Purification and Transplantation of Myogenic Progenitor Cell Derived Exosomes to Improve Cardiac Function in Duchenne Muscular Dystrophic Mice

Animals were handled according to approved protocols and animal welfare regulations of the Institutional Animal Care and Use Committee of the Medical College of Georgia at Augusta University.

1. Isolation and Purification of MPC-derived Exosomes

1. Seed 5 x 10⁶ C2C12 cells in a 15 cm cell culture dish with 20 mL complete Dulbecco’s modified Eagle’s medium (DMEM) containing 10% fetal bovine serum (FBS), 100 U/mL penicillin G and 100 μg/mL streptomycin. Incubate at 37 °C and 5% CO₂.
2. Prepare exosome-depleted medium by ultracentrifugation of FBS at 100,000 x g for 18 h at 4 °C using a swinging bucket rotor. Discard the pellet.
3. Replace the complete DMEM with exosome-depleted medium when monolayer cells reach 80% confluence in the culture dish.
4. Use a transfer pipette to collect the supernatant from the cell culture dish every 48 h for a total of three times. Collect the supernatant in 50 mL centrifuge tubes, and centrifuge at 150 x g for 10 min to remove the remaining cells.
5. Filter the supernatant through a 0.22 μm filter to eliminate cell debris. Ultracentrifuge the filtered medium in ultra-clear tubes using the swinging bucket rotor at 100,000 x g for 120 min at 4 °C to precipitate exosomes.
6. Resuspend the exosome-containing pellet in phosphate buffered saline (PBS) and fill the entire ultra-clear tube with PBS. Perform ultracentrifugation of this suspension at 100,000 x g for 120 min at 4 °C to eliminate contaminating proteins.
7. Discard the supernatant and resuspend the exosome pellet in 100 µL PBS. Store at -80 °C for future use.
8. For electron microscopy analysis, place 3 µL of exosomal suspension on a 200 mesh formvar-coated copper electron microscope grid for 5 min at room temperature. Continue with standard uranyl acetate staining²³. Examine the semi-dry grid by transmission electron microscopy.

2. Intramyocardial Exosome Delivery

NOTE: We used six mice in each group.

1. Anesthetize DBA/2J-DMD^MDX mice (male; age > 10 months; body weight (BW) > 18 g) with intraperitoneal (i.p.) injection of 100 mg/kg BW of ketamine combined with 10 mg/kg BW of xylazine, and confirm the depth of anesthesia by toe pinch.
2. Fix each mouse in supine position on a surgical platform by using tape to secure each limb and place a 3-0 suture horizontally below the upper teeth to hold the upper jaw in place.
3. Apply the depilatory cream to the left side of the mouse chest to remove the fur from the skin.
4. Perform endotracheal intubation via oral cavity using a 24 G catheter, and ventilate the mouse with room air at a rate of 195 breaths/min using a rodent ventilator (see the Table of Materials).
5. Disinfect the skin with 75% alcohol and 10% povidone iodine.
6. Make a 10‒15 mm oblique incision from the left sternal edge to the left armpit, cut the pectoralis major and pectoralis minor by a scissor, then make a left thoracotomy through the fourth intercostal space. Gently insert the retractor bands to spread the thoracic cavity to a width of 10 mm. Take care to avoid damage to the left lung.
7. Use two straight tweezers to remove the pericardium, pull them apart, and place them behind the retractor tips to expose the heart. Inject MPC-Exo (50 μg in 30 µL PBS) or 30 μL PBS (as a control) intramyocardially into the anterior wall of the left ventricle using a 31 G insulin needle at one site.
8. Use 6-0 nylon sutures to close the thoracic cavity, pectoralis muscles and skin in sequence.
9. Recover mice. Once rhythmic, spontaneous breathing is present, remove the mice from the ventilator, and extubate. Observe and record the mice condition after surgery.

3. Echocardiography

NOTE: A single observer blinded to the experimental groups performs echocardiography and data analysis.

1. Two days after PBS/exosome transplantation, anesthetize the mice with 1‒2% isoflurane.
2. Fix a mouse in supine position using tape, and apply the preheated acoustic gel on the left chest area.
3. Assess left ventricular function by echocardiography as previously described^16,24.
   1. Obtain the parasternal long axis view of the left ventricle (LV) in two dimensions (B mode), and then rotate the ultrasound probe 90° to obtain a LV short-axis view at papillary muscle level.
   2. Record M-mode echocardiographic images and measure left ventricular end-diastolic diameter (LVEDD), left ventricular end-systolic volume (LVESD), left ventricular end-diastolic volume (LVEDV), and left ventricular end-systolic volume (LVESV).
      NOTE: Fractional shortening (FS%) = [(LVEDD − LVESD) / LVEDD] x 100, and left ventricular ejection fraction (LVEF%) = [(LVEDV − LVESV) / LVEDV] x 100.

4. Immunohistochemistry

1. Two days after PBS/exosome transplantation, anesthetize mice (see step 2.1) and cut the sternum open to expose the thoracic cavity. Dissect the inferior vena cava, and perfuse the heart through the left ventricle apex with 3 mL PBS, followed by 3 mL of 4% paraformaldehyde at room temperature, using a butterfly catheter with a 25 G needle attached to a 5 mL syringe.
2. Embed fixed hearts into paraffin, and cut into 5 μm thick sections.
3. Deparaffinize slides in xylene and rehydrate in ethanol solutions in following order: 100% xylene (3 min), 100% xylene (3 min), 100% ethanol (1 min), 100% ethanol (1 min), 95% ethanol (1 min), 80% ethanol (1 min), H₂O (1 min).
4. Fix tissue sections with 4% paraformaldehyde for 8 min at room temperature.
5. Immerse slides in sodium citrate (0.01 M, pH 6.0) and perform antigen retrieval using an antigen retriever.
6. Permeabilize sections for 1 h at room temperature with 1% polyethylene glycol tert-octylphenyl ether in PBS.
7. Block sections with 5% goat serum in PBS for 1 h at room temperature.
8. Incubate sections with diluted primary antibody (anti-dystrophin antibody) in PBS (1:100) overnight at 4 °C, then wash three times with PBS for 5 min.
9. Incubate sections with diluted secondary antibody (Alexa Fluor 488 conjugated goat anti-rabbit IgG) in PBS (1:400) for 45 min at room temperature.
10. Control autofluorescence using an autofluorescence quenching kit (see the Table of Materials).
11. Use mounting medium containing DAPI (see the Table of Materials) to mount the slides.
12. Observe slides under a confocal microscope.