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Kinyoun Acid Fast Staining

Kinyoun Acid Fast Staining

Although no definative evidence has been presented, the high lipid content (especially the mycolic acid component ) of mycobacteria is thought to be related to the mechanism of acid fastness. It has long been known that even light mechanical injury to the cell wall will cause them to lose the acid fast characteristic, suggesting that permeation through cell membranes might be an improtant part of the mechanism. Carbol fuchsin is used to stain the slide. Acid alcohol is used to decolorize the slide. It has been sugested that the dye replacement power of the counter stain could be used to decolorize and counter stain at the same time. It has been shown that acid alcohol increases the differentiation obtained, and gives superior results. After decolorization, the slide may be counter stained with methylene blue or brilliant green. The acid fast organisms will appear red while non-acid fast organisms will stain blue or green.

483A-8oz Kinyoun Carbol Fuchsin 8 oz.
483A-1gl Kinyoun Carbol Fuchsin 1 gallon
311A-8oz Acid Alcohol Decolorizer 8 oz.
311A-1gl Acid Alcohol Decolorizer 1 gallon
675A-8oz Methylene Blue 1% 8 oz.
675A-1gl Methylene Blue 1% 1 gallon
460A-8oz Brilliant Green 1% 8 oz.
460A-1gl Brilliant Green 1% 1 gallon

Specimen Collection
Organisms being stained by an acid fast method are usually taken from a solid or liquid medium on (in) which they have been cultured from their original source (e.g. wounds, throat, swabs, sputum, etc.). An aqueous suspension is made, in the case of the solid medium, by taking a small amount of the material and suspending it in a drop of distilled water on a microscope slide. Care should be taken not to make the smear too thick. In the case of a liquid medium, a drop is used directly from the culture container. However, due to the solids from the medium, this method is not always satisfactory. The suspension made by either method is air dried, then “fixed” by passing rapidly through a Bunsen burner flame two or three times. Allow the smear to cool before staining.

  1. Place the “fixed” smear on a staining rack and flood slide with Kinyoun stain for 2-3 minutes.
  2. Wash off the stain with distilled water.
  3. Decolorize with acid alcohol until no more color runs from the smear.
  4. Rinse thoroughly with distilled water.
  5. Flood slide with methylene blue or brilliant green for 1-2 minutes.
  6. Rinse thoroughly with distilled water and air dry.
  7. Examine under high dry magnification and verified under oil immersion.

Note: Staining times may vary to suit the individual.

Sources of Error

  1. Overheating (burning) during fixation can be avoided by just touching the back of the slide to the back of the hand each time the slide has been passed though the flame.
  2. Do not stain smears which have only been air dried. Smears must also be “fixed”.
  3. Smears should not be too thick. After air drying, examine under a microscope. If there are no areas of bbacteria separation, more water should be added to dilute the smear.
  4. After staining it is essential that the back surface of the slide is wiped clean.
  5. If washing with distilled water is not done adequately, crystallization of the stain may appear on the slide.