You may use the Scotch tape method. Just touch the fungal growth with a piece of scotch tap and place it on a microscopic slide containing small drop of lactophenol cotton blue

https://www.youtube.com/watch?v=ktENJEkopXI

This video is useful

Thanks, but that's not really a fixation protocol. Also, I prefer to avoid plastic tape where possible as it's not fully optically clear, especially if I'm using an imaging method like DIC.

There is essentially no worthwhile documentation for making fungal mounts, and it mostly boils down to trial and error with a sharps bin full of failures before you start making progress.

Do not use Scotch tape, it decreases the amount of light which makes it to your camera and reduces the dynamic range. Also it barely picks up anything worth photographing and rips apart relevant structures (e.g. you pick up a conidiophore of an Aspergillus but the stipe got torn and there is no foot cell)

What I typically do is slice out a 1 cm^2 piece of agar with a sterile scalpel. Make 2 series of parallel cuts, perpendicular to each other (essentially make a # pattern). Gently remove the square (containing the leading edge of a colony) and place onto a microscope slide (with the side containing growth face up).

I use DIC as well, and I just place a drop of sterile water on top of the agar square, place a cover slip down, and mount onto the microscope stage.

Now I typically work with Fusarium species which spread out on a clear water agar surface, and the philiades are dispersed enough that this method works very conveniently for me. If you are working with Aspergillus or Penicillia the above method will probably result in a large, opaque, knotted mess under the microscope. The nice thing about DIC is that the depth of field is super small so you may be able to focus in on a conidiophore while the hyphal mess is mostly out of focus.

The alternative is to make 3 cuts at the leading edge of a colony. Make one cut perpendicular to the leading edge behind where you see sporulation (think cutting the crust off a pizza) like ---------. Then make 2 cuts around 1 cm apart perpendicular to the first into the agar like --|---------|--. Then you need to make another cut parallel to the first one (--------) at like a 45 degree angle towards the colony. Instead of a square like above you are extracting a triangle-ish rectangle with the least amount of agar as possible.

From there you extract the agar piece containing the colony and place onto a microscope slide. Add a couple drops of sterile water and use needles to "tease" out the knotted mess that the hyphae is in. Do not poke repeatedly, do not rip to oblivion, just try to loosen the mess a bit and try to flatten everything.

If possible, cut off any excess agar. Then place a cover slip and try to crush the hyphae the best you can. If you picked up too big of a piece the cover slip may snap and break. You can try shearing the slide a bit to spread out the hyphae, but too much will ruin everything worse than Scotch tape.

The goal is to have as flat a layer of hyphae as possible before you begin looking for structures under the microscope. Do not waste your time looking around the center of the knotted mess, instead look around the edges of the clumps you teased apart. If all goes well you will see some conidiophores free from the mess and suspended loosely enough to photograph.

If you end up with a slide which is essentially an ocean of spores, my recommendation is to hold the agar piece you just extracted onto the microscope slide with a teasing needle and rinse the colony with sterile water before teasing to remove the bulk of the spores. There will still be plenty left, but hopefully you could now see if that Aspergillus isolate has metulae or if that Penicillium isolate is biverticillate without all the spores in the way.

-Best, someone who has had to make microscopic mounts of fungal food contaminants for the past 5 years.

P.S. If you want to preserve a slide you can seal the edges with clear nail polish to try to seal in the moisture. I just save the photographs, I throw away the slides after I am done.

Use a scotch tape, add few drops of formaldehyde and allow to stand for 1min. Overturn to decant. Allow to dry. Add lactophenol-in- cotton blue and observe.

The clear scotch tape is great for routine investigation, but it will cloud over time and is not resistant to heating (if you ever need to heat). The best way I know of is to use the slide mount method where you grow the fungus directly on the slide.