Cardiopulmonary Complex Decellularization: A Technique for Decellularizing Heart and Lungs from Murine Model

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Post-mortem Microsurgery

1. Euthanize the mouse using a CO₂ chamber.
   1. Place mouse into a 4 L chamber, and begin to fill with 100% CO₂, starting at 0.2 L/min for 2 min, increasing until reaching a flow of 0.8 L/min after 3 min. The mouse should fall unconscious during the first 2 min, and then respiration should cease (usually around 5 min, but flow can be maintained as necessary). Confirm death prior to proceeding to the next step.
2. Shave the thorax, abdomen and back of the mouse with the hair clipper and disinfect with 70% ethanol. Shaving greatly reduces the number of artifacts due to the presence of hair either on samples for imaging or biochemical analysis.
3. Pin the mouse to a polystyrene tray, extending its fore- and hindlimbs, as well as its head and tail. Place it under the microsurgery microscope.
4. Using a Mayo straight pattern scissors establish surgical access with a cutaneous incision running from the submandibular region to the lower abdomen and dissect subcutaneously to expose the thoracic wall and peritoneum.
5. Using microsurgical scissors, cut the pectoralis major and pectoralis minor muscles along the sixth intercostal space on both sides of the thoracic wall.
6. Using straight-pattern scissors, cut the sternum along the previous incisions, and then complete a sternotomy by cutting the sternum along its long axis, then elevate and pin both sides of the thoracic wall to expose the cardiopulmonary complex.
7. Using round-tipped micro-forceps (or Dumont micro-forceps), excise the thymus and surrounding adipose tissue by delicately pulling them off their attachments. This will reveal the major vessels.
8. Using the cautery, cauterize the descending cava vein and, using straight pattern scissors, cut the oesophagus.
9. Using sharp micro-forceps, separate the brachiocephalic veins and the brachiocephalic, left common carotid and left subclavian arteries from the underlying tissue to facilitate ligation and cauterization.
10. Using micro-needle holder, sharp micro-forceps and 9-0 suture place stitches above the emergence of the brachiocephalic, left common carotid and left subclavian arteries.
11. Cauterize the brachiocephalic veins.
12. Separate the submandibular salivary glands along the midline to expose the neck muscles and the trachea. Separate the muscles to expose the cricothyroid ligament. Using micro-scissors, open an entrance by sectioning the ligament.
13. Introduce a 27 G catheter in the trachea and delicately push until the trachea branches into the bronchi (i.e., until resistance to the catheter is met, then retreat 3 mm). Be careful not to disrupt the bronchi. Using a 6-0 suture, place 3 stitches around the trachea to secure the catheter.
14. Section the mouse at the height of the 12th thoracic vertebra. The descending aorta runs anteriorly to the spine and should be sectioned here along with the spine. Set the lower half apart.
15. Retrogradely catheterize the aorta and push the catheter until it reaches the aortic arc. Using 9-0 suture, place 4 stitches around the aorta, beginning 5 mm below the catheter tip.

2. Decellularization

1. Connect the mouse to a pump system using silicone tubing and Luer connectors. Perfuse with deionized water at 200 µL/min for 15 min. Maintain this flow rate during decellularization.
2. Change the perfusion agent to 0.5% sodium deoxycholate (DOC) diluted in deionized water and perfuse overnight.
3. Change the perfusion agent to 0.1% sodium dodecyl sulphate (SDS) diluted in deionized water and perfuse for 8 hours.
4. Perfuse with deionized water overnight to wash away SDS and DOC for 24 h.
5. Resect the decellularized heart and lungs by sectioning its attachments to the thorax using a curved scissors and store in a sterile cryo-tube with deionized water with 1% (v/v) penicillin-streptomycin and 0.3 µM sodium azide at 4 °C. ECM scaffolds can be stored for at least for 12 weeks. If the scaffold will be used for biochemical analysis (e.g., mass spectrometry), snap freeze in liquid nitrogen.