MYCOLOGY – Page 2

GRAM STAINING PROCEDURE FOR FUNGAL DETECTION

Stains walls containing protein sugar complexes called peptidoglycan. Typically have low lipid content. Gram positive fungi stain violet. Gram negative fungi decolourise and stain pink with safranin or neutral red counter stain.

1. Label a clean grease free slide with sample identification number.

2. Place a drop of normasaline and emulsify a small sample to make a thin smear

4. Apply crystal violet stain to the smear for 60 seconds. Drain stain and wash with water.

3. Heat-fix the smear.

5. Add Gram’s Iodine for 60 seconds and the drain excess.

6. Decolourise with acid alcohol or Acetone for 30 seconds.

7. Counterstain with safranin or neutral red for 60 seconds and then drain excess.

8. Rinse and allow to air dry.

9. Apply immersion oil
Examine under microscope at X100.
Record the findings

LACTOPHENOL COTTON BLUE (LPCB) FOR FUNGAL DETECTION

Used for visualisation of fungal elements and structures. Mainly performed from fungal colonies and used for identification.
The phenol in the stain will kill any organisms while the lactic acid preserves fungal structures. Cotton blue stains the chitin in fungal cell walls. The same structures may be seen as with wet mount; however LPCB preparation can be made permanent.

How to prepare the LPCB stain

Concentrated phenol………………………20 mls
Lactic acid………………………………….20 mls
Glycerol………………………………………40 mls
Cotton blue(china blue)………………… …0.05g
Distilled water…………………………..20 mls
Dissolve cotton blue in distilled water, then add the rest of the ingredients. Mix well.

Procedure of Using the LFCB stain

1. Place 1 drop of LPCB on clean grease free slide.

2. Immerse fungal colonies with help of swab or Scotch tape or tease mount.

3. Apply coverslip gently.

4. Visualize under microscope at X10 then X40.

5. Record the findings

SALINE MOUNT FOR FUNGAL DETECTION

Fungal elements that may be observed are budding yeast, hyphae and pseudohyphae, conidia, thin branching filaments resembling bacteria(fungus like bacteria),granules ,or coccidioides immitis spherules.

Preparation of reagent

Normal saline:
Nacl 0.85 g in 100 mls distilled water.

The procedure of saline mount

1. Place one drop of specimen on a glass slide and add one drop of saline.

2. Place coverslip and observed under low and high power under microscope.

Organism appear refractile, shinny or slightly green. Differentiate yeast cells from RBCs; yeast contain inclusion while others do not.

Look for spores,branching hyphae

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10% KOH MOUNT

USING 10% KOH FOR FUNGAL DETECTION

If the specimen is skin, hair or nail, the background cellular material may mask fungal elements. Potassium hydroxide will dissolve the keratin in these specimens, thus making any fungi more visible.

If there are a lot of cellular material in sputum or vaginal secretions, KOH mount may be prepared to dissolve the background thus making any yeast more visible.

However a saline mount must be additionally done to quantitate epithelial cells and WBCs and note presence of Trichomonas.

How to prepare 10% KOH

Potassium hydroxide……….…………10 g
Glycerol…………………………….20 mls
Distilled water………………………80 mls

Dissolve KOH in water, and then add glycerol. Glycerol prevents crystallization of the reagent and allows KOH preparations to be maintained for two days before drying up.

The procedure of using 10% KOH

1. Place specimen on clean grease free slide.

2. Add a drop of 10% KOH.

3. Apply coverslip and gently warm the slide over a flame.

4. Gently press the coverslip to spread the already softened tissue evenly on the slide.

5. Examine under a microscope X10 then X40.

6. Record the findings

Potassium hydroxide is a highly corrosive deliquescent chemical, therefore handle it with great care and make sure the stock bottle of chemical is tightly stoppered after use’.

TYPES OF FUNGAL MEDIA

1. Czapek’s agar:

It is used for the subculture of Aspergillus species for their differential diagnosis. It contains sucrose as C-source and nitrate as the sole source of nitrogen, useful for the general cultivation of fungi, yeasts and soil bacteria.

2. Potato Dextrose Agar (PDA):

It is a relatively rich medium for growing a wide range of fungi. Many standard procedures use a specified amount of sterile tartaric acid (10%) to lower the pH of this medium to 3.5 +/- 0.1, inhibiting bacterial growth. Chloramphenicol acts as a selective agent to inhibit bacterial overgrowth of competing microorganisms from mixed specimens, while permitting the selective isolation of fungi.

3. Sabouraud’s dextrose agar (SDA): Sabouraud Dextrose Agar (SDA) is a selective medium primarily used for the isolation of dermatophytes, other fungi and yeasts but can also grow filamentous bacteria such as Nocardia. Sabouraud agar is used to culture fungi and has a low pH that inhibits the growth of most bacteria; it also contains the antibiotic gentamicin to specifically inhibit the growth of Gram-negative bacteria. Hay infusion agar is specific for the culturing of slime moulds. The acidic pH of this medium (pH about 5.0) inhibits the growth of bacteria but permits the growth of yeasts and most filamentous fungi. Antibacterial agents (with antibiotics) can also be added to augment the antibacterial effect.

4. Brain-heart infusion agar (BHIA). Malt extract and less commonly brain heart infusion medium. To prevent contamination of the medium by bacteria, chloramphenicol is used, but prevents the growth of Actinomyces, which others grows well on Sabouraud dextrose agar.

STERILE TECHNIQUES FOR TRANSFERRING FUNGAL CULTURE

1.Take an inoculating needle, usually a thin needle or wire at the end of a long pencil-like handle, and heat it in an alcohol or gas flame until it glows bright red

2. Allow the needle to cool for about 15 seconds. (A hot needle will kill the mould that is to be transferred).

3. Open the Petri dish containing the culture just wide enough to allow entry of the needle.

4. With the heat-sterilized needle, cut out a small portion of the colony margin. Hyphal tip transfers work best as they are usually the most active parts of the culture; in addition, transfers from the heavily sporulating central portions will result in spores being spread into the air. Especially in medical work, hyphal tip transfers are essential. The excised colony margin should be only about 1 mm square.

5. Transfer the square of colony margin to the sterile plate, making sure that the lid is opened only wide enough to admit the needle and make the transfer. Place the block at the centre, withdraw the needle and flame it until it is red hot, to kill all adhering spores and hyphae.

6. Close the lid; label the plate with a marking pen, including name of culture and date. We usually wrap a thin strip of paraffin film around the sides of the plate to cover the opening, but this is not absolutely necessary; just a couple of pieces of masking tape to hold the lid down will do.

7. Leave the culture to grow in a protected place that has as little air movement as possible.

HAIR SAMPLE FOR FUNGAL DETECTION

• A Wood’s light, if available, may be helpful in selecting specimens.

• Pluck broken or lustreless hairs from periphery of lesion. Scrape scalp from edge of hair loss area.

• Do not cut hair.

• If hair are broken off (endothrix), it often may be necessary to scrape the coiled hair stubble from the scalp with a sterile blade or slide edge, rather than plucking with tweezers.

• Submit specimen to laboratory in sterile petri dish.

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SKIN SCRAPING FOR FUNGAL DETECTION

• Clean skin with 70% alcohol.

• Scrape edges of lesion (as edge has greatest amount of viable fungus) with a blunt scalpel blade.

• Collect skin scales in a sterile petri dish or similar wide-mouthed container or alternatively, skin scrapings may be collected in a clean, dry piece of paper folded securely with Scotch tape and labelled properly.

• If a skin scraping does not yield sufficient material, then a swab or Scotch tape could be pressed on the lesion

NAIL SAMPLE FOR FUNGAL DETECTION

• Clean nail with 70% alcohol.

• Examine for damaged, discoloured, brittle or dystrophic area.

• Material should be taken from the affected areas.

• Entire thickness of the damaged nail should be cut as far back as possible. Any crumbly material or material under the nail should be collected and sent in a sterile container.

• If skin lesions are present they should be scraped and the material collected should be sent separately.

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EYE SWAB AND SCRAPING FOR FUNGAL DETECTION

• Pus and discharge samples can be collected with a cotton wool swab and sent using standard precautions to the lab.

• Due to the sensitivity of the region and serious consequences of error, only an experienced ophthalmologist collects eye samples (e.g. corneal scrapings, intraocular fluid aspirate and swabs).

• For corneal scrapings, inoculate media plates and prepare slides at the patient’s side. Always contact the laboratory to obtain suitable media prior to the procedure. Alternatively, send corneal scrapings directly to the laboratory.

• Intraocular fluid (vitreous and aqueous) is collected using specialised equipment in the operating room.

• If sample is insufficient to perform both smear and inoculation of plates, give priority to culture.

• In case of contact lens-related infections, send contact lenses, case and cleaning solution to the lab for culture as well. Ear Swab and Scraping

• A physician collects samples from outer and middle ear. Skin scrapings from the external auditory canal are preferred. Use a sterile swab stick to collect exudates or debris.

• For deeper ear infections and to avoid damage to the ear drum, an ENT surgeon or experienced physician should use a speculum to draw specimen.

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