Embryo Fixation

1. Collect eggs from healthy, well-fed flies on apple or grape juice agar plates smeared with yeast paste. Flies placed on a consistent, light-regulated circadian rhythm are best for collecting eggs, and will lay eggs in bursts around their perceived dawn and dusk. It is not recommended to store embryo collections on plates at 4° C. for fixation at a later time.
2. Squirt some embryo wash buffer on the plate, and gently brush the embryos off the agar with a paint brush and pour them into small mesh embryo basket. Wash away the yeasty suspension with the embryo wash buffer.
3. Dechorionate the embryos in ~50% bleach for 2-3 minutes. Dunk the basket in the bleach, and periodically squirt some from outside the basket onto the embryos to provide some gentle stirring.
4. Wash the dechorionated embryos thoroughly to remove all traces of bleach. Alternate washes between double distilled (dd)H₂O, which causes the embryos to clump together, and the embryo wash buffer, which breaks the clumps. Perhaps the exposed vitelline membranes of the embryos are somewhat hydrophobic and cause them to associate in the ddH₂O. These clumps are easily dissociated with the wash buffer, however, and any residual bleach in pockets between embryos will be removed. Alternate washes between ddH₂O and the wash buffer two or three times, washing for a total of 2-3 minutes. Do a final wash in ddH₂O, making the embryos clump, which are then quite easy to be picked up with the wet paint brush.
5. Using the paint brush, transfer the embryos to the fix buffer in a 20 ml scintillation vial. After the transfer, add the formaldehyde and heptane. The embryos should now float at the interface between the two phases. Cap the vial and tape it on its side to an orbital platform shaker. Shake it hard for 25 minutes at 220-230 rpm. The original Tautz and Pfeifle paper on whole-mount RNA in situ hybridization states that it is necessary to maintain an effective emulsion between the organic and aqueous phases during fixation. There is a choice of fixation buffers and formaldehyde solutions, specified in Reagents and Recipes. The optimum range of embryo volume to add to one scintillation vial yields 20-75 µl fixed, devitellinized embryos settled in methanol in an eppendorf tube. Loading too many embryos in a vial is detrimental to the quality of the fixed embryos. Also, loading too few (<10 µl) seems to create some difficulty at the devitellinization step (see below).
6. After shaking, let the bubbles at the interface pop. It is possible to disrupt the bubbles with a glass pasteur pipette. Having a very small amount of methanol in the pipette touching the bubbles as the pipette is dragged through them can also help for very stubborn bubbles. One cause of these stubborn bubbles between the phases is if some small pieces of agar from the collection plate get into the vial, so try to avoid gouging the surface of the agar plate when collecting the embryos at the beginning. Normally, the bubbles will pop by themselves within a few minutes. Completely remove the bottom, aqueous phase with a pipette, avoiding pulling up the embryos. To get the very last bit of the bottom phase, a narrow-tipped pulled pasteur pipette or a p200 micropipette can be used.
7. Add 8 ml methanol, cap the vial and shake vigorously by hand for 20-30 seconds, swirl and place it on the bench. Watch the two phases separate and the fixed, devitellinized embryos settle to the bottom in methanol. There still should be an upper phase of heptane, and all the burst vitelline membranes and non-devitellinized embryos remain in a cloudy layer at the interface. First remove the heptane, then all the debris at the interface, and lastly most of the methanol, leaving the embryos covered in methanol to a few millimeters depth. Rinse the embryos in 1 ml fresh methanol, trying not to rinse the embryos stuck to the side of the vial down onto the settled ones, so as to not contaminate the devitellinized embryos with vitelline debris. Finally, tilt the vial so that the embryos sink to one corner, and using a clean short pasteur pipette gently squirt the embryos with the methanol, then take the embryo slurry up into the pipette and transfer them in methanol to a 1.5 ml eppendorf tube. Wash the embryos with 3 changes (1 ml) of methanol, then with 4 changes of ethanol. The embryos can be stored long term under ethanol at -20° C. or in methanol at -20° C., although methanol is a stronger fixative than ethanol. Also, it is claimed (Sullivan et al., 2000) that embryos that have spent some time in cold storage make better stains. We have found the same improvement.

One potential problem with the devitellinization step is that a significant percentage of the embryos remain at the organic/aqueous interface. This seems to be more common when working with small numbers of embryos and is probably caused by not removing enough of the aqueous phase after the shaking fixation. Perhaps the embryos find refuge in the remaining aqueous pockets instead of entering the methanol. If one encounters this problem, make sure that the aqueous phase is completely removed. Also, try shaking the vial for a longer time after the addition of methanol, 1-2 minutes instead of 20-30 seconds. When working with small numbers of embryos, an alternative is to do the fixation in a 2 ml eppendorf tube with scaled-down volumes of reagents, although it is important that a good emulsion of the phases is still achieved during shaking. In the protocol presented here, our experience has been that more embryos work better, which necessitates more female flies in the embryo collection cages. This will result in doing fewer and better fixations at the cost of sorting extra flies.

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