hES cells, as mouse ES cells, require a feeder layer of mitotically inactivated fibroblast cells or other cell types in order to remain pluripotent in culture. The hES cells that we routinely use, BG01 and BG02 cell lines (5,8), were established on a MEF feeder layer. The MEF cells are made by standard procedures using 13.5-d post coitus mouse embryos and are frozen immediately after monolayer formation (see Note 6). The MEF cells are frozen immediately after they form a monolayer at passage one or two. The performance of MEF as feeders will be compromised by multiple passage, so it is advisable not to expand MEF before use as feeders.
1. Sacrifice pregnant mice by asphyxiation with CO2. Place the mouse on its back. Pin the animal down onto a covered Styrofoam box lid by inserting needles through its legs.
2. Spray with 70% ethanol and swab with sterile gauze. Using scissors make a cut through the belly skin (at the median line just over the diaphragm).
3. Grasping the skin on both sides of the incision, pull in opposite directions to expose the untouched ventral surface of the abdominal wall (see Note 7).
4. Cut longitudinally along the median line of the exposed abdominal wall with sterile scissors, revealing the viscera. The uteri filled with embryos should be seen in the posterior abdominal cavity.
5. Dissect out the uterus with sterile forceps and scissors and place into a 50-mL screw-capped tube containing 20 mL sterile PBS +/+.
6. If more than one animal is being used, place the tube containing the uterus in the refrigerator or on ice until all uteri have been removed.
7. After the uterus has been isolated, place the dissected female carcass into an autoclave bag for proper disposal. Take the intact uteri to the tissue culture laboratory, and transfer them to a fresh dish of sterile PBS.
3.2.2. Dissection of Embryos
1. Place uterus in a 100-mm polystyrene Petri dish containing 10 mL PBS (+/+) (see Note 8).
2. Using watchmaker forceps and working under a dissection scope, tear the uterus with two pairs of sterile forceps, keeping the points of the forceps close together to avoid distorting the uterus and bringing too much pressure to bear on the embryos.
3. Release embryos from embryonic sacs and transfer to a new Petri dish containing fresh PBS (+/+) (see Note 9).
4. Under a dissecting microscope, remove the embryo heads and livers, intestines, heart, and all viscera using two watchmaker forceps.
3.2.3. Irypsinization of Cells
1. Transfer the embryo carcasses to a fresh Petri dish (no PBS). Carefully mince the embryos with a curved surgical scissors.
2. Add 5 mL 0.25% trypsin/EDTA per 10 fetuses and triturate through a 10-mL pipet.
3. Transfer the embryo/trypsin solution from the Petri dish with a 10-mL pipet to the barrel of a 10-mL syringe with an attached 18-G needle. Replace the syringe plunger and slowly and gently push the embryo/trypsin solution through the needle. Collect in a 50-mL conical tube.
4. Gently pass the embryo/trypsin suspension through the needle a second time.
5. Incubate the tissue suspension for 15 min at 37°C, triturating briefly every 5 min in a biosafety cabinet through a 10-mL pipet to dissociate the tissue.
6. Add an equal volume of MEF medium. Triturate the suspension vigorously with a pipet.
7. Plate one embryo equivalent per 175-cm2 flask and add complete medium to make up a final volume of 30 mL per flask (see Note 10).
9. The next day, change the medium with an equal volume of fresh complete medium. On d 3-4, when the cells are 90% confluent, the MEF p0s are ready to be frozen.
3.2.4. Harvesting and Freezing of MEFs
1. Rinse the 90% confluent cells once with 20 mL PBS (-/-).
2. Detach cells using 3 mL 0.05% trypsin/EDTA solution per flask.
3. Tap the flask after 1 min to detach cells.
4. Inactivate the trypsin solution by adding 5 mL complete medium.
5. Pool cells from all flasks into 50-mL tubes ensuring even distribution among various tubes.
6. Count the number of viable cells using trypan blue and a hemacytometer (see Note 12).
7. Centrifuge the cell suspension at room temperature for 4 min at 200g.
8. Resuspend the cells in complete medium at two times the desired final freezing concentration (see Note 13).
9. Slowly and drop-wise, add an equal volume of 20% DMSO in FBS (see Note 14).
10. Dispense 1 mL cell mixture into prelabeled cryovials and place cryovials into a freezing container (which is prefilled with 250 mL room temperature isopropanol) and freeze at -80°C overnight (see Note 15).
11. After overnight incubation, rapidly remove the cryovials from the freezing container and place in liquid nitrogen (-196°C) for long-term storage.
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