This procedure centers around the humane utilization of amphibian models in a manner that maximizes efficiency. Standardization practices can be applied to samples taken and preserved at the originating colony, mitigating biosecurity risks and eliminating animal transport stress. Profusion can be completed quickly, maximizing the time available to sample tissues while they are still fresh.
This freshness is especially important for proteomic and transcriptomic profiling. This protocol is detailed and explicit, making it usable for individuals with a wide variety of backgrounds. This facilitates the sharing of high-quality samples between Xenopus labs.
Along with this profusion technique, a similarly comprehensive protocol is being worked on for the standardized sampling of all major organs. Following 15 minutes of submersion in the euthanasia solution, ensure the Xenopus is anesthetized by pinching the foot of the frog. Record the weight and any other parameters needed at this time.
Using a 31-gauge needle, inject laevis in each forelimb muscle with 250 microliters of heparin in PBS. Inject Xenopus tropicalis with 100 microliters of heparin in PBS. Trim the tip off a perfusion needle with wire cutters.
Attach the trimmed needle to the pump, and circulate 54 units per milliliter of heparinized PBS perfusate. Purge all air bubbles from the tubing. Place the dissection tray at an incline within a secondary container to facilitate blood drainage.
After completing the primary euthanasia, check the loss of pain response by pinching the foot. Place the frog on its back. Pin down each limb.
Cut through the midline of the skin. Then cut laterally to create two skin flaps. Grab the linea alba with forceps and pull it away from the cell cavity.
Cut through the musculature. Cut or pin all the flaps out of the way. Cut through the coracoid bones with the help of dissection scissors, and remove any excess tissue to gain better access to the heart.
Next, gently shift the stomach of the dissected Xenopus to the top of the left liver lobe. Gently grasp a lung tip using tissue forceps, and pull it outside the cell cavity. Then pin the lung through the tip.
Pull the thin pericardium tight with tissue forceps. Using the tip of a pair of iridectomy scissors, gently perforate the pericardium without cutting any underlying tissue. Peel the pericardium away from the heart chambers.
Gently grasp the ventricle apex with forceps. Without perforating the ventricle, insert the needle through the closure of the forceps into the ventricle chamber. Clamp the tissue forceps in place using a needle holder or a hemostat.
Start the flow of the pump at approximately five milliliters per minute. This will engorge the heart chambers and the arterial trunk. Use scissors to make a nick in the right auricle.
Continue the pump flow until the vasculature of the stomach blanches. Lance the left heart auricle. Increase the flow rate of the pump to 10 milliliters per minute.
Using a transfer pipette, rinse the coelomic cavity in perfusion media. If the organs shift during rinsing, gently shift them back to maintain visibility of the stomach and lungs. To help visualize the perfusate flowing out of the heart, the auricles may further be reduced.
Keep the needle in place until the stomach vasculature and lungs have blanched, and the perfusate flowing from the heart appears clear. Successful perfusion was observed in all tissues, except the liver, making them distinctly lighter and less saturated with blood. This is most visible in albinos.
