Acid-Fast Stain

Members of the bacterial genus Mycobacterium contain large amounts of lipid (fatty) substances within their cell walls. These fatty waxes resist staining by ordinary methods. Because this genus contains species that cause important human diseases (the agent of tuberculosis is a Mycobacterium), the diagnostic laboratory must use special stains to reveal them in clinical specimens or cultures.

When these organisms are stained with a basic dye, such as carbolfuchsin, applied with heat or in a concentrated solution, the stain can penetrate the lipid cell wall and reach the cell cytoplasm. Once the cytoplasm is stained, it resists decolorization, even with harsh agents such as acid-alcohol, which cannot dissolve and penetrate beneath the mycobacterial lipid wall. Under these conditions of staining, the mycobacteria are said to be acid fast (see colorplate 9). Other bacteria whose cell walls do not contain high concentrations of lipid are readily decolorized by acidalcohol after staining with carbolfuchsin and are said to be nonacid fast. One medically important genus, Nocardia, contains species that are partially acid fast. They resist decolorization with a weak (1%) sulfuric acid solution, but lose the carbolfuchsin dye when treated with acid-alcohol. In the acid-fast technique, a counterstain is used to demonstrate whether or not the fuchsin has been decolorized within cells and the second stain taken up.

The original technique for applying carbolfuchsin with heat is called the Ziehl-Neelsen stain, named after the two bacteriologists who developed it in the late 1800s. The later modification of the technique employs more concentrated carbolfuchsin reagent rather than heat to ensure stain penetration and is known as the Kinyoun stain. A more modern fluorescence technique is used in many clinical laboratories today. In this method, the patient specimen is stained with the dye auramine, which fluoresces when it is exposed to an ultraviolet light source. Because any acid-fast bacilli take up this dye and fluoresce brightly against a dark background when viewed with a fluorescence microscope, the smear can be examined under 400 × (high-dry) magnification rather than 1,000 × (oil-immersion) magnification. As a result, the slide can be screened more quickly for the presence of acid-fast bacilli (see colorplate 9).

Purpose

To learn the acid-fast technique and to understand its value when used to stain a clinical specimen

Materials

A young slant culture of Mycobacterium phlei (a saprophyte) 24-hour broth culture of Bacillus subtilis A sputum specimen simulating that of a 70-year-old man from a nursing home, admitted to the hospital with chest pain and bloody sputum Gram-stain reagents Kinyoun’s carbolfuchsin Acid-alcohol solution Methylene blue Slides Diamond glass-marking pencil Marking pencil or pen 2 Χ 3-cm filter paper strips Slide rack Forceps

Procedures

1. Prepare two fixed smears of each culture and two of the simulated sputum. In practice, the smears are fixed with methanol for one minute or are heat-fixed at 65 to 75°C to be certain any tuberculosis bacilli present are killed. To make smears of the agar slant culture, first place a drop of water on the slide, and then emulsify a small amount of the colonial growth in this drop.
2. Ring and code one slide of each pair with your marking pencil or pen, as usual.
3. The other slide of each pair must be ringed and coded with a diamond pencil. This device scratches the glass indelibly, so that the marks remain even during the prolonged staining process.
4. Gram stain the set of slides marked with the pencil or pen in step 2.
5. Stain the diamond-scratched slides by the Kinyoun technique:
1. Place the slides on a slide rack extended over a metal staining tray, if available.
2. Cover smear with a 2 Χ 3-cm piece of filter paper to hold the stain on the slide and to filter out any undissolved dye crystals.
3. Flood the slide with concentrated carbolfuchsin solution and allow to stand for five minutes.
4. Use forceps to remove filter paper strips from slides and place the strips in a discard container. Rinse slides with water and drain.
5. Cover smears with acid-alcohol solution and allow them to stand for two minutes.
6. Rinse again with water and drain.
7. Flood smear with methylene blue and counterstain for one to two minutes.
8. Rinse, drain, and air dry.
6. Examine all slides under oil immersion and record observations under Results see colorplate 9 for examples.

Results
*Note: Some saprophytic mycobacteria may stain weakly gram positive or appear beaded in Gram-stained smears.