Embryo Collection

Abstract

Before beginning, add yeast paste to enough new apple juice plates for all of your cages (use juice plates that have been pre-warmed to room temp if possible). 1. Remove the apple juice plate from the cage and replace with a fresh plate. 2. Dechorionate embryos in 50% bleach (squirt directly onto egg lay plate; no need to remove yeast) for 2-5 min. 3. Prepare fixative in glass scintillation vials: H2O 4.5ml 37% formaldehyde 0.5ml Heptane 5ml 4. Wash dechorionated embryos into collection net with H2O. Rinse embryos and sides of collection tube well. 5. Unscrew collection cap, pick out net with forceps, blot off H2O, dunk into fix vial and shake off embryos. If desired, collection tube/cap may be rinsed a second time onto net to transfer any remaining embryos. 6. Screw cap onto scintillation vial and fix on nutator for 12-15 min (longer fixation up to 20 min produces stiffer/more durable embryos, but these may be harder to dissect). 7. Remove fix (aqueous bottom phase) with glass pasteur pipet. Remove as much as you can without aspirating embryos. Keep separate beaker for heptane/formaldehyde waste; dispose of in waste container in hood. 8. Add 10ml 100% methanol, or until scintillation vial is nearly full. Devitellinize (“crack”) embryos by shaking for 30 sec. Devitellinized embryos fall to bottom; undevitellinized embryos and membranes stay at interphase. (If you add enough methanol there won’t be an interphase and all will fall to bottom.) 9. Aspirate upper phase (heptane), interphase and most of methanol. 10. Wash embryos twice with methanol in scintillation vial, then transfer to 1.5ml tube for storage. Be sure to label your tubes so that you know what cross or stock was used to produce the embryos.