Login processing...

Trial ends in Request Full Access Tell Your Colleague About Jove

Developmental Biology

Preparation of 3D Decellularized Matrices from Fetal Mouse Skeletal Muscle for Cell Culture

Published: March 3, 2023 doi: 10.3791/65069


The extracellular matrix (ECM) plays a crucial role in providing structural support for cells and conveying signals that are important for various cellular processes. Two-dimensional (2D) cell culture models oversimplify the complex interactions between cells and the ECM, as the lack of a complete three-dimensional (3D) support can alter cell behavior, making them inadequate for understanding in vivo processes. Deficiencies in ECM composition and cell-ECM interactions are important contributors to a variety of different diseases.

One example is LAMA2-congenital muscular dystrophy (LAMA2-CMD), where the absence or reduction of functional laminin 211 and 221 can lead to severe hypotony, detectable at or soon after birth. Previous work using a mouse model of the disease suggests that its onset occurs during fetal myogenesis. The present study aimed to develop a 3D in vitro model permitting the study of the interactions between muscle cells and the fetal muscle ECM, mimicking the native microenvironment. This protocol uses deep back muscles dissected from E18.5 mouse fetuses, treated with a hypotonic buffer, an anionic detergent, and DNase. The resultant decellularized matrices (dECMs) retained all ECM proteins tested (laminin α2, total laminins, fibronectin, collagen I, and collagen IV) compared to the native tissue.

When C2C12 myoblasts were seeded on top of these dECMs, they penetrated and colonized the dECMs, which supported their proliferation and differentiation. Furthermore, the C2C12 cells produced ECM proteins, contributing to the remodeling of their niche within the dECMs. The establishment of this in vitro platform provides a new promising approach to unravel the processes involved in the onset of LAMA2-CMD, and has the potential to be adapted to other skeletal muscle diseases where deficiencies in communication between the ECM and skeletal muscle cells contribute to disease progression.


The extracellular matrix (ECM) is a major constituent of tissues, representing their non-cellular component. This three-dimensional (3D) structure not only provides physical support for cells, but also plays a crucial role in the biochemical processes involved in the development of organisms1. The formation of a tissue-specific ECM occurs during development, as a result of the complex interactions between cells and their niches, influenced by various intra- and extracellular stimuli. The ECM is a highly dynamic structure that undergoes chemical and mechanical rearrangements in a temporal-spatial manner and directly impacts cell fate2. One of the most notable characteristics of the ECM is its functional diversity, as each tissue ECM displays a unique combination of molecules that provide different topologies and properties that are tailored to the cells it contains1.

ECM signaling and support are crucial for development and homeostasis, and when disrupted can lead to multiple pathological conditions3,4. One example is LAMA2-deficient congenital dystrophy (LAMA2-CMD), which is the most common form of congenital muscular dystrophy. The LAMA2 gene encodes for the laminin α2 chain, which is present in laminin 211 and laminin 221, and when mutated can lead to LAMA2-CMD5. Laminin 211 is the main isoform found in the basement membrane surrounding skeletal muscle fibers. When laminin 211 is abnormal or absent, the link between the basement membrane and muscle cells is disrupted, leading to the onset of the disease6. Patients with LAMA2-CMD show a mild to severe phenotype depending on the type of mutation in the LAMA2 gene.

When the function of the laminin α2 protein is affected, patients can experience severe muscle hypotonia at birth and develop chronic inflammation, fibrosis, and muscle atrophy, leading to a reduced life expectancy. To date, no targeted treatments have been developed and therapeutic approaches are limited to alleviating the symptoms of the disease7. Therefore, understanding the underlying molecular mechanisms involved in the onset of this disease is crucial for developing appropriate therapeutic strategies6,8. Previous work using the dyW mouse9, a model for LAMA2-CMD, suggests that the onset of the disease starts in utero, specifically during fetal myogenesis10. A better understanding of how the fetal myogenesis defect emerges would be a game changer in generating novel therapeutic approaches for LAMA2-CMD.

In vitro systems provide a controlled environment for studying cell-cell and cell-ECM interactions, but 2D culture models lack the complexity of native tissues. Decellularization of tissues produces tissue- and developmental stage-specific acellular ECM scaffolds that more accurately mimic the natural cell microenvironment compared to 2D models and engineered/synthetic scaffolds. Decellularized matrices (dECMs) have the potential to preserve the molecular and mechanical cues of the host tissue, making them better alternative models for understanding in vivo processes11.

There are various techniques, reagents, and conditions that can be used for decellularization12,13. In this study, a decellularization protocol for the fetal mouse heart, described by Silva et al.14,15, is adapted to fetal mouse skeletal muscle and found to retain all tested ECM components (laminin α2, total laminins, fibronectin, collagen I, and collagen IV). The protocol includes three steps: cell lysis by osmotic shock (hypotonic buffer), plasma membrane dissolution and protein dissociation (0.05% sodium dodecyl sulfate [SDS]), and enzymatic destruction of DNA (DNase treatment). To our knowledge, this is the first established protocol for decellularizing mouse fetal skeletal muscle.

To use this 3D in vitro system for studying LAMA2-CMD, it is crucial to maintain the laminin α2 chain after decellularization. Therefore, an optimization protocol was implemented where different detergents (SDS and Triton X-100) and concentrations (0.02%, 0.05%, 0.1%, 0.2%, and 0.5%) were tested (data not shown). The optimal choice for cell removal and preservation of the laminin α2 protein was found to be 0.05% SDS. C2C12 cells, a well-established myoblast cell line16,17, were used to seed the dECMs. These cells invade the dECM, proliferate, and differentiate inside these scaffolds, synthesizing new ECM proteins. The successful production of this 3D in vitro model offers a new approach to understanding the molecular and cellular processes involved in fetal myogenesis, the onset of LAMA2-CMD, and can be extended to other muscle diseases where the communication between the ECM and skeletal muscle cells is disrupted.

Subscription Required. Please recommend JoVE to your librarian.


All the methodologies described were approved by the Animal Welfare Committee (ORBEA) of the Faculty of Sciences, University of Lisbon, and Direção Geral de Veterinária (DGAV; ref. 0421/000/000/2022), and are in accordance with the European Directive 2010/63/EU.

1. Preparation of decellularization buffers and reagents

NOTE: All solutions used during the decellularization protocol should be sterilized by autoclaving and stored for up to 3 months unless stated otherwise.

  1. Prepare 10x phosphate-buffered saline (10x PBS) by adding sodium chloride (NaCl) at 137 mM, potassium chloride (KCl) at 2.68 mM, potassium-dihydrogen phosphate (KH2PO4) at 8.1 mM, and disodium-hydrogen-phosphate dihydrate (Na2HPO4) at 1.47 mM to demineralized water (dH2O) and adjust the pH to 6.8. Add 100 mL of 10x PBS to 900 mL of demineralized H2O to generate 1x PBS. Autoclave and store at room temperature (RT). Add sterile penicillin/streptomycin stock solution (10,000 U/mL penicillin and 10 mg/mL streptomycin [pen/strep]) to a final concentration of 1% prior to use.
  2. Prepare the hypotonic buffer by dissolving 1.21 g of Tris(hydroxymethyl)aminomethane (Tris base) and 1 g of ethylenediaminetetraacetic acid (EDTA) in 1 L of dH2O (10 mM Tris base 0.1% EDTA). Use a magnetic stirrer until dissolved and adjust the pH to 7.8 with NaOH/HCl. Sterilize by autoclaving and store at RT. Add pen/strep to a final concentration of 1% prior to use.
  3. Prepare the hypotonic wash buffer by dissolving 1.21 g of Tris base in 1 L of dH2O (10 mM Tris base). Use a magnetic stirrer until dissolved and adjust the pH to 7.8 with NaOH/HCl. Autoclave and store at RT. Add pen/strep to 1% prior to use.
  4. Prepare the anionic detergent treatment by adding 0.05 g of sodium dodecyl sulfate (SDS) to 100 mL of hypotonic wash buffer to produce a 0.05% SDS treatment solution. Filter through a 0.22 µm membrane filter. Store at RT for up to 1 month.
    NOTE: The SDS solution should not be autoclaved, as SDS precipitates and degrades upon autoclaving.
  5. Prepare the DNase treatment solution by dissolving 1.21 g of Tris base in 0.5 mL of dH2O and add 1 mL of 1 M magnesium chloride (MgCl2). Adjust the pH to 7.8 with NaOH/HCl. Autoclave and store at RT. Add DNase I prior to use (50 U/mL).

2. Sample collection

NOTE: Wild-type C57/BL6 mice were utilized in the study. All techniques were conducted in a laminar flow hood under sterile conditions.

  1. Euthanize adult pregnant female mice at embryonic day (E) 18.5 of pregnancy using isoflurane inhalation followed by cervical dislocation. Subject the mice to terminal dissection to extract the fetuses.
    1. Spray the abdomen of the mouse with 70% ethanol.
    2. Using surgical forceps and scissors, lift a fold of skin in the lower abdomen and make a U-shaped incision to expose the abdominal cavity18.
    3. Collect the uterine horns containing the E18.5 fetuses by cutting the oviducts and cervix and immediately transfer them to a Petri dish with ice-cold 1x PBS with 1% pen/strep18.
    4. Quickly remove the fetuses from the uterus and extraembryonic tissues and euthanize them by decapitation using scissors18.
  2. Transfer one fetus at a time to a Petri dish with ice-cold 1x PBS. Using a stereo microscope, remove the skin and cut off the limbs. From the ventral side, cut through the ribcage and remove the sternum and the underlying organs.
  3. Flip the fetal trunk dorsal side up. Remove the cervical portion of the vertebral column, the dorsal fat deposits, and the connective tissue on top of the deep back muscles.
  4. Use surgical forceps to anchor the ribcage and carefully scrape and detach the deep back muscles from the surrounding tissues using a micro scalpel10. This procedure results in the isolation of two muscle pieces per fetus (left and right), both primarily composed of longissimus and iliocostalis muscles, with some of the transversospinalis and levator costarum muscles included.
  5. Store the muscle samples in 1x PBS with 1% pen/strep at 4 °C for short-term storage (<1 week), or at -80 °C for longer periods.
    ​NOTE: Use freshly collected samples for best results. Samples frozen for extended periods (>3 months) show lower decellularization efficiency and more protein degradation. Epaxial muscle masses were used for the decellularization experiments, but other muscles (e.g., limb muscles) can be processed for decellularization using the same protocol.

3. Fetal skeletal muscle decellularization

NOTE: All techniques were conducted in a laminar flow hood under sterile conditions. For a detailed schematic representation, see Figure 1A. All steps were performed with agitation in an orbital shaker with a diameter of 120 mm (165 rpm) at 25 °C unless stated otherwise. Add 1% pen/strep to solutions before use. When removing the solutions, aspirate carefully to avoid sample entrapment in the pipette.

  1. Day 1 - Prepare approximately 2 mm x 1 mm x 1 mm (~3.5 mg) tissue fragments from the epaxial muscle masses. Add 3 mL of hypotonic buffer with 1% pen/strep to each well of a 12-well plate and add one whole muscle tissue fragment per well.
    1. Incubate the tissue fragments in hypotonic buffer with agitation for 18 h (overnight) (O/N).
  2. Day 2 - Aspirate the hypotonic buffer using a fine-tip pipette and wash the samples 3 x 1 h with 3 mL of 1x PBS each time with agitation.
    1. Incubate the samples for 24 h with 3 mL of 0.05% SDS detergent solution with agitation.
      NOTE: (CHECK POINT) After SDS incubation, the samples should be transparent in appearance. The samples display a gelatin-like consistency and are, therefore, more prone to adhere to the pipette when removing the liquids.
  3. Day 3 - Remove the SDS detergent solution using a fine-tip pipette and wash the tissue fragments 3 x 20 min with 3 mL of hypotonic wash buffer each time with agitation.
    NOTE: (PAUSE POINT) The samples can be maintained in hypotonic wash buffer at 4 °C for 18 h (O/N). Ensure SDS is well removed during washing steps, as residual SDS can be cytotoxic.
    1. Incubate the tissue fragments for 3 h with 2 mL of DNase solution with agitation at 37 °C.
    2. Remove the DNase solution using a fine-tip pipette and wash the tissue fragments 3 x 20 min with 3 mL of 1x PBS each time with agitation. Finally, wash O/N with agitation (60 rpm).
      NOTE: After the DNase treatment, the dECMs become sticky due to the presence of DNA residues. Therefore, careful washing is necessary.
    3. Store the dECMs in 1x PBS with 1% pen/strep at 4 °C until recellularization (<1 week). For other uses, store at -80 °C.

4. Decellularization quality assessment

NOTE: DNA quantification, DAPI/methyl green staining, and phalloidin staining were performed to evaluate the presence of residual cell contents after decellularization. Immunohistochemistry and western blot analyses were conducted to assess the retention of key ECM proteins after decellularization.

  1. Quantification of the DNA present in the dECM
    NOTE: The dECMs should be compared to the native tissue to detect the presence of DNA.
    1. Prior to decellularization, weigh a 1.5 mL microcentrifuge tube on a high precision digital scale. Before transferring the muscle sample to the tube, remove all the remaining 1x PBS using a paper towel. Weigh the tube with the sample. Calculate the wet weight of the samples using equation (1):
      ​Samplewet weight = weighttube with samples- weightempty tube      (1)
    2. Decellularize the samples following the protocol described in section 3.
    3. Place the samples in 2 mL microcentrifuge tubes.
      NOTE: The samples can be kept at -20 °C until DNA extraction and quantification.
    4. Perform DNA extraction of the dECMs and native tissue samples using a spin column-based kit. Add the digestion buffer according to the manufacturer's instructions.
    5. Homogenize the samples in a bead mill twice for 2.5 min each time (flipping the mounts) using one tungsten carbide bead per tube (use chilled tube mounts).
    6. Add proteinase K and incubate O/N with slow agitation at 56 °C.
    7. Proceed with the protocol as described in the manufacturer's instructions.
    8. Elute with the buffer provided by the manufacturer and centrifuge 2 x 1 min at ≥6,000 x g, each time at 4 °C to increase the DNA yield.
      NOTE: DNA can be stored at -20 °C until quantification.
    9. Quantify the DNA present in the samples using a fluorescent dsDNA detection kit following the manufacturer's instructions.
    10. Normalize the DNA content as nanograms of DNA per milligram of original wet weight of sample.
    11. Analyze the data using a two-tailed Student's t-test and express as the mean ± standard error of the mean (SEM).
  2. Immunohistochemistry
    NOTE: Solutions 1, 2, and 3 and the fixative solution should be prepared before use and can be kept frozen for up to 6 months. All antibodies and dyes used and respective dilutions are listed in Table 1.
    1. Preparation of solutions
      1. Prepare fixative solution by adding 2 g of paraformaldehyde (PFA), 8 g of sucrose, and 24 µL of 1 M CaCl2 to 77 mL of 0.2 M Na2HPO4 and 23 mL of 0.2 M NaH2PO4, to a final volume of 200 mL by adding dH2O. Adjust the pH to 7.4.
      2. Prepare 0.12 M phosphate buffer by adding 13.5 g of Na2HPO4 and 3.2 g of NaH2PO4 to 1 L of dH2O. Adjust the pH to 7.4.
      3. Prepare solution 1 by adding 4 g of sucrose to 100 mL of 0.12 M phosphate buffer.
      4. Prepare solution 2 by adding 15 g of sucrose to 100 mL of 0.12 M phosphate buffer.
      5. Prepare solution 3 by adding 15 g of sucrose and 7.5 g of gelatin to 100 mL of 0.12 M phosphate buffer. Heat at 37 °C until the gelatin dissolves.
      6. Prepare methyl green stock solution (2% methyl green powder dissolved in dH2O)19.
    2. Immunodetection
      1. Fix the samples using the fixative solution for at least 4 h at 4 °C.
      2. Wash the samples 2x for 10 min with 1x PBS.
      3. Keep O/N, or over 1 day, in solution 1 at 4 °C.
      4. Wash and keep O/N or over 1 day in solution 2 at 4 °C. Let the samples warm to RT.
      5. Incubate for 3 h in solution 3 at 37 °C. Keep extra solution 3 at 37 °C to be used later.
      6. Mold small (2 cm x 1 cm x 1 cm) containers in aluminum foil. Place a thin layer of solution 3 in the molded container and let it set. Place the samples on the solidified solution 3 and cover with warm solution 3. Orientate the samples and let them solidify. Mark the location on the solidified solution 3 with a color pen.
      7. Freeze by placing the containers on the surface of dry ice-chilled or liquid nitrogen-chilled isopentane.
      8. Using optimal cutting temperature (O.C.T.) compound, fix the frozen gelatin cubes containing the samples to the cryostat mount.
      9. Section the frozen gelatin cubes and transfer the tissue sections to the slides. Let them dry for 60 min.
      10. Use a hydrophobic marker to trace a line around the sections.
      11. Wash the slides 3 x 10 min in 1x PBS.
      12. Cover the sections with 1% bovine serum albumin (BSA), 1% goat serum, and 0.05% Triton X-100 diluted in 1x PBS (blocking solution) for 30 min (blocking step).
      13. Dilute the primary antibodies in blocking solution and cover the sections. Incubate O/N at 4 °C in a closed box, with moist paper, to prevent the sections from drying.
      14. Wash the slides 3 x 10 min with 1x PBS.
      15. Dilute the secondary antibodies in blocking solution and cover the sections. Incubate for 1.5 h at RT in the dark.
        NOTE: Avoid light exposure to prevent fluorophore degradation.
      16. Wash the slides 3 x 10 min with 4x PBS.
        NOTE: Higher concentration of PBS can be used when a strong background is present.
      17. Submerge the slides in DAPI solution (5 µg/mL 1,4-diazabicyclo-2,2,2-octane in 0.1% Triton X-100/1x PBS) for 30 s each.
      18. Rinse in 1x PBS.
      19. Mount the sections in anti-fading medium (50 mg/mL n-propyl-gallate in 1x PBS:glycerol [1:9]) and seal with a coverslip. Store at 4 °C until observation and image acquisition in a fluorescence microscope20.
        NOTE: For the in toto immunohistochemistry experiments, the same protocol (see steps was used, except the samples were previously fixed in 4% PFA diluted in 1x PBS for 3 h at RT. DNA staining was performed by adding methyl green to the secondary antibody solution. All antibodies and dyes used, and their respective dilutions, are listed in Table 1. Samples were then mounted in anti-fading medium between two coverslips separated by steel rings, glued together with beeswax, and 100 µm image stacks were acquired using a confocal microscope21.
  3. Western blot analysis
    NOTE: All antibodies used and respective dilutions are listed in Table 1.
    1. Preparation of solutions
      1. Prepare the 2x SDS-PAGE loading buffer by adding 20 mL of glycerol, 4 g of SDS, and 0.2 mL of bromophenol blue to 80 mL of 100 mM Tris base solution. Adjust the pH to 6.8. Add fresh dithiothreitol (DTT) prior to use.
      2. Prepare the Tris buffered saline wash buffer with Tween 20 (TBST) solution by adding 2.4 g of Tris base, 8.8 g of NaCl, and 1 mL of Tween-20 to dH2O to a final volume of 1 L. Adjust the pH to 7.4-7.6 using HCl.
      3. Prepare the 10x running buffer (RB) by adding 30.2 g of Tris base, 144.2 g of glycine, and 10 g of SDS to 1 L of dH2O. Before use, add 100 mL of 10x RB to 900 mL of dH2O to prepare 1x RB (working solution).
        NOTE: While 10x RB can be stored at RT for several months, 1x RB can be reused on different runs.
      4. Prepare the transfer buffer (TB) by adding 5.82 g of Tris base, 2.93 g of glycine, and 0.5 g of SDS to 800 mL of dH2O. After the components are dissolved, add 200 mL of methanol. Chill at 4 °C.
        NOTE: The TB should be prepared freshly before every use.
    2. Protein extraction
      1. Collect epaxial muscles from E18.5 mice and immediately place them in a microcentrifuge tube (2 mL) containing 2x SDS-PAGE loading buffer with freshly added DTT (100 mM DTT). Process E18.5 dECM in the same way.
      2. Add one tungsten carbide bead per tube. Homogenize 2x in a bead mill for 2.5 min each time (flip the mounts).
      3. Sonicate the samples for 5 min in an ultrasound bath.
      4. Heat the samples for 10 min at 50 °C.
      5. Centrifuge the samples at 12,000 × g for 15 min at 4 °C.
      6. Transfer the supernatant into a fresh 1.5 mL microcentrifuge tube.
      7. Quantify the protein using a microvolume spectrophotometer.
      8. Store at -20 °C until further use.
    3. Polyacrylamide gel electrophoresis
      1. Use precast acrylamide gradient gel (4%-20%)
      2. Mount the gel in the electrophoresis tank. Remove the comb carefully. Add 1x RB until the line displayed in the electrophoresis tank. Make sure the wells are filled with RB.
      3. Load 100 µg of protein per sample in the wells. Load 12 µL of high molecular weight protein standard.
      4. Run for a total of 100 min with constant voltage (10 min at 150 V and 90 min at 185 V).
    4. Transfer
      1. Soak the filter pads and sponges with chilled TB (4 °C) while the gel is running.
      2. Cut the polyvinylidene difluoride membranes to the size of the gel. Activate them with methanol and wash with dH2O. Soak in the TB.
      3. Mount the transfer cassette with the sponges, filter pads, gels, and activated membranes according to the manufacturer's instructions.
      4. Place the cassette in the electrophoresis tank containing the chilled TB (on a bed of ice). Add the cooling unit. Run for 90 min at 100 V.
      5. After the transfer, stain with a Coomassie blue substitute to assess the transfer quality.
    5. Immunodetection
      1. Prepare the blocking solution (TBST with 5% low-fat milk). Incubate the membranes with agitation for 1 h at RT.
      2. Rinse 3 x 5 s with TBST.
      3. Incubate the membranes O/N with the primary antibodies diluted in TBST with 2% BSA and 0.02% sodium azide (3 mL per membrane) in a cold chamber (4 °C) with agitation.
      4. The next day, wash 3 x 5 min with TBST.
      5. Incubate the membranes with horseradish peroxidase (HRP)-conjugated secondary antibodies diluted in TBST with 5% low-fat milk (5 mL for each membrane) for 1 h at RT.
      6. Wash the membranes with TBST 3 x 5 min. Keep in TBST until the detection step.
      7. Visualize the protein using a commercially available developing kit. Add detection reagents according to the manufacturer´s instructions. Acquire images of the bands.
      8. To test more than one antibody per membrane, after the detection step, strip the former antibodies with three washes (5 min each) using TBST and repeat steps

Table 1: Antibodies and dyes used in immunohistochemistry and western blot analysis and the respective dilutions. Please click here to download this Table.10,22

5. Cell culture in decellularized matrices

NOTE: All the techniques were performed under sterile conditions in a laminar flow hood. All incubations were performed at 37 °C and with 5% CO2.

  1. Prepare the cell culture medium, high glucose Dulbecco's Modified Eagle Medium with stable glutamine and with sodium pyruvate (DMEM), supplemented with 10% fetal bovine serum (FBS) and 1% pen/strep (complete culture medium).
  2. Place the dECMs in a Petri dish in 1x PBS with 1% pen/strep in a laminar flow hood and separate them into pieces of approximately 500 µm x 500 µm x 250 µm, using a micro scalpel and tweezers.
    NOTE: dECMs have a soft texture, and therefore it is often easier to use tweezers to separate them into approximately same sized pieces instead of cutting them with the micro scalpel.
  3. Transfer the fragments into a 96-well plate (three or four pieces per well) with 200 µL of complete culture medium, previously warmed to 37 °C. Incubate for 2 h in the incubator.
  4. Add 500 µL of trypsin to a T25 flask seeded with sub-confluent (~70%) C2C12 cells. Resuspend in 1 mL of complete medium.
  5. Add 10 µL of the resuspension to 10 µL Trypan blue dye and load into a hemocytometer. Count and calculate the cell number and confirm cell viability.
  6. Aspirate the medium from the 96-well plate containing the dECMs and add 200 µL of complete culture medium containing 50,000 viable C2C12 cells.
  7. Incubate for 2 days and then, using tweezers, transfer the dECMs (with the cells) to a 48-well plate with 400 µL of complete culture medium. Every 2 days, carefully aspirate the medium with a micropipette to prevent the dECMs from detaching from the bottom of the well and add fresh medium until day 8.
    NOTE: Matrices are kept for 2 days in 96-well plates to promote C2C12 cell adhesion to the dECMs, and are then transferred to a 48-well plate to allow access to a larger volume of medium with nutrients. dECMs should adhere to the bottom of the wells.
  8. For the differentiation experiment, replace the complete culture medium with differentiation medium (DMEM supplemented with 2% horse serum and 1% pen/strep) on day 8, followed by incubation in differentiation medium for 4 days until day 12.

Subscription Required. Please recommend JoVE to your librarian.

Representative Results

The goal of the decellularization protocol is to produce dECMs that closely resemble the composition of native tissue. To determine the effectiveness of the decellularization process, various methods were employed, including examination of tissue morphology, measurement of DNA levels, staining for F-actin, and analysis of key ECM components using immunohistochemistry and western blotting techniques. Specifically, five major ECM components of skeletal muscle tissues were analyzed.

Throughout the protocol, samples change in appearance (Figure 1B1-4). After isolation, the muscle tissue appears reddish due to the presence of myoglobin (Figure 1B1). After incubation in the hypotonic buffer, which lyses cells and removes most of the cytoplasmatic proteins, the samples turn white (Figure 1B2). SDS, an anionic detergent commonly used in tissue decellularization12, effectively removes cytoplasmic and nuclear material, as well as membranes. As a result, the samples become more transparent after SDS treatment (Figure 1B3). However, high concentrations or prolonged exposure to this detergent can cause protein denaturation, loss of glycosaminoglycans, and disruption of collagen fibers12. The optimal balance between cell removal and preservation of the ECM is achieved using 0.05% SDS for 24 h, as shown in Figure 2 and Figure 3. Finally, DNA is removed through DNase treatment, resulting in transparent dECM scaffolds that are slightly smaller than after SDS treatment (Figure 1B4).

The success of the decellularization protocol is indicated by the amount of residual DNA present in the matrices after the process. DNA quantification reveals a nearly 100% decrease in the dECMs compared to the native tissue (Figure 1C). DAPI staining also confirms the absence of DNA in the dECMs (Figure 2B,D,F,H,J).

The presence of five major ECM proteins was evaluated by immunohistochemistry and by western blot after decellularization and compared to the native tissue. Immunostaining for the laminin α2 subunit (Figure 2A,B) and total laminins (Figure 2C,D) shows that the dECMs display a tubular staining for these proteins (arrows in Figure 2B,D), similar to the laminin matrix surrounding myotubes in the native tissue (arrows in Figure 2A,C). Western blot analysis for laminin α2 subunit reveals the presence of two bands in the native tissue (Figure 2A'), while three bands, with a smaller molecular weight than expected, can be detected in the dECM (Figure 2B'). This may be the result of protein degradation or removal during the decellularization process. Western blot analysis for total laminins shows some fragmentation of these proteins in the dECM compared to samples from the native tissue (Figure 2C',D').

Figure 1
Figure 1: Fetal skeletal muscle decellularization procedure and evaluation of tissue morphology and DNA content. (A) Schematic representation of the decellularization protocol. (B) Tissue morphology throughout the protocol, from freshly collected to decellularized tissue. Scale bar = 1 mm. (C) Quantification of DNA present in the dECMs compared to the native tissue. Data expressed as mean ± SEM. Student's t-test, two-tailed **p < 0.01. The decellularization protocol efficiently removes nuclear content producing acellular dECMs. Abbreviations: dECMs = decellularized matrices; PBS = phosphate-buffered saline; SDS = sodium dodecyl sulfate; NT = native tissue. Please click here to view a larger version of this figure.

Immunostaining for fibronectin (Figure 2E,F) and collagen I (Figure 2G,H) shows the presence of these proteins in the interstitial space between cells in the native tissue (arrows in Figure 2E,G) and a similar staining in the dECMs (arrows Figure 2F,H). Western blot analysis shows similar bands for fibronectin in both conditions (Figure 2E',F'), indicating that this protein is not particularly affected by the decellularization process. However, in the case of collagen I (Figure 2G',H') there are fewer bands in the dECMs, indicating some degree of degradation. Immunostaining for collagen IV shows a tubular staining pattern in the native tissue (arrow in Figure 2I) and a similar staining in the dECM, although the tubular structures are narrower (arrow in Figure 2J). Western blot analysis for collagen IV reveals the presence of three bands in the native tissue (Figure 2I'). While the same three bands are observed in the dECMs (Figure 2J'), the molecular weight of these is lower than expected. Similarly to laminin α2, this can be the result of protein degradation or removal during the decellularization.

dECMs are then seeded with C2C12 myoblasts and cultured for 8 days in complete culture medium. C2C12 cells colonize the dECMs and proliferate, as shown by phospho-histone 3 nuclear staining (arrows in Figure 3A). After an additional 4 days of culture in differentiation medium, C2C12 cells differentiate and fuse into multinucleated (blue arrows in Figure 3B) myotubes expressing a myosin heavy chain (dashed line in Figure 3B). Interestingly, intracellular and/or pericellular staining for the laminin α2 chain (yellow arrows in Figure 3C,D), total laminins (magenta arrow in Figure 3C), and fibronectin (magenta arrow in Figure 3D) can be detected in C2C12 cells cultured in complete culture medium, suggesting that these cells are able to synthesize these ECM proteins de novo, therefore contributing to the formation of their niche. These results demonstrate that the decellularization protocol generates a dECM microenvironment where C2C12 myoblasts can proliferate, differentiate, and form multinucleated myotubes.

Figure 2
Figure 2: Assessment of five ECM proteins in native versus decellularized E18.5 fetal muscle tissue. Immunohistochemistry on sections of native tissue (A,C,E,G,I) and staining of dECMs (B,D,F,H,J) with DAPI, a DNA marker, phalloidin detecting F-actin, and for ECM proteins. Scale bar = 15 µm. In the native tissue, immunostaining for laminins and collagen IV is present surrounding the myofibers (A,C,I; yellow arrows) and staining for fibronectin and collagen I is detected in the interstitial space between myofibers (E,G; yellow arrows). In the dECMs, staining for laminins and collagen IV (B,D,J; yellow arrows) show a tubular pattern, surrounding the spaces left by the myofibers after decellularization. Fibronectin and collagen I immunostaining of dECMs is consistent with their presence in the interstitial space (F,H; yellow arrows). (A'-J') Western blot analysis for five ECM proteins in native tissue (A',C',E',G',I') and in dECM samples (B',D',F',H',J'). Both approaches show protein preservation after decellularization. Antibodies used and respective dilutions are listed in Table 1. Abbreviations: LNα2 = laminin α2 chain; LNp = pan-muscle laminins; FN = fibronectin; Col I = collagen I; Col IV = collagen IV; MHC = myosin heavy chain. Please click here to view a larger version of this figure.

Figure 3
Figure 3: dECM recellularization with a myoblast cell line (C2C12 myoblasts). (A) Maximum intensity projection of a confocal image of a 100 µm stack of recellularized matrix, colonized with C2C12 cells labelled with methyl green staining DNA, phalloidin detecting F-actin, and anti-pH3 antibody, a proliferation marker (magenta arrows). Scale bar = 35 µm. (B) Maximum intensity projection of a confocal image of a 100 µm stack showing a multinucleated (blue arrows indicate the nuclei) myosin heavy chain-positive myotube (dashed line) formed during culture. Scale bar = 35 µm. (C,D) Immunohistochemistry confocal image showing intra- or peri-cellular staining for laminin α2 chain (yellow arrows), total laminins (magenta arrows in C), and fibronectin (magenta arrows in D) in myoblasts, suggesting de novo protein synthesis. Scale bar = 10 µm. Antibodies used and respective dilutions are listed in Table 1. Abbreviations: pH3 = phospho-histone 3; LNα2 = laminin α2 chain; LNp = pan-muscle laminins; FN = fibronectin; MHC = myosin heavy chain. Please click here to view a larger version of this figure.

Subscription Required. Please recommend JoVE to your librarian.


The ECM is a complex network of macromolecules that is present in all tissues and plays a crucial role in regulating cell behavior and function2. The ECM acts as a physical scaffold for cells to attach to and provides cues that actively modulate cellular processes such as proliferation, motility, differentiation, and apoptosis. Thus, proper formation and maintenance of the ECM is essential for both development and homeostasis1.

While 2D cell culture models have been widely used, they are increasingly being replaced by more advanced 3D platforms. This is because 2D cultures lack the chemical and physical cues that affect cell behavior, while 3D cultures are considered a more realistic alternative for studying molecular and cellular dynamics in native tissues11. Decellularization of tissues results in the production of scaffolds that more closely mimic the microenvironments of biological tissues, as demonstrated in a number of studies across various tissue or organ systems14,15,23,24,25. The ability of dECMs to replicate native tissue microenvironments holds great potential for research on normal development, various disease states, and the effect of drugs or toxins on tissues11.

The protocol used in this study builds on the protocol developed by Silva et al. for decellularizing fetal heart tissue14 as a starting point. It involves a combination of a hypotonic buffer, treatment with an anionic detergent (SDS), and a DNase treatment. One of the main challenges in decellularization protocols is finding a balance between removing cells and preserving the ECM protein composition. Given our focus on using this 3D cell culture system to study the early stages of LAMA2-CMD, special attention was paid to preserving laminin 211 during decellularization. The protocol of Silva et al.14 led to the loss of laminin α2 chain immunoreactivity in the decellularized fetal muscles; this could be due to the concentration of SDS used. Therefore, alternatives to the 0.2% SDS detergent solution step were tested, such as lower concentrations of SDS (0.1%, 0.05%, and 0.02%) and the substitution of SDS with different concentrations of Triton X-100 (0.5% and 0.2%). The best results were achieved by using 0.05% SDS detergent treatment for 24 h. This concentration effectively removed cell contents while preserving laminin α2 chain immunoreactivity after decellularization. This protocol reproducibly produces acellular dECMs that are free of cellular residues, including DNA.

The protocol used in this study preserves both the interstitial matrix proteins (fibronectin and collagen I) and basement membrane proteins (laminins and collagen IV). Future studies should assess whether collagen VI is also preserved, as it is also a player in muscular dystrophies26. It is known that SDS can disrupt protein ultrastructure and damage collagens12; for fetal skeletal muscle, it was important to use a low concentration of SDS (0.05%) to maintain laminin α2 immunoreactivity. However, the western blot results show that the decellularized samples display more bands after immunodetection of laminins and collagens compared to the native tissue, indicating that some protein degradation occurred as a result of the decellularization process27.

Importantly, the recellularization experiments demonstrate that these matrices are reliable scaffolds that consistently support cell adhesion, proliferation, and differentiation. SDS has been reported to be cytotoxic28, and therefore the washing steps included in the protocol are crucial if the matrices are to be used for recellularization. These scaffolds were effectively colonized by C2C12 cells, indicating their suitability as a model system for 3D culture of cells. The observation of intracellular and pericellular staining for ECM proteins surrounding the C2C12 cells further suggests that the cells are actively contributing to their microenvironment within the dECMs. Additionally, when placed in differentiation medium, the C2C12 cells differentiated, fused, and formed myotubes within the dECMs.

A significant challenge in this procedure is the manipulation of the samples throughout the protocol. The samples are very small and soft, requiring care and skill in handling to prevent entrapment in the fine-tip pipette and loss of the samples. The best results are obtained when starting the protocol with fresh tissue. Using frozen, stored samples can impede cellular content removal and lead to increased protein degradation, hindering protein detection and reducing decellularization efficiency.

Only a few studies have reported the decellularization of fetal tissues15,29,30,31. Specifically, regarding the decellularization of fetal skeletal muscle, only one previous study has reported on the decellularization of composite samples of dermis, subcutaneous tissue, and panniculus carnosus31. To the best of the authors' knowledge, this is the first time that a decellularization protocol for isolated fetal mouse skeletal muscle has been established. This protocol can serve as a foundation for the creation of similar protocols for fetal muscle tissues of other species, such as pigs and humans13.

The present protocol for decellularizing E18.5 mouse skeletal muscle is very similar to the procedure of Silva et al.14, who applied it to an E18 mouse heart. The only difference is the concentration of SDS used, which is considerably lower than the one used for the fetal heart (0.05% vs. 0.2%), possibly due to different physical properties of these two fetal tissues.

The development of this in vitro model not only allows for study of the processes involved in normal fetal muscle development, but also enables the parallel investigation of early onset muscular dystrophies and myopathies, such as LAMA2-CMD, which manifests as a myogenesis defect at E18.5 in the dyW mouse model10. However, it should be noted that this system is limited in that it only includes the ECM and muscle cells and does not contain other cell types such as neurons, endothelial cells, and fibroblasts. The relevance of these additional cell types may vary depending on the disease, and modifications to the culture system may be needed to include them. Overall, the use of dECMs as described in this study can be applied in the study of various early onset muscular dystrophies and myopathies.

Subscription Required. Please recommend JoVE to your librarian.


The authors have no conflicts of interest to disclose.


This work was funded by the Association Française contre les Myopathies (AFM-Téléthon; contract no. 23049), the MATRIHEALTH project, and cE3c unit funding UIDB/00329/2020. We would like to thank our donor Henrique Meirelles who chose to support the MATRIHEALTH Project. This work benefitted from the infrastructures of the Faculty of Sciences Microscopy Facility, a node of the Portuguese Platform of BioImaging (reference PPBI-POCI-01-0145-FEDER-022122), and we thank Luís Marques for his assistance with image acquisition and processing. Finally, we thank Marta Palma for technical support and our research team for their generous contributions.


Name Company Catalog Number Comments
12 Well Cell Culture Plate, Flat, TC, Sterile Abdos Labware P21021
4′,6-Diamidino-2-phenylindole dihydrochloride Merck D8417
4–20% Mini-PROTEAN TGX Precast Gel Bio-Rad 4561093
48 Well Cell Culture Plate, Flat, TC, Sterile Abdos Labware P21023
96 Well Cell Culture Plate, Flat, TC, Sterile Abdos Labware P21024
Bovine Serum Albumin, Fraction V NZYtech MB04601
BX60 fluorescence microscope Olympus
Cryostat CM1860 UV Leica
Dithiothreitol ThermoFisher R0862
DMEM high glucose w/ stable glutamine w/ sodium pyruvate Biowest L0103-500
DNase I PanReac AppliChem A3778
DNeasy Blood & Tissue Kit Qiagen 69506
Ethylenediaminetetraacetic acid (EDTA) Merck 108418
Fetal bovine serum Biowest S1560-500
Fine tip transfer pipette ThermoFisher 15387823
Goat serum Biowest S2000-100
Hera Guard Flow Cabinet Heraeus
Heracell 150 CO2 Incubator Thermo Scientific
HiMark Pre-stained Protein Standard Invitrogen
Horse Serum, New Zealand origin Gibco 16050122
HRP-α- Rabbit IgG abcam ab205718
HRP-α- Rat IgG abcam ab205720
HRP-α-Mouse IgG abcam ab205719
ImageJ v. 1.53t
Methyl Green Sigma-Aldrich 67060
MM400 Tissue Lyser Retsch
NanoDrop ND-1000 Spectrophotometer ThermoFisher
Paraformaldehyde, 16% w/v aq. soln., methanol free Alfa Aesar 043368-9M
Penicillin-Streptomycin (100x) GRiSP GTC05.0100
Phalloidin Alexa 488 Thermo Fisher Sci. A12379
Polystyrene Petri dish 60x15mm with vents (sterile) Greiner Bio-One 628161
Qubit dsDNA HS kit Thermo Scientific Q32851
Qubit™ 3 Fluorometer Invitrogen 15387293
S6E Zoom Stereo microscope Leica
Sodium Dodecyl Sulfate Merck 11667289001
SuperFrost® Plus adhesion slides Thermo Scientific 631-9483
SuperSignal West Pico PLUS Chemiluminescent Substrate Thermo Scientific 15626144
TCS SPE confocal microscope Leica
Tris-(hidroximetil) aminometano (Tris base) ≥99% VWR Chemicals 28811.295
Triton X-100 Sigma-Aldrich X100-100ML
Trypan Blue Solution, 0.4% Gibco 15250061
Trypsin-EDTA (0.05%) in DPBS (1X) GRiSP GTC02.0100
TWEEN 20 (50% Solution) ThermoFisher 3005
WesternBright PVDF-CL membrane roll (0.22µm) Advansta L-08024-001
α-Collagen I abcam ab21286
α-Collagen IV Millipore AB756P
α-Collagen IV Santa Cruz Biotechnology sc-398655
α-Fibronectin Sigma F-3648
α-Laminin α2 Sigma L-0663
α-MHC D.S.H.B. MF20
α-Mouse Alexa 488 Molecular Probes A11017
α-Mouse Alexa 568 Molecular Probes A11019
α-pan-Laminin Sigma L- 9393
α-phospho-histone 3 Merk Millipore 06-570
α-Rabbit Alexa 568 Molecular Probes A21069
α-Rabbit Alexa 488 Molecular Probes A11070
α-Rat Alexa 488 Molecular Probes A11006



  1. Mecham, R. P. Overview of extracellular matrix. Current Protocols in Cell Biology. 10, 10.1.1-10.1.6 (2012).
  2. Frantz, C., Stewart, K. M., Weaver, V. M. The extracellular matrix at a glance. Journal of Cell Science. 123 (24), 4195-4200 (2010).
  3. Lamandé, S. R., Bateman, J. F. Genetic disorders of the extracellular matrix. Anatomical Record. 303 (6), 1527-1542 (2020).
  4. Ahmad, K., Shaikh, S., Ahmad, S. S., Lee, E. J., Choi, I. Cross-talk between extracellular matrix and skeletal muscle: implications for myopathies. Frontiers in Pharmacology. 11, 142 (2020).
  5. Tome, F. M., et al. Congenital muscular dystrophy with merosin deficiency. Comptes Rendus de l'Academie des Sciences. Serie III, Sciences de la Vie. 317 (4), 351-357 (1994).
  6. Gawlik, K. I., Durbeej, M. Skeletal muscle laminin and MDC1A: pathogenesis and treatment strategies. Skeletal Muscle. 1 (1), 9 (2011).
  7. van Ry, P. M., et al. ECM-related myopathies and muscular dystrophies: pros and cons of protein therapies. Comprehensive Physiology. 7 (4), 1519-1536 (2017).
  8. Gawlik, K. I., Durbeej, M. A family of laminin α2 chain-deficient mouse mutants: advancing the research on LAMA2-CMD. Frontiers in Molecular Neuroscience. 13, 59 (2020).
  9. Kuang, W., et al. Merosin-deficient congenital muscular dystrophy. Partial genetic correction in two mouse models. Journal of Clinical Investigation. 102 (4), 844-852 (1998).
  10. Nunes, A. M., et al. Impaired fetal muscle development and JAK-STAT activation mark disease onset and progression in a mouse model for merosin-deficient congenital muscular dystrophy. Human Molecular Genetics. 26 (11), 2018-2033 (2017).
  11. McCrary, M. W., Bousalis, D., Mobini, S., Song, Y. H., Schmidt, C. E. Decellularized tissues as platforms for in vitro modeling of healthy and diseased tissues. Acta Biomaterialia. 111, 1-19 (2020).
  12. Crapo, P. M., Gilbert, T. W., Badylak, S. F. An overview of tissue and whole organ decellularization processes. Biomaterials. 32 (12), 3233-3243 (2011).
  13. Philips, C., Terrie, L., Thorrez, L. Decellularized skeletal muscle: A versatile biomaterial in tissue engineering and regenerative medicine. Biomaterials. 283, 121436 (2022).
  14. Silva, A. C., et al. Comparable decellularization of fetal and adult cardiac tissue explants as 3D-like platforms for in vitro studies. Journal of Visualized Experiments. (145), (2019).
  15. Silva, A. C., et al. Three-dimensional scaffolds of fetal decellularized hearts exhibit enhanced potential to support cardiac cells in comparison to the adult. Biomaterials. 104, 52-64 (2016).
  16. Yaffe, D., Saxel, O. Serial passaging and differentiation of myogenic cells isolated from dystrophic mouse muscle. Nature. 270 (5639), 725-727 (1977).
  17. Burattini, S., et al. C2C12 murine myoblasts as a model of skeletal muscle development: morpho-functional characterization. European Journal of Histochemistry. 48 (3), 223-233 (2004).
  18. Murphy, D. Caesarean section and fostering. Methods in Molecular Biology. 18, 177-178 (1993).
  19. Prieto, D., Aparicio, G., Machado, M., Zolessi, F. R. Application of the DNA-specific stain methyl green in the fluorescent labeling of embryos. Journal of Visualized Experiments. (99), e52769 (2015).
  20. Lichtman, J. W., Conchello, J. -A. Fluorescence microscopy. Nature Methods. 2 (12), 910-919 (2005).
  21. Jonkman, J., Brown, C. M., Wright, G. D., Anderson, K. I., North, A. J. Tutorial: guidance for quantitative confocal microscopy. Nature Protocols. 15 (5), 1585-1611 (2020).
  22. Paulsson, M. A. T. S. Biosynthesis, tissue distribution and isolation of laminins. The Laminins. , 1-26 (1994).
  23. Fedecostante, M., et al. Recellularized native kidney scaffolds as a novel tool in nephrotoxicity screening. Drug Metabolism and Disposition: the Biological Fate of Chemicals. 46 (9), 1338-1350 (2018).
  24. Wagner, D. E., et al. Comparative decellularization and recellularization of normal versus emphysematous human lungs. Biomaterials. 35 (10), 3281-3297 (2014).
  25. Brouki Milan, P., et al. Decellularization and preservation of human skin: A platform for tissue engineering and reconstructive surgery. Methods. 171, 62-67 (2020).
  26. Lamandé, S. R., Bateman, J. F. Collagen VI disorders: Insights on form and function in the extracellular matrix and beyond. Matrix Biology. 71-72, 348-367 (2018).
  27. Baptista, P. M., et al. The use of whole organ decellularization for the generation of a vascularized liver organoid. Hepatology. 53 (2), 604-617 (2011).
  28. White, L. J., et al. The impact of detergents on the tissue decellularization process: a ToF-SIMS study. Acta Biomaterialia. 50, 207-219 (2017).
  29. Nakayama, K. H., Batchelder, C. A., Lee, C. I., Tarantal, A. F. Renal tissue engineering with decellularized rhesus monkey kidneys: age-related differences. Tissue Engineering. Part A. 17 (23-24), 2891-2901 (2011).
  30. Ma, X., et al. Development and in vivo validation of tissue-engineered, small-diameter vascular grafts from decellularized aortae of fetal pigs and canine vascular endothelial cells. Journal of Cardiothoracic Surgery. 12 (1), 101 (2017).
  31. Wu Young, M. Y., et al. Decellularized fetal matrix suppresses fibrotic gene expression and promotes myogenesis in a rat model of volumetric muscle loss. Plastic and Reconstructive Surgery. 146 (3), 552-562 (2020).
This article has been published
Video Coming Soon

Cite this Article

Gameiro dos Santos, P., Soares, A. R., Thorsteinsdóttir, S., Rodrigues, G. Preparation of 3D Decellularized Matrices from Fetal Mouse Skeletal Muscle for Cell Culture. J. Vis. Exp. (193), e65069, doi:10.3791/65069 (2023).More

Gameiro dos Santos, P., Soares, A. R., Thorsteinsdóttir, S., Rodrigues, G. Preparation of 3D Decellularized Matrices from Fetal Mouse Skeletal Muscle for Cell Culture. J. Vis. Exp. (193), e65069, doi:10.3791/65069 (2023).

Copy Citation Download Citation Reprints and Permissions
View Video

Get cutting-edge science videos from JoVE sent straight to your inbox every month.

Waiting X
Simple Hit Counter