All the methods start by preparing 'fixed' smears on the microscope slides

An aqueous suspension is first made, in the case of a non liquid specimen, by taking a small amount of the sample and suspending it in a single drop of distilled water on a 25 mm x 75 mm slide. Do not make the smear too thick. If you have a liquid sample, a single drop is used directly from the culture container. The suspension made by either method is air dried, then "fixed" by passing rapidly through the Bunsen flame two or three times. Allow the smear to cool before staining.

GRAM STAINING

Reagents needed...

Crystal Violet
Grams Iodine or Stabilized Gram's Iodine
Grams Decolouriser
Safranin or basic fuchsin counter stain

Procedure

1. Place the 'fixed' smear in the staining rack and immerse completely with crystal violet for between 30 and 60 seconds.
2. Wash off the surplus stain using distilled water.
3. Cover the slide with iodine for 30 seconds.
4. Rinse off using distilled water.
5. Bleach with Gram decolouriser for between 10 and 15 seconds.
6. Wash thoroughly using distilled water.
7. Cover completely with safranin (or fuchsin) for between 30 and 60 seconds.
8. Finally wash with distilled water and air dry.

AURAMINE 'O' STAINING

Identification of Mycobacteria with auramine 'O' is due to the affinity of the mycolic acid for the fluorchrome which occurs in cell walls. The Mycobacteria are observed as luminous yellow rods against a dark background. Potassium permanganate can help suppress non-specific fluorescence.

Slides stained with auramine 'O' may be re-stained later with Ziehl-Neelsen or Kinyoun stain directly, providing that any immersion oil has been removed.

Reagents needed...

Auramine 'O'
Fluorescent Decolourising bleach
Potassium Permanganate solution

Procedure

1. Place the 'fixed' smear in a staining rack and flood the slide with auramine 'O' for 15 minutes. Do not allow the surface dry out.
2. Wash off the stain with distilled water.
3. Flood slide with the fluorescent decolouriser for between 30 and 60 seconds.
4. Rinse thoroughly with distilled water.
5. Flood slide with potassium permanganate solution for 2 minutes. Do not allow the surface dry out.
6. Wash thoroughly with distilled water and air dry.

Excitation

To illuminate the slide, use the same light source as used for fluorescent microscopy. These are usually either powerful mercury or xenon arc-discharge (burner) lamps that contain a combination of dichroic mirrors and filters capable of exciting fluorescent chromophores and filtering out the excitation light from the viwed image.

Filter combinations
K530 excitation filter with BG 12 barrier
G-365 excitation filter and an LP 420 barrier filter
Mercury lamps have peaks of intensity at 313, 334, 365, 406, 435, 546, and 578 nanometers

AURAMINE-RHODAMINE STAINING

In this method the Mycobacteria appear bright yellow or orange against a greenish background.

Reagents needed...

Rhodamine-Auramine
Fluorescent Decolourising bleach
Potassium Permanganate solution

Procedure

1. Insert the 'fixed' smear in the staining rack and immerse the slide in rhodamine-auramine for about 15 minutes. Do not allow the surface to dry out.
2. Wash off the stain with distilled water.
3. Flood the slide with fluorescent decolouriser for between 2 and 3 minutes.
4. Wash with distilled water.
5. Flood the slide with potassium permanganate solution for 3 or 4 minutes. Do not allow the surface to dry out.
6. Rinse completely using distilled water and dry in warm air.

KINYOUN (FUCHSIN) ACID FAST STAINING

Reagents needed...

Kinyoun Carbol Fuchsin
Acid Alcohol

Procedure

1. Place the 'fixed' slide on a staining rack and flood it with Kinyoun stain for 2 to 3 minutes.
2. Wash off the stain using distilled water.
3. Decolourise the specimen with acid alcohol until no more colour runs from the smear.
4. Rinse thoroughly using distilled water.
5. Counterstain the slide with methylene blue or brilliant green for 1-2 minutes.
6. Rinse thoroughly with distilled water and dry in air.

NILE RED FLUORESCENCE STAINING

Nile Red is hydrophobic dye that is highly selective for lipid vesicles in cells. The excitation and emission maxima of nile red fluorescence varies over the range... Excitation 490-550 nm and emission 530-610 nm. Tissue lipids fluoresce yellow or gold to red. _{[Greenspan J. Cell Biol. 100, 965-973, 1985]}

Procedure

Stain sections for 1-5 minutes.

There is a modified Nile Red method that uses a 4% potassium hydroxide solution after the Nile red staining stage.

PHENOLIC ACRIDINE ORANGE STAINING

Bacteria appear as bright orange luminous rods against a dark background.

Reagents needed...

Phenolic Acridine Orange Stain
Hcl destaining acid in reagent alcohol, with counterstaining methylene blue

Procedure

1. Put the 'fixed' smear in a staining rack and immerse the slide with Phenolic acridine orange for 15 minutes.
2. Wash with distilled water.
3. Flood slide with methylene blue destaining acid for 2 minutes.
4. Rinse with distilled water.
5. Air dry only. (blotting is contra indicated)

Examine microscopically using a fluorescent microscope

ZIEHL-NEELSON ACID FAST STAINING

Acid fast organisms will appear red while non acid fast organisms will stain blue if methylene blue is used as the counterstain or green in the case of brilliant green stain.

Reagents needed...

Carbol Fuchsin (ZN)
Acid Alcohol Decolouriser
Methylene Blue 1% or Brilliant Green 1%

Procedure

1. Place the 'fixed' slide on a staining rack and flood copiously with Ziehl-Neelson stain. Apply heat underside of slide for 3 minutes, but do not allow the stain to boil.
2. Wash off surplus stain with distilled water.
3. Destain with acid alcohol until no more colour runs from the smear.
4. Rinse thoroughly with distilled water.
5. Flood slide with methylene blue or brilliant green for 1 or 2 minutes.
6. Rinse thoroughly with distilled water and dry in air.

Examine using high non oil magnification and finally verify under oil immersion.