Onion epidermis: fixed by mild heat then stained

[Hobbyst46]

This is only my 3rd or 4th attempt to make semi-permanent stained onion skin slides. There are many variants of this preparation in the literature.
Previously I stained the epidermis as is, fresh from the onion, mounted in water on the slide under a coverslip, with variable success. Often the epidermis would curl or even roll into a narrow "tube" during staining, failing the specimen. Also, the epidermis would float on the stain solution, the top layer appearing to be hydrophobic and dry.

So I tried the following protocol, after W. Dioni (more or less), as follows:
1. Throw the peeled epidermis immediately into a mix of 30ml tap water+45ml boiling water, for 5 min. The temperature starts at about 66 and falls to about
49C.
2. Transfer the epidermis into a 0.05% (approx.) solution of the (powder) dye in distilled water (DW). No buffers or co-solvents. Wait 3-5 min.
Stains: Neutral Red; Methylene Blue; Safranin; Toluidine Blue.
3. Transfer to DW (rinse excess dye) for about 10 min. Repeat the rinse.
4. Transfer to a 40% solution of fructose in DW. Wait 1 hour.
5. Mount in 85% fructose.

Results: (1) The epidermis remains fairly flat after the heat fixation, yet the hydrophobicity of the top layer remains, and it floats in the stain bath.
(2) The epidermis still floats, or curls and rolls into a tube especially in the Neutral Red bath but also in other baths. Diluting the stain helps
somewhat, at the expense of the final color intensity of the specimen appearing to be hydrophobic and dry.
(3) I straightened the curled pieces with a tooth pick, but some of the specimen is physically damaged and tears down easily. It is difficult to
obtain a well-spread specimen and uniform staining of its area.
(4) After 2 days: almost no leaching of the stain from the specimen into the mounting media.
(5) Very few air bubbles in the slide (might be related to the mounting medium, not the fixation).

Some photos are shown.
Fields of view are chosen from best parts, excluding folded parts as much as possible.
Single frames, no post processing.
Please ignore the small artifacts of near the top corners.
Comments are welcome!

[Hobbyst46]

(continued from above)

[mrsonchus]

Great set, really nicely-done and a nice description of your method - thanks for posting.

Personally I think your depth of staining is just-right, optimizing detail without the over-staining that is so easy to fall-foul of when considering the staining-level by looking at the slide with the naked-eye rather than through the 'scope.

This is especially true with the two related stains, Methylene-blue and Toluidine-blue - and your use here of those two is excellent to my eyes.
Nicely balanced and very good detail, both nuclear and cytoplasmic (skeleton). Safranin is also a favourite of mine - and this too has performed nicely for you - in fact, I like them all for slightly different reasons.

Is this epidermis from the 'inside surface' of the onion's 'layers'? This is the upper surface of each layer, which are I'm sure you know leaves. If you use the 'outer' epidermis you'll find many stomata in your sample - these are really nice to see stained also.

Great start, lovely results - please show us more as you go on your adventures - your posts are great to read and very interesting.

John B.

[Hobbyst46]

@mrsonchus:
Thanks a lot! You removed my concern. I am relieved to know that the staining level is acceptable.
These are indeed the lower epidermis from the leaves inside the bulb, the ones WITHOUT stomata. I have already inspected the upper epidermis and stomata
but thought that they would not be stained well. Having just now seen your 2015 post, I change my mind and try it as well, and maybe get some Fast Green too.
BTW, I noticed that you used a dye in cellosolve solution - I prefer water solutions for home use.
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[mrsonchus]

Oh - yes, back then I bought my stains 'off the shelf' - and the cellosolve was the solvent that's commonly used by suppliers of pre-mixed Safranin and indeed Fast-green. I can't get hold of any cellosolve to copy the formulas, and now use only aqueous Safranin - 1% w/v is just-right (Safranin in water/alcohol is I find pretty useless - the alcohol inhibits the Safranin far too much.

On the other hand, using Fast-green in anything below about 85% alcohol with de-ionised water (or of course toe completely anhydrous cellosolve formula) will be pretty hopeless - any water following staining with Fast-green will very rapidly and completely wash out the Fast-green.
This is why it's used after aqueous Safranin staining and a water-rinse, then movement to alcohol - alcohol does indeed remove (differentially) Safranin - but slowly enough to make it's (alcohol's that is) use with post-Safranin stained tissue fine (desirable in fact when used as a differentiator for Safranin) as a preparation for a following Fast-green stain - which is then washed with alcohol (any water in the rinse will remove the Fast-green)...

I'd suggest,
Stain with 1% aqueous Safranin for about 2 minutes,
water (DI) rinse away excess stain until stain stops coming out of tissue (epidermis) (about 30 sec - 1 minute will do fine)
move tissue into alcohol at 25% for about 1 minute,
move to 50% alcohol again for about 1 minute,
move to 75% alcohol for another minute,
move to the highest concentration of alcohol available to you (I use Isopropanol, and it's maximum is 95% - which may be considered 'pure' I find) for another minute - now the tissue is ready for Fast-green.

Using say 0.5% (w/v) Fast-green in 85% alcohol (isopropanol mixed with DI water calculated as though the 95% alcohol is actually 'pure') you will only need to stain very briefly - Fast-green lives up to it's name! Stain for about 5-10 seconds then rinse a couple of times with pure alcohol - then mount in an alcohol-based mountant - this will give you a pretty long-lasting mount if you seal it with nail-polish for example after a day or so.

To 'keep things aqueous' Buy some Alcian-blue (powder) and mix a 1% aqueous solution as stock.
Mix the 1% Safranin and the 1% Alcian-blue 1:1 and keep as your working-solution.

This is like a 'magic mixture' and will stain your tissue differentially - for your onion epidermis you should get bright blue non-lignified cell walls (the epidermis cell walls) with red nuclei!
Simply stain in the mixture for say 5 minutes (you may need to try different times as live tissue isn't quite the same as the fixed and sectioned tissue I work with now) then rinse away excess stain in DI water - about 2 minutes dipping or flooding-and-draining repeatedly.

If the tissue is too red add a little vinegar (for the acetic acid content) to water (say 20% vinegar to 80% DIW) and rinse tissue - this should remove Safranin from cell walls and reveal the bright blue that should still be in the tissue as the Safranin masks it but doesn't displace it....

Rinse to DIW and mount in an aqueous mountant - perhaps fructose or gelatin etc...

I may give this a run-through myself tomorrow if I get the chance.

Good luck!

[Hobbyst46]

Thanks John B. for the very comprehensive info!
Yes, for the time being I prefer the water solution so the Alcian sounds a better choice.

In the past I found that glycerol (in which I mounted) leached stains (neutral red, methylene blue, congo red) from the plant tissue (although perhaps it was simply a high excess of dye in the tissue). Nail polish contains a powerful solvent, so I keep to mounting in fructose. So less transitions of the specimen into water->alcohols->water->alcohol etc.

What I still dislike about the onion is the waxy hydrophobic layer on the fresh epidermis. You kept your onion epidermis in 70% IPA for weeks. I consider using either pure (95+%) IPA, or 70/30 (v/v) IPA/Lactic acid, for an hour, say, or perhaps octane for just a few minutes, to partially break the wax. Might work or not work...and the epidermis of many plants is waxy, after all.

[mrsonchus]

Sounds worth a go. Perhaps the tiniest spot of wetting-agent may do the trick, perhaps washing-up liquid?

Interesting to see how you get on - I remember pushing the epidermis under too!

Keep us posted.