Homebrew Resources for Bryology
Yesterday I found myself lingering in the stool-softener section of the pharmacy trying to find the product that contained docusate sodium. Turns out this is the second product in the laxative aisle that is helpful for looking at bryophytes. Many of these tools and techniques are described in two great resources for home bryology:
(1) @david1945wagner 's Tools, Tricks & Techniques section of his website. Here you can get tips for simplifying microscopy, microscope photography, specimen hydration (stool softener!, special needlepoint applicator bottles), staining, slide preservation and more. Dr. Wagner has also put out two really illustrative videos on making herbarium packets and botanical illustration.
(2) Janice Glime's Lab Techniques chapter (cowritten with David Wagner) from her fantastic five volume open access Bryophyte Ecology book. This chapter gets into the nitty gritty of specimen cleaning, staining, mounting and preserving for a variety of uses. This is where you can learn about
- Cleaning specimens: I use a cheap ultrasonic bath intended for jewelry cleaning
- Staining and Clearing bryophytes (eg. Methylene Blue, a readily available fish medication, can be used for staining sphagnum and spores, KOH for clearing and staining leaves)
- How to section (freehand with drug store razor blades)
- Microscope "hacks"
The one technique not mentioned in these two resources that I find very useful is another product from the laxative aisle-- Polyethylene Glycol 3350 (PEG). I learnt about this in @mccuneb & Martin Hutten's Common Mosses of Western Oregon and Washington book. The use of PEG can feel illicit on account of it being a fine white powder that you put on glass and hold over a flame, but it is really helpful for making sections of tiny or delicate leaves. I simply put my leaves or shoots or sporophytes on a glass slide, cover it with the dry PEG granules and then hold a lighter under the slide until the granules melt and surround my specimen. The slide is then left to cool for about a minute, leaving me with an "embedded" specimen without the trouble of parafin and dehydration. The trimmed specimen block is then ready for sectioning.
Bulk Sectioning of Bryophytes
It is crude, but here is a video demonstrating my bulk sectioning technique. It is one way to get good stem and leaf cross sections, best applied when positional accuracy is of no great consequence.
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Thanks for summarizing this @rambryum. I've used many of those resources but still struggle with leaf sections as I come to bryophytes late in life :) Tips help, even if they do involve lingering in the stool-softener section of my local pharmacy.
Who knew! I will have to check these out at my pharmacy. By the way, I have found the Microscope Preparation techniques appendix of Sphagnum Mosses of Eastern Canada (G. Ayotte and L. Rochefort, 2021, Bibliotheque et archives nationales du Quebec and Archives Canada) to be extremely helpful, with step by step instructions and illustrations.
@bstarzomski Definitely try different ways of sectioning to see what works best for you. Personally I go for a bit messier of an approach, stripping a bunch of leaves and stacking them, sandwiching them between slides, and then using the top slide edge as a guide for the fresh razor blade. Generally with care you can get many informative sections with variation somewhat represented. When sectioning leaves where the position of the section on the leaf is informative (ex Racomitrium sl) this may not be the ideal approach as you have to judge the position of the section on the leaf by the relative widths of other sections. I do this without using a dissecting scope and it seems to suffice (for IDs), but for teeny tiny leaves or if I want to be confident about the position of the section, I use a similar approach but with just one leaf/shoot under the dissecting scope with careful hands.
@cwardrop @bstarzomski I have now made and added a video of my bulk sectioning technique to this post. Video production is not my specialty, as you will see, but this covers the bases. My technique is different than Conner's only in that I use the wood block which I have found gives me more traction than two slides and may allow my razor blades to survive for a few extra slices.
https://www.youtube.com/watch?v=DBL5ceqqJns&t=6s
That is super, @rambryum. Feel free to make more! For Sphagnum, I feel I have a lot to learn about staining. The first directions I had said the stain should be intense, so I used undiluted crystal violet. My tissues were so stained they barely let any light through. After noticing that other people's slides looked a lot less gunky, I tried various dilutions, some too weak, but I am still experimenting with what shows structure the best (I give up on aesthetically pleasing). I know other stains are used as well. A video of staining would be great.
@rambryum I never aspired to be a regular guy, but have you ever tried playing with Rain-X? It is freaky stuff, and recently made waves in the world of microscopy for marine microinverts. I have often wondered if it might work for some mosses (or even more likely, liverworts or hornworts) that have similar properties. Available in the auto section of any hardware store.
See https://pubmed.ncbi.nlm.nih.gov/34118945/.
@jstraka you have no idea how excited I am to try this. Thanks for the tip.
Please report on it if you try it!
@janetwright RE Sphagnum staining; The process I do is 1) Fully rehydrate the branches 2) Place an end of a branch in a drop of undiluted crystal violet, waiting for the stain to wick through the rest. 3) Place the stained branch in a drop of clean water and shake it around with forceps, (maybe repeating with another drop of clean water). 4) Strip the leaves on a slide in another drop of clean water.
I think rinsing after staining rather than initially diluting the dye may help.
You could also try making a normal wet mount and then putting a drop of dye on one side of the coverslip and a very absorbent wipe on the other to draw the dye through the sample. Repeating this afterwards with clean water will help clear things up.
Cheers.
Thanks, @cwardrop, will try it!
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