COMPARISON OF SKELETAL MUSCLE DECELLULARIZATION PROTOCOLS AND RECELLULARIZATION WITH ADIPOSE-DERIVED STEM CELLS FOR TISSUE ENGINEERING

Abstract

Decellularization is a novel technique employed for scaffold manufacturing, as a strategy for skeletal muscle (SM) tissue engineering applications. However, poor decellularization efficacy is still a problem for the use of decellularized scaffolds as truly biocompatible biomaterials. For recellularization, adipose-derived stem cells (ASCs) are a good option, due to their immunomodulatory and pro-regenerative capacity, but few studies have described their combination with muscle-decellularized matrices (mDMs). This work aimed to evaluate the efficiency of four multi-step decellularization protocols to produce mDMs and to investigate in vitro biocompatibility with ASCs. Here, we described the different efficacies of muscle decellularization methods, suggesting the need for stricter standardization of the method, considering the large range of applications in SM tissue engineering, which is also a promising platform for preclinical studies with rat disease models using autologous cells.

1. Introduction

Muscle regeneration is a complex organized process depending on the activation of satellite cells that undergo asymmetric division, proliferation, and terminal differentiation [1]. Despite great regenerative capability, extensive muscle injuries, such as volumetric muscle loss (VML), impair appropriate tissue recovery, leading to excessive adipose tissue deposition and fibrosis, compromising muscle function [2]. The standard available treatment for VML is transferring autologous muscle flaps, which often results in donor site morbidity, poor engraftment and vascularization and low functional recovery [3].

Skeletal muscle (SM) tissue engineering arises as a novel alternative strategy to improve muscle regeneration. It is based on the use of specific stem cells combined with biomimetic scaffolds, which must be capable of reproducing biochemical and physical properties, providing an environment for cell adhesion, growth, migration, survival and differentiation [3].

Decellularization is being extensively used to produce tissue engineering scaffolds, since it removes the immunogenic components of the tissue while preserving bioactive extracellular matrix (ECM) compounds, such as collagen type I, laminin, fibronectin and glycosaminoglycans [4]. SM native ECM is a highly organized network that confers structural integrity and mechanical support for muscle contraction, coupling the entire tissue into a functional unity [5]. ECM interacts with cell integrins, dystroglycans and sarcoglycans, retains growth factors and bioactive molecules, regulates progenitor cells quiescence or activation, and provides an environment for revascularization and reinnervation, acting on tissue development, homeostasis and regeneration after injury [5, 6].

In order to obtain muscle decellularized matrices (mDMs), many decellularization protocols have been published [7–11], which include chemical (detergents, hypotonic or hypertonic solutions), physical (agitation, temperature) and enzymatic methods [12, 13]. The proposed criteria for decellularized scaffolds are the following: no remaining visible nuclear material; less than 50 ng of DNA per milligram of tissue; DNA length less than 200 bp; preserved ECM components and cytocompatibility [14]. However, despite these established parameters, there have been controversies regarding the reproducibility and efficiency of some published decellularization protocols [10]. Traces of cytoplasmatic components and DNA elicit host immune response in vivo, highlighting the relevance of investigating these methodologic issues for the future applicability of mDMs as medical devices [14,15]. Circumventing these issues is extremely relevant because ECM is evolutionarily conserved among species, opening the possibility of developing acellular ECM-based scaffolds from different organisms without the risk of rejection [12,16].

Another challenge in the tissue engineering field is choosing a suitable cell type for mDMs recellularization, which is also an important way to evaluate in vitro cytocompatibility, a measure of possible impacts of the scaffolds in the survival and morphology of cultured cells [17]. Mesenchymal stem cells (MSCs) are a promising cell type for mDMs recellularization, because of their strong immunomodulatory properties and the ability to secrete trophic antiapoptotic bioactive factors that can contribute to tissue regeneration [18]. Adipose-derived stem cells (ASCs) are a very attractive type of MSCs for clinical application because high amounts of cells can be obtained from the patient by liposuction, and also because of their multilineage in vitro differentiation capability [19, 20]. ASCs transplantation is already a safe treatment for many diseases with promising outcomes [21].

The combination of mDMs with ASCs is an attractive option for muscle injury [22]; however, this approach has been poorly explored so far. Therefore, we aimed to evaluate the efficiency of different decellularization protocols (P1, P2, P3 and P4) to produce mDMs from rat tibialis anterior muscles, to characterize mDMs and to analyze the capacity of ASCs to survive and colonize them, evaluating their potential for SM tissue engineering application.

2. Material and Methods

All reagents were acquired from Thermo Fisher Scientific (USA) except where indicated.

2.1. Animals

Lewis rats 4 – 8 months of age were used in this study. Experiments were approved by the Ethics Committee in the use of animals of the Federal University of Minas Gerais (CEUA/UFMG, protocol 60/2017) and guidelines of the National Council for Animal Experimentation Control (CONCEA) for the care and use of laboratory animals were followed. Animals were maintained in appropriate cages and with access to food and water ad libitum.

2.2. SM decellularization

Rats were euthanized and tibialis anterior muscles (n=3, per protocol) were collected aseptically. Muscles were rinsed in 0.15% sterile phosphate-buffered saline (PBS), followed by fascia and tendons removal, and treatment with four different protocols termed P1, P2, P3 and P4. Protocols P1, P2, and P3 were performed at 4 °C, and suffered minor modifications from Porzionato and collaborators [10]; Stern and colleagues [9] and Conconi and collaborators [7], respectively. P4 was carried out at room temperature, except where indicated, and was adapted mainly from Wang and colleagues [23]; Triton X-100 solution composition, concentration and time of treatment in P4 was applied as described by Porzionato and collaborators [10]. Mechanical agitation was applied in all steps, except where indicated. Samples for protocol P1 were cut into 1 cm². Samples of P2, P3 and P4 protocols were previously frozen at −80 °C and cut into thin slices. The detailed multi-step protocols are described in Table 1.

Table 1. Detailed multi-step decellularization protocols.

	Protocol
Step	P1	P2	P3	P4
1	Ultrapure water (UW) for 24 h	UW for 48 h (5 changes)	UW for 72 h	UW for 48 h
2	0.05% trypsin-0.02% EDTA for 1 h at 37 °C (without agitation)	0.05% trypsin-0.02% EDTA for 1 h (without agitation)	4% sodium deoxycholate for 4 h	NaCl 0.5 M for 4 h
3	PBS wash (3x)	DMEM overnight	DNase I (5 U/ml) for 3 h at 37 °C	NaCl 1 M for 4 h
4	2% Triton X-100–0.8% NH4OH for 72 h	1% Triton X-100 for 5 days (1 change per day)	Repeated steps 1–3	Deionized water (dH2O) for 1 h
5	UW for 48 h	UW for 48 h (1 change)		Repeat steps 2–4
6	0.15% PBS (3x)	0.15% PBS for 24 h		0.25% trypsin/EDTA at 37 °C for 2 h (with agitation)
7				dH2O for 1 h
8				2% Triton X-100–0.8% NH4OH for 3 days
9				UW for 24 h
10				DNase I (5 U/ml) for 3 h at 37 °C
11				0.15% PBS for 48 h

UW: ultrapure water; PBS: phosphate-buffered saline; EDTA: Ethylenediamine tetraacetic acid; DMEM: Dulbecco’s Modified Eagle Medium; dH2O: deionized water

2.3. Evaluation of decellularization efficiency

Control SM and mDMs were histologically processed, sectioned into 6 μm slices, and submitted to Hematoxylin and Eosin (H&E) and Masson’s Trichrome staining according to standard protocols.

To further evaluate cell removal, it was performed scanning electron microscopy (SEM). Samples were fixed overnight with 2.5% glutaraldehyde-2% paraformaldehyde, and subjected to secondary fixation with 1% osmium tetroxide, dehydration, critical point drying followed by platinum metallization. Images were captured with SEM Quanta FEG 3D FEI at the Center of Microscopy/UFMG.

2.4. DNA content and integrity after decellularization

Total DNA was extracted with DNAzol^® reagent (Invitrogen; USA) according to manufacturer instructions. Control SM and muscles decellularized with P1, P2, P3 and P4 were treated with 1 ml of DNAzol and mechanically homogenized. Then they were subjected to a series of centrifugation and ethanol treatment. Quantification was performed with spectrophotometer SkanIt MultilaserGO (Thermo Fisher Scientific, USA). To evaluate the genomic DNA integrity, a 248 bp genomic sequence of the Actb gene was amplified by PCR (F: CCATAGTGGGGTGTGGTCAG / R: CCTAGAAGCATTTGCGGTG). About 50 ng of DNA for each sample was used in PCR reactions and amplicons were visualized in 2% agarose gel with 10 μg/ml ethidium bromide.

2.5. Preservation of ECM components on mDMs

In order to check whether ECM components were preserved after decellularization protocol, immunofluorescence was performed with mDMs previously considered efficiently decellularized, according to results from histology, SEM and DNA quantification. Samples were fixed with paraformaldehyde 4% for 20 min, incubated with a blocking solution containing 1% albumin bovine serum for 30 min, primary antibodies against laminin (anti-rabbit 1:200; Abcam, UK) and collagen I (anti-mouse; Abcam, UK) for 2 h, followed by secondary antibody Alexa Fluor^® 555 (anti-rabbit and anti-mouse IgG, 1:500, respectively) and 1 μg/ml Hoechst 33259 (Invitrogen, USA) for 1 h. Images were captured in confocal microscope LSM 880 from ZEISS, using ZEN software (Zeiss, DE) at the Center of Image Processing and Acquisition (CAPI-ICB/UFMG).

2.6. ASCs isolation and characterization

Isolation and characterization of ASCs were performed as previously described [19]. Rat adipose tissue was collected immediately after death and transferred to Dulbecco’s Modified Eagle Medium (DMEM) high glucose with antibiotics. The tissue was washed with PBS and treated with 0.15% collagenase type I at 37 °C for 1 h. The homogenate was centrifuged at 300 ×g for 10 min and the pellet was resuspended in basal medium: DMEM with 5 mM sodium bicarbonate, 100 U/ml penicillin, 0.1 mg/ml streptomycin, 0.25 μg/ml B amphotericin (PSA; Sigma-Aldrich, USA), 60 mg/ml gentamicin and 10% fetal bovine serum (FBS; Cripion, BR). Cells were expanded in culture plates and the medium was changed every other day. Subculture was performed after reaching 80% confluence with 0.05% trypsin treatment for 5 min at 37 °C. ASCs were used in passages 3–4. For ASCs functional characterization, in vitro osteogenic and adipogenic differentiation potential was evaluated. For adipogenic differentiation, cells were plated in 6-well plates in a density of 5×10⁴ cells per well and cultivated in basal medium containing 0.1 μM dexamethasone, 50 μM indomethacin, 100 UI insulin (Eli Lilly, USA) and 0.5 mM 3-Isobutyl-1-methylxanthine (IBMX; Sigma-Aldrich, USA) [19]. After 21 days, cells were stained with Oil Red O. For osteogenic differentiation, 1×10⁵ cells were plated in 6-well plates and cultivated in basal medium with 50 μg/ml ascorbic acid, 10 mM β-glycerophosphate and 1 μM dexamethasone. Von Kossa staining was performed after 21 days [19].

For phenotypical characterization, ASCs were incubated with antibodies against CD11b/c, CD54, CD73, CD90, MHCI and CD45 (all from BD Pharmingen^™, USA), CD34 (Santa Cruz Biotechnology, USA) and MHCII (Abcam, USA) [24]. Cells were then treated with anti-mouse IgG Alexa Fluor^® 647 (Invitrogen, USA) for 30 min. Cells incubated only with secondary antibodies were used as negative control and unmarked cells were utilized for population establishment. Data were collected in Guava EasyCyte 6–2L (Millipore, USA), 15,000 events were acquired and analyses were performed using FlowJo (BD Biosciences, USA).

2.7. Recellularization of mDMs and in vitro biocompatibility evaluation

To perform recellularization, mDMs considered decellularized were treated with PBS and antibiotics for 24 h at room temperature with agitation. Then, mDMs were exposed to UV light for 1 h, treated with DMEM for 24 h at 4 °C, and incubated in basal media for 1 h at 37 °C. ASCs were seeded in a small volume (approximately 20 μl) and incubated for 2 h at 37 °C. The medium was completed and replaced every other day. Cell viability was evaluated utilizing Calcein-AM staining and MTT assay. A total of 5–10×10⁴ ASCs were seeded per mDM and after 1 or 2 weeks they were stained with 5 μM CellTrace^™ Calcein red-orange AM (Invitrogen, USA) or 5 mg/ml of (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT; Invitrogen, USA), respectively, following manufacturer instructions. The controls consisted of ASCs cultivated on culture plates. Images of viable cells were captured in a confocal microscope Zeiss LSM 880 with ZEN software (for Calcein-AM staining) or Olympus IX70 Microscope (Olympus Corporation, JP) (for MTT staining). MTT formazan crystals were solubilized with 10% SDS-HCl at 37 °C overnight and absorbance at 595 nm was measured with a spectrophotometer.

Cell morphology and capacity of cells to adhere and colonize mDMs were evaluated by SEM. ASCs were plated at a density of 5–10×10⁴ cells per mDM and cultivated for 14 or 21 days. Then, the recellularized matrices were processed for SEM as described above.

2.8. Statistical analysis

Data were plotted using GraphPad Prism 5.0 software (Graph Pad Software, USA). For DNA quantification, One Way ANOVA and Bonferroni’s post-test were performed, considering significant p<0.05. For MTT analysis, an unpaired t-test was applied, considering significant p<0.05.

3. Results

3.1. Evaluation of decellularization efficacy and preservation of ECM components

Decellularization was performed with rat tibialis anterior muscles. The capacity of removing cellular components and DNA was compared among 4 protocols: P1, P2, P3 and P4 (Figure 1A). Analyzing gross macroscopic appearance after decellularization, P3-derived matrices were reddish, suggesting incomplete cell removal (Figure 1B), and muscles decellularized with protocol P1, P2 and P4 turned to white translucent.

Figure 1. SM isolation and decellularization to produce mDMs.

A: Schematic representation of rat SM isolation and decellularization protocols to eliminate cells and DNA in order to produce mDMs. B: Macroscopic aspect of SM after treatment with protocols P1, P2, P3 and P4. Scale: 1000 μm.

Histological analysis of control SM cross-sections showed typical rounded muscle fibers organized in bundles surrounded by ECM (Figure 2A, F). Muscles decellularized with P1 (Figure 2B, G) and P4 (Figure 2E, J) showed no visible remaining myofibers and a tissue composed mainly of ECM, as evidenced by Masson’s trichome staining. Meanwhile, treatment with P2 (Figure 2C, H) and P3 (Figure 1D, I) protocols resulted in many preserved myofibers, despite some differences in tissue organization, when compared to native SM.

Figure 2. Histological analysis of SM after decellularization with P1, P2, P3 and P4.

Histological evaluation with H&E staining (A-E) and Masson’s trichome staining (F-J) of rat SM before (A, F) and after decellularization with protocols P1 (B, G), P2 (C, H), P3 (D, I) and P4 (E, J). Scale: 100 μm.

To further evaluate cell removal and to evaluate tissue architecture, we have performed SEM (Figure 3). Surprisingly, in P1 decellularized muscles, many rounded cells attached to the ECM were identified (Figure 3B). Confirming histological findings, muscles treated with P2 (Figure 2C) had some myofibers remnants, and samples treated with protocol P3 (Figure 2D) had many intact organized elongated myofibers, corroborating low decellularization efficiency of these methods. Tissue decellularized with P4 showed many tubular cavities, suggesting preservation of the three-dimensional architecture of SM, and no trace of cellular components were visualized (Figure 2E).

Figure 3. Analysis of mDMs ultrastructure by scanning electron microscopy.

Representative scanning electron micrographs of control SM (A) and muscles treated with protocols P1 (B), P2 (C), P3 (D) and P4 (E). The right column represents higher magnifications of the central column. White arrows point to the remaining cells. Scale: 100 μm.

Genomic DNA quantification was performed to analyze the amount of DNA remaining in the ECM, an important criterium for decellularized scaffolds. Results revealed that P1-treated muscles were found to contain 138.55 ± 36.13 ng DNA/mg in dry weight, similar to control SM, whose amount was 154.52 ± 32.42 ng/mg. A significant reduction of DNA quantification was observed in muscles treated with P2 (6.75 ± 2.78 ng/mg), P3 (54.07 ± 20.11 ng/mg) and P4 (38.80 ± 8.97 ng/mg) compared to both native SM and muscles treated with P1 (p<0.05) (Figure 4A). Accordingly, protocols P2 and P4 reduced DNA content below the recommended 50 ng per mg of dry tissue. Corroborating quantification results, gel electrophoresis after amplification of a 248 bp fragment showed visible PCR product from SM and P1 DNA samples (Figure 4B), suggesting, at least partially, DNA integrity preservation. No visible bands were observed for P2, P3 and P4-derived samples.

Figure 4. Genomic DNA content analysis of mDMs.

A: Genomic DNA quantification of dry tissue of control SM before and after treatment with protocols P1, P2, P3 and P4. Data are presented as the mean ± standard error; n = 3. * Compared to each protocol and # compared to SM (* or # represents p<0.05, ** or ## represents p<0.005, One Way ANOVA and Bonferroni’s post-test). B: Agarose gel electrophoresis corresponding to Actb gene amplification from DNA extracted of mDMs and SM. NTC: No Template Control.

Taken together, these results showed that muscles decellularized with P4 were the only ones that passed all criteria for decellularized scaffolds. Therefore, the complete characterization and all subsequent experiments were performed with matrices produced with protocol P4, named here as mDMs.

To characterize mDMs, we have performed immunofluorescence staining for ECM components, collagen I and laminin. We observed that collagen I and laminin were still present in mDMs (Figure 5A e 5B), despite a different spatial distribution of these proteins compared to native SM. Hoechst staining confirmed the complete absence of a nucleus on mDMs, contrary to what is observed for fresh SM (Figures 5A and 5B).

Figure 5. Presence of ECM components in mDMs.

A: Representative confocal images of SM and mDMs decellularized with P4 showing the preservation of collagen I (red). B: Representative confocal images of SM and mDMs decellularized with P4 showing the preservation of laminin (red). Hoechst staining evidenced nuclei in SM and the absence of nuclei in mDMs (Hoechst, blue). Scale: 50 μm.

3.2. ASCs isolation and recellularization of mDMs

In order to characterize mDMs biocompatibility after treatment with decellularizing agents, we performed in vitro recellularization using MSCs. As a source of MSCs, we choose ASCs, which were isolated from Lewis’s rat inguinal fat. Cells were cultivated, presented a fibroblastic morphology, adherence to plastics and proliferative capacity maintained for at least 6 passages (data not shown). To confirm the specific isolation of ASCs, we performed functional characterization through in vitro adipogenic or osteogenic differentiation. After 21 days of treatment with differentiation medium, lipid droplets were evidenced with Oil Red O (Figure 6B) and the mineralized matrix was visualized through Von Kossa staining (Figure 6D), while no staining was observed in ASCs treated with basal media (Figure 6A and 6C). Immunophenotyping was performed through flow cytometry for MSCs and hematopoietic surface markers. As expected, more than 95% of the cells expressed CD90, CD73, CD54, and MHCI, while less than 2% were positive for hematopoietic markers CD45, CD34 and CD11 and also for MHCII (Figure 6E).

Figure 6. Characterization of rat adipose-derived stem cells.

Functional and immunophenotypic characterization of rat ASCs isolated from inguinal fat. A, B: Adipogenic differentiation was evidenced by Oil Red, with lipid droplets stained in red. A: Control group was maintained in basal medium. B: ASCs differentiated into adipogenic lineage. C, D: Osteogenic differentiation was demonstrated by von Kossa staining. C: Control group, cultivated in basal medium. D: ASCs differentiated into osteogenic lineage. Scale: 50 μm. E: Flow cytometry analysis showing expression of MSC surface markers MHCI, CD90, CD73 and CD54 and nearly absence of expression of hematopoietic markers MHCII, CD45, CD34 and CD11 by rat ASCs. Data are expressed as the mean percentage of positive cells ± standard error (n=3).

The in vitro recellularization of mDMs was performed by seeding ASCs on mDMs, and cell viability of recellularized mDMs and controls was checked by Calcein staining. The red fluorescent cells on mDMs and control were observed, suggesting that mDMs support the culture of ASCs and present in vitro biocompatibility after 7 days of culture (Figures 7A and 7B). In addition, higher magnification of Calcein staining micrographs of recellularized mDMs (Figure 7B1) showed that cells not only adhered, but also acquired an elongated morphology.

Figure 7. Recellularization of mDMs with ASCs and in vitro cytocompatibility.

A, B: ASCs were cultivated on 2D culture plates or on mDMs for 7 days and stained with Calcein-AM. Representative confocal images showing viable cells both in control (A) and recellularized mDMs (B). B1: Higher magnification of Calcein staining of ASCs cultivated on mDMs, showing elongated cell morphology. C-E: After 14 days, ASCs cultivated on 2D culture plates and recellularized mDMs were stained with MTT. Dark formazan crystals indicate viable and metabolic active cells both in control (C) and recellularized mDMs (D). Scale: 100 μm. E: Quantification of MTT optical density (OD) from ASCs cultivated on mDMs or control. Unpaired t test; *** is p<0.0005; n=3. F: Representative scanning electron micrographs reveal ASCs adherence into mDMs after 14 and 21 days. White arrows indicate cells. Scale: 100 μm (A-D) and 25 μm (F).

To confirm ASCs viability, recellularized mDMs were treated with MTT, and formazan crystals were observed inside mDMs, showing viable and metabolic active cells (Figures 7D) after 14 days of culture, similar to control (Figure 7C). Quantification of MTT optical density (OD) revealed that cell viability was statistically higher on scaffolds compared to control bidimensional (2D) culture plates (Figure 7E). To analyze cell adherence and morphology of ASCs on mDMs, SEM was performed after 14 and 21 days. As shown in Figure 7F, in 14 and 21 days we could observe that ASCs adhered to mDMs, colonize the scaffolds, and emitted cytoplasmic extensions, especially after 21 days of culture.

Taken together, these data indicate that decellularization protocol P4 was capable of successfully removing cells from SM, and protocols P1, P2 and P3 did not reach the minimal criteria for decellularized matrices. The mDMs showed well in vitro biocompatibility since cells were capable of adhering and surviving into mDMs for as long as 21 days.

4. Discussion

Previously considered as an inert tissue constituent, ECM has been shown to have an active role in both tissue homeostasis and regeneration after injury [25]. SM ECM acts not only as a mechanic structural reinforcement but also as a signalling environment, cell fate regulator and regenerative orchestrator. The notion of its complex functional role has led to the development of decellularization techniques to produce muscle ECM-based scaffolds. Harnessing ECM biological properties, biodegradability and biocompatibility without the risk of immunogenic rejection may help the development of functionalized tissue-specific engineered scaffolds, for instance, SM matrices for muscle regeneration [25, 26].

In the present work, we have tested four different decellularization protocols for rat SM. Protocols involving the use of hypotonic solutions, enzymes and detergents that differed on reagent type, concentration and exposure time. Among the four tested protocols, only P4 was efficient, which proved to promote decellularization by all tested methods, satisfying the minimum criteria defined elsewhere [14].

Triton X-100 is a non-ionic detergent known to interact with lipids, disrupting their contact with other lipids and proteins [15]. Considering that P1, P2 and P4 protocols included Triton X-100, we propose that a higher concentration of Triton X-100, in this case, is more important than the time of exposition, since we used 1% and 2% Triton X-100 for 5 and 3 days in P2 and P4 decellularization protocols, respectively. In fact, despite P2 having demonstrated the capacity to decellularize quadriceps and hamstring rat muscles with 1% Triton X-100 [9], this protocol was modified in other studies, that used 3% Triton X-100 instead [27]. Under our conditions and in another study [28], tibialis anterior muscle decellularization was not achieved with P2. This may be due to different types of muscle and their size, or small nuances of the protocol. Variations in tissue composition and density as well as concentration and exposure time of decellularizing agents can modify the effectiveness of cell removal [12].

Another chemical compound commonly used in decellularization is sodium deoxycholate, an ionic surfactant used in protocol P3. In this work, however, sodium deoxycholate treatment was not able to remove myofibers, as shown in our histology and SEM results, despite DNA amount was reduced compared to control SM samples. This protocol, with minimal modifications, has been successfully applied [7, 8] for rat abdominal muscles; nevertheless, in other work, resulted in incomplete cell removal of rat, human or rabbit tibialis anterior and abdominal rectus muscles [10].

In decellularization, it is not rare to obtain discordant results after reproducing published protocols [10, 28, 29]. Such variability reflects the relevance of more detailed analysis and standardized procedures to design and apply tissue-engineered decellularized scaffolds. Additionally, reveals an urgent need to establish regulatory standards with quantifiable measures for specific dangerous agents [16].

Decellularization efficiency depends on tissue composition, density and the source from which it was extracted [12]. As proceeded for protocols P2, P3 and P4, cutting muscle tissue in thin slices may improve cell removal efficiency since it allows more exposure to decellularizing agents [9]. Another strategy could be perfusion, which has been employed for the heart [30], skeletal muscle [31] and even the whole human limb [32].

We believe that P4 gathered efficient methodologies of decellularization, which explains the good quality of our results. For instance, freeze-thawing muscle samples may have helped in disrupting cell membranes; and P4 was the only protocol that included the NaCl treatment step, which contributes to osmotic shock and cell lysis [12].

Compared to other studies that showed the absence of cellular components basically through one or two methods [7, 27], here we performed many different experiments to characterize our decellularized matrices. The investigation of decellularization efficiency is a crucial step because remnant cellular debris may result in low in vitro biocompatibility and host rejection in vivo [15]. Damage Associated Molecular Pattern molecules and other cell remnants released because of cell membrane disruption can influence the immune response to ECM scaffolds [16]. Also, compared to completely decellularized muscles, anuclear scaffolds with remaining myofibers are shown to have fewer satellite cell pool restoration, possibly because these myofibers hinder cell migration and correct orientation in vivo [11].

In general, all decellularization protocols cause changes in tissue’s original structure. The ideal protocol is the one that removes all cells and causes the least possible damage to the ECM [14]. We have demonstrated that mDMs from the P4 protocol preserved two main muscle ECM components, collagen I and laminin, as described in other works with other decellularization protocols [26, 29, 33], and partially the tubular endomysium structure, as shown by SEM.

Compared to commercially available ECM matrices, such as porcine urinary bladder matrix, or isolated ECM components, muscle-derived ECM supports better muscle regeneration, because of its tissue-specific physical and chemical cues [34, 35]. mDMs alone or seeded with specific cells have been used to successfully promote myogenesis and muscle regeneration in animal models for VML [35]. Also, recellularized scaffolds augment the number of cells and secretion of exogenous factors into the injury site [36].

In this work, we have performed mDMs recellularization with rat ASCs. Cells were isolated from rat inguinal fat and presented fibroblastic morphology, adherence to plastic culture plates, expression of characteristic surface markers and in vitro differentiation capacity, as described for human ASCs [20]. Recellularization of mDMs with ASCs showed good cytocompatibility with cell survival and adherence, as evidenced by MTT assay, Calcein staining and SEM after 7, 14 or 21 days of culture. ASCs cultivated in mDMs showed higher viability than control 2D culture, possibly because of the porosity and three-dimensional structure of mDMs, which increases cell growth surface area. Similarly, pig musculofascial ECM was demonstrated to be a compatible niche for human ASCs proliferation [23].

According to SEM analysis, ASCs adhered, emitted cytoplasmic extensions and were able to interact intimately with mDMs, especially at 21 days of culture. Similarly, other studies showed adherence to ASCs [23] and myoblasts [7] in mDMs.

Considering that, ASCs have potent paracrine activity, angiogenic and anti-inflammatory properties, and have multilineage potential in vitro, it is highly speculated their application for the treatment of musculoskeletal disorders [22]. Furthermore, ECM scaffolds induce a Th2 response [37], and have immunomodulatory and anti-inflammatory properties in vivo [38]. Fibroblasts co-cultured with satellite cells on mDMs have an important role in scaffolds colonization and in myogenic differentiation towards mature multinucleated myofibers [11]. Further studies are necessary to pursue whether ASCs in decellularized scaffolds could similarly cause these effects, which seems to be a good strategy for improving muscle regeneration.

In conclusion, one important and poorly explored approach is to recellularize mDMs with rat autologous ASCs directing them for preclinical studies; using rat models for VML or other muscle disorders. We have shown the manufacturing and characterization of rat-derived mDMs that were capable of supporting rat ASC’s survival. Our results highlight the need for an urgent standardization of decellularization methods in terms of technical reproducibility and characterization, which is particularly important before initiating preclinical studies. Although many issues still need to be clarified, this might be a great avenue for future applications using SM tissue engineering principles.

Abbreviations

ASCs

adipose-derived stem cells

ECM

Extracellular matrix

mDMs

muscle-decellularized matrices

MSCs

mesenchymal stem cells

SEM

scanning electron microscopy

SM

Skeletal muscle

VML

volumetric muscle loss