Embryo Fixation

Descriptions of all solutions can be found in Reagents and Recipes.
  1. Collect eggs from healthy, well-fed flies on apple or grape juice agar plates smeared with yeast paste. Flies on a solid, light-regulated circadian rhythm lay eggs best, and will lay in bursts around their perceived dawn and dusk. I don't store embryo collections on the plates at 4° C. and then pool them into one 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 get them into your favorite embryo mesh 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. But the clumps are easily dissociated with the wash buffer and any residual bleach hiding in pockets between embryos will be removed. I usually alternate three times between the two, and wash for a total of 2-3 minutes. Your nose is a reliable sensor of residual bleach. Do a final wash in ddH2O, making the embryo clumps which are then quite easy to pick 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: I usually use 220-230 rpm. The original Tautz and Pfeifle paper on whole-mount RNA in situ hybridization states the need to maintain an effective emulsion between the organic and aqueous phases during fixation. You have 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, in my opinion. 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. You can try to disrupt the bubbles with a glass pasteur pipette. Having a very small amount of methanol in the pipette touching the bubbles as you drag it through them can also help if they are very stubborn. One reason you might get many stubborn bubbles between the phases is that you've put some small pieces of agar from the collection plate into the vial, so try to avoid gouging the surface of the agar plate when you're brushing up the embryos at the beginning. Normally, the bubbles will pop by themselves within a couple of minutes. Completely remove the bottom, aqueous phase with a pasteur pipette, avoiding pulling up the embryos. To get the very last bit of the bottom phase, you can use a narrow-tipped pulled pasteur pipette or a p200 micropipette.
  7. Then 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. You should have maintained an upper phase of heptane, and all the burst vitelline membranes and non-devitellinized embryos will have remained in a cloudy layer at the interface. First remove the heptane, then all the debris at the interface, then most of the methanol, leaving the embryos covered in a few millimeters of methanol. Rinse the embryos in 1 ml fresh methanol, trying not to rinse the embryos stuck to the side of the vial down onto the nicely settled ones. Why contaminate your beauties with the defective rejects and random debris? Finally, tilt the vial so that the embryos sink to one side, and using a clean short pasteur pipette gently blow the embryos with the methanol, then take this 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. That's it! Now the embryos can be stored long term under ethanol at -20° C. Many people store their embryos in methanol at -20° C.; I don't know if it makes much difference, but methanol is a stronger fixative than ethanol. Also, it's said (Sullivan et al., 2000) that embryos that have spent some time in cold storage make nicer stains. I agree, and in practice rarely have enough energy to fix embryos and set up a hybridization on the same day, so that storage time happens by default.

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 popping out into the methanol. If you encounter this problem, make sure removal of the aqueous phase is complete, but 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, a good alternative is to do the fixation in a 2 ml eppendorf tube with scaled-down volumes of reagents, although it's important that a good emulsion of the phases is still achieved during shaking. In the protocol presented here, my experience has been that more embryos work better, so get more females to start with to make more eggs. You'll end up doing fewer and better fixations at the cost of sorting some more flies.

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