Part III: Culture
culture examination and identification

Examination schedule
All cultures should be examined 72 hours after inoculation to check that liquid has completely evaporated, to tighten caps in order to prevent drying out of media and to detect contaminants. Thereafter, cultures are examined weekly, or if this is not operationally feasible, on at least three occasions, viz

• after one week to detect rapidly growing mycobacteria which may be mistaken for M. tuberculosis
• after three to four weeks to detect positive cultures of M. tuberculosis as well as other slow-growing mycobacteria which may be either harmless saprophytes or potential  pathogens
• after eight weeks to detect very slow-growing mycobacteria, including M. tuberculosis,  before judging the culture to be negative

It is useful to label containers with cultures with the dates necessary for examination and to place containers in the incubator in chronological order.

Should contaminated cultures be found during the examination, those where the surface has been completely contaminated or where medium has been liquefied or discoloured should be sterilised and discarded. Certain contaminating organisms produce acid from constituents of the medium and the lowering of pH unbinds some of the malachite green from the egg (indicated by the medium changing to dark green). Tubercle bacilli will not grow under these conditions and cultures should be discarded. Cultures with partial contamination should be retained until the eighth week. Late contamination does not exclude the presence of M. tuberculosis; it is therefore advisable to prepare a smear from the surface of the medium. Should microscopy indicate the presence of acid-fast bacilli, an attempt could be made to re-decontaminate and re-inoculate the culture.

Reading of cultures
Typical colonies of M. tuberculosis are rough, crumbly, waxy, non-pigmented (cream coloured) and slow- growers, ie. only appearing three weeks after inoculation.

With doubtful cultures or when less experienced staff read cultures, the acid-fastness should be confirmed by Ziehl-Neelsen (ZN) staining. A very small amount of growth is removed from the culture using a loop and gently rubbed into one drop of sterile saline on a slide. At this point the ease with which the organisms emulsify in the liquid should be noted: Tubercle bacilli do not form smooth suspensions, unlike some other mycobacteria. The smear is allowed to dry, fixed by heat and stained by the ZN method.

For preliminary identification of tubercle bacilli the following characteristics apply:

• Tubercle bacilli do not grow in primary culture in less than one week and usually take  three to four weeks to give visible growth
• The colonies are buff coloured (never yellow) and rough, having the appearance of bread  crumbs or cauliflower
• They do not emulsify in the saline used for making smears but give a granular suspension
• Microscopically they are frequently arranged in serpentine cords of varying length or show district linear clumping. Individual cells are between 3Fm and 4Fm in length

Differentiation of M. tuberculosis
Although a presumptive diagnosis of tuberculosis may be made by an experienced laboratory technologist on the basis of the characteristics of tubercle bacilli described before, it is best to do confirmatory tests. Unfortunately there is no completely reliable single test that will differentiate M. tuberculosis from other mycobacteria. Nevertheless, the following tests, when used in combination with the characteristics described before will enable the precise identification of >95% of M. tuberculosis strains.

NIACIN TEST
Niacin (nicotinic acid) plays a vital role in the oxidation-reduction reactions that occur during metabolic processes in all mycobacteria. Although all mycobacteria produce niacin, comparative studies have shown that, because of a blocked metabolic pathway, M. tuberculosis accumulates the largest amount of nicotinic acid and its detection is useful for its definitive diagnosis. Niacin negative M. tuberculosis strains are very rare, while very few other mycobacterial species yield positive niacin tests.

Cultures grown on egg medium yield the most consistent results in the niacin test and LJ medium is therefore recommended. A culture must be at least three to four weeks old and must have sufficient growth of more than 50 colonies. Because M. tuberculosis excrete niacin into the growth medium, cultures with confluent growth may give a false-negative niacin reaction because the extracting fluid cannot come in contact with the culture medium. When this occurs, expose the underlying medium surface by either scraping away or puncturing through some of the culture growth.

Aeration of cultures intended for niacin testing is very important. Caps should be loose on slants throughout the entire incubation period and special Cap-o-Test stoppers are recommended.

Niacin test with chemical reagents
Controls
Control the reagents by testing the extract from an uninoculated tube of medium (negative control) and use an extract from a culture of M. tuberculosis H37Rv as positive control.

Reagents
Aniline solution, 4%

• Aniline is oncogenic and penetrates through the skin. Work with gloves and be very careful
• Aniline may change colour on exposure to air and light; prepare a fresh solution when  necessary

Fresh, clear colourless aniline 4ml

Ethanol 95% 96ml

Mix aniline with ethanol in an amber bottle and store in the dark in the refrigerator. Discard if solution turns yellow.

Cyanogen bromide solution, 10%

• Cyanogen bromide is a severe lacrimator and toxic if inhaled; work in a  well-ventilated fume hood when preparing the solution and in a biological safety cabinet when testing cultures
• Cyanogen bromide is oncogenic and penetrates through the skin. Work with gloves and be very careful
• In acid solution, cyanogen bromide hydrolyses to hydrocyanic acid, which is extremely toxic. Discard all reaction tubes into a disinfectant solution made alkaline by the addition of sodium hydroxide

Cyanogen bromide crystals 5g

Distilled water 50ml

• Add cyanogen bromide crystals to distilled water in a glass beaker
• Cover the beaker with foil and leave in the fume cupboard at room temperature. The crystals take approximately 24 hours to dissolve at room temperature
• Do not heat the solution over a Bunsen flame
• Pour into a tightly capped amber bottle and store in the refrigerator
• Warm to room temperature to dissolve any precipitate formed upon cooling
• Prepare small amounts because cyanogen bromide is volatile and loses strength on  storage. Weak solutions give false-negative results

To avoid unnecessary prolonged exposure to the cyanogen bromide while weighing the following procedure may be followed:

• Write down the weight of an empty beaker closed with a piece of aluminium foil
• Remove the approximate quantity (eg. approximately ¹/₁₀ of the  contents of a 100g bottle for a 10% solution) of the white cyanogen bromide crystals into the beaker, cover it and record the weight
• Calculate the difference between the two readings to obtain the exact weight of crystals in the beaker
• Add the required amount of distilled water to give a final concentration of 10%

In some countries a 4% aqueous potassium cyanide solution containing bromide is used and is prepared as follows:

• Bromine water is highly corrosive and volatile and should be stored away from other chemical reagents
• KCN is very poisonous and should be handled in a fume hood
• Break an ampoule of bromine (50ml) in a 1 000ml capacity dark glass flask (with glass  stopper) containing 150ml cold distilled water
• Prepare a 4% aqueous potassium cyanide solution by dissolving 4g KCN in 100ml distilled  water. The KCN must be pure and not hydrated
• With a pipette, remove 1ml of the bromine layer beneath the surface of the bromine water and transfer it to the bottom of a 250ml Erleynmeyer flask. Rapidly add, drop by drop, the  potassium cyanide solution, shaking by rotation until total decolorisation is obtained

Procedure
Refer to Diagram 2.

Results and interpretation

Negative : No colour

Positive : Yellow colour appearing within 5 minutes. The colour appears as a ring at the interface of the two reagents, or if the tube is shaken, as a yellow column of liquid.

Niacin test with paper strips

Paper test strips for the detection of niacin are commercially available. They compare well to the chemical reagents in detecting niacin production. A paper-strip method obviates the need to prepare and store the unstable and toxic chemicals used to demonstrate the presence of niacin, but is much more expensive.

Procedure
Refer to Diagram 3.

Results and interpretation

Negative : No colour

Positive : Yellow liquid in the bottom of the tube. Discard any colour on the stip itself; this may occur because of oxidation of chemicals, especially at the top of the strip

Precautions

• Always check the expiry date of commercial test strips
• To prevent false-negative results promptly reseal tubes  after inserting paper strip; if tubes are left unsealed the gas evolved as chemical mix on  the strip may escape into the atmosphere

NITRATE REDUCTION TEST
M. tuberculosis is one of the strongest reducers of nitrate among the mycobacteria, which allows for this test to be used in combination with the niacin test in differentiating M. tuberculosis from the other mycobacteria.

Cultures to be tested for nitrate reduction should be four weeks old and have abundant growth Löwenstein Jensen egg medium is recommended.

Classical method with liquid reagents
Reagents
Sodium nitrate substrate in buffer

Prepare 0.01M sodium nitrate in 0.022M phosphate buffer, pH 7.0 as follows:

KH₂PO₄ 3.02g
Distilled water 1000ml

Dissolve potassium phosphate in distilled water to provide an 0.022M solution      Solution 1

Na₂HPO₄ 3.16g
Distilled water 1000ml

Dissolve sodium phosphate in distilled water to provide an 0.022M solution          Solution 2

Add 611ml of solution ã to 389ml of solution â and mix well.
Check pH to be 7.0          Solution 3

Complete sodium nitrate substrate buffer

NaNO₃ 0.85g
Solution ä 1000ml

Dissolve the sodium nitrate in the buffer and dispense in 100ml aliquots. Sterilise by autoclaving at 121EC for 15 minutes. When needed, aliquots of the substrate solution are aseptically dispensed into sterile screw-capped tubes in 2ml quantities.

Hydrochloric acid solution

Concentrated HCI 10ml
Distilled water 10ml

Slowly add concentrated HCI to distilled water (never the reverse) to obtain a 1:1 dilution. Store in an amber bottle in the dark in the refrigerator.

Sulfanilamide solution, 0.2%

Sulfanilamide 0.2g
Distilled water 100ml

Dissolve sulfanilamide in distilled water and store in an amber bottle in the dark in a refrigerator.

N-naphthylethylene-diamine solution, 0.1%

N-naphthylethylene-diamine 0.1g
Distilled water 100ml

Dissolve naphthylethylene-diamine in distilled water and store in an amber bottle in the dark in a refrigerator.

Controls
Control the reagents by testing the extract from an uninoculated tube of medium (negative control) and use an extract from a culture of M. tuberculosis H37Rv as positive control.

Procedure
Refer to Diagram 4.

Results and interpretation
Negative : No colour. If no colour develops, the test is either negative or the reduction has proceeded beyond nitrite. Add a small amount of powdered zinc to all negative tests by tipping the end of a slightly moistened applicator stick into dry zinc and shaking into the liquid.

1. If nitrate is still present, it will be catalysed by the zinc and a red  colour will develop, indicating a true negative
2. If no colour develops the original reaction was positive but the nitrate  was reduced beyond nitrite. Repeat the test to confirm the observation

Positive : Red colour, which vary from pink to very deep red-crimson:

Faint pink = +/-
Clear pink = 1+
Deep pink = 2+
Red = 3+
Deep red = 4+
Purplish red = 5+

Only 3+ to 5+ is considered positive.

Method with crystalline reagent
The dry crystalline reagent is easy to prepare, has a shelf-life of a least six months and has the added advantage that only one reagent is needed to detect nitrate rather than the three liquid reagents used in the conventional chemical test.

Reagents
Sodium nitrate substrate in buffer

Prepare as described in classical method, page 60.

Crystalline reagent
Sulfanilic acid 1 part
N-(1-naphthyl)-etylenediamine dihydrochloride 1 part
L(+) - tartaric acid10 parts

Put the chemicals in an amber bottle and mix by vigorous manual shaking about 30 times. (Tartaric acid is much more crystalline than the other two chemicals and may have to be ground using a mortar and pestle to ensure good mixture of the reagents). The dry mixture has a heterogeneous crystalline appearance. Store in the amber bottle at room temperature.

Controls
Control the reagents by testing the extract from an uninoculated tube of medium (negative control) and use an extract from a culture of M. tuberculosis H37Rv as positive control.

Procedure
Refer to Diagram 5.

Results and interpretation
Negative : No colour. If no colour develops, the test is either negative or the reduction has proceeded beyond nitrite. Add a small amount of powdered zinc to all negative tests by tipping the end of a slightly moistened applicator stick into dry zinc and shaking into the liquid.

1. If nitrate is still present, it will be catalysed by the zinc and a red  colour will develop, indicating a true negative
2. If no colour develops the original reaction was positive but the nitrate  was reduced beyond nitrite. Repeat the test to confirm the observation

Positive : Red colour, which vary from pink to very deep red-crimson:

Faint pink = +/-
Clear pink = 1+
Deep pink = 2+
Red = 3+
Deep red = 4+
Purplish red = 5+

Only 3+ to 5+ is considered positive.

Nitrate test with paper strips

Paper test strips for the detection of nitrate following nitrate reduction are commercially available. The paper strip test method yields most consistent results with mycobacteria that vigorously reduce nitrate, such as M. tuberculosis. It therefore provides reliable results and is much less labour-intensive than the chemical method, but is much more expensive.

Procedure
Refer to Diagram 6.

Results and interpretation
Negative : No colour change

Positive : Top portion of the strip changes to light or dark blue

Precautions

• Always check the expiry date of commercial test strips
• Because the strips are sensitive to sunlight, excess heat  and moisture, they should be stored between 2EC and  8EC in the original container, tightly capped
• Discard strips if they become discoloured, for this  indicates deterioration of the reagent
• Do not rely on results of test strips if the positive  control culture gives weak or negative reactions

NITRATE REDUCTION STANDARDS
In order to ensure consistency in interpreting nitrate reduction reactions it is recommended that a series of standards depicting the colour intensity from " to 5+ be prepared. These keep indefinitely and should be used whenever nitrate tests are done.

Reagents
Stock solution

1. 0.067M disodium phosphate (9,47g of  anydrous Na₂HPO₄ per 1 000ml)
2. 0.067M monopotassium phosphate (9.07g of KH₂PO₄ per 1 000ml)
3. 0.067M trisodium phosphate (25.47g of Na₃PO₄C12H₂O per 1 000ml)
4. 1% phenolphthalein (1g in 100ml 95% ethyl  alcohol)
5. 1% bromthymol blue (1g in 100ml 95% ethyl  alcohol)
6. 0.01% bromthymol blue: prepare by mixing 1.0ml of no.Ò above in 100ml of distilled  water.

Working buffer solution

Mix 35ml of stock solution Î, 5ml of stock solution Ï and 100ml of solution Ð.

Procedure

• Place eight clean test tubes (number 1-8) in a rack. Use the same size tubes as used to perform the nitrate reduction test.
• Put 2ml of working buffer solution into tubes 2 through 8.
• To 10ml of working buffer solution, add 0.1ml of Ñ and 0.2ml of Ó ___________ Solution Ô
• Add 2ml of solution Ô to the tube numbered 1. This is the 5+ colour standard.
• To the tube  number 2 in the series, add 2ml of solution Ô.  Mix well and transfer 2ml to the next tube (number 3). Continue to make serial dilutions  of 2ml, discarding 2ml from the 8th tube.
• The colour standards:
• tube 1 = 5+
• tube 2 = 4+
• tube 3 = 3+
• tube 5 = 2+
• tube 6 = 1+
• tube 8 = +/-
• Autoclave  tubes, seal and store at 5EC.

CATALASE TEST
Catalase is an intracellular, soluble enzyme capable of splitting hydrogen peroxide into water and oxygen, ie. 2H₂O₂6 2H ₂O + O ₂. The oxygen bubbles into the reaction mixture to indicate catalase activity. Virtually all mycobacteria passes catalase enzymes, except for certain isoniazid-resistant mutants of M. tuberculosis and M. bovis.

Mycobacteria posses several kinds of catalase that vary in heat stability. Quantitative differences in catalase activity can be demonstrated by one or more of the following tests:

• Room temperature or drop method  (indicates the presence of catalyse)
• Semiquantitative test (indicates level of catalyse production)
• 68EC  test at pH7 (indicates loss of catalyse activity due to heat)

Drug susceptible strains of M. tuberculosis do form catalyse as indicated by the drop method, produce less than 45mm of bubbles in the semiquantitative test and lose catalase activity when heated to 68EC for 20 minutes. For these tests 14 day-old cultures on LJ butts should be used, ie. the media tubes should be inspissated in an upright position to provide a butt and should not be slanted. The tubes must have stoppers which permit exchange of air, eg. Cap-o-Test stoppers. The cultures should be incubated in a well-humidified incubator at 35E-37EC, with loose caps, for 14 days.

Reagents
0.067M phosphate buffer solution, pH 7.0

Na₂HPO₄ anhydrous 9.47g
Distilled water 1000ml

Dissolve disodium phosphate in distilled water to provide an 0.067M solution ____ Solution 1

KH₂PO₄ 9.07g
Distilled water 1000ml

Dissolve monopotassium phosphate in distilled water to provide an 0.067M solution __ Solution 2

Hydrogen peroxide, 30%

30% hydrogen peroxide (H₂O₂), also known as Superoxol (Merck) is stored in the refrigerator.

• Ensure that the H₂O₂ used is 30% and not the 3% kind obtained from pharmacies
• Wear rubber or plastic gloves and a  protective eye shield when handling superoxol

Tween 80, 10%

Tween 80 10ml
Distilled water 90ml

Mix Tween 80 with distilled water and autoclave at 121EC for 10 minutes. The Tween may settle during autoclaving and may be resuspended by swirling immediately after autoclaving and during cooling. Store in the refrigerator.

Complete catalase reagent (Tween-peroxide mixture)

Immediately before use, mix equal parts of 10% Tween 80 and 30% hydrogen peroxide. Allow 0.5ml reagent for each strain to be tested.

Controls
Drop method

Use an uninoculated tube of medium as negative control and an LJ butt of M. tuberculosis H37Rv as positive control

Semiquantitative and 68EC tests

Use an uninoculated tube of medium as negative control and an LJ butt of M. terrae as positive control.

Procedures
Drop method

Examine the 14 day-old LJ slant to ascertain that growth has occurred. Add one to two drops of the freshly-prepared Tween-peroxide mixture to the slant with the culture growth. Observe for a period of 5 minutes for the formation of bubbles.

Results and interpretation

Negative : No bubbles formed

Positive (slow) : Few slowly forming bubbles

Positive (rapid) : Immediate copious formation of bubbles

Semiquantitative test

Examine the 14 day-old LJ butt to ascertain that growth has occurred. Add 1ml of the freshly-prepared Tween-peroxide mixture, replace caps loosely and allow to stand at room temperature for 5 minutes. A column of foam will form.

Measure the height of the foam column (ie. from the top of the liquid on the LJ medium to the top of the foam).

Results and interpretation

Low or no catalase activity : Less than 31mm of foam

Inconclusive result : Between 31 and 45mm of foam

High catalase activity : More than 45mm of foam

'68EC, pH7.0 test

Refer to Diagram 7.

Results and interpretation

Positive : Bubbles

Negative : No bubbles

On rare occasions, bubbles may be seen rising from the sedimental cells in such small quantity that foam does not form at the surface of the fluid. This is still recorded as a positive reaction.

GROWTH ON MEDIUM CONTAINING p-NITROBENZOIC ACID (PNB)
In laboratories where facilities and reagents for niacin and nitrate testing are not available, identification of tubercle bacilli may be done by a combination of one or more of the catalase tests described previously together with growth at 25EC on LJ medium and growth on LJ medium containing p-nitrobenzoic acid at 37EC. Problems with incubation at 25EC may be encountered in tropical regions. A refrigerated incubator should be used where available; as an alternative, a water bath within a refrigerator or cold room should be used.

Procedure

• Inoculate two slopes of LJ medium containing glycerol and  one tube of LJ medium containing p-nitrobenzoic acid (PNB) at a concentration of 500mg/litre
• Incubate one LJ slope and the PNB slope at 37EC in an internally illuminated incubator and examine at 3, 7, 14 and 21 days. When growth is evident on the LJ slope examine it for pigment. If an internally illuminated incubator is not available, remove slopes from the dark incubator as soon as growth is evident, loosen the caps to  admit some oxygen and expose them to daylight (but not direct sunlight) or place 1m from a  laboratory bench lamp for 1 hour. Reincubate and examine for pigment the following day
• Incubate the other LJ slope at 25EC and examine at 3, 7, 14 and  21 days

Results and interpretation
M. tuberculosis does not grow within three days at 37EC and does not grow at all at 25EC or on PNB medium. It also does not produce yellow or orange pigment in the dark or after exposure to light.