Perfusion of Mouse
From Bridges Lab Protocols
- 0.9 % Saline solution
- 10% formalin solution
- 30 gauge needle
- 1 mL syringe
- Peristaltic pump
- Collecting dish
- Surgical stage/platform
- Make sure this is level with the edge of the collecting dish to allow for the need to rest during the procedure
- Dissecting scissors
- Clamp scissors
- Standard scissors
- Taken from: http://web.jhu.edu/animalcare/rdf/avertin.html
- Stock Solution (1.6 g/ml)
- Mix 25 g avertin and 15.5 ml tert-amyl alcohol at room temperture for ~12 hours in a dark bottle.
- Can store stock solutions at room temperature for one year.
- Working solution (20 mg/ml)
- Mix 0.5 ml avertin stock solution and 39.5 ml 0.9% saline in dark/foil covered container.
- Filter solution through 0.2 micron filter into a dark/foiled covered container.
- Store solution at 4 degree C.
- Replace working solution each month.
- Avertin is light sensitive.
- Degradation products are lethal.
- Always store at 4 degree C
- Do NOT use a solution that is yellow or contains a precipitate.
- Avertin is lipid soluble and may require a larger dose in an obese animal.
Perfusion Pump Set Up
- Place needle on end of tubing
- Insert one tube into 0.9% saline solution, and one tube into 10% formalin solution
- Turn pump on to a flow rate of 0.72 ml/min
- Have stopcock turned to allow for flow of saline
- Fill line with 0.9% saline until you have a steady stream of 0.9% saline from the end of the needle.
- Turn off peristaltic pump and turn stopcock to allow for formalin solution to flow
- Turn on peristaltic pump and pump 10% formalin solution until flow fluid is before the stopcock
- Flow has not entered the main line
- Turn off peristaltic pump and turn stopcock to allow for saline to flow
- Turn on pump and pump saline through the main line at stated flow rate
- Place mouse in isofluorane chamber and lightly anesthetize the animal.
- Remove mouse from chamber
- For mouse anesthesia, administer 0.8 ml/20 g (of mouse body weight) Avertin through intraperitoneal injection with 30 ½ gauge needle.
- Wait for 3 minutes or until the mouse no longer responds to painful stimuli, such as paw pinch before proceeding.
- Lay the mouse on its back.
- Using tweezers and operating/dissecting scissors open up the skin and expose the chest cavity.
- Cut open the diaphragm using standard scissors.
- Be careful to not pierce the heart.
- Using clamp scissors, grab at the base of the sternum, cut through the ribcage and lift to expose the heart.
- Transfer the mouse to the procedural stage/platform.
- Insert the 30 gauge needle (from the tubing with saline/10% formalin solution) into the apex of the left ventricle, being careful to keep the tip of the needle in the lumen of the ventricle.
- Immediately after inserting the needle into the left ventricle, cut the right ventricle using standard scissors.
- This allows for the blood to flow out of the mouse and drain into collecting dish.
- Perfuse with saline solution for 10 minutes.
- After 10 minutes, switch the stopcock to allow for flow of formalin solution, at same flow rate as saline.
- Perfuse for 10-12 minutes with 10% formalin solution.
- With the pump still on, remove needle from left ventricle.
- Mouse is now fixed for tissue collection.
- When collecting tissues, keep tissues in 10% formalin solution for ~24 hours at 4 degree C.
- Transfer tissues to PBS after for long-term storage at 4 degree C.