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Sectioning stained embryos.

The protocols for plastic and wax sections as used by the Vize lab. These protocols work, but they have not been optimized. If anyone has better protocols please let us know.


Stain samples strongly, remember that in a section only the very strongest stain will be visible, even quite strong background will not be visible. BCIP/NBT is the only stain we have successfully used, we have tried Red new fuschin but it was not stable, others have not been tested.

Fix in Bouins (recipes) or in formalin (60 minutes) to stabilize stain. Rinse Bouins fixed sample extensively in 70% ethanol, rinse formalin fixed samples twice with TBS (recipes). Rinse and store in 100% methanol or 100% ethanol.
For best sections use samples that have not been previously been cleared in BB:BA, it makes them too fragile.


Standard protocol.

1) take samples in 100% methanol, remove methanol and replace with 100% ethanol, leave 5'

2) exchange ethanol with fresh 100% ethanol, leave 5'

3) repeat step 2 (note it is important to remove all water it will interfere with wax infiltration)

4) replace half of ethanol with HemoD (a less toxic xylene substitute) leave 10'

5) replace all of solution with HemoD, leave 10'

6) repeat step 5

7) warm bottle in 55 deg C oven containing molten paraplast. Replace 50% of HemoD with molten wax. Let it sit in the oven for 5 to 10'

8) Remove as much as possible of the HemoD/wax and replace with 100% wax. Mix and leave in the oven for one hour.

9) repeat step 8 twice more, each change left in oven for at least 30 minutes. Leave in final change in oven overnight.

10) pour a shallow layer of wax into a mould, when it has partly set transfer in an embryo USING A WARM glass pipette, and orientate such that its head is facing one side of the block.

11) Once the wax holding the embryo has hardened a little, fill mould with molten wax and put it aside to set.

Leave blocks until the following day. Section at 10 microns (thicker is often useful for faint stain)


Remember plastic sections work best when very thin (1 to 2 microns), so only strongly stained samples will work. The advantage of this method over wax is in resolution, which is much better. We use the Drosophila fly eye protocol of Tomlinson and Ready (1983) as supplied by Janice Fischer Vize and Kathleen and it works very well.

1) dehydrate sample through to 100% ethanol

2) replace 100% ethanol with propylene oxide (wear gloves and use a fume hood)

3) after 5 minutes, repeat step 2

4) remove propylene oxide and replace with 1:1 mixture of propylene oxide: durcupan (see below). You will need a volume of about 1 ml of each to get them to mix. Vortexing may be required! If this is damaging your sample replace the prop ox with durcupan more slowly, e.g. 10 to 20% increments

5) incubate overnight in 1:1 mix

6) replace 1:1 mix with straight durcupan. Once again, if mixing is too difficult, try slower steps

7) leave in staright durc for at least 4 hours, overnight is OK but no longer, as the durc will get too sticky

8) place the embryo in a small mould and orient.

9) bake in vacuum over at 80 deg C for 48 hours

10) slice at 1 to 2 microns, we use a fresh glass knife.

11) we have had little luck with counterstains, but have found them unnecessary with phase optics.

Other plastics can be easily substituted. See the Ted Pella catalog for a number of epoxy alternatives.

How to make durcupan.

1. Buy a kit from Fluka (cat# 44610)
2. Read the instructions!

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