You will need:
13.5 day pregnant mouse (we use MTK NEO inbred white mice)
2 sets sterile instruments
one containing a pair of curved forceps and a pair of iris scissors
one containing two pairs curved forceps, one pair iris scissors and a #3 size scalpel handle
Phosphate buffer saline (PBS)
Sterile medium size petri dishes (tissue culture standard)
18 gauge needle
luer lock syringe (about 6cc should suffice)
#11 size flat-edged scaple blade
trypsin / EDTA
Dulbecco's Modification of Eagles Medium (with 10% fetal calf serum, 1% penicillin / streptomycin, 1% L-glutamine 0.2% 0.1 m BME)
large flasks (tissue cultured standard - about 154cm2 area)
Class II Laminar Flow hood
Before starting, pour out 2 x petri dishes of PBS in the hood. Pregnant mouse is killed by cervical dislocation. (This is not done in the hood but on clean benchcote). Lay mouse out on its back and swab belly with 70% ethanol. With a pair of scissors (not sterile) nip a small cut across the belly. Grasping the skin above and below the nip with your finger, tear the skin apart and draw back over the head and hind legs to expose the viscera of the gut. This method is cleaner than cutting through the fur and enable you to reach the uterus with no risk of touching the fur (cutting through dry fur creates a bacterial aerosol).
Using sterile forceps and iris scissors dissect out the uterus, taking care not to touch the fur or the benchcote with the uterus or instruments. Place the uterus into a petri dish of sterile PBS and swirl around to remove blood. Transfer uterus to second petri dish of sterile PBS and move dish to hood.
Using the second set of sterile instruments and a fresh sterile petri dish, isolate the embryos. Be sure to remove the placement and embryonic sacs. Using the scalpel handle with the #11 blade on it, cut off embryo heads and scoop out the liver with a pair of forceps. The head and forelimb should be cut off as shown below.
Discard the head and liver and leave the bodies in fresh PBS in a fresh petri dish.
Take 6cc luer lock syringe with 18 gauge needle attached and remove plunger. KEEP PLUNGER STERILE. Drop the embryo bodies inside the syringe and add 3ml tyrpsin/EDTA. Put plunger back in syringe and squirt contents of syringe into a large tissue culture flask. Place the flask onto a warming tray (37¡C) for 2-3 minutes. Back to the hood and add 20ml DMEM. When adding the DMEM, try to wash any tissue off the walls of the flask. Pipette the tissue / medium up and down a few times to help break up the tissue. Transfer flask to an incubator at 37¡C with 5% CO2. Do not put less than 7 embryo in one large flask.
IMPORTANT: Loosen lid of flask in incubator to allow gas exchange in medium.
This is the primary isolation or passage one. PMEF's should attach and begin to divide in 1-3 days. During this time do not disturb, so as to allow PMEF's to settle and attach.
After 2 days change the medium. It will be very acidic. After 3-4 days the culture will need splitting. Remove media and gently wash the monolayer with 2 X 10ml PBS. Add 2ml trypsin EDTA and split 1:4. After a further 2-4 days the culutre will be ready for freezing. The number obtained from each flask will be between 5-10 x 106 cells. Freeze cells in 10% DMSO at 3 X 106/ ampule.
When recovering the cells from LN2 put all the cells into a medium flask. When confluent these are split into 1 medium and one large flask (1:3). The large flask can be treated with mitomycinC and the medium flask split again. Do not passage beyond P6.
