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Contact Information
Office of
Public Safety
Westmont College
955 La Paz Road
Santa Barbara, CA 93108
805.565.6043
safe@westmont.edu

Chemical Hygiene Plan

Appendix B. Sample SOP

Perfusions

Materials:

- styrofoam box with trash bag lining the inside and a hole on top that allows for fluid to drain down into bag

- 2 60ml syringes

- tubing with valve

- 23 gauge butterfly needle, connected to tubing

- 1 pair of forceps, serrated tips

- small scissors to cut atrium

- mid-sized scissors to cut through skin/rib cage

- large scissors to cut head off

- at least six 18 gauge needles

- 0.9% NaCl solution

- 4% paraformaldehyde

- Nembutal

- IP syringes

Procedure:

1. Inject 0.1ml of Nembutal into the IP cavity of the mouse.

2. Fill one of the 60ml syringes with the NaCl solution and the other with paraformaldehyde. Attach both to the tubing valve. Run enough saline through the tubing that you can be assured that there is nothing else in the line.

3. Once the mouse is determined to be down by a tail test, pin it on top of the styrofoam box with 18 gauge needles, one needle through each limb. Pinch skin right over the bladder and make the first cut with the mid-sized scissors. Cut up each side of the abdominal cavity, up through the rib cage, being extremely careful to not snip any organs. Cut through the tissues to expose the heart, then pin the large flap of skin up to keep the entire cavity exposed.

4. Poke the tip of the butterfly needle into the left ventricle, (mouse’s left, your right) being careful not to push too far and puncture other chambers. While holding the butterfly needle in place, use the small scissors to make a small cut in the right atrium. You will know when a cut has successfully made by the blood that will pour out.

5. Slowly pump saline through the line, watching to see that the liver turns a yellowish color and that the blood flowing out of the atrium steadily turns nearly clear. Ideally, this will take about 15-20ml of saline, but more is often necessary.

6. Switch the knob on the valve to open the para line, and slowly pump all 60ml of para through the mouse’s circulatory system. You should see the body move, (tail curl, fists clench, etc.) as the tissue is fixed. Record all observations on the perfusion chart.

7. Cut the head off with the large scissors, being careful not to cut the ears off. Place the head upside down in a head jar filled partially with paraformaldehyde, making sure that both eyes are submerged. Store this in the refrigerator until dissection.

8. Pump saline through the line to clear out all para before beginning the next perfusion. When finished with all perfusions, rinse all tools thoroughly with ddH20.