Fixation and permeabilisation

  1. Collect embryos on grape juice agar collection plates, and rinse them onto a Nitex mesh sieve, rinsing them well with water containing 0.05% Triton-X-100 (optional) to remove excess yeast.

  2. Dechorionate in 50% household bleach (2.5% sodium hypochlorite final conc.), by dipping the whole Nitex sieve into a beaker of bleach for 2-4 minutes and swirling the embryos once or twice. Rinse well in PBS/0.05% Triton-X-100 to remove the bleach. Do not let the embryos dry out at this stage.

  3. Transfer the embryos with a soft paintbrush to a vessel with a tight-fitting lid containing:

    PEM-formaldehyde is:

  4. The size of the vessel, and the quantity of fixative should be adjusted according to the number of embryos used. For about 50-500 emryos I use a 5ml glass vial with a screw-cap; for 0.5-1g of embryos I use a 50ml polypropylene tube with a screw cap. It is a good idea to use enough fix/heptane to fill the container to at least 70-80%, as this stops too many embryos being lost by sticking to the sides.

  5. This fix is a good all-purpose one, but there may be some antigens which are destroyed by it. The most likely source of problems is the formaldehyde solution, which contains up to 10% methanol as a stabiliser. Some antigens (for example 24B10 antigen) are methanol sensitive, in which case, either buy expensive and unstable formaldehyde without methanol, or make it by dissolving paraformaldehyde in water. To do this heat the water to near boiling and add a drop or two of 1M NaOH. You can substitute PBS or Balanced Salt Solution (for recipe see Wilcox, in Roberts: Drosophila, A Practical Approach. IRL Press. 1986) for the PEM, but I like PEM the best for most cases. Other fixes are possible: anything you know preserves your antigen should work, though there might be penetration problems if you use too high a glutaraldehyde concentration.

  6. Vigorously shake the embryo/heptane/fix for 15-30 minutes at room temperature.

  7. With a pipette, remove as much of the aqueous (bottom) layer as possible, without taking the embryos at the interface.

  8. Add 1 volume of 100% methanol to the embryos/heptane, and shake very hard for 30 seconds-1 minute. Allow the two phases to separate. The devitellinised embryos will sink to the bottom, and the embryos with their vitelline membranes still attached and the empty vitellines will remain at the interface. This step is not always very efficient: anything over 50% sinking is fine, and on a good day it should be 80 or 90%.

  9. Remove the heptane (top layer), all the interface, and the methanol above the embryos with an aspirator.

  10. All the subsequent steps are carried out in the same tube, which can be filled with a solution, the embryos allowed to settle, and the solution removed.

  11. Treat the embryos with:

    	100% methanol 		10 mins  (at this stage, embryos can be
    	PBS/0.2% saponin	10 mins    stored at -20C for several monthes)
    	PBS/0.2% saponin	10 mins

    The embryos can be stored at 4C like this for a week or two.

    Before staining:

    PBS/0.2% saponin/5% serum (normal goat or foetal calf)	10 mins

    As discussed above, some antigens are methanol sensitive, and in this case the vitelline membranes have to be removed by hand. In this case, substitute the following for 6-9 above:

    6a) Remove the aqueous (lower) phase. Take some embryos in heptane with a pipette, and spread them dropwise onto a microscope slide covered with double sided sticky tape. Let the heptane evaporate, and just as it does, immerse the slide in PBS/0.2% saponin in a petri dish; do not let the embryos dry out on the slide. Using a fine tungsten needle, gently tease the embryos out of their (already cracked) vitelline membranes; this step is made much easier if the embryos are initially fixed for longer than normal (about 45 minutes). Transfer the embryos to an Eppendorf tube of PBS/0.2% saponin/5% serum, using a Gilson pipette with a yellow tip cut off to make a wide enough opening.

Antibody Staining

Stain the embryos in Eppendorf tubes or, for large numbers of embryos, 12x75mm polypropylene tubes (Falcon 2063).

  1. Incubate the fixed embryos in primary antibody diluted in PBS/0.2% saponin/5% serum. Typically monoclonal supernatant is diluted about 1:5, polyclonal serum about 1:500, and ascites even more, but you should determine the best for your antibody. Incubate for 2 hours at room temperature, or overnight at 4 (which is probably a little more sensitive).

  2. Rinse in 3 changes of PBS/0.2% saponin for 10 minutes each (longer washing may reduce background).

  3. Incubate in secondary antibody diluted in PBS/0.2% saponin/5% serum for 2 hours to overnight. In most cases, I use peroxidase conjugated 2nd antibody, and do a DAB reaction; this is the most sensitive detection system available, and can be made even more so with silver-gold intensification. However, a fluorescent conjugated 2nd antibody may be used, in which case, I prefer rhodamine or Texas red to fluoroscein, since they are less prone to quenching under epifluorescent illumination.

  4. Rinse in 3 changes of PBS/0.2% saponin for 10 minutes each, and one rinse of PBS. For fluorescent labelled embyos mount in 85% glycerol containing 2.5% .i.isopropyl gallate,; and view with epifluorescent illumination. An optional .i.Hoechst 33258; stain can be added to the penultimate rinse (1g/ml; caution - mutagen); this will strongly label all nuclei, and is viewed with ultraviolet epifluorescence. Slides mounted in glycerol dry out after a few days unless the edge of the coverslip is sealed with nail varnish.

  5. For the DAB reaction, when using peroxidase-conjugated 2nd antibody, incubate the embryos in 400ul of 0.5mg/ml diaminobenzidine in PBS. After 10 minutes, add 2ul of 3% hydrogen peroxide and watch the development of the brown reaction product under a stereomicroscope. When the reaction is sufficient (usually 5-15 minutes), stop it with 2 or 3 rinses in PBS. (DAB can be made up in 10X aliquots (5mg/ml in water) and stored at -70. USE GREAT CARE WHEN WORKING WITH DAB. IT MAY BE A VERY POTENT CARCINOGEN. Use copious amounts of bleach to inactivate it (leave overnight), and then rinse it away with large amounts of water. Treat gloves, tips, and tubes in the same way.)

  6. Either take the embryos through .i.silver-gold intensification; (see Liposits et al. Neuroscience 13, 513-525. 1984 and the previous edition of the Rubin Lab Manual) or mount them in 85% glycerol, or dehydrate them through an ethanol series, clear them in methyl salicylate until they sink, and then mount them in the methyl salicylate (this gives the best results, but is less straightforward than aqueous glycerol).


In this protocol, I have suggested using 0.2% saponin throughout the staining. This is a good, very mild detergent which should not extract many antigens. However, if it does not work, you could try using no detergent (which makes the embryos sticky, and reduces the ease of antibody penetration), or substituting another detergent. If your antigen is nuclear, 0.1% Triton-X-100 might be better.


The best overall discusssion of these, and many other, techniques is: Immunocytochemistry: Theory and Practice. Lars-Inge Larsson. CRC Press (1988)

original text by Matthew Freeman