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Transgenic Core Facility
Protocols
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1. Superovulation The use of hormones in females prior to mating, to stimulate the release of greater numbers of eggs, has been used successfully in this facility on the FVB strain. It does increase two-to threefold the number of embryos obtained, but this is somewhat countered by an increase in the percentage of eggs that are not fertilized. Nonetheless, it is an economical way to maximize yield. It is most effective when young (4-6 week) females and only the most productive males are used. Aliquots of pregnant mares’ serum gonadotropin (PMSG) and human chorionic gonadotropin (HCG) are made up at the appropriate concentrations and stored at -80˚ C. It is important to allow the hormone to thaw gently at room temperature - accelerating the thawing seems to reduce the efficacy of the hormones. 1) Using a 1 cc syringe with a 30 G x 1/2” needle, inject each FVB female with 0.1cc of PMSG intraperitoneally (the female’s estrus cycle can be ignored when using the hormones). Make a note of the injection time. 2) After 46-48 hours, inject the same mice with 0.1 cc of HCG intraperitoneally. Aliquots of HCG, need to be diluted 1:10 with sterile saline before injection. 3) Once they are injected with HCG, place females with intact FVB males, as with natural matings. 4) Check females for “plugs” the following morning. Back to Top Back to Protocol Page
2. Isolation of Single-Cell Embryos from Donor Prepare the following 5 dishes (35mm petris) before beginning : 2.5 ml M2 media (for hyaluronidase) 2 x 1.5 ml M2 1.5 ml M16 media Drop culture of M16 under 3 ml mineral oil 1) Remove the oviducts of donor females on the same morning that they were found to be plugged, placing them in a dish of M2. 2) Transfer oviducts to a dish containing 2.5 ml of M2 and 60 µl of hyaluronidase. 3) Under a dissection scope, use two forceps to break open the bulge in each oviduct, releasing the eggs. Discard oviducts. 4) The eggs will be in a cloudy mass which will gradually be broken up by the hyaluronidase. Use a mouth pipet with a drawn transfer pipet to collect the eggs in a clear area of the dish. 5) When eggs are free of debris, transfer them and rinse them in the last dish of M2, then in M16, then transfer to the drop culture. 6) Place the drop culture and the rinse dishes of M2 and M16 in an incubator set at 5% CO2 and 37˚ C. Back to Top Back to Protocol Page
3. DNA Injection into Pronucleus 1) Make a drop culture on a clean, siliconized glass depression slide: Overlay a 10 µl drop of M2 media with just enough mineral oil to cover the drop without overflowing the depression. Place the slide on the microinjection microscope stage. 2) Fill the holding pipet with M2 media and mount on the micromanipulator apparatus. Using the angular control of the micromanipulator, slowly lower the pipet into the drop (watching through the microscope) until the tip is at the bottom of the injection chamber. 3) Using a mouth pipet, transfer fertilized eggs into the injection chamber. 4) Load DNA into an injection needle: Using a mouth pipet, insert a clean sterilized DNA filler into the DNA tube. Draw up a small amount of liquid, and insert the filler into the blunt end of the injection needle. Gently transfer DNA into the needle and withdraw the filler. Rest the needle horizontally to allow the liquid to travel to the pulled tip. This may be accelerated by gently flicking the needle with the tip pointing down. 5) Mount the loaded injection needle on the other micromanipulator apparatus and lower into the injection chamber until the tip is in approximately the same focal plane as the holding pipet. 6) Break the tip of the needle against the holding pipet: The tip of the holding pipet is brought into focus under highest magnification, then the needle tip is brought into the same focal plane using the micromanipulator instead of the focus knobs. The needle is then nudged gently against the holding pipet until it can be seen to be slightly blunted. 7) Use the holding pipet to position and hold a fertilized egg by adjusting the suction of the pipet with a 60 ml syringe. The egg can be ‘bounced’ off and recaptured until it is in a suitable orientation. The egg should be positioned so that the larger of the two pronuclei is located on the outer edge of the egg closest to the injection pipet. Pull back slightly on the syringe controlling the holding pipet, to be sure the egg will not rotate when the needle is introduced. You should see the outer membrane being pulled slightly into the holding pipet, but be careful not to pull in the inner mass of the egg. 8) Under the highest magnification, using the ‘40’ filter, focus on the pronucleus. Bring the injection needle into the same focal plane using the micromanipulator controls. Push the tip of the needle gently into the pronucleus while applying gentle pressure to the syringe. The needle may have to overshoot the leading side of the pronucleus in order to break through the membrane. When the tip is inside the pronucleus, apply pressure to the corresponding syringe (more than you’d expect) until the pronucleus swells. Once the pronucleus has been injected, quickly remove the needle. Back to Top Back to Protocol Page
4. Implantaion of Embryos into Surrogate Mother This procedure takes place entirely in the cubicle room located in the animal facility. You will need: Sterilized surgical pack Mouth pipet Several transfer pipets 30 G 1/2” needle Embryos in the original drop culture of M16 media 35mm petri dish containing 1.5 ml of M2 media 1) Use Nembutol to anesthetize a 0.5 day pseudo-pregnant female. Ketamine/Xylazine may also be used. Inject 0.3 - 0.5 cc IP, depending on the weight of the foster mother. This provides 20 - 30 minutes of surgical plane anesthesia. 2) Transfer embryos into M2. Discard the transfer pipet used for this, as the mineral oil may be somewhat toxic to the surrogate mother. 3) Thoroughly wet down the back of the mouse with 95% EtOH, sponge off excess with a kimwipe. Lay the mouse on her stomach on a clean, dry Kimwipe, which will then be used to transfer her back and forth. Locate the hump in the spine and the hollow immediately caudal to it. Make a transverse incision in the skin in this hollow, 1.0 to 1.5 cm long. 4) Before making an incision in the peritoneal wall, visually locate the ovary; it will be located just under the wall. It is a 3 - 4 mm round red organ which may show white spots on its surface. Using a small pair of surgical scissors and a pair of rat-tooth forceps, make a small (5 mm) incision in the wall over the ovary. Use a small clamp attached to a fat pad to keep the ovary in position. 5) Load the injected embryos into a mouth pipet. First draw up M2 media to fill the pipet to the ‘shoulder’ of the pipet, then two small air bubbles, then the embryos, and finally one more small air bubble. The number of eggs implanted in each mouse depends on the number, but 24 per mouse (12 per oviduct) are average. 6) Use a dissection scope to focus on the oviduct of the mouse. Find the swollen bulge in the oviduct. Use a 30 G needle to make a hole between the ovary and this bulge. Insert the pipet into this hole and gently blow the eggs into the oviduct until air bubbles appear in the bulge. 7) Remove the clamp and allow the ovary to return to the body cavity. Close the skin incision using surgical staples. 8) Visually check for pregnancy in 10 days. Back to Top Back to Protocol Page
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