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The aggregation method for generating chimaeras, as opposed to the microinjection technique, is useful as it does not require expensive microinjection apparatus or sophisticated manipulative skills, and utilizes outbred blastocysts which are easier to obtain in numbers. The ES cells are intorduced into the developing embryo by the adherance of the cells to a dezonulated 8 cell stage embryo. The embryos are cultured to blastocysts and then transfered into a pseudopregnant hybrid (C57 X CBA) F1 recipient.
Preparation of the aggregation plate.
Aggregation and culture of the embryos with the ES cells is performed in microwells prepared in a plastic tissue culture plate using a darning needle.
1. In a 6cm petri dish add up to 10 drops of M16 media (ca. 3-4mm, larger allows too muh turbulence when the palte is moved).
2. Cover the entire plate with parafin or Dow Corning Fluid 200 (viscosity 50cs).
3. Sterilize a darning needle by dipping in ethanol and ignite. Note be very careful not to burn the plastic layer off (ie. don't place in a bunsen burner). Just sterilizing in ethanol is probably sufficient and more"needle friendly".
4. Press the darning needle through the oil and media into the palstic and make a circular movement with the free end of the needle but DO NOT twist the needle (it scores the sides). The well should be ca. 300um dia with a smooth wall. NOTE. In our experience it is critical to make the wells through the oil and media rather than on a dry plate, presumably to make smooth (lubricated) wells. When we made wells on dry plates the embryos always died.
5. Make 5-6 wells in each drop and then return the plate to the incubator.
Removal of the Zona pellucida from the embryo.
After recovering 2.5 day mouse embryos (see Flushing Morula protocol) the zona is removed in acid Tyrode's solution.
Make to 90ml with Travenol water and dissolve components. Adjust pH to 2.5 with 5M HCl and make to 100ml with Travenol water. Filter sterilize and make aliquots of 1ml. Store at -20¡C.
Warm to room temperature before use.
After removing the zona the embryos become extremely sticky and therefore siliconized pipettes are required for handling them.
Siliconizing Capillary Glass
Embryos that have had the zona pellucida (jelly-like outer coating) removed are "sticky" to handle, often getting stuck inside the transfer pipette during handling. This problem is overcome by siliconizing the capillary glass (inside only) before it is used to fashion transfer pipettes.
NOTE: This procedure must be carried out wearing gloves in a fume hood!
Use a 27 gauge needle and 3 cc syringe (filled) to run a small amount of silicone (Sigmacote) through the inside of a piece of capillary glass, allowing excess Sigmacote to drip into a 10ml tube. This is repeated with each piece of glass and the glass placed in a measuring cylinder. The excess Sigmacote in the 10ml tube may be used to refill the syringe when empty. Once 50-80 pieces have been siliconized the glass is rinsed no less than eight times with Milli-Q water, shaking the water out of the glass by hand after each rinse. The glass is then rinsed six times with "Travenol" water. The cylinder is then sealed with Parafilm which is punctured several times with a needle to allow evaporation with minimal dust collection. The cylinder is placed in a hot air oven to speed evaporation. Once completely dry, the glass is ready to use.
Sigmacote is made by Sigma Chemicals. The catalogue number for Sigmacote is SL-2
1. Put several drops of M16 and also acid Tyrodes into a small petri dish (do not mix).
2. Transfer all of the embryos to one of the drops of M16. We use Random bred Swiss embryos.
3. Collect up to 50 embryos (less if unable to handle that no. quickly) with as little media as possible and transfer to a drop of acid Tyrodes, then transfer immediately to a fresh drop of acid Tyrodes.
4. Agitate the embryos while watching them under the scope.
5. Transfer the embryos to a drop of M16 as soon as the zona dissolves. This should take around 90 seconds.
6. Rinse in a second drop of M16 immediately and then transfer to the aggregation plate, placing one embryo in each well. Return plate to the incubator.
7. We include a control of 6-8 unmanipulated embryos during the culture period.
Preparation of ES cells for aggregation.
We use small clumps of ES cells rather than single ES cells as these are much easier to place in the wells. The cells are harvested essentially as described in "Passaging and Spliting ES cells" except that at the end of the tyripsinization step they are GENTLY pipetted up and down 6-8 times rather than the vigourous motion usually used to create a single cell suspension. This yields clumps of cells of 4-12.
1. Aspirate media from the plate, wash with 5ml PBS and with 5ml PBS/EGTA. Aspirate and place plate on 37¡C warming tray.
2. After 2-5min. remove the flask and checl under the dissecting scope. Individual cells should be apparent in the collonies.
3. Add 1ml trypsin/EDTA and rock flask back and forward for ca. 1min. until the cells lift off the plate. Gently pipette the cells up and down with a 1ml pipette 6 times and then add 1ml of ES cell media. Wash gently up and down several times.
4. Add cells to 2ml of ES cell media to dilute to an appropriate concentration.
1. Using a pasteur pipette, place a drop of suspended ES cell clumps in a small petri dish.
2. Under the dissecting scope and using a very narrow siliconized transfer pipette, select clumps of 5-10 cells. Transfer one clump to each well and watch to be sure that it comes to rest adjacent to the embryo. Very carefully return the plate to the incubator once all the embryos and wells are filled.
3. Our embryos take 24 hours to develop to expanded blastocysts at which point they are transfered into psuedopregnant (C57 X CBA) F1 mice using a siliconized, flame polished pipette.
Before attempting to remove expanded blastocysts, make sure that the pipette is wide enough. To remove blasts from wells, fill a transfer pipette with M16 up to the end of the narrow pulled section then position the end over each well and blow the embryos out. Arrange them in groups of 12 for easy collection during transfer.
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