1.7 General Methods
for Mycobacterial
Growth
Most mycobacteria will grow on fairly simple substrates that
include glycerol as a carbon source, mineral salts and ammonia or
amino acids for nitrogen. Some species, including M. avium subsp.
paratuberculosis, M. haemophilum, and M. genavense, are fastidious
and require supplements including hemin, hemoglobin, or other
iron compounds. M. leprae has not yet been cultured on axenic
media and its growth requirements will be discussed separately.
Carbon dioxide and fatty acids such as those in egg yolk can
stimulate mycobacterial growth. Optimal growth temperature can
range from 28 to 45
C, with visible colonies being produced by
most species between 7 days and 6 weeks. However, most species
grow best between 35 and 37
C, although M. chelonae and
M. ulcerans have lower optimal growth temperatures. RGM produce visible colonies between 3 and 7 days [8].
For optimal growth and especially primary isolation, CO 2 supplementation is necessary. 5–10% CO 2 during incubation is recommended. Candle jars do not lower the oxygen tension sufficiently
for mycobacteria to grow and are not recommended. If CO 2 incubators are not available, a commercial anaerobic system with tablets
or sachets that produce CO 2 can be used.
1.8 Staining
of Mycobacteria
Microscopic examination of mycobacteria is still widely used not
only to confirm cell morphology and purity but also in clinical
settings to provide an initial bacteriologic evidence of the presence
of mycobacteria in a clinical specimen. The high concentration of
lipids in the mycobacterial cell wall makes them waxy and hydrophobic, which provides an impermeable layer to common routine
stain such as the Gram Stain. Another key characteristic of this cell
wall is its resistance to acid and alcohol, defining these cells as AFB
or Acid Alcohol Fast Bacilli (AAFB). The two most widely used
methods for acid-fast staining are the carbol fuchsin-based methods, such as Ziehl-Neelsen and Kinyoun (that rely on light/brightfield microscopes) and the fluorochrome-based procedures that use
fluorescent dyes such as auramine-O or auramine-rhodamine (that
rely on fluorescent microscopes).
The two main carbol fuchsin-based methods rely on two different approaches to stain the cells. In the “hot” Ziehl-Neelsen technique, the phenol-carbol fuchsin stain is heated to enable the dye to
penetrate the waxy mycobacterial cell wall. In the “cold” Kinyoun
method, the stain penetration is enhanced by increasing the concentration of basic fuchsin and phenol and incorporating a “wetting
agent” chemical. Once the fuchsin binds to the mycolic acid in the
mycobacterial cell wall, an acid decolorizing solution is applied to
remove the red dye from the background cells, tissue fibers, and
only mycobacteria will retain the dye. Then malachite green or
methylene blue is used as counterstaining reagent to provide contrast color against which the red AFB can be seen. Kinyoun staining
is recommended for culture staining, especially under BSL3
8
Elizabeth Wallace et al.
for Mycobacterial
Growth
Most mycobacteria will grow on fairly simple substrates that
include glycerol as a carbon source, mineral salts and ammonia or
amino acids for nitrogen. Some species, including M. avium subsp.
paratuberculosis, M. haemophilum, and M. genavense, are fastidious
and require supplements including hemin, hemoglobin, or other
iron compounds. M. leprae has not yet been cultured on axenic
media and its growth requirements will be discussed separately.
Carbon dioxide and fatty acids such as those in egg yolk can
stimulate mycobacterial growth. Optimal growth temperature can
range from 28 to 45
C, with visible colonies being produced by
most species between 7 days and 6 weeks. However, most species
grow best between 35 and 37
C, although M. chelonae and
M. ulcerans have lower optimal growth temperatures. RGM produce visible colonies between 3 and 7 days [8].
For optimal growth and especially primary isolation, CO 2 supplementation is necessary. 5–10% CO 2 during incubation is recommended. Candle jars do not lower the oxygen tension sufficiently
for mycobacteria to grow and are not recommended. If CO 2 incubators are not available, a commercial anaerobic system with tablets
or sachets that produce CO 2 can be used.
1.8 Staining
of Mycobacteria
Microscopic examination of mycobacteria is still widely used not
only to confirm cell morphology and purity but also in clinical
settings to provide an initial bacteriologic evidence of the presence
of mycobacteria in a clinical specimen. The high concentration of
lipids in the mycobacterial cell wall makes them waxy and hydrophobic, which provides an impermeable layer to common routine
stain such as the Gram Stain. Another key characteristic of this cell
wall is its resistance to acid and alcohol, defining these cells as AFB
or Acid Alcohol Fast Bacilli (AAFB). The two most widely used
methods for acid-fast staining are the carbol fuchsin-based methods, such as Ziehl-Neelsen and Kinyoun (that rely on light/brightfield microscopes) and the fluorochrome-based procedures that use
fluorescent dyes such as auramine-O or auramine-rhodamine (that
rely on fluorescent microscopes).
The two main carbol fuchsin-based methods rely on two different approaches to stain the cells. In the “hot” Ziehl-Neelsen technique, the phenol-carbol fuchsin stain is heated to enable the dye to
penetrate the waxy mycobacterial cell wall. In the “cold” Kinyoun
method, the stain penetration is enhanced by increasing the concentration of basic fuchsin and phenol and incorporating a “wetting
agent” chemical. Once the fuchsin binds to the mycolic acid in the
mycobacterial cell wall, an acid decolorizing solution is applied to
remove the red dye from the background cells, tissue fibers, and
only mycobacteria will retain the dye. Then malachite green or
methylene blue is used as counterstaining reagent to provide contrast color against which the red AFB can be seen. Kinyoun staining
is recommended for culture staining, especially under BSL3
8
Elizabeth Wallace et al.