Collection location: Stockbridge, Henry Co., Georgia, USA [Click for map]
Very small, apothecia with raised margins. Thallus K-; apothecial discs K+reddish (maybe?)
Spores large, smooth and elliptical to amygdaliform; septate, with hour-glass like oil drops. Very large, ~17-24 × 7.8-13.7 μm; asci clavate, 8-spored.
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sum(score * weight) /
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I chalk it up to lack of prior experience or formal training, but I go around and around with the testing details. I get it that confidence grows with experience, but dang, sometimes I feel like it should “just work”!
For me, it refuses to diffuse only when I’ve done a wash with K first. Or if I squash it pretty hard. I’ve taken to using a trick of Frank Bungartz: he holds the far side of the coverslip, e.g. with forceps, and slips the edge of a razor blade under the side with the drop of lugol’s and pries up the coverslip a little. Sections go skittering every which way, but not as much as you would expect. And it can be very effective at causing the lugol’s to diffuse evenly over the entire coverslip.
With all due respect to Kerry, I haven’t yet observed my lugol’s failing to react no matter how old it is. His claim to me in personal emails was that he found that old lugol’s would still react I+ blue but then not change to red (hemiamyloid reaction is I+ blue turning red). It’s possible that was the mode of failure for his reagent. Perhaps my reagent fails the other way: reacting too strongly giving false positives? But I have yet to see a false negative which is what Kerry warns against. (I keep my stock solution in the dark, in temperature-controlled areas, and well-sealed, so I really don’t see why it should have deteriorated, anyway. And I go through it quickly, so my lab aliquot never gets very old.)
I sometimes wonder why I’m apparently the only one (maybe in good company with you, Jason!) that struggles so much with nitpicky details of these standard chemical tests. Everyone else seems to just do it, and never second guess it. I feel like I spend my whole life second guessing every last detail of all of my macro morphology, microscopy, chemistry, etc.!
I have very poor luck stacking compound scope shots. That’s only been effective a couple of times, despite repeated attempts. (You know, those sections that are just so perfect and crisp that you want to keep them somehow, whatever it takes!)
The dissecting scope is much better for stacks. I just hold the camera (Casio Exilim EXZ1080, discontinued make) against the eyepiece by hand(!) I use a 2 second shutter delay. And I advance the focus very slightly between each shot. Super low-tech, but I’ll be darned if it doesn’t work. It will never compete with what a “real” solution like Jason’s can do, or what a fancy rig with rail and bellows or super macro lens can do with controlled lighting, automated focus advance and shutter release, etc. can do. But it gets the job done.
Like Jason, I use CombineZP. I’ve tried using enblend etc. on linux, but I get really bad halo effects, and I haven’t figured out how to adjust the configuration to eliminate that yet. I would much prefer the linux solution, because then I could automate it completely. And I could finally get rid of my ancient copy of Windows XP that I run on Virtualbox within Linux, the sole reason for having is to run CombineZP!
I don’t remember what brand I have, it’s 2% Iodine and 4% Potassium Iodide (according to my notes). It just won’t easily go under the slip, always spreads around the perimeter, so frustrating! I’ve read Knudsen’s Acarospora testing notes touting fresh Lugol’s, which mine isn’t, so have always wondered if that was an issue in my results, Aspicilia also, as we have mentioned in other posts.
It was probably $100 a couple years ago, and works ok. It attaches like a lens to the camera, and fits into a sleeve that you put into the eyepiece after removing the actual eyepiece lens. The D3100 has a flip screen on the back which makes it work. I put it on a cheapo Amscope, and when I shoot a dozen or so pics and stack them using the freeware Combine ZP it works decently. All depends on how much care I take when shooting the different focal planes. Most of my closeups are with this method for the last couple years (see_observation 333012_), up until this winter when I just haven’t had time to do this and began shooting through the eyepiece with my Samsung, which is less than decent but does the job for basic representation. Jason has great shots that he has stacked apparently shooting through the eyepiece, I’d love to know how he got those so clean!
My partner and I tried a few microscope cameras. Most were worse than my point-and-shoot Casio through the eye piece, which is what I use. We finally got an OMAX microscope camera, and that, I have to admit, really is pretty good. Just a pain to use because I don’t run Mac or Windows. Blah. :(
I thought your micro photos were great. It doesn’t get much better than that. You’re always going to really struggle to get the focus just right. The depth of field is infinitessimal. That’s where the microscope camera with good software really helps. You get what you see. With the iPhone and point-and-shoot camera (DSLR won’t work because the lens is way too big, has to be cheap-o camera!) you just sort of take a bunch of photos at a bunch of random focus settings and hope for the best. Hope the spores are still there when you finally accidentally get the right focus!
re: heating — Yes, I just hold the slide over a bic lighter for 4-5 seconds. Too close to the flame and it gets soot on it. Too far and obviously it doesn’t heat enough. You’ll get the knack quickly. Super easy. Try a Caloplaca especially. Take a look at it first, unheated, then heat it and look again. World of a difference.
Note that old specimens (> several years) don’t require any tricks to clear the internal structure. Not sure why. Maybe they’ve had time to slowly diffuse the contents of the spores and cells so that everything is at the same index of refraction. That might be how heating works, too. It might force the process to happen instantaneously. Maybe it makes the membranes more porous. I’m not a biochemist! :)
I’ve had the scope about a year, but just now trying to start looking at some lichens. I have KOH that I actually think was used to mount this specimen. I also have Melzer’s, (which is iodine based, correct?). I was able to see structure pretty decent in person, but I’m only using my eye phone to take pics through the eye piece, of which mine happens to be very filthy by the way (any tips on better micro photos w/o breaking the bank and buying an eye piece camera?). I do have one eye piece camera thing, but the quality is very subpar, 1.3 megapixels I think, but I’ll have to play around with it again. Haven’t really used it since I’ve had it. Mostly because I’m also having a hard time trying to calibrate the camera via the software on my computer. I find it much easier to just make measurements using my eye piece reticle that was a much more straightforward process to calibrate.
R. maculans keyed out pretty well (my opinion of well at least, lol), the definitive features were the hourglass shaped lumina, thick-walled appearance and overall measurements. I find the most difficult part is trying to get a decent mount that allows the cover to remain relatively flush. That continues to be my struggle.
Regarding the heat application, what does that entail? Is it simply holding a flame under the prepared slide prior to scoping, or is there more to it? Thanks for info/tips thus far. It’s much easier to scope Basidiomycota, both spore deposits and tissue/hyphae sections. Easy to get hooked on this type of stuff!!
What’s up with that? Are you using my stock of lugol’s or did you find your own supplier? Mine is really old at this point, wondering if I should replace it.
Those Rinodina spore characters will be so clear you will be amazed. And if you need to see the spore septum swelling or not you can easily add K after heat.
Works great for Caloplaca too, esp to measure spore isthmus. I generally do it before photographing any spores now. And Iodine us such a pain to get under the cover slip for some reason…
Congrats on the microscope! This is a hard genus to start with, but I think you’re right. The “oil drops” are the cell lumina, as you say. They are hard to see in fresh specimens. There are a few tricks which help clarify the internal structure better.
1. iodine — It’s great but often stains everything so dark for me I can’t see the spores. Different people get different mileage out of that.
2. KOH — Definitely does a great job separating and clearing the spores… but it also can change the size and even shape of spores. (Some spores swell around the middle in KOH, e.g. the “dirinaria-type” spores.) But this can work in a pinch. Just subtract maybe ~10% from the measurements to ballpark it.
3. heat — This is my favorite by far. I think J-Dar is a convert as well. And John Sheard himself wrote a short communication for the British Lichen Society about this method, too. (He’s the Rinodina expert in North America.) What you do is gently heat the slide under the section by holding the slide over the flame of a bunsen burner or bic lighter. Just for a few seconds. If it boils, you’ve done it too long! If the spores aren’t clear afterward, then heat it some more. This has the advantage of not changing colors or size or shape at all. Or getting the slide dirty.
Your spores look like they could well be pachysporaria-type (R. maculans has that kind of spore). That is, the spores have thick walls all around, leaving only small round lumina in the center clustered around the septum.