My lab is trying to eliminate alcohol steps from our H/E protocol in order to minimize tissue shrinkage for stereology purposes. We are using 80um free-floating dog brain tissue (no paraffin), so the initial deparaffinization/rehydration is not necessary. The steps we are having trouble with are the ethanol rinse before the eosin, and the eosin stain itself. The problem is that A) we want to eliminate alcohol, and B) the eosin stain dissolves out upon coverslipping using our aqueous mount PVA-DABCO (as opposed to permount, the organic mount).
Does anyone have any protocols that don't involve the use of alcohol with eosin? Does such a thing exist? Does it stain better/worse? Are there specific types of eosin we must purchase?
As stated by Matthew, aqueous eosin solution should work, but the problem he highlights would be the leaching of the stain from the tissue into the mounting medium. So why not try the following? I have never tried it, but I see no reason why it should not work. Stain your tissue with haematoxylin as usual and depending on the type of haematoxylin used, either 'blue' or treat with a mordant. Bring sections to water. A semi-permanent preparation could then be prepared by mounting in glycerin jelly which has already been pre-stained with eosin. In this case, the eosin should diffuse from the mountant into the tissue to produce double-stained sections. Of course, you would need to ring the coverslip and keep slides horizontal. Just a thought, but it might just work.
Matthew mentioned the most widely used aqueous solution of Eosin. I have used it in past to counterstain free-floating 50 micrometre human punch skin biopsies (fixed in paraformaldehyde). The tissue was immunostained, then counterstained and coverslipped with DAKO aqueous mountant and secured on the outside edges of the coverslip with permanent mountant (to protect against evaporation). My goal was to have a strong immunstaining and weak counterstaining, so leaching of Eosin into the aqueous mountant was not important - I had the sections photographed as soon as the coverslips were secured in place. This particular mountant 'gellifies' (s0lidifies) after initial fluid phase, so the leaching is slowed down, then stops. If you contact me directly - If you would wish to contact me direcly - I might be able to help you with some more details. I was, the same as you, concerned with eliminating tissue shrinkage, so I was using an aqueous mountant. My tissue was'already shrunked during fixation, but alcohol step would shrink it further, so my problem was partially similar to yours - therefore using aqueous eosin and solidifying aqueous mountant might be a good solution - possibly even with experimenting with a mountant with a small amount of added eosin. A word of warning, though - the mountant has to be kept free of air bubbles, so after mixing with a drop of eosin you would need to stand a bottle in upside-down position (in a small beaker). 80 um sections are quite thick, so you need to use quite diluted Eosin if you do not want to have too dark staining obscuring your images.
I can recommend eosin Y made by Fluka to you. I used it a few years ago and I dissolved it in destilled water.
Concerning mounting media: in our lab people use glycerin or protein glycerol (catalogue number P049.1 on http://www.carlroth.com ).
All the best.
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