Observation 45826: Galerina sphagnorum (Pers.) Kühner
When: 2010-05-24
Collection location: Maine, USA [Click for map]
No herbarium specimen
0 Sequences


Copyright © 2010 Erlon Bailey
Copyright © 2010 Erlon Bailey
Copyright © 2010 Erlon Bailey
Copyright © 2010 Erlon Bailey

Proposed Names

Please login to propose your own names and vote on existing names.

Eye3 = Observer’s choice
Eyes3 = Current consensus


Add Comment
Oh, wait, another thing…
By: Douglas Smith (douglas)
2010-05-25 15:57:56 CDT (-0400)

Scraped a gill? No, don’t do that. Just grab some micro-fine tweezers/forceps, and just pluck out a whole gill, and lay that down on the slide. I usually try for 2-3 so I can make sure I get a good gill with a good edge to see the cystidia.

Ooh, eeek, no…
By: Douglas Smith (douglas)
2010-05-25 15:55:57 CDT (-0400)

Using dried material is fine, that how most people do it actually, since if you are careful, dried material will last for many years.

But you need some sort of way of re-hydrating the material for view. And immersion oil is only to be put between the cover slide and the 100x lens, not on the material. That is only going to make the material worse really. Also don’t use the oil on anything other than the 100x lens, or it will mess up the other lenses…

Anyway, using dried material, I re-hydrate with alcohol. This is a popular method, there are others. I like this one, with using 95%-97% alcohol it dries quickly, and also it kills bacteria, so you don’t have to watch your spores get kicked around. Just one tiny drop per mount, and let it dry off. Takes 1-2 mins, if that. I start taking notes while that is going on. (Date, sample, type of section, reagent used, yadda, yadda…)

Once that dries, then you need some medium to put the material in under the cover slip. Water can work fine, but you can have troubles with surface tension making air bubbles around the material. KOH is good, since it has a low surface tension, and a few light taps after putting on the cover slide break up and drive away the bubbles (but this can disolve gelatin if that is what you are looking for, then use water). One small drop, put on cover slip from one edge, let it drop, light tap-tap. Ready to go.

So, now you have re-hydrated material, in water or KOH, under a cover slip, and into the scope. Look around at 10x until you got a good spot, check the edges of the gill for cystidia, take a few photos of those at 40x, get a spot with lots of spores, put on a drop of oil and go to 100x, and a photo there…

Ok, that might be getting a little far, but go for it!

Gill fragments
By: Erlon (Herbert Baker)
2010-05-25 15:35:19 CDT (-0400)

These spores are from the gills of the fungi i let dry over night, i scrapped the gills as i held the fungi above the slide.
Not all the spores are collapsed, why are they so concave after only one night?
I’ll try a fresh specimen next time, would this change the smoothness of the spore?

I put a drop of immersion oil on the gill fragments then placed the cover glass and another drop of oil on top of that, is that the correct procedure?

Oooh, yeah…
By: Douglas Smith (douglas)
2010-05-25 15:27:36 CDT (-0400)

Yeah, those are dried out and horrible. How did those spores get on the slide? I don’t think you can tell much from those as they are.

No, how did the spores get on the slide?
By: Douglas Smith (douglas)
2010-05-25 13:42:44 CDT (-0400)

No, I meant how did the spores get onto the slide? Did you take spores from a gill? The stipe? The cap surface? Spore drop onto the slide? What reagents did you use? Did you prepare the material with alcohol first, and then add reagent? Water, KOH, Meltzer’s… stuff like that. Wish you had Meltzer’s, do you? Except that doesn’t matter, my notes tell me that G. sphagnorum and G. paludosa both have dextrinoid spores, so there isn’t much to learn there.

But no need for a camera mount, I don’t have one. (Actually I do, but I don’t use it.) Hold the camera up to the eye piece, and take a photo from there. That is how all my micro-shots get taken. Give it a try and see how it works for you, you might have to turn the scope light up or down until you get a good image, but it usually works pretty well without much hassle. Try to use “incandescent” light setting on the white balance if you can, that usually works better on scope lights. But if you can’t do that, no worries, just give it try, and take a photo through the eye piece.

No photos
By: Erlon (Herbert Baker)
2010-05-25 12:59:45 CDT (-0400)

I’m still working on getting a camera attachment for my microscope.
By collapsing i mean concave.., i figure they become that way as they dry out?
Some of them are practically folded in half.
I prepared the slide with Type A immersion oil, with a cover glass.

I’m sorta new at using the scope, please forgive my ignorance.

Do you have photos?
By: Douglas Smith (douglas)
2010-05-25 11:31:04 CDT (-0400)

Do you have photos of the spores? What do you mean by collapsing? How was the slide prepared where you looked at the spores?

By: Erlon (Herbert Baker)
2010-05-25 10:39:30 CDT (-0400)

Galerina paludosa spores are roughened, these are smooth and collapsing.

Created: 2010-05-24 18:18:13 CDT (-0400)
Last modified: 2014-06-03 17:25:06 CDT (-0400)
Viewed: 173 times, last viewed: 2017-06-13 10:52:13 CDT (-0400)
Show Log