Buruli ulcer - Diagnosis of Mycobacterium ulcerans disease
Annex 9 - Biochemical and culture tests used for the identification of slow-growing mycobacteria
The next two tubes were inoculated with a scotochromogenic strain (pigmentation produced in the dark and in the light).
The last two tubes were inoculated with an African M. ulcerans strain (pigmentation produced in the dark and in the light).
Most of these tests are based on studies carried out by the International Working Group on Mycobacterial Taxonomy which described highly reproducible tests for use in systematics of the genus Mycobacterium (Wayne et al., 1974, 1976). The tests recommended as minimal standard tests for the identification of slow growing mycobacteria were described in 1992 by Vincent Lévy-Frébault and Portaels.
Pigmentation and photoreactivity
Mycobacteria may synthesize carotenoid pigments which confer a yellow-to-red pigmentation to colonies. Carotenogenesis is achieved in the absence of light by scotochromogenic mycobacteria. For these mycobacteria, the influence of light is essential, and prolonged incubation in light can only intensify the pigmentation. Photochromogenic mycobacteria require exposure to light and oxygen for carotenogenesis. Other mycobacteria do not develop pigmentation under any light conditions. However, a few nonphotochromogenic strains may develop some pigmentation as they age (weak to bright yellow or pink to coral). Because of these variable characteristics of pigment formation, pigmentation must be determined as follows: a cell suspension, properly diluted to obtain isolated colonies, is inoculated into two tubes containing the appropriate solid medium. It is important to obtain isolated colonies to ensure the maximal oxygenation necessary for carotenogenesis. One of the tubes is wrapped or placed in any device that ensures total darkness. The tube protected in this way and the unshielded tube are incubated at the optimal temperature until growth is plainly visible in the unshielded tube.
The test has to be performed as soon as well-developed colonies are observed because photoinduction of carotenogenesis may not occur in old cultures and, as mentioned above, pigmentation can develop slowly while the control remains non-pigmented. The tube incubated in the dark is placed at a distance of 20 cm from a 60 W fluorescent lamp (or equivalent tungsten bulb) for 1 to 2 hours. Good aeration of the culture, which is properly ensured by loosening the cap, is critical as carotenogenesis is oxygen dependent. The cultures are then incubated again and examined for pigmentation 24 hours after illumination.
A comparison of the pigmentation of the shielded tube that was exposed once to light with the unshielded tube determines whether the mycobacterium is scotochromogenic, photochromogenic, or nonphotochromogenic on the basis of the criteria as described in Figure 43.
Resistance to isoniazid, thiophene-2-carboxylic hydrazide, hydroxylamine, p-nitrobenzoic acid and NaCl
Resistance to inhibitory agents is assessed by incorporating each inhibitory agent into separate tubes of L-J medium. Most compounds are added into the uncoagulated solution prepared as shown in Table 7.
Table 7: Preparation of inhibitory agents*
From the standard culture suspension, 10-2 and 10-4 dilutions are prepared, and 0.2 ml portions of the 10-2 dilution are inoculated into tubes of medium containing inhibitory agents. In addition, control tubes are inoculated with 0.2 ml of the 10-2 and 10-4 dilutions. The colonies are counted after no fewer than 3 weeks incubation or when growth is visible in the control tubes. If growth on a drug-containing medium is less than the growth of the 10-4 dilution control (i.e. less than 1% resistance), the culture is reported as susceptible; growth on drug-containing medium equal to the growth of the 10-2 dilution control (i.e. more than 1% resistance) is reported as resistant. Colonies in the 10-2dilution control tubes must be numerous (more than several hundred colonies or confluent growth); otherwise, interpretation of the test may be inaccurate.
Catalase activity
Tube 2 and 3 were inoculated with two strains producing less than 45 mm foam.
Tube 4 was inoculated with a catalase positive strain producing more than 45 mm foam.
Catalase activity. The test used to detect catalase activity is semiquantitative and reflects differences in enzyme
kinetics (Wayne et al, 1976). For this test, butts (not slants) of L-J medium are used (Fig. 44).
The flat circular surface of the medium is inoculated with 3 drops of an undiluted suspension of mycobacteria,
prepared as described above for growth rate determinations.
The tubes are incubated at the optimal temperature for growth. After the medium is examined to ensure that there is growth, 1 ml of a freshly prepared assay reagent is added. The assay reagent consists of a mixture of 0.5 ml of 30% H2O2 and 0.5 ml of 10% Tween 80 in H2O. The tube is placed in an upright position, and the height of the foam column generated after exactly 5 minutes is measured. A column of foam that rises more than 45 mm above the surface of the medium is recorded as positive. If the column is less than 31 mm, the result is recorded as negative. If the foam column is between 31 and 45 mm, the test should be repeated. Agar media may not be substituted for L-J medium, as they yield erratic results.
Tween 80 hydrolysis
The Tween 80 hydrolysis test (Wayne et al., 1974) is based on the fact that neutral red is orangish-yellow, even at a pH corresponding to the pH of the red form, when it is complexed with Tween 80 (a polyethylene derivative of sorbitan monooleate). Hydrolysis of Tween 80 leads to a change in the colour of the substrate to pink. This colour change is not due to a pH shift caused by the production of oleic acid but to the destruction of Tween 80.
The substrate solution consists of 0.5 ml of Tween 80 in 100 ml of 0.067 M phosphate buffer (pH 7.0) to which 2 ml of a 1% aqueous solution of neutral red is added. The solution is dispensed into screw-cap tubes (16 by 125 mm) in 4 ml amounts, and the preparations are sterilized by autoclaving. After cooling, a loopful of bacteria is suspended in 4 ml of substrate, and each tube incubated at 30–33°C for up to 10 days. A change in colour from amber to pink or red is recorded after 24 h and 5 and 10 days of incubation as a positive reaction. The pink colour must develop in the fluid itself, so the suspension should not be shaken until after each reading is made. Otherwise, the fact that the cells themselves may be neutral red-positive and take up the pink dye may lead to a false-positive reading (Wayne, 1985).
Urease activity
The method of Steadham (1979) is used. A loopful of actively growing culture on L-J medium is emulsified in 1 ml of urea broth prepared as shown in Table 8.
Table 8: Urease activity (the Steadham method)
The tubes are incubated at 30–33 °C and are read after 1 and 7 days.
A change in the colour of the broth from bright yellow to dark pink or red is an indication of the breakdown
of urea and is read as a positive reaction.
Niacin detection
Some mycobacteria have a block in the nicotinamide-adenine dinucleotide (NAD) scavenging pathway and accumulate extracellular niacin. The standard method used for the niacin assay requires 3 to 6 weeks incubation on L-J medium. A culture is covered with 1 ml of sterile distilled water, and the surface of the culture is broken with a spatula to improve extraction of the niacin, which is excreted and accumulates in the medium. The tube is placed horizontally to ensure maximal contact of the water with the culture.
After a 20-minute extraction period, 0.5 ml of the liquid is transferred to a screw-cap tube; then 0.5 ml of a solution containing 4% aniline in 95% ethanol and 0.5 ml of a 10% cyanogen bromide solution are added. The tube is tightly closed immediately, for safety reasons. If niacin is present, a yellow colour develops within 5 minutes.
To avoid any misinterpretation because of yellow pigments extracted from chromogenic mycobacteria, aniline can be replaced by a solution containing 3% benzidine in absolute ethanol. This method produces a red colour for positive reactions.
Note: Benzidine is carcinogenic and its use may be forbidden in some countries. This compound, as well as cyanogen bromide, must be handled with special care; a worker should wear a mask and gloves to weigh the powder and should handle the solution in a safety cabinet.
Paper strips impregnated with reagents are commercially available, and the instructions of the manufacturer must be followed. The colour developed in the liquid, and not the colour on the paper strip, denotes the reaction.
Nitrate reductase activity. The nitrate reductase test usually employs simple buffered substrate solution containing 0.01 M NaNO3 in M/45 phosphate buffer (pH 7.0) prepared with 0.085 g of NaNO3, 0.117 g of KH2PO4, and 0.485 g of Na2HPO4 12H2O in 100 ml of distilled water. A loopful of cells is suspended in 2 ml of the substrate solution, and the preparation is incubated at 37 °C for 2 hours. Nitrite formation is detected by adding 1 drop of an aqueous hydrochloric acid solution (1/1, vol/vol) and then 2 drops of a 0.2% sulfanilamide solution and 2 drops of a 0.1% naphthylethylenediamine solution. Colour intensity, which may range from pale pink (+/–) to deep red (5+), is determined by comparison with the colour standards described below. On this scale a positive reaction must be at least a 2+.
The substrate solution and all test reagents must be stored in the dark at 4 °C. If a precipitate forms or if a reagent changes colour, the solution should be discarded, and a fresh one prepared. Colour standards for interpretation are prepared as described below. A working buffer (solution 1) is prepared by combining 35 ml of 0.067 M disodium phosphate (9.47 g of anhydrous Na2HPO4 per litre), 5 ml of 0.067 M monopotassium phosphate (9.07 g of KH2PO4 per litre), and 100 ml of 0.067 M trisodium phosphate (25.47 g of Na3PO4 – 12 H2O per litre). To 10 ml of solution 1, 0.1 ml of 1% ethanolic phenolphthalein and 0.2 ml of 0.01% bromthymol blue (1 ml of 1% ethanolic bromthymol blue in 100 ml of distilled water) are added to make solution 2. A 2 ml portion of solution 2 in an appropriate tube (tube 1) represents the 5+ colour standard. Then 2 ml of solution 1 is placed into seven additional tubes (tubes 2 to 8), and 2 ml of solution 2 is added to tube 2. A 2 ml portion is transferred to the next tube, and serial dilutions of 2 ml are made in the remaining tubes using distilled water as diluent (2 ml is discarded from tube 8). Tube 2, the tube containing equal amounts of solutions 1 and 2, corresponds to the 4+ colour standard; tubes 3, 5, 6 and 8 correspond to the 3+, 2+, 1+ and +/– colour standards, respectively.
Acid phosphatase activity
The substrate solution for the acid phosphatase activity test (Wayne, 1985) consists of 100 mg of the pyridine salt of phenolphthalein phosphate dissolved in 100 ml of 0.2 M acetic acid-sodium acetate buffer (pH 5.2). The buffer should be steamed at 100 °C for 30 minutes and cooled to room temperature before the phosphate substrate is added. A tube containing 1 ml of substrate is inoculated with a loopful of cells and the preparation is incubated at 37 °C for 4 hours. Then 1 ml of 10% Na2CO3 in H2O is added to stop the reaction and develop the colour.
Suitable dilutions of a colour standard stock (1 mg of free phenolphthalein per ml of 95% ethanol) are made in H2O to yield phenolphthalein concentrations of 2.5, 5, and 10 µg/ml. Aliquots are treated with sodium carbonate solution in the same manner as the test solutions. A test preparation that is colourless or a test preparation that is pink and has an intensity less than that of the 2.5 µg/ml standard is scored as negative. Reaction mixtures that have intensities corresponding to the intensity of the 5 or 10 µg/ml standard are recorded as positive.