- Clear the fix and wash bottle lines of any air bubbles. Sufficient perfusion pressure is obtained with the bottles 100-120 cm above the animal (but no more!).
- Run wash solution through cannula to remove any fix from lines. Set at a slow drip.
- Place the heavily anesthetized animal on its back on a rack or paraffin block over a sink. Note: A barbiturate type of anesthesia is highly preferred to gaseous types such as Fluothane, with which vessels tend to constrict resulting in poor perfusion.
- Spread the forelimbs and secure each paw to the rack with a surgical towel clamp or a pin if on a paraffin block. Start running faucet water.
- Make a cut along the sternum ~8 cm long, low enough to expose the sternum’s end.
- Grasp end of sternum with a 5″ hemostat. Use sharp scissors to cut diaphragm laterally on both sides and cut toward the head across ribs and parallel to lungs.
- If brain tissue is all that is desired, clamp the descending aorta (not necessary for mice). With a mosquito hemostat, reflect one lung and locate the descending aorta as it runs along the spinal column. Clamp it and allow the lung to return.
- Release the hose clamp on the perfusion tube so that the fluid is just barely passing out of the cannula tip.
- With left hand, use a small pair of rat tooth forceps to grasp ventral tip of the heart.
- With right hand, use a #11 scalpel with blade edge out (away from heart) and pierce the left ventricle, allowing the blade to penetrate far enough to make the slit large enough for the cannula tip.
- Quickly remove blade and lower heart to prevent spurts from the long incision.
- With right hand, insert the cannula and direct it up through the left ventricle into the ascending aorta. Stop when the tip of the cannula is visible within the aorta.
- Maintain placement of the cannula in the left hand, holding the cannula and forceps together, side by side. With the right hand free, use the rubber-tipped hemostats to clamp around the aorta, holding the tip of the cannula in place. (Clamping is difficult or not feasible in mice so the cannula/ needle must be held while perfusing.)
- With left hand, release the cannula and forceps from heart and close the wash line clamp. Or you may choose to allow more wash before fix.
- Fully open the fix line clamp and perfusion tube clamp.
- With left hand, use the teeth of the rat tooth forceps or scissors to puncture the right auricle, allowing the escape of return circulation.
- For rats, allow 200 to 250 cc of fix to flow. For mice, allow 50-100 cc of fix to flow. Consider the perfusion a dialysis process, exchanging formaldehyde and water, rather than a ‘flushing’ process.
- Close both clamps and remove perfusion instruments. Remove skin from head and decapitate at a level even with forelimbs. Place head in a container filled with perfusion fix solution. Removal of the skin is preferred.
For degeneration staining, wait at least 24 hours before extracting brain, to allow brains to fully harden and to prevent the induction of artifacts during staining.
Perfusion Pump Calibration
When switching from gravity perfusion to a peristaltic pump, calibration of the pump can be confusing.
Use this simple method to ensure adequate pressure:
- Using an IV bag or other container from which tubing can be attached and allowed to ﬂow, position the container 1-1.2 meters above the table to be used for perfusion.
- Use just water in the container.
- Place the same cannula that will be used for the perfusion on the end of the tubing.
- Attach the cannula horizontally on the edge of the table with the tip pointed out over the ﬂoor.
- Open the clamp and allow the stream to ﬂow.
- Mark where the stream strikes the ﬂoor.
- Now transfer the tubing to the pump that will be used for perfusion.
- Turn on the pump (connected to a water reservoir) and adjust the pressure such that the stream strikes the ﬂoor in the same place as before with the elevated container.
- This will ensure that the pump pressure is calibrated to the correct systolic/ diastolic range.
Other helpful notes:
• Start each perfusion with perfusion wash solution. For mice and rats, use 15 mL of solution. This will separate blood serum from the formaldehyde, which can “ﬁx” red blood cells in the vasculature.
• The number of mL’s of solution used for each animal is important. Don’t use time to determine this ﬂow rate, use marks on your perfusion solution bottles to ensure adequate volumes have passed through the animal. For mice, use 150 mL of perfusion solution. For rats, use 250 mL of solution.
• DO NOT exsanguinate the animal ﬁrst.
• See the Perfusion Fix and Solutions section of our website to determine which solution would best suit your experiment.