and complementary assays. In line with this rationale, whole pig psoas major muscles were first incubated in glycerol buffer, then dissected into long fiber bundles and decellularized via freeze–thaw cycles followed by SDS treatment, yielding translucent scaffolds suitable for characterization. The dissection step was central to the method, providing its defining advantage. The goal was to isolate long, thin, parallel-aligned bundles while preserving the native 3D architecture. The psoas major was selected for its favorable anatomy: a fusiform muscle with parallel fibers ⁴⁶ over 20 cm long and low fat and connective tissue content, making it a reliable source of aligned muscle bundles. Glycerol incubation, as described by Rome, ⁴⁷ permeabilized membranes, relaxed the tissue, and maintained structural integrity, thereby facilitating clean separation of fiber bundles. ⁴⁸ Through this approach, using only fine tweezers and scalpels, thin fiber bundles exceeding 10 cm in length could be reproducibly isolated while preserving intact ECM surfaces. Preserving the structure achieved during dissection, including bundle length and ECM integrity, required careful selection of the decellularization protocol. Although a 2% SDS treatment for 24 h detergent-based method was identified as the most widely applied strategy for skeletal muscle, ¹⁴ preliminary trials showed that this protocol caused severe damage to the bundles, rendering them unsuitable for further exploration. To minimize disruption, lower SDS concentrations and shorter incubation times were combined with freeze–thaw cycles, which are known to enhance detergent penetration and clearance of cellular debris by disrupting cell membranes through ice-crystal formation. ⁴⁹ All steps were carried out at low temperature with gentle agitation to preserve ECM proteins and improve detergent diffusion. This strategy yielded translucent scaffolds that retained their bundle length and submillimeter diameter, placing them within the physiologically relevant diffusion limit for oxygen and nutrients and thereby enabling culture without prevascularization. The scaffolds also maintained visible fiber alignment, discernible in the translucent bundles to the naked eye and confirmed by SEM imaging. This anisotropy is critical for guiding muscle cells to align into myotubes capable of coordinated contraction. ⁵⁰ Notably, the impact of SDS concentration on scaffold integrity was directly observable: Higher concentration visibly disrupted ECM organization, with the most extensive structural damages occurring in scaffolds treated with 1% SDS. These results demonstrate that the combined approach of glycerination, bundle dissection, and decellularization through freeze–thaw cycling and SDS treatment produces scaffolds that are promising candidates for SMTE and warrant further characterization.

Decellularization efficiencie was assessed by quantifying residual DNA, with all conditions yielding values below the accepted threshold of 50 ng/mg ECM dry weight. ⁴³ Treatment duration had a stronger influence than SDS concentration, with 24 h incubations producing the most pronounced DNA reduction. Because residual DNA can trigger inflammatory responses and graft rejection, ⁵¹ this enhanced clearance suggests a safer scaffold profile. Consistent with this biochemical assessment, qualitative morphological confirmation of decellularization was provided by complementary histological (H&E) and fluorescence (DAPI) analyses. While scattered or poorly defined residual nuclear material was observed following shorter treatments, improved clearance was observed with extended exposure, even at low SDS concentration. Notably, although many published protocols rely on higher SDS concentrations, harsher detergents, or prolonged perfusion,^8,14 these results indicate that milder conditions can achieve comparable outcomes. This may be attributed to a combination of factors, including the prepermeabilization effect of glycerination, ⁴⁷ as well as the relatively small diameter of the fiber bundles, which likely facilitates reagent penetration and enhances decellularization efficiency. µCT imaging confirmed effective removal of cellular material, revealing empty tubular cavities aligned along the fiber axis. These structures correspond to former myofiber locations and demonstrate preservation of ECM architecture and anisotropy throughout the bundle core. This organization was maintained in scaffolds treated with lower SDS concentrations, whereas higher concentrations led to progressive disruption of structural features and, in the harshest condition (1% SDS 24 h), collapse of the construct, indicating loss of structural integrity, consistent with observations from both SEM and transverse H&E staining. As an aligned tubular architecture provides essential structural cues for myotube alignment and coordinated contractility,^10,50 these findings support the use of the less aggressive decellularization conditions. The resulting pore sizes in noncollapsed scaffolds appear suitable for cellular infiltration, with the potential to accommodate a range of relevant cell types, including C2C12 myoblasts (∼25 µm ⁵² ), which mature into fused myotubes (∼30 µm ⁵³ ) and large human myocytes (∼100 µm ⁵⁴ ). As skeletal muscle cells are the largest mammalian cells, ⁵⁵ insufficient pore size is a limitation for achieving deep colonization in SMTE. ¹⁰ The large pores observed here should facilitate cell infiltration and broaden applicability across both experimental and translational contexts in SMTE. Alongside the preserved architecture, the biochemical profile of the scaffolds remained consistent with that of native ECM. Collagen and GAG staining were performed to evaluate this as they represent the major structural and biochemical components of native skeletal muscle ECM. ⁵⁶ Collagen types I and III can account for up to 10% of ECM dry weight ⁵⁶ and respectively provide tensile strength and elasticity to muscle tissue. ⁸ Proteoglycans, including GAGs, support matrix hydration ⁵⁷ and growth factor binding. ⁵⁶ Both collagen and GAGs were detected throughout the constructs, consistent with previous works on SDS-based muscle decellularization protocols. ⁵⁷ Together, these results support that the scaffolds retain key structural and biochemical features of native skeletal muscle ECM.

Mechanical testing showed that the scaffolds retained properties relevant to functional muscle constructs. All samples withstood strains exceeding 40%, a level comparable to the upper physiological range of native muscle in vivo and known to promote myotube maturation in vitro.^5,58 Maximum force remained unchanged, consistent with preserved ECM integrity. Although scaffolds subjected to longer SDS exposure exhibited higher maximum stress, this reflected scaffold thinning and associated reduced cross-sectional area rather than improved material strength, as overall force capacity was constant. Scaffold thinning in the 24 h samples may therefore reflect higher cellular clearance, consistent with DNA quantification results, which increased porosity while leaving mechanical performance intact. Analysis of stress–strain behavior revealed a nonlinear, J-shaped curve response, in agreement with previous reports for similar biological tissues.^59–61 This behavior reflects the characteristic mechanical response of fibrous soft tissues and supports the identification of two distinct regimes, with trends in each region in agreement with prior studies.^59–61 In the low-strain (toe) region, decellularized scaffolds exhibited reduced stiffness, likely reflecting increased compliance associated with a more porous and hollow structure, as well as enhanced fiber straightening at initial loading. ⁶⁰ At higher strains, however, these scaffolds displayed increased stiffness, indicating that once the matrix is engaged, resistance to deformation is preserved or enhanced. The apparent Young’s modulus measured in this higher-strain region, which is the parameter most commonly reported for skeletal muscle, fell within the lower range of values described for native tissue.^44,45 Notably, the higher modulus observed in the 24 h condition places these scaffolds within a stiffness range reported to support myogenic differentiation.^44,45

Biocompatibility was assessed using PrestoBlue metabolic assays and live/dead staining. Metabolic activity increased across all scaffold types, although this alone cannot confirm biocompatibility as metabolic upregulation is also a hallmark of cellular stress. Live/dead staining provided the necessary confirmation, showing predominantly viable cells with minimal death after seven days. The relatively low number of visible cells in these images, particularly at early time points, is likely influenced by both the initial seeding conditions and imaging limitations. C2C12 cells were seeded at low density, as is appropriate for this cell type, ⁶² and only ∼30% of the initially seeded cells attached to the scaffold, resulting in few cells detectable at day 1. These seeding efficiencies are consistent with previously reported values for cell attachment to engineered scaffolds, particularly in the absence of dynamic seeding.^63,64 In addition, the use of widefield fluorescence microscopy may underestimate cell presence within the 3D scaffold due to limited optical sectioning, potentially obscuring cells located deeper within the material. Despite these limiting factors, a clear increase in live cell numbers between day 1 and day 7 was observed. Together, these results indicate that scaffolds supported cell survival and proliferation, confirming their biocompatibility and validating the decellularization approach. Detergent removal is a key concern in SDS-based methods, since residual detergent can induce cytotoxicity. ⁶⁵ Here, extended wash cycles proved sufficient to eliminate cytotoxic residues, as evidenced by robust cell growth across all conditions. Interestingly, scaffolds subjected to longer SDS treatments supported greater cell infiltration, likely due to subtle structural changes such as increased porosity or reduced bundle diameter that facilitate cell migration into deeper regions. Achieving deep infiltration is particularly relevant, as limited cell penetration remains a challenge in SMTE. ⁶⁶ The retained anisotropy of the scaffolds also provided directional cues for cell orientation. By day seven, particularly under the 0.5% SDS 24 h condition, cell alignment with the internal fiber structure was observed. Rather than forming a single continuous alignment, cells organized into local domains broadly oriented in the same direction, though not fully colinear across the scaffold. This pattern likely reflects minor fiber disorganization introduced during handling, but such regional alignment nonetheless represents a positive step toward coordinated myotube formation as even domain-level organization can promote functional maturation. Prestretching or mechanical conditioning before seeding may further enhance global fiber alignment and strengthen the scaffolds’ instructive potential.

Following identification of the 0.5% SDS 24 h protocol as the most promising preparation strategy, a differentiation assay was performed on this condition as a proof-of-concept. The results were consistent with observations from the proliferation stage, with clear cell alignment along the scaffold. The scaffolds supported the formation of elongated, multinucleated myotube-like structures after 5 days in differentiation medium. Phalloidin staining revealed that these cellular structures displayed preferential alignment along the scaffold fiber axis, consistent with the known influence of anisotropic topography on myoblast orientation. ⁵⁰ While more specific myogenic markers, such as MF20 or tropomyosin, would provide a more comprehensive characterization of differentiation, the morphological features observed here support the conclusion that the scaffold environment permits progression toward a differentiated phenotype. Further quantitative analysis and marker-based validation will be required to fully characterize the extent of myogenic progression.

Altogether, these in vitro assays provide important validation by demonstrating that the scaffolds are noncytotoxic, support cell colonization, and present an architecture conducive to alignment. It should be noted, however, that in vitro compatibility does not guarantee in vivo performance. Factors such as immune response, vascularization, and long-term integration will need to be evaluated before considering clinical applications. Moreover, while this study provides initial qualitative evidence of myotube formation and early differentiation, comprehensive functional assessments, including quantification of fusion, maturation, and contractility, remain to be investigated to determine the scaffolds’ full regenerative potential.

The degradation profile of the selected scaffold was then examined to assess its temporal stability under extended in vitro culture conditions, where maintaining structural integrity over time is essential to properly support cell proliferation, alignment, and maturation. In this context, a 1-month time frame was selected to represent the upper range of durations typically explored in the field, ¹ providing a more stringent benchmark for scaffold persistence. Despite modest weight loss, scaffolds maintained their mechanical performance over 1 month, with no significant reduction in maximum forces, stress, or strain. This stability suggests that the preservation of fiber alignment and bundle architecture contributes to maintaining structural integrity and resilience during prolonged incubation. The initial ∼20% dry-mass reduction within the first week likely reflects loss of loosely bound ECM components, after which weight stabilized, suggesting retention of the core fibrillar matrix. The gradual degradation profile observed after this initial drop achieves the desired balance between preserving mechanical competence during early tissue infiltration and permitting eventual resorption.

Together, these findings strengthen the case for the 0.5% SDS 24 h protocol as a robust foundation for SMTE, balancing effective decellularization with preserved architecture, mechanical strength, and stability. Placing these findings in a broader context, this work addressed a key structural bottleneck in the field: the limited size of engineered muscle constructs. By maintaining native fiber alignment over physiologically relevant lengths exceeding 10 cm, these scaffolds overcome the dimensional constraints that have traditionally restricted engineered muscle to millimeter- or centimeter-scale constructs.

Although thickness remains limited by the absence of intrinsic vascularization, the ability to generate long, structurally coherent scaffolds represents a step toward reproducing native muscle structure. Given that force generation in engineered muscle has been shown to scale with myotube length, ⁶⁷ access to extended fiber dimensions may be relevant for investigating functional performance at more physiologically representative scales. In this context, such constructs open promising avenues for applications including bio-actuated systems and mechanobiological studies that benefit from extended, aligned muscle segments. They also present opportunities for disease modeling in contexts where spatial scale and alignment are important. In particular, these scaffolds may enable investigation of cell behavior over dimensions that more closely approximate native human muscle fibers, which can extend up to multiple centimeters in vivo. ⁴⁰ This is relevant for studying processes such as myofiber maturation and pathological changes that propagate along the fiber length, as observed in conditions such as muscular dystrophies.^68,69 In addition, although the present scaffold format is not intended for direct regenerative use, its geometry may lend itself to modular strategies in which multiple aligned units are combined to generate constructs with increased cross-sectional mass and functional capacity. This approach could provide an alternative route to larger-scale engineered muscle tissues compared with whole-muscle decellularization approaches. Future work will be required to explore these possibilities and determine the extent to which such systems can support functional maturation at larger scales.

Authors’ Contributions

M.Z.C.: Writing—original draft (lead); writing—review and Editing (equal); investigation (lead); conceptualization (equal); formal analysis (lead); and methodology (lead). G.B.: Writing—review and Editing (equal); investigation (supporting); and formal analysis (supporting). S.J.B.S.: Writing—review and editing (equal) and supervision (supporting). P.-A.M.: Writing—review and editing (equal); supervision (lead); conceptualization (equal); and methodology (supporting).

Supplemental Material