1. Inject two to four adult female frogs in the dorsal lymph sac with 500-800 U HCG, and incubate at 15°C for 12-16 h before transplantations.
2. Set up injection area: Coat inside of transplantation needles with Sigmacote to prevent shearing of sperm nuclei flowing through the needle (needles can be coated 10 min to several months before use). Attach approx 1 cm Tygon tubing (R-3603 1/32 in.; Fisher, cat. #: 14-169-1A) to the end of a plastic pipetman (200 |L) tip, and use the pipetman to draw up Sigmacote; then attach the other end of the tubing to the injection needle. Depress the pipetman plunger to force Sigmacote through the needle until a few drops emerge from the tip, and then release the pipetman plunger to withdraw most of the solution. Rinse needle with water before using for transplantations.
3. Adjust the transplantation apparatus. Fill a Petri dish with water, attach a needle to the tubing and micromanipulator, and establish a very low positive needle pressure through the needle. To do this, set the air flow valve to partially closed (so that it can be opened further), and then open the vacuum valve until liquid is drawn into the needle. When the needle is filled to the wide bore, partially close the vacuum until the flow is either stopped or just slightly outward owing to the partial air flow pressure. The pressure should be so low that it should not be possible to see the meniscus moving at all.
When you are finished adjusting the system and are ready to load for transplants, put a finger over the exhaust tube to discharge the liquid from the needle, back-load the needle, and add positive pressure to the needle just slightly to begin injecting. This is done either by increasing the air flow or screwing done on the clamp fitted on the exhaust tube.
4. Fill agarose-coated injection dishes with 0.4X MMR + 6% Ficoll.
5. Set up a reaction. Sample reaction (~1:10 dilution of sperm stock): 4 |L sperm stock (~4 x 105 nuclei) and 5 ||L linearized plasmid (150-250 ng/|L). Incubate for 5 min.
6. Add: 0.5 |L of an ~1:5 dilution of XbaI or NotI, 2 |L 100 mMMgCl2 (add to 5 mM final at all steps to aid enzyme action), and 25 | L high-speed extract.
7. Mix the reaction by gentle pipeting (using a clipped yellow tip). Incubate for 10 min at room temperature; sperm will now be visibly swelled if diluted into Hoechst as before and observed with a 10X-20X objective.
8. While sperm are swelling in reaction mixture, collect eggs from individual frogs and dejelly them in 2.5% cysteine hydrochloride in 1X MMR (pH 8.0 with NaOH).
9. Under the dissecting microscope, inspect the eggs released from each frog for general health (eggs with even pigmentation and that remain round after dejellying). Draw the healthiest eggs into a wide-bore Pasteur pipet and transfer them to the square space in the injection dish. We generally fill the square space with eggs such that no space is left between the eggs. After about 5 min in 0.4X MMR + 6% Ficoll, the eggs will pierce easily.
10. Dilute the sperm into sperm dilution buffer (SDB) at 1:25-1:100 (such that the final dilution is 1:250-1:1000 or a concentration of 1-2 sperm/10-15 nL injection volume). For some enzymes, such as NotI or XbaI, add MgCl2 to 5 mM to aid enzyme action.
Before removing sperm from the stock tube or from the dilution used for injection, always mix thoroughly with a cut yellow tip, since sperm will rapidly settle out of the suspension.
11. Use a piece of Tygon tubing attached to a yellow tip (as previously described for Sigmacoting needles) to draw up the dilute sperm suspension and back-load the needle. Reattach the needle to the micromanipulator, and turn the air pressure up just slightly so that solution begins to flow from the needle tip (seen under the microscope as a schlearing solution of a different density). Owing to the low air pressure, solution will flow out of the needle only when the tip enters the liquid.
12. Transplant sperm nuclei into unfertilized eggs. The rate of flow should be robust enough that the needle does not reverse flow or clog with cytoplasm during injections and slow enough to be manageable. At the flow and injection rates we generally use, about 10-nL vol is delivered in each injection, so a 1:500-1:1000 dilution of the original sperm stock allows approximately one sperm to be injected in that volume. Move the needle fairly rapidly from egg to egg, piercing the plasma membrane of each egg with a single, sharp motion and then drawing the needle out more slowly. The angle of the needle should be perpendicular to the membrane surface (rather than glancing) to avoid tearing the plasma membrane.
A hole about the diameter of the needle tip should be visible on the egg and should remain open for about 5 s after injection; when the flow is too low, the hole created in the egg by the needle instantly closes over after injection and little or no volume is delivered. When the flow is too rapid, the surface of the egg near the injection site may ripple or the site of injection may expand in size significantly. If the needle becomes clogged with cytoplasm, bring the tip to the airliquid interface of the dish. Sometimes the surface tension of the interface removes the cytoplasm plug in the end of the needle. If a needle tip is too narrow, or if it becomes partially clogged with debris during transplantations, the injected nuclei will be damaged during transplantation, and haploid embryos will result.
Haploid tadpoles have shortened trunks and tails, are thicker than normal throughout the trunk region with a "pigeon-chested" appearance, and often have heads and tails that curl toward the dorsal side; these tadpoles will live for a while, but usually become edemic and die around the time of feeding (21).
You can determine whether your sperm dilution and the flow rate used for injections were appropriate by watching the first cleavage of the transplanted eggs. If few of the eggs received a nucleus, the frequency of cleavage will be low; one-fifth to one-third of our transplantations typically result in normally cleaving embryos. If too much volume was injected into the eggs, they may also fail to cleave; in this case, the animal hemisphere pigmentation may appear mottled or "marbleized," or have other signs or unhealthiness owing to overinjection. Eggs that were injected with more than one nucleus will divide at the time of first cleavage abnormally into three or four (or more) cells. Many of these embryos will develop to blastula stages, but most fail during gastrulation; in some, a region of the embryo will fail to cellularize and die. Eggs injected with multiple nuclei that do gastrulate usually do so abnormally; typically, blastopore closure is incomplete, resulting in embryos that form two wings of somites and neural tissue on each side of the exposed yolky tissue lying in the center of the trunk. This type of gastrulation failure is common to stressed or unhealthy embryos (particularly embryos derived from "soft" eggs).
13. When the cleaving transplant embryos have reached the 4- to 16-cell stage, gently separate them from uncleaved eggs and move with a wide-bore Pasteur or Spemann pipet to a separate dish of 0.1X MMR + 6%Ficoll + 50 |g/mL gentamycin. We commonly culture transplanted embryos in 6- or 12-well tissue-culture dishes with about 10-embryos/well, since culturing embryos at high density can compromise their health. It is also important to remove dying embryos promptly, since they also can compromise the health of their siblings.
14. When embryos are around stage 12, media is replaced with 0.1X MMR + 50 |g/ mL gentamycin without Ficoll. Because of the large needle tip used for transplantations, embryos often develop large blebs at the site of injection. These blebs occur when cells are forced out of the hole left in the vitelline membrane at the injection site, but they generally do not affect development. The blebs usually fall off on their own at the neurula or tailbud stages, but they can be removed manually once the embryos have reached the late blastula stage.
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