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

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):
      ​Sample_{wet weight} = weight_{tube with samples}– weight_{empty 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 4.2.2.12-4.2.2.18) 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 4.3.5.2-4.3.5.7.

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.