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Mycobacterium tuberculosis

Minggu, 18 Maret 2012


Part III: Culture
media
The definitive diagnosis of tuberculosis demands that M. tuberculosis be recovered on culture media and identified using differential in vitro tests. Many different media have been devised for cultivating tubercle bacilli and three main groups can be identified, viz egg-based media, agar-based media and liquid media.
The ideal medium for isolation of tubercle bacilli should (a) be economical and simple to prepare from readily available ingredients, (b) inhibit the growth of contaminants, (c) support luxuriant growth of small numbers of bacilli and (d) permit preliminary differentiation of isolates on the basis of colony morphology. For the culture of sputum specimens, egg-based media should be the first choice, since they meet all these requirements. These is increasing evidence that liquid media may give better results with other specimens. While cost prevents their routine use with sputum specimens, it is recommended that both egg-based and liquid medium be used for non-repeatable specimens, eg. cerebrospinal fluid and biopsy material.
It is recommended that all sputum specimens submitted for culture also undergo microscopic examination as outlined in the Technical Series on Microscopy.
Advantages and disadvantages of egg-based media
Advantages
•    it is easy to prepare
•    it is the least expensive of all media available and supports good growth of tubercle bacilli
•    it may be stored in the refrigerator for several weeks provided it was made from fresh eggs and culture bottle caps are tightly closed to minimise drying by evaporation
•    contamination during preparation is  limited because it is inspissated after being placed in bottles. In addition, the  malachite green added to the media suppresses the growth of non mycobacterial organisms
Disadvantages
•    it may take as long as eight weeks before  cultures become positive, especially if specimens contain few bacilli or if  decontamination procedures have been overly harsh
•    when contamination does occur, it often  involves the total surface of the medium and the culture is usually lost
Precautions during media preparation
For media of the best quality, chemicals of certified purity, clean glassware and freshly distilled and sterilised water should be used. Directions for preparing media must be followed precisely and without modification. A few general points to obtain good quality media and avoid contamination of reagents and media are as follows:
•    Keep the environment as clean as possible. Swab the work surface with a suitable disinfectant (eg.  5% methylated spirits) before dispensing sterile reagents and media. Clean the floor with a wet mop to limit dust
•    Use sterile  glassware and equipment
•    Use reagent  grade chemicals and reagents unless otherwise specified
•    Check the  temperature of inspissators and hot air ovens
•    Follow strict  aseptic techniques when preparing media, eg. flaming flasks and tubes
•    When preparing egg-based media, carefully clean egg shells before breaking
•    Do not overheat medium during inspissation
•    Do not leave prepared media exposed to light (including ultra-violet light), but store in the refrigerator in the dark when not in use
•    Do not skimp on the volume of medium. Place 6-8ml of egg medium in each bottle or 20ml into each test tube

Preparation of egg-based media


LÖWENSTEIN-JENSEN MEDIUM

Löwenstein-Jensen (LJ) medium is most widely used for tuberculosis culture. The modification of the International Union Against Tuberculosis and Lung Disease (IUATLD) is recommended and will be described in detail. LJ medium containing glycerol favours the growth of M. tuberculosis while LJ medium without glycerol but containing pyruvate encourages the growth of M. bovis. Both should be used in countries or regions where patients may be infected with either organism.
Ingredients
Mineral salt solution
Potassium dihydrogen phosphate anhydrous (KH2PO4) 2.4g
Magnesium sulphate (MgSO4. 7H2O) 0.24g
Magnesium citrate 0.6g
Asparagine 3.6g
Glycerol (reagent grade) 12ml
Distilled water 600ml
Dissolve the ingredients in order in the distilled water by heating. Autoclave at 121EC for 30 minutes to sterilise. Cool to room temperature. This solution keeps indefinitely and may be stored in suitable amounts in the refrigerator.
Malachite green solution, 2%
Malachite green dye 2.0g
Sterile distilled water 100ml
Using aseptic techniques dissolve the dye in sterile distilled water by placing the solution in the incubator for 1-2 hours. This solution will not store indefinitely and may precipitate or change to a less-deeply coloured solution. In either case discard and prepare a fresh solution.
Homogenised whole eggs
Fresh hens' eggs, not more than seven days old, are cleaned by scrubbing thoroughly with a hand brush in warm water and a plain alkaline soap. Let the eggs soak for 30 minutes in the soap solution. Rinse eggs thoroughly in running water and soak them in 70% ethanol for 15 minutes. Before handling the clean dry eggs scrub the hands and wash them. Crack the eggs with a sterile knife into a sterile flask and beat them with a sterile egg whisk or in a sterile blender.

Preparation of complete medium

The following ingredients are aseptically pooled in a large, sterile flask and mixed well:
Mineral salt solution 600ml
Malachite green solution 20ml
Homogenised eggs (20-25 eggs, depending on size) 1000ml
The complete egg medium is distributed in 6-8ml volumes in sterile 14ml or 28ml McCartney bottles or in 20ml volumes in 20 x 150mm screw-capped test tubes, and the tops are securely fastened.
Inspissate the medium within 15 minutes of distribution to prevent sedimentation of the heavier ingredients.
Coagulation of medium
Before loading, heat the inspissator to 80EC to quicken the build-up of the temperature. Place the bottles in a slanted position in the inspissator and coagulate the medium for 45 minutes at 80E-85EC (since the medium has been prepared with sterile precautions this heating is to solidify the medium, not to sterilise it). Heating for a second or third time has a detrimental effect on the quality of the medium.
The quality of egg media deteriorates when coagulation is done at too high a temperature or for too long. Discolouration of the coagulated medium may be due to excessive temperature. The appearance of little holes or bubbles on the surface of the medium also indicates faulty coagulation procedures.

Poor quality media should be discarded.

Sterility check
After inspissation, the whole media batch or a representative sample of culture bottles should be incubated at 35E-37EC for 24 hours as a check of sterility.
Storage
The LJ medium should be dated and stored in the refrigerator and can keep for several weeks if the caps are tightly closed to prevent drying out of the medium. For optimal isolation from specimens, LJ medium should not be older than 4 weeks.
For the cultivation of M. bovis, LJ medium is enriched with 0,5% sodium pyruvate. Glycerol is omitted and 8.0g sodium pyruvate is added to the mineral solution.

OGAWA MEDIUM

This medium is cheaper than Löwenstein-Jensen because it is made without asparagine.
Ingredients
Mineral salt solution
Potassium dihydrogen phosphate anhydrous (KH2PO4) 3.0g
Sodium glutamate 3.0g
Distilled water 300ml
Dissolve the ingredients in distilled water by heating. Autoclave at 121EC for 30 minutes to sterilise. Cool to room temperature. This solution keeps indefinitely and may be stored in suitable amounts in the refrigerator.
Malachite green solution, 2%
Malachite green dye 2.0g
Sterile distilled water 100ml
Using aseptic techniques dissolve the dye in sterile distilled water by placing the solution in the incubator for 1-2 hours. This solution will not store indefinitely and may precipitate or change to a less-deeply coloured solution. In either case discard and prepare a fresh solution.
Homogenised whole eggs
Fresh hens' eggs, not more than seven days old, are cleaned by scrubbing thoroughly with a hand brush in warm water and a plain alkaline soap. Let the eggs soak for 30 minutes in the soap solution. Rinse eggs thoroughly in running water and soak them in 70% ethanol for 15 minutes. Before handling the clean dry eggs scrub the hands and wash them. Crack the eggs with a sterile knife into a sterile flask and beat them with a sterile egg whisk or in a sterile blender.
Preparation of complete medium
The following ingredients are aseptically pooled in a large, sterile flask and mixed well:
Mineral salt solution 300ml
Malachite green solution 18ml
Whole hens? eggs (12-16 eggs, depending on size) 600ml
Glycerol 18ml
The resulting pH of the medium is 6.8. The medium is mixed well and distributed in 6-8ml volumes in sterile 14ml or 28ml McCartney bottles or in 20ml volumes in 20x150mm screw-capped test tubes.
Coagulation of medium
Before loading, heat the inspissator to 80EC to quicken the build-up of the temperature. Place the bottles in a slanted position in the inspissator and coagulate the medium for 45 minutes at 80E-85EC (since the medium has been prepared with sterile precautions this heating is to solidify the medium, not to sterilise it). Heating for a second or third time has a detrimental effect on the quality of the medium.
The quality of egg media deteriorates when coagulation is done at too high a temperature or for too long. Discolouration of the coagulated medium may be due to excessive temperature. The appearance of little holes or bubbles on the surface of the medium also indicates faulty coagulation procedures.
Poor quality media should be discarded.
Sterility check
After inspissation, the whole media batch or a representative sample of culture bottles should be incubated at 37EC for 24 hours as a check of sterility.
Storage
The medium should be dated and stored in the refrigerator and can keep for several weeks if the caps are tightly closed to prevent drying out.
In laboratories where centrifuges are not available, a simple culture technique could be employed as follows: Sputum specimens are decontaminated with equal volumes of 4% NaOH and inoculated directly onto modified or acid-buffered Ogawa medium. This technique shows a fairly comparable oase yield when compared with concentrated culture techniques.

ACID-BUFFERED OGAWA MEDIUM

Ingredients
Modified Ogawa Acid-buffered Ogawa
Potassium dihydrogen phosphate 2g 3g
Magnesium citrate (KH2PO4) 0.1g -
Sodiumglutamate 0.5g 1.0g
Glycerol 4ml 6ml
Distilled water 100ml 100ml
Homogenised whole eggs 200ml 200ml
2% Malachite green solution 4ml 6ml
Final pH 6.4 6.2
Preparation
Dissolve the ingredients in the distilled water and boil for 30 minutes. Cool to room temperature and add the homogenised eggs and malachite green solution. Transfer 6-8ml volumes to suitable bottles and inspissate at 85EC for 45-60 minutes.
A variety of more expensive and labour intensive culture methods1are available to countries with the required financial and human resources. Some of these will be discussed briefly:
OPTIONS

HERMAN KIRCHNER LIQUID MEDIUM

This medium is most useful and least expensive of the liquid media for culture and tubercle bacilli. It has the additional advantage that it can support a  large inoculum.

DUBOS OLEIC ACID-ALBUMIN LIQUID MEDIUM
This medium is recommended for the cultivation of tubercle bacilli from cerebrospinal, pleural and peritoneal fluid. It may be prepared from basic ingredients or may be obtained commercially as a ready-to-use base to which sterile albumin or serum is added.

MIDDLEBROOK 7H-10 AND 7H-11 AGAR MEDIUM
Middlebrook 7H-10 may be made from basic ingredients or may be prepared from commercially available 7H-10 agar-powdered base and Middlebrook oleic  acid-albumin-dextrose-catalase (OADC) enrichment. 7H-11 is a 7H-10 agar enriched by the  addition of enzymatic digest of casein. It is best to prepare 7H-10 and 7H-11 medium in  small quantities of 200 to 400ml to minimise the amount of heat needed to melt the agar.  Boiling the basal medium before autoclaving (either to solubilise the agar or to provide stocks of prepared base that may be stored and boiled for later use) should be avoided  because the repeat heating produces medium of inferior quality.
When Middlebrook 7H-10 or 7H-11 medium is used for isolation cultures must  be incubated in an atmosphere of 10% CO2. Exposure of Middlebrook 7H-10 or  7H-11 agar to either daylight of heat results in the release of formaldehyde in sufficient concentration to inhibit the growth of mycobacteria.
   
 
OPTION
SELECTIVE MEDIUM
Specimens which are excessively contaminated may be inoculated onto selective antibiotic-containing media. Use may be made of antibiotics to which mycobacteria are not sensitive but which are capable of destroying the contaminants, eg.  penicillin (50-100 units/ml), nalidixic acid (35Fg/ml) or polymyxin (20-25Fg/ml).
The antibiotics may be:
•    Added to egg medium before inspissation
•    Added to the surface of the medium slant or
•    Mixed with the inoculum
Mycobactosel medium contains several antibiotics, eg. cycloheximide  (0.4mg/ml), lincomycin (0.002mg/ml) and nalidixic acid (0.035mg/ml), while mycobactosel agar is commercially available. Antibiotic enriched medium should be stored in the  refrigerator in the dark for a maximum of four weeks.
1 Kent PT, Kubica GP. Public health mycobacteriology: Guide for the Level III Laboratory. US Department of Health and Human Services, Centres for Disease Control, USA, 1985.  
    
OPTION
RADIOMETRIC METHOD FOR TUBERCULOSIS CULTURE
Recent development in the diagnosis of tuberculosis include an automated system for detecting early growth of mycobacteria by a radiometric method (BACTEC: Beckton Dickinson). Sputum or other homogenates are decontaminated as necessary  and added to vials containing Middlebrook 7H12 medium, an antibiotic mixture (to avoid the  growth of other organisms) and 14C-labelled palmitic acid. The medium is  prepared commercially (BACTEC 12B: Beckton Dickinson) in rubber-sealed bottles  and inoculated with a syringe and hypodermic needle. If mycobacterial growth occurs, 14C  palmitic acid is utilised and 14CO2 is produced. The air space above  the medium in each bottle is sampled automatically by the BACTEC machine at fixed  intervals and the amount of radioactive gas is estimated and recorded. Infectious aerosols  are contained in the apparatus and captured in HEPA filters before the air is exhausted.
Growth of mycobacteria may be detected within 5-7 days, but positive results require further testing to distinguish between tubercle bacilli and other  mycobacteria. In the BACTEC machine, p-nitro-a-acetylamino-ß-priophenone (NAP) is used and tubercle  bacilli can be differentiated within five days. NAP inhibits the growth of M. tuberculosis and usually does not affect the growth of MOTT bacilli.
Comparative tests have shown that the method is very successful and  reliable and that confirmatory results for M. tuberculosis can be obtained within  two weeks. However, the BACTEC machine is very expensive to purchase and to operate. In addition, two hazards must be considered if the machine is to be used for routine tuberculosis bacteriology: the use of hypodermic needles for the inoculation of media  carries the risk of needle-stick injury, while the culture media is radioactive and presents a problem in terms of waste disposal.
In summary, the BACTEC method is invaluable for the detection of tubercle bacilli in material such as cerebrospinal fluid where rapid results are crucial in the management of the patient. However, the high cost of both the apparatus and the  radio-labelled medium prohibits its routine use in most high tuberculosis prevalence countries.

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Part III: Culture
homogenisation and decontamination
The usual microbiological techniques of plating clinical material on selective or differential culture media and subculturing to obtain pure cultures cannot be applied to tuberculosis bacteriology. M. tuberculosis requires special media not used for other organisms and grows slowly, taking three to six weeks or longer to give visible colonies. Cultures are usually made in bottles rather than in petri dishes because tubercle bacilli are present in relatively small numbers in most specimens; this necessitates large inocula which are spread out over the surface of the media. Because of the long incubation time required, the bottles are tightly stoppered to prevent drying of the cultures (which would occur in petri dishes).
The majority of clinical specimens submitted to the tuberculosis culture laboratory are contaminated to varying degrees by more rapidly growing normal flora organisms. These would rapidly overgrow the entire surface of the medium and digest it before the tubercle bacilli start to grow. Most specimens must, therefore, be subjected to a harsh digestion and decontamination procedure that liquefies the organic debris and eliminates the unwanted normal flora.
All currently available digesting/decontaminating agents are to some extent toxic to tubercle bacilli; therefore, to ensure the survival of the maximum number of bacilli in the specimen, the digestion/decontamination procedure must be precisely followed. In order for enough tubercle bacilli to survive to give a confirmatory diagnosis, it is inevitable that a proportion of cultures will be contaminated by other organisms. As a general rule, a contamination rate of 2%-3% is acceptable in laboratories that receive fresh specimens; if specimens (especially sputum) take several days to reach the laboratory then losses due to contamination may be as high as 5%-10%. It is also important to note that a laboratory which experiences no contamination is probably using a method that kills too many of the tubercle bacilli.
When culturing tubercle bacilli, three important aspects should be borne in mind:
•    Specimens must be homogenised to free the bacilli from the mucus, cells or tissue in  which they may be embedded. The milder this homogenisation the better the results
•    Neither homogenisation nor decontamination should unnecessarily diminish the viability of tubercle bacilli
•    The success of homogenisation and decontamination depends on:
o    the greater resistance of tubercle bacilli to strongly alkaline or acidic digesting solutions
o    the length of exposure time to these agents
o    the temperature build-up in the specimen during centrifugation
o    the efficiency of the centrifuge used to sediment the tubercle bacilli
Many different methods of homogenisation and decontamination of sputum specimens for culturing have been described but there is no universally recognised best technique. The choice of a suitable method is to a large extent determined by the technical capability and the availability of staff in a laboratory, as well as the quality and type of equipment available. Each method has its limitations and advantages and it is recommended that regional/central laboratories standardise on one method only. Methods which consistently yield the highest percentage of positive cultures are those which require:
•    well trained staff
•    relatively expensive equipment (eg. centrifuges) and related supplies
•    continued maintenance of equipment and of good staff performance
Any method which require the use of a centrifuge present some problems which must be considered:
•    The centrifuge must be fast enough to attain a relative centrifugal force (RCF) of 3 000 x g. If the RCF is not high enough, many tubercle bacilli remain in suspension following  centrifugation and are poured off with the discarded supernatant fluid. Recent studies  have shown that 3 000 x g for 15 minutes would sediment 95% of mycobacteria in a digested sputum specimen. The specific gravity of tubercle bacilli ranges from 1.07 to 0.79, making centrifugal concentration of specimens ineffective if the RCF is not 3 000 x g
•    Precautions must be taken to minimise the potential for staff infection in the event of  tube breakage during centrifugation. These include:
o    using a floor model centrifuge with lid and a fixed angle rotor. The mass of the fixed  angle rotor permits centrifugation of tubes with small weight differences without causing  vibration and possible tube breakage
o    always ensuring that tubes in the centrifuge are balanced. The weight of centrifuge tubes can be balanced by adding sterile saline to specimens or by inserting tubes with sterile water or 70% ethanol among the tubes containing sputum (using 70% ethanol in stead  of water in the tubes used as balances will reduce the risk in the event of breakage)
o    using aerosol-free safety cups if available
o    enclosing the centrifuge in a specially ventilated cabinet if possible
Digestion and decontamination procedures
Sputum specimens
Sputum specimens should not be pooled because of the risk of cross-contamination. Since the exposure time to digestants/decontaminants has to be strictly controlled it is best to work in sets equivalent to one centrifuge load (eg. eight specimens at a time).
Always digest/decontaminate the whole specimen, ie. do not attempt to select portions of the specimen as is done for direct microscopy. If the sputum will pour, it should be gently decanted from the specimen container into the centrifuge tube. If the specimen is too viscuous to pour, an equal volume of digestant/decontaminant could be added to the sputum in the specimen container and the mixture poured carefully into the appropriate centrifuge tube.
Since sputum specimens are the most common clinical specimens submitted for tuberculosis culture, homogenisation and decontamination procedures have been largely targeted towards their processing. Specimens other than sputum demand even more care during processing because of the low numbers of tubercle bacilli present in positive specimens.
SODIUM HYDROXIDE (MODIFIED PETROFF) METHOD
This method is used widely in developing countries because of its relative simplicity and the fact that the reagents are easy to obtain.
NaOH is toxic, both for contaminants and for tubercle bacilli; therefore, strict adherence to the indicated timing is required
Reagents
4% sodium hydroxide (NaOH) solution
Sodium hydroxide pellets (analytical grade) 4g
Distilled water 100ml
Dissolve NaOH in distilled water and sterilise by autoclaving at 121EC for 15 minutes.
Sterile saline
Sodium chloride pellets (analytical grade) 0.85g
Distilled water 100ml
Dissolve NaCI in distilled water and sterilise by autoclaving at 121EC for 15 minutes.
An alternative method for preparing 4% NaOH is as follows:
Add the contents of a 250g bottle NaOH pellets to 500ml distilled water and fill water to the 625ml mark. Be careful since heat is released in this reaction. When needed, prepare a fresh 4% NaOH solution by adding 40% NaOH to sterile distilled water in the proportion 1:10.
Procedure
Refer to Diagram 1.
Advantages
•    The NaOH method is simple and inexpensive and provides fairly effective control of contaminants
•    The time needed to process a single specimen is approximately one hour; 20 specimens would take  approximately two hours, with centrifuge capacity being the limiting factor
•    Sterilised NaOH solution will keep for several weeks
Limitations
•    The specimen exposure times must be strictly followed to prevent over-kill of tubercle bacilli
•    The NaOH procedure is very robust and may kill up to 60% of tubercle bacilli in clinical specimens.  This initial kill is independent of additional contributory factors such as heat build-up in the centrifuge and centrifugal efficiency
A variety of more expensive and labour intensive homogenisation and decontamination methods1 are available to countries with the required financial and human resources. Some of these will be discussed briefly.
1Kent PT, Kubica GP. Public health mycobacteriology: Guide for the Level III Laboratory. US Department of Health and Human Services, Centres for Disease Control, USA, 1985.
OPTION
N-ACETYL-L-CYSTEINE-SODIUM HYDROXIDE (NALC-NaOH) METHOD
The mucolytic agent NALC (used for rapid digestion of sputum) enables the decontaminating agent (NaOH) to be used at a lower final concentration of 1%. Sodium citrate is included in the digestant mixture to bind the heavy metal ions which may be present in the specimen and could inactivate the acetyl-cysteine.
•    Properly performed, this method provides more positive cultures than other methods, resulting in the killing of approximately 30%  of tubercle bacilli
•    The time needed to process a single  specimen is approximately 40 minutes; 20 specimens would take approximately 60 minutes
•    Acetyl-cysteine loses activity rapidly in  solution, so the digestant should be made fresh daily
•    The indicated specimen exposure time must  be strictly adhered to and a 1:10 dilution of resuspended sediment must be made to  decrease the concentration of any toxic components that may inhibit growth of tubercle bacilli
•    Reagents such as bovine albumin and the  required filters are expensive
      
OPTION
ZEPHIRAN-TRISODIUM PHOSPHATE (Z-TSP) METHOD
The use of trisodium phosphate and Zephiran (benzalkonium  chloride) to homogenise and decontaminate specimens results in a more gentle digestion  procedure.
•    The procedure need not be as critically  timed as NaOH digestion procedures
•    The method results in the killing of approximately 30% of tubercle bacilli
•    The time required for one specimen is  nearly two hours; 20 specimens would require four hours
Excessive contamination is sometimes encountered in clinical material from certain patients, from certain areas or at certain times, and may present a difficult problem. For these problem specimens alternative decontamination methods such as 5% oxalic acid or 4% sulphuric acid may be used.1
OPTIONS
OXALIC ACID METHOD
This method is often helpful for specimens consistently  contaminated with Pseudomonas species.
SULPHURIC ACID METHOD
This method is sometimes helpful for urine and other thin  watery body fluids that consistently yield contaminated cultures when processed with one  of the alkaline digestants.
Other specimens
Gastric lavage
These specimens should be processed within four hours of collection since their acidity is damaging to tubercle bacilli. Usually, gastric lavage does not need to be decontaminated, provided it has been collected aseptically in a sterile container. Centrifuge the total volume at 3 000 x g for 30 minutes. If contamination is suspected the sediment should be mixed with 2ml of 4% sulphuric acid and allowed to stand for 15 minutes, after which 15ml sterile saline is added. Centrifuge this mixture at 3 000 x g for 15 minutes and neutralise the sediment with 4% NaOH containing a phenol red indicator. Inoculate the sediment immediately onto culture medium.
Urine
Centrifuge the total volume at 3 000 x g for 15 minutes. Discard the supernatant fluid and add 2ml of 4% sulphuric acid to the sediment. Let stand for 15 minutes, add 15ml sterile saline and centrifuge at 3 000 x g for 15 minutes. Neutralise the sediment with 4% NaOH containing a phenol red indicator. Inoculate the sediment immediately onto culture medium.
1Kent PT, Kubica GP. Public health mycobacteriology: Guide for the Level III Laboratory. US Department of Health and Human Services, Centres for Disease Control, USA, 1985.
Laryngeal swabs
Cover the swab (in its original tube) with 5% oxalic acid and allow to act for 15 minutes. Remove the swab to another tube containing sterile saline. Lift after a few minutes, allow to drain and use to inoculate culture media.
For optimal results the oxalic acid (which might contain tubercle bacilli washed off from the swab) should be transferred to a centrifuge tube and centrifuged at 3 000 x g for 15 minutes. Wash the sediment once with sterile saline, centrifuge at 3 000 x g for 15 minutes and inoculate immediately onto culture medium.
Tissue
Lymph nodes, biopsies and other surgically resected tissue should be cut into small pieces with a sterile scalpel or scissors. Homogenise the specimen in a sterile porcelain mortar or tissue grinder, using 0.5-1ml sterile saline and a small quantity of sterilised sand (if necessary in mortar). This suspension can be directly inoculated onto culture media if the sterility measurements described before have been met; if not, decontaminate using 4% sulphuric acid as described for urine.
Mortars, pestles and tissue grinders must be cleaned and sterilised thoroughly to prevent false positive results or contamination due to organisms left over from previous specimens
Pus
This may be treated in the same way as aspirated fluids. If the material is very thick, it should be treated in the same way as sputum.
Cerebrospinal fluid
Cerebrospinal fluid should be concentrated by membrane filtration, by high speed centrifugation or by precipitation methods. Precipitation can be achieved by adding 0.1ml sterile rabbit serum or an equivalent albumin solution for every 10ml of cerebrospinal fluid. Mix until uniformly cloudy, centrifuge at 3 000 x g for 15 minutes and culture the sediment if contamination is not suspected.
If no sediment can be obtained by centrifugation, a sterile 20% solution of sulpho-salicylic acid may be added drop by drop until turbidity sets in. This precipitate is then more easily spun down to form a sediment.
If contamination of cerebrospinal fluid is likely, the sediment is mixed with 2ml of 4% sulphuric acid and allowed to stand for 15 minutes. Add 15 ml of sterile saline and centrifuge at 3 000 x g for 15 minutes. Inoculate sediment onto culture media.
Clots
In the case of specimens that form large clots, eg. pleural and ascitic fluids, it is recommended that clot formation be avoided by the addition of sodium citrate at the time of specimen collection. Add two drops of 20% sodium citrate for every 10ml fluid collected.
When clots are present, they can be digested with Petroff?s NaOH method after homogenisation as described for tissue.
Other body fluids (including pleural fluid)
Mucopurulent fluid : Treat as for sputum when volume is 10ml or less.
Clear fluid : If collected aseptically centrifuge at 3 000 x g for 15 minutes and inoculate sediment directly onto culture media. If volume is more than 10ml treat as for gastric lavage.
Tubercle bacilli may adhere to glass or plastic surfaces. To optimise recovery, containers could be rinsed with sterile saline. Centrifuge the saline at 3 000 x g for 15 minutes and inoculate 2-3 drops onto culture media.
Specimens differ greatly in their degree of contamination and decontaminants should be selected to suit the nature of the specimens. The need for decontamination is also determined by the freshness of the specimen and by the efficiency of refrigeration before processing.
The following specimens usually do not need decontamination when aseptically collected into sterile containers:
•    Spinal, sinovial or other internal body fluids
•    Bone marrow
•    Pus from cold abscesses
•    Surgically resected specimens  (excluding autopsy material)
•    Material obtained from pleural, liver  and lymph nodes as well as biopsies (if not fistulised)
Whenever doubt exists about the contamination of specimens, an untreated portion may be inoculated onto nonselective bacteriological media, eg. nutrient agar, and incubated for 24 hours to check for the presence of fast-growing nonmycobacterial organisms. The remaining portion of the specimen is kept untreated and refrigerated until the absence of contaminants is confirmed. Should this not be the case the remaining specimen can then be appropriately decontaminated.
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