BASIC MYCOLOGICAL TECHNIQUES

R. T. Moore, University of Ulster, UK

  I. Stains
 II. Slide Preparation
III. Permanent Mounting Methods
IV. Preserving Spore Prints
 V. References

I. STAINS (Dring, 1971; Stevens, 1974)

A. Amann's lactophenol + stain:
   20g phenol (crystals):
   20g (16ml) lactic acid;
   40g (31ml) glycerol;
   20g/ml distilled water.
   Warm the phenol in the water to dissolve it, then add lactic acid & glycerol;
   Add 0.05(-1.0)g Poirrier's (cotton) blue or acid fuchsin.

AQUEOUS
B. Phloxine (particularly good for Homobasidiomycetes & filamentous fungi):
   *Stain: 1% aqueous phloxine.
   *Accompaniments: 70% ethyl alcohol; 5% aqueous KOH.
C. Melzer's iodine reagent (specifically for amyloid reaction):
   100   g chloral hydrate;
     5   g potassium iodide;
     1.5 g iodine;
   100   ml distilled water.

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II. SLIDE PREPARATION

TECHNIQUE. Best results can be achieved by viewing the specimen under a dissecting microscope to select & pick up a small amount of material from the substrate; use a pointed scalpel or a flattened needle (remains of tungsten filaments from old light bulbs are a good source for making dissecting needles).

A. General method:
   *Tease a small amount of tissue or hyphae into a few drops of alcohol (removes air); blot near edge of material or allow to evaporate
   *Add a drop of KOH (expands dry material)
   *Gently rinse preparation with distilled water; blot near edge of material
   *Follow with a drop of stain
   *Add a drop of mounting medium (water for temporary; Karo or Hoyer's for permanent - see below)
   *Carefully place a coverslip on the material for microscopic examination

B. Clear polyester adhesive tape method (particularly good for showing the natural orientation of dry spores)
   *Note. Transparent polyester film tape 850 is available from 3M. It is very thin, unaffected by water, and optically clear; cellophane tapes are generally unsatisfactory on all of these counts.
   1. Put a small drop of either distilled water or clear permanent mounting medium on a clean slide
   2. Using scissors and forceps, cut off a small piece of tape (a couple of mm's square), from the roll of tape: BE CAREFUL NOT TO TOUCH THE TAPE WITH YOUR FINGERS
   3. Working under the dissecting microscope, hold the small piece of tape with the forceps and touch the sticky side gently to the fungal colony
   4. Next, place the piece of tape sticky side up on the drop on the slide
   5. Add a small drop of stain to the fungal material;
   6. Add a larger drop of distilled water or permanent mounting medium
   7. Carefully lay a cover slip on the preparation

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III. PERMANENT MOUNTING METHODS

A. Lactic acid/cotton blue (ringing)
   1. Put a small drop of lactic acid/cotton blue stain on a slide (IT MUST NOT LEAK OUT BEYOND THE COVER SLIP
   2. Using a dissecting microscope, place a smidgen of material in the stain
   3. If you have problems getting the material off the needle or scalpel tip use an edge of your mounting cover slip (held by the edges). (A round cover slip is preferable.)
   4. GENTLY heat by passing over a flame to remove residual air bubbles or soften agar that may be present
   5. Ring cover slip with nail polish; let dry until hard; repeat several times (until edge of cover slip is blurred). (Clear polish is preferable, but coloured can be used for coding.)
B. Polyvinyl alcohol (PVA) permanent mounting medium (Omar et al., 1978/79)
   1. Ingredients: 1.66 g Polyvinyl alcohol (available from Sigma); 10.0 ml distilled water; 10.0 ml lactic acid; 1.0 ml glycerine
   2. Formulation: [a] dissolve PVA crystals in distilled water [b] add PVA solution to lactic acid while stirring vigorously; [c] add glycerine - filter if necessary; [d] let stand 24 hours to mature
   3. Using: [a] immediately under microscope; [b] for oil immersion, heat at 40° C for 10 minutes to set; [c] completely hardens, on its own, after 24 - 36 hours
   *Notes.
      ~medium is a colourless oily liquid that has a low refractive index (ca 1.39)
      ~can be used directly on lactophenol cotten blue preparations (its viscous nature causes excess stain to move to the edge of the coverslip)
      ~alternatively, 0.01-0.1% w/v cotton blue stain can be added during preparation

AQUEOUS STAINED PREPARATIONS (do not need to be ringed)
C. Hoyer's fluid (see Cunningham, 1972)
   1. Ingredients: 15g gum arabic or guiac; 100 g chloral hydrate; 10g glycerin; 25ml distilled water.
   2. Formulation: [a] soak the gum in the water for ~24 hrs (to prevent bacterial growth add a crystal of chloral hydrate about the size of a small pea); [b] add the rest of the chloral hydrate and let the solution stand several days until all material is dissolved; [c] add the glycerin.
   *Notes.
      ~Use like water to mount materials,
      ~Tends to remove stains used for hyaline materials,
      ~Excellent for phase contrast microscopy,
      ~Lasts indefinitely, i.e., years.
D. Karo syrup (Stevens, 1974; Johansen, 1940; Patrick, 1936).
  Note. This is an American product made from maize that is a mixture of dextrose, dextrin, and maltose and whose sugars, unlike those of cane syrup (treacle), do not crystallize on drying; when hard it is as firm as balsam and ringing is not necessary except in very humid climates.
   1. Formulation: 100ml Karo syrup (white); 300ml distilled water; 4ml (40%) formalin (to prevent molding).
   2. Use in combination with aqueous stains (e.g., phloxine)
E. Archive quality slides (See Volkmann-Kohlmeyer & Kohlmeyer, 1996).

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IV. PRESERVING SPORE PRINTS (Hadzic, 1992)

A. Cut rectangles from acetate sheets: 8 X 3.75 inches (20 X 9.5 cm)
B. Score a fold line across the middle of the width.
C. Make spore print by placing a cap on one half.
D. Fold in half and store in an envelope.

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V. References

Cunningham, J. L. 1972. A miracle mounting fluid [Hoyer's] for permanent wholemounts of microfungi. Mycologia 64: 906-911.

Dring, D. M. 1971. Techniques for microscopic preparation. in C. Booth, ed. Methods in Microbiology, vol. 4, pp. 95-11. Academic Press.

Hadzic, I. 1992. How to preserve a spore-print. The Mycologist 6: 188-9.

Johansen, D. A. 1940. Plant Microtechnique. McGraw-Hill.

Omar, M. B., Bolland, L. & Heather, W. A. . A permanent mounting medium for fungi [PolyVinylAlcohol+lactic acid/cotton blue].
   Stain Technology 53(1978): 293-294;
   Bulletin of the British Mycological Society 13(1979)31-32.

Patrick, R. 1936. "Karo" as a mounting medium. Science 83: 85-86.

Stevens, R. B., ed. 1974. Mycology Guidebook. University of Washington Press.

Twomey, D. G. 1977. The rapid preparation of micro-fungi for microscopic observations. Bulletin of the British Mycological Society 11: 148-149.

Volkmann-Kohlmeyer, B. & Kohlmeyer, J. 1996. How to prepare truly permanent microscope slides. Mycologist 10: 107-108.

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