I didn’t succeed with the clone, as it got some nasty contams and stalled..
But from the faint spore print I succeeded to germinate and now I have some
clean culture growing on agar. I already have some grain that is colonizing and will try
further to see what will happen.

Btw, two very experienced BG hunters told me they are sure this is P.semilanceata

links are very helpful, especially Stamets’ book. I will do some agar to agar transfers,
but I will also put some spores on agar. Will see how it goes.
This is the phone adapter I got btw: https://www.amazon.com/...

Stage micrometer is next on my list too. Fungifun link you have sent me, states that aafeurope has Bulgarian office, I might try contacting them to see if I can get some big enough HEPA filter in order to build laminar flow hood.

https://fungi.com/products/antibiotic-malt-extract-agar-1lb

….do some agar/mycelium clean-up work (by cutting out small areas of healthy, un-contaminated mycelium and transferring them to new petri dishes). It can be done successfully with effort and endurance.

More Links:
https://www.amscope.com/accessories/micrometer.html
https://www.amazon.com/...
https://www.shroomery.org/...
https://www.amazon.co.uk/...
https://www.amazon.co.uk/...
https://www.amazon.co.uk/...
https://www.amazon.co.uk/...
https://www.amazon.co.uk/...
http://www.uspehfilter.com/...
https://www.environmental-expert.com/...

• Those links are for ideas/insights – not necessarily ideal products.

I use self made still air box, gloves, 70% etanol and flame heating of instruments.
Agar is prepared at home with malt extract, then sterilized in PC at 0.8 bar – that is the pressure I measured, I plan to put stiffer springs on valves and manometer.

I still have the print I made, I might give it another chance, but I made it on paper.

Do you utilize a laminar flow hood for your studies?

More Links:
http://www.fungifun.org/English/Flowhood
https://www.shroomery.org/...
http://library.uniteddiversity.coop/...

two out of three petris covered with some kind of white mold,
the other one I don’t know what to think, is it mold or myc, looks more like mycelium,
but grow seems stalled, see the last pic how it looks now.

…Promising to call this a “Psilocybe” in general based on what I can see. How is the culture on agar going so far?

found some time to do this. Not sure if quality is good enough to see anything else apart from spores, but I really don’t know what to look for, and my microscope is probably not good enough.

This is a single gill piece, laying, rehydrated in 96% etanol, then destilled water + one drop liquid hand soap. Then stained with 1% Congo red. Pictures taken with the phone with adapter (16 MPix camera vs. sub 2 MPix that comes with the scope).

I uploaded just way too much pictures, not sure which are good enough, but I could remove some if requested. Thank you.

P.S.
Two guys from Bulgaria told me independently that this is just a large P.Semilanceata. So, they could be right maybe.

If you need answers for something, feel free to ask…

10 gr. powder, I will have to prepare 1g./100ml solution. I got NaOH (instead of KOH), 95% etanol,and distilled water.
I think with these I will be able to make somewhat decent pictures this weekend. Unfortunately I don’t have yet smartphone adapter, so I will use 720p camera that came with the scope. I don’t have stage micrometer too, so I will post only objective magnification.

A Common List of Stains & Mounting Liquids
☑ CONGO RED 1.0% AQUEOUS SOLUTION, 4 oz. (CH3130-4) (If this is not available a very good alternative is: Cat. No. 2250-4 Ricca Congo Red Indicator 0.1% (w/v) aqueous solution. CAS No. 573-58-0 reagent). – used as a good staining agent for cell walls, basidia, cystidia, basidioles, and hyphae (including septa)
☑ 2%, 3%, or 4% KOH (This is more of a mounting liquid but can cause color changes in some cells) – This is a very good mounting liquid used very frequently ◊
☑ 10% Ammonium Hydroxide can be used with some species with greater clarity of vision. It also does not crystallize and eventually become blurry as seen with KOH, although this chemical does give off a noteworthy fume and some mycologists may want to use a fume hood in its company
☑ A drop of Softsoap (Liquid hand soap) in distilled water (2oz dropper bottle) – used as a general mounting liquid when KOH is unavailable
Additional List of Stains & Uses
Use with caution and read the MSDS for each item thoroughly beforehand.
☑ “GSM” Abbreviated for: 20g Glycerol Conc. + 1g Sodium Hydroxide (NaOH) + 20ml Methyl Cellosolve + 60ml Distilled Water – noted by some as the best general mounting liquid of all
☑ Melzer’s Reagent – used to check spores for amyloid, nonamyloid, and dextrinoid presence. (If this is not available a fair alternative is: Lugol’s Dilute Solution) – used for coloring white spores dark enough for visibility
☑ Ammonia (Also called NH3 and Ammonium Hydroxide) 1%-25% with Water (25% is a typical amount) Mounting agent used in place of KOH. 웃
☑ Brilliant Cresyl Blue – observe spore walls with more detail
☑ 1%-10% Sodium Hydroxide (NaOH) with Water * – Can be used in place of KOH
☑ Combine a 5% KOH solution with a 10% Glycerol solution to prevent KOH precipitation and extend the slide’s life by several hours
☑ Abel’s Fluid: 25ml Ethanol 96% + 15ml or 18.9g Glycerol Conc. + 25ml Ammonia Conc. + 35ml Distilled Water **
☑ 20% Glycerol with 2-4% NaOH in Distilled Water – used as a mounting liquid after staining with Congo Red but not SDS Congo Red
☑ 1% Phloxine – Makes hyphae more visible (This is another red stain). It’s also used to see cell contents. Mix 1 drop Phloxine, 1 drop Congo Red, and 1 drop of 3% KOH.
☑ 1% Congo Red (1%) + 1% Boric Acid + 10% Ethanol + 15% Glycerol + Distilled Water – lasts several years
☑ Safranin – Can color nuclei red
☑ 10% Ammonium Hydroxide (NH4OH) with 90% Water ☼ ∂ – for viewing spore ornamentation, internal structure of spores, some chrysocysitidia, incrusted hyphae 웃
☑ Water with Dilute Glycerol – mounting liquid
☑ Calcofluor White with 10% KOH -
☑ Potassium Hydroxide (KOH) with Chlorazol Black
☑ KOH-DMSO Preparation
☑ Mucicarmine
☑ Periodic acid-Schiff (PAS) and PAS Digest stain
☑ Grocott’s Methenamine Silver (GMS) stain
☑ SDS Congo Red – Do not use with KOH. With fresh collections it can show walls and septa. With dried collections it can be used with ammonia or with DWAEG.
☑ Toluidine Blue O – Used for spore wall layers
☑ Trypan Blue
☑ Cresyl Blue – Used for spore wall layers
☑ Ruthenium Red
☑ Patent Blue V – stains deutoroplasm of chrysocystidia, some pigmentations
☑ Congo Red with Ammonia
☑ Sudan III – lipids/resins
☑ Sudan IV – same as Sudan III
☑ Carmine – stains nuclei, siderophilous granules, deuteroplasm of some secretory hyphae and cystidia
☑ Iron-Acetocarmine – Shows nuclei, basidium granules, some siderophilous walls
☑ DWAEG (Partial rehydration – moisture-adding liquid composed of 16ml Distilled Water, 4ml of 25% Ammonia, 80ml of 96% Ethanol, 1g Glycerol). Used for sections but still requires mounting medium.
☑ Carbol Fuchsin
☑ Chloral Hydrate
☑ Chlorovanillin
☑ Acetocarmine
☑ Erthrosin
☑ Fuchsin
☑ Guaiac
☑ Sulphobenzaldehyde (Sulfobenzaldehyde)
☑ Lactophenol Cotton Blue
☑ Lactoglycerol in its place

Caution: Many of these substances are toxic (poisonous) if inhaled, touched, or consumed in any way.
KOH and NaOH can alter the structure of cell walls and may impact microscopy (including swelling, color changes, and even the dissolving of ornamentations)
◊KOH and NaOH can slowly corrode glass dropper bottles over the course of a few months (Replace solutions accordingly).
*Methyl Cellosolve = Ethylene Glycol Monomethyl Ether.
***Abel’s Fluid can shrink physalohyphae while also making cytoplasm become more granular.
☼ Spore color can be altered by Ammonium Hydroxide in some species.
∂ Some mycologists will use a 3% or 5% Ammonium Hydroxide solution instead of 10%.
웃 Ammonia and Ammonium Hydroxide are toxic substances (They should not be exposed to skin (or any part of the body) and they should not be inhaled).

Note: Proper disposal of liquids, glass slides, cover slips, and razor blades is necessary and environmentally intelligent. A “sharps container” can be used for glass and razor blades.

Rehydration Materials
With patience, GSM or Abel’s Fluid can be used (See formulas above)
2-10% KOH in H2O (KOH = Potassium Hydroxide). Requires appropriate handling. Safety sheet: http://fscimage.fishersci.com/msds/19431.htm
10% Ammonium Hydroxide (NH4OH) with 90% H2O
Ethyl Alcohol (=Ethanol) 95-96% (approximately) in 4-5% H2O

Note: 1% or 0.1% Congo Red can be used exactly as it is in order to stain a section. One or two drops, sometimes more depending on the size of a section, is all that’s required. Let the stain soak in for 1-4 minutes (depending on the fungus).

These online stores are helpful. I will see what else I can find locally first, and will order otherwise.

☑ I would discourage the use of food dyes/coloring for this.

➜ 1% Phloxine is another red stain that may work just as well as Congo Red. Lactophenol Cotton Blue or another blue stain can be used, too. GSM may be good enough to differentiate cells microscopically as well, but it will look more clear, and less colored. Personally, I think GSM is one of the best tools for mycologists until the next generation of fluids is developed. It might be hard to find during the pandemic though.

Check out: https://micro-science.co.uk/...

Also, definitely consider contacting the following website for vendor resources if the area you live in does not have a reliable store: https://www.myko-shop.de/...

More: https://www.emsdiasum.com/...

KOH and isopropanol is easy, but I have troubles finding congo red.
Can it be substituted with some food grade colour die? Thank you!

so much for your time and valuable tips. I will see what I can do and will post back when I have results. Not sure for alder, but I can find beech and oak chips. Will see.

✓ I recommend Alder for your wood chips if they are available.

For working with dried material:

It can be hydrated in 5-10% KOH and then stained with Congo Red or another stain of your choosing (Cotton Blue, etc). If KOH is not available, 70% Isopropanol Alcohol can be used in its place, followed by water.

Rehydrating A Single Gill After It’s Fully Or Partially Dried

There are multiple methods you can try, and sometimes it can take a bit of patience and dilligence…

Starting with a whole, dried mushroom: Break off the stem then break the cap in half so you have full access to the gills. Using a razor blade, slowly press in between two gills and break off a whole gill. Once placed onto a slide, 2-3 drops of 5% KOH are added. After two minutes or more the KOH should rehydrate the specimen. Once it is rehydrated you may now absorb the excess KOH with a napkin corner/edge. Now a prepare cut-out can be made just as you would with a freshly picked specimen. A little more KOH should be added, if necessary, before adding a cover slip so it seen throughout the entire cover slip area.

Note: If you want to stain the gill after rehydrating it with KOH, just absorb the excess KOH with a napkin corner, then stain with congo red or another type of stain for a 1-2 minutes, absorb the excess congo red (or stain), then add more KOH and a cover slip.

Other liquids used for rehydration that are recommended: Ethyl alcohol (95% or 96%), isopropanol alcohol (91%), GSM, Abel’s Fluid, and Clémençon’s Solution.

Rehydrating A Complete Cap With Gills

In addition to working with a single gill, it’s good to also work with a gill cross section using the entire pileus and lamella. The easiest method to this follows.

First, break off the stipe so only the pileus and lamella remain. In a small beaker, add enough 91% rubbing alcohol to allow the cap to be submerged completely in the alcohol. Depending on the size of the mushroom and species, it should take between 10 and twenty minutes for the tissue transition from a dense, hard mass into soft, rehydrated tissue.

After submerging in rubbing alcohol, fill a separate small beaker with distilled (or DI) water. Transfer the mushroom tissue into the H2O and soak for ten additional minutes.

The material is removed with curved forceps (also known as jeweler’s curved forceps) and cut in half so we’re working with exactly one half of the mushroom cap.

From the above section, extremely thin cross sections can now be made. This will provide a view of the pileipellis, pileus trama, gill trama, subhymenium, and limited areas of the gill face.

⇨ Alternatively, you can simply place a mushroom cap and gills into a little GSM which can be used both as a rehydrating fluid and a mounting fluid.

- https://web.archive.org/...

..and mycelium seems to be healthy. In a couple of days I will transfer it to grain and if all goes well I will have an opportunity to try cultivating it in grow chamber.
The substrate I am thinking will have some wood, compost and dung, so that it have a chance whatever it is.

This weekend I will get my microscope and will try taking some pictures, but not sure what I can do the with dried mushroom. Should I soak it in some water or chemical solution?

Although Psilocybe semilanceata has different appearance at times, I think you have a species that grows on wood and not on dung or dung-rich soil and grass.

Any chance this to be huge mutant P. semilanceata?

✔ You might get better images of your microscopy with a cell phone, but it depends on the phone.

✔ Try to measure the length and width of 10-30 spores.

✔ Also see if you can tell us about the cystidia, and see if you can obtain some good images with measurements.

LINKS

Page 366: http://mykoweb.com/...

https://www.mushroomexpert.com/microscope.html

https://images.mushroomobserver.org/microscopy.pdf

https://images.mushroomobserver.org/Fungal%20Microscopy.pdf

https://web.archive.org/...

http://czechmycology.org/_cm/CM60202.pdf

http://www.micobotanicajaen.com/...

https://www.shroomery.org/...

http://www.davidmoore.org.uk/...

microscope is what I have. Not sure if I can make any good pictures and what exactly to look for, but I can try. The camera eyepiece gives less magnification than optical and also produces some chromatic aberration. I’ve posted some (bad) pics of spores on shroomery.

Would like to hear some opinions also on these as I found them on exactly same place two weeks later:
https://mushroomobserver.org/438378?q=1WtCd
https://mushroomobserver.org/438379?q=1WtCd

…is necessary to help others confidently identify this collection. Do you have access to a light compound scope?

I’ve been today at the same place.. Found around some similar mushrooms, always solitary, always dead leaves and some grass around – none of them was on dead wood, so maybe not P.serbica. Some specimens bruised blue very visible.
Will add new observations tomorrow with my new findings.

P. fimetaria is described as dung lover, no dung at the place though. Also, shouldn’t they have an annulus? This one does not seem to have any, neither any remnants visible.